Dendritic Cells: Biology and Clinical Applications

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Contributors

Luciano Adorini Istituto Superiore di Sanità, Roma, and Roche Milano Ricerche, Milano, Italy Matthew L. Albert The Rockefeller University, New York, NY, USA Paola Allavena Istituto di Ricerche Farmacologiche Mario Negri, Milan, Italy Francesca Aloisi Istituto Superiore di Sanità, Roma, and Roche Milano Ricerche, Milano, Italy Sebastian Amigorena INSERM, Institut Curie, Paris, France Fabrice André Institut Gustave Roussy, Villjuif, France Jonathan M. Austyn University of Oxford, John Radcliffe Hospital, Oxford, UK Jacques Banchereau Baylor Institute for Immunology Research, Dallas, TX, USA Penelope A. Bedford Imperial College School of Medicine, Northwick Park Institute for Medical Research, Harrow, UK Donna Beer Stolz University of Pittsburgh, Pittsburgh, PA, USA Nina Bhardwaj The Rockefeller University, New York, NY, USA C. Allen Black Magee Womens Research Institute, Pittsburgh, PA, USA Yuri V. Bobryshev St. Vincent’s Hospital, Sydney, NSW, Australia Francine Brière Schering-Plough Laboratory for Immunology Research, Dardilly, France Jozef Borvak Baylor Institute for Immunology Research, Dallas, TX, USA

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Christophe Caux Schering-Plough Laboratory for Immunology Research, Dardilly, France Jonathan Cebon Ludwig Institute for Cancer Research, Melbourne Tumor Biology Branch, Austin & Repatriation Medical Centre, Heidelberg, Victoria, Australia Vito R. Cicinnati University of Pittsburgh School of Medicine, Pittsburgh, PA, USA S. Citterio University of Milano-Bicocca, Milano, Italy Georgina J. Clark Mater Medical Research Institute, Mater Misericordiae Hospitals, South Brisbane, Queensland, Australia L. Cochand Hôpitaux Universitaires de Genève, Genève, Switzerland Sandra Columba-Cabezas Istituto Superiore di Sanità, Roma; and Roche Milano Ricerche, Milano, Italy Tyler J. Curiel Baylor Institute for Immunology Research, Dallas, TX, USA Ian Davis Ludwig Institute for Cancer Research, Melbourne Tumor Biology Branch, Austin & Repatriation Medical Centre, Heidelberg, Victoria, Australia Anthony J. Demetris University of Pittsburgh Medical Center, Pittsburgh, PA, USA Xin Dong University of Pittsburgh School of Medicine, Pittsburgh, PA, USA Bertrand Dubois INSERM U404 ‘Immunité et Vaccination’, Lyon, France Louis D. Falo Jr University of Pittsburgh School of Medicine, Pittsburgh, PA, USA Lian Fan The Scripps Research Institute, La Jolla, CA, USA Jerome Fayette Schering-Plough Laboratory for Immunology Research, Dardilly, France Nadine C. Fernandez Institut Gustave Roussy, Villejuif, France I. Frank The Rockefeller University, New York, NY, USA S. Schlesinger Frankel Division of Retrovirology, Walter Reed Army Institute of Research and the Henry M. Jackson Foundation, Rockville, MD; and Armed Forces Institute of Pathology, Washington, DC, USA Lawrence Fong Stanford University School of Medicine, Palo Alto, CA, USA Jean-François Fonteneau The Rockefeller University, New York, NY, USA John V. Forrester University of Aberdeen, Aberdeen, Scotland, UK John J. Fung University of Pittsburgh Medical Center, Pittsburgh, PA, USA

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Thomas F. Gajewski University of Chicago, Chicago, IL, USA Anne Galy Karmanos Cancer Institute, Wayne State University, Detroit, MI, USA Andrea Gambotto University of Pittsburgh School of Medicine, Pittsburgh, PA, USA Wendy S. Garrett Yale University School of Medicine, New Haven, CT, USA Angela Granelli-Piperno The Rockefeller University, New York, NY, USA F. Granucci University of Milano-Bicocca, Milano, Italy A.M. Gudin School of Medicine, Southampton University, Southampton, UK Derek N.J. Hart Mater Medical Research Institute, Mater Misericordiae Hospitals, South Brisbane, Queensland, Australia S.T. Holgate School of Medicine, Southampton University, Southampton, UK J.A. Holloway School of Medicine, Southampton University, Southampton, UK Fang-Ping Huang Sir William Dunn School of Pathology, University of Oxford, Oxford, UK R. Ignatius The Rockefeller University, New York, NY, USA Tatyana Isaeva Baylor Institute for Immunology Research, Dallas, TX, USA Ronald Jaffe University of Pittsburgh Medical Center, Pittsburgh, PA, USA Nori Kadowaki DNAX Research Institute, Palo Alto, CA, USA Pawel Kalinski University of Pittsburgh, Medical Center, Pittsburgh, PA, USA Martien L. Kapsenberg University of Amsterdam, Amsterdam, The Netherlands Kristine Kikly GlaxoSmithKline Pharmaceuticals, King of Prussia, PA, USA Stella C. Knight Imperial College School of Medicine, Northwick Park Institute for Medical Research, Harrow, UK Marie H. Kosco-Vilbois Serono Pharmaceutical Research Institute, Plan-les-Ouates, Switzerland Adriana T. Larregina University of Pittsburgh School of Medicine, Pittsburgh, PA, USA Marie Larsson The Rockefeller University, New York, NY, USA Andrew Lee The Rockefeller University, New York, NY, USA Li Ming Liu Sir William Dunn School of Pathology, University of Oxford, Oxford, UK

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Yong-Jun Liu DNAX Research Institute, Palo Alto, CA, USA David Lo Digital Gene Technologies Inc., La Jolla, CA, USA Michael T. Lotze GlaxoSmithKline Pharmaceuticals, King of Prussia, PA, USA Lina Lu Thomas E. Starzl Transplantation Institute, and Department of Surgery, University of Pittsburgh Medical Center, Pittsburgh, PA, USA Thomas Luft Ludwig Institute for Cancer Research, Melbourne Tumor Biology Branch, Austin & Repatriation Medical Centre, Heidelberg, Victoria, Australia Kelli MacDonald Mater Medical Research Institute, Mater Misericordiae Hospitals, South Brisbane, Queensland, Australia G. Gordon MacPherson Sir William Dunn School of Pathology, University of Oxford, Oxford, UK Thomas C. Manning University of Chicago, Chicago, Illinois, USA Alberto Mantovani Istituto di Ricerche Farmacologiche Mario Negri, Milan, Italy Eugene Maraskovsky Ludwig Institute for Cancer Research, Melbourne Tumor Biology Branch, Austin and Repatriation Medical Centre, Heidelberg, Victoria, Australia Florentina Marches Baylor Institute for Immunology Research, Dallas, TX, USA Carole Masurier Institut Gustave Roussy, Villejuif, France Hiroyuki Matsue University of Texas Southwestern Medical Center, Dallas, TX, USA Dieter Maurer University of Vienna Medical School, Vienna, Austria Paul G. McMenamin University of Western Australia, Nedlands, Western Australia, Australia Ira Mellman Yale University School of Medicine, New Haven, CT, USA D. Messmer The Rockefeller University, New York, NY, USA Michael Murphey-Corb University of Pittsburgh, Medical Center, Pittsburgh, PA, USA Noriko Murase University of Pittsburgh Medical Center, Pittsburgh, PA, USA Laurent P. Nicod Hôpitaux Universitaires de Genève, Genève, Switzerland Karen A. Norris University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, USA Peta J. O’Connell Thomas E. Starzl Transplantation Institute and Department of Surgery, University of Pittsburgh, Pittsburgh, PA, USA Glenn D. Papworth University of Pittsburgh, Pittsburgh, PA, USA

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Karolina Palucka Baylor Institute for Immunology Research, Dallas, TX, USA Melissa Pope The Rockefeller University, New York, NY, USA Bali Pulendran Baylor Institute for Immunology Research, Dallas, TX, USA Paul Racz Bernhard Nocht Institute for Tropical Medicine, Germany Gwendalyn J. Randolph Institute for Gene Therapy and Molecular Medicine, Mount Sinai School of Medicine, New York, NY, USA Graça Raposo CNRS, Institut Curie, Paris, France Will Redmond The Scripps Research Institute, La Jolla, CA, USA Armelle Regnault INSERM, Institut Curie, Paris, France Christina R. Reilly The Scripps Research Institute, La Jolla, CA, USA M. Rescigno University of Milano-Bicocca, Milano, Italy Paola Ricciardi-Castagnoli University of Milano-Bicocca, Milano, Italy M. Rittig University of Erlangen, Erlangen, Germany Paul D. Robbins University of Pittsburgh Medical Center, Pittsburgh, PA, USA Russell D. Salter University of Pittsburgh School of Medicine, Pittsburgh, PA, USA Amanda E. Semper School of Medicine, Southampton University, Southampton, UK Barbara Serafini Istituto Superiore di Sanità, Roma; and Roche Milano Ricerche, Milano, Italy Vassili Soumelis DNAX Research Institute, Palo Alto, CA, USA Silvano Sozzani Istituto di Ricerche Farmacologiche Mario Negri, Milan, Italy Hergen Spits Division of Immunology, Netherlands Cancer Institute, Amsterdam, The Netherlands C. Stahl-Hennig German Primate Center, Göttingen, The Netherlands Ralph M. Steinman The Rockefeller University, New York, NY, USA Raymond J. Steptoe Walter and Eliza Hall Institute of Medical Research, Melbourne, Victoria, Australia Georg Stingl University of Vienna Medical School, Vienna, Austria Akira Takashima University of Texas Southwestern Medical Center, Dallas, TX, USA

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K. Tenner-Racz Bernhard Nocht Institute for Tropical Medicine, Hamburg, Germany Magali Terme Institut Gustave Roussy, Villejuif, France Clotilde Théry INSERM, Institut Curie, Paris, France Ranjeny Thomas Centre for Immunology and Cancer Research, University of Queensland, Princess Alexandra Hospital, Brisbane, Queensland, Australia Angus W. Thomson Thomas E. Starzl Transplantation Institute, and Departments of Surgery and Molecular Genetics and Biochemistry, University of Pittsburgh Medical Center, Pittsburgh, PA, USA M. Urbano University of Milano-Bicocca, Milano, Italy B. Valzasina University of Milano-Bicocca, Milano, Italy Stephane Vandenabeele The Walter and Eliza Hall Institute of Medical Research, Post Office Royal Melbourne Hospital, Melbourne, Victoria, Australia Slavica Vuckovic Mater Medical Research Institute, Mater Misericordiae Hospitals, South Brisbane, Queensland, Australia Simon C. Watkins University of Pittsburgh, Pittsburgh, PA, USA Shuang Wei Baylor Institute for Immunology Research, Dallas, TX, USA Jeffrey Weber USC/Norris Comprehensive Cancer Center, Los Angeles, CA, USA Cara C. Wilson University of Colorado Health Sciences Center, Denver, CO, USA Joseph Wolfers Institut Gustave Roussy, Villejuif, France Li Wu The Walter and Eliza Hall Institute of Medical Research, Melbourne, Victoria, Australia L. Zhong The Rockefeller University, New York, NY, USA Laurence Zitvogel Institut Gustave Roussy, Villejuif, France Weiping Zou Baylor Institute for Immunology Research, Dallas, TX, USA

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Foreword to the Second Edition Nossal J.V. The University of Melbourne, Victoria, Australia

First, therefore, we must seek what it is that we are aiming at; then we must look about for the road by which we can reach it most quickly, and on the journey itself, if only we are on the right path, we shall discover how much of the distance we overcome each day, and how much nearer we are to the goal toward which we are urged by a natural desire. But so long as we wander aimlessly, having no guide, and following only the noise and discordant cries of those who call us in different directions, life will be consumed in making mistakes – life that is brief even if we should strive day and night for sound wisdom. Let us, therefore, decide both upon the goal and upon the way and not fail to find some experienced guide who has explored the regions towards which we are advancing; for the conditions of this journey are different from those of most travel. On most journeys some well-recognized road and inquiries made of the inhabitants of the region prevent you from going astray; but on this one all the best beaten and the most frequented paths are the most deceptive. Nothing, therefore, needs to be more emphasized than the warning that we should not like sheep follow the lead of the throng in front of us, travelling, thus, the way that all go and not the way that we ought to go. Seneca, On the Happy Life (AD 58).

Drs Lotze and Thomson are to be congratulated on the tremendous success of the first edition of this book, which has necessitated a second edition after such a short time. It is indeed a pleasure to provide this brief foreword from the perspective of one who has followed the progress of cellular immunology at close quarters for 45 years, and thus is truly an eye-witness to history. It is amazing to me to think of how little the cellular basis of immune responses preoccupied the early giants of our discipline. For example, Paul Ehrlich (1900), though highlighting the possible importance of cell surface molecules in the antibody response, scarcely mentioned which cell he thought might be doing the job. To the extent that people worried about the question at all, the macrophage, well known for its role in phagocytosis of invading microorganisms, was presumed to be the antibody-former, for example by Jerne (1955). It was not until the beautiful studies by Fagraeus (1948) on the emergence of proliferating plasmablasts and plasma cells that the full spotlight was put on these specialized cells as fabricating antibody, as later proven by Leduc et al. (1955) and the author (Nossal, 1959). Similarly, the role of the lymphocyte in cell-mediated immunity became clear during the 1950s and early 1960s. As these cells were in no way implicated in antigen capture, there was a real gap in understanding needing to be filled. During the 1960s, a golden age in cellular immunology, the notion that three types of cells collaborated in immune responses gradually began to hold sway. These were the macrophage, the T cell and the B cell. The dendritic cell had not yet come into full focus. xv

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Strangely enough, it was the author’s collaboration with Gordon Ada (Nossal et al., 1964, 1968) that first brought a peculiar kind of dendritic cell into prominence when we reported the extraordinary antigen-capturing potential of primary lymphoid follicles and the fact that germinal centres soon developed in close proximity to the antigen depot. Structural studies revealed that the antigen was actually captured on the surface of long intertwining dendritic processes, and furthermore that antigen was retained in this location for prolonged periods. Dr John Tew and his colleagues, particularly Andras Szakal and Marie Kosco subsequently deepened our understanding of follicular dendritic cells and their role in immunological memory (Tew et al., 1990). Further clarity entered the field when Tew, Thorbecke and Steinman (1982) presented a sensible nomenclature for dendritic cells in immune responses. However, nothing can eclipse the tremendous role that Ralph Steinman (1991) has played in bringing dendritic cells into focus and prominence. It was his methods of identification and isolation of the ‘regular’ dendritic cells that allowed a subsequent veritable tidal wave of experimentation which has now led to the rich diversity of structure and function which makes the present volume such fascinating reading. The reason why DCs are so prominent in current discussions in immunology relates to their extraordinary and perhaps unique ability to activate and maintain the survival of T lymphocytes. It first became clear that, on a quantitative basis, they were by far the most powerful stimulators of allogeneic T-cell responses. In the field of T-cell activation in response to recall antigens, they are also paramount. In regulation of antibody formation, dendritic cells pre-pulsed with antigen are by far the most immunogenic cells that can be found. In vivo, it is now clear that so-called interdigitating dendritic cells are also present at the initiation of the germinal center reaction. The sequence appears to be that DCs, perhaps after migrating from peripheral sites of antigen uptake, firstly activate T cells which in turn activate B cells, some of which migrate into the primary lymphoid follicle. The germinal center reaction is initiated when, in turn, these B cell blasts interact with follicular dendritic cell-bound antigen. Twenty-five years after our initial interest in follicular dendritic cells, our attention was caught again by the germinal center reaction (summarized in Nossal, 1994). Technology had advanced to the stage where molecular biological and cell culture methods allowed the process of somatic mutation in germinal center B cells to be studied at the single cell level. This quickly confirmed the prior hypothesis of Kocks and Rajewsky (1989) that the somatic hypermutation which eventually leads to affinity maturation of the antibody response is not a feature of either primary or secondary B cells giving rise to a clone of antibody-forming cells. Rather, the process occurs during the separate clonal development of memory cells in germinal centers. The germinal center reaction permits an iterative process whereby only B cells with heightened affinity for the antigen in question gain access to antigen bound on follicular dendritic cells, permitting further stimulation and clonal proliferation, with multiple rounds of mutation and selection finally leading to greatly changed and higher affinity antibodies. There are many respects in which the germinal center reaction resembles the original process of generation of primary B cells. Suppression and then re-emergence of lg receptors are noted on the centrocyte surface followed by generation of new patterns of immunoglobulin expression conferred by high rates of mutation and not as originally believed by V gene translocations. A phase of substantial susceptibility to tolerance follows (Pulendran et al., 1995). Indeed, if B cells are ‘caught’ in transit between the deeper layers of the germinal center, where cells are proliferating rapidly, and the areas of antigen deposition on the more superficial aspects of the germinal center encounter soluble antigen, they undergo apoptotic death. This resembles in principle what happens in the bone marrow if a B cell encounters ‘its’ antigen prior to being released from the marrow as a mature B cell. Presumably this mechanism prevents survival of B cells which could hypermutate, generating high-affinity anti-self reactivity. If any further evidence of the great importance of DCs in the immune responses was required, it was provided by the work of Boyle et al. (1998) on DNA vaccines. One of the real problems hindering the

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more rapid development of DNA vaccines has been the relatively weak nature of the immune responses they induce in many model systems. If, however, the gene construct encoding an antigen is modified so as to coexpress the molecule CTLA4, both B- and T-cell responses are tremendously enhanced. CTLA4 is a counterstructure to two receptors, CD80 and CD86, which are constitutively expressed on DCs. The strategy is therefore one which targets the product of DNA immunization directly to DCs. This strategy may well prove to have important practical applications. No student of cellular immunology will be surprised to learn that there are different subsets of DCs beyond the variations we have briefly described. This aspect is at a fairly early stage of exploration and it is fitting that the present state of knowledge will be summarized by Dr Li Wu.

CONCLUSIONS The editors are to be congratulated on having assembled a fascinating and authoritative series of chapters which summarize so many aspects of the anatomy, physiology, biochemistry and pathology of dendritic cells in considerable detail. The fact that this second edition follows so quickly after the first reflects the vigor and excitement of the field. Very few of us pondering the intricacies of the three cell interactions in immunology, which we thought to rotate around macrophages, T cells and B cells, would have thought as recently as 25 years ago that a book of these dimensions, with no fewer than 45 chapters, could be devoted to dendritic cells. A new, exciting, important and practical series of paradigms has been created in this short period. Let us hope that this book will stimulate still further effort, and that the third edition some years hence will also bring forth powerful new insights.

REFERENCES Boyle, J.G., Brady, J.L. and Lew, A.M. (1998). Enhanced responses to a DNA vaccine encoding a fusion antigen that is directed to sites of immune induction. Nature 392, 408–411. Ehlrich, P. (1900). On immunity, with special reference to cell life. Proc. R. Soc.B 66, 424. Fagraeus, A. (1948). The plasma cellular reaction and its relation to the formation of antibodies in vitro. J. Immunol. 58, 1–13. Jerne, N.J. (1955). The natural selection theory of antibody formation. Proc. Natl Acad. Sci. USA 41, 849–857. Kocks, C. and Rajewsky, K. (1989). Stable expression and somatic hypermutation of antibody V regions in B-cell developmental pathways. Annu. Rev. Immunol. 7, 537–559. Leduc, E.H., Coons, A.H. and Connolly, J.M. (1955). Studies on antibody production; primary and seconary responses in popliteal lymph node of rabbit. J. Exp. Med. 102, 61–72. Nossal, G.J.V. (1959). Antibody production by single cells, III: the histology of antibody production. Br. J. Exp. Pathol. 40, 301–311. Nossal, G.J.V. (1994). Differentation of the secondary B-lymphocyte repertoire.: the germinal center reaction. Immunol. Rev. 137, 173–183. Nossal, G.J.V., Ada G.L. and Austin, C.M. (1964) Antigens in immunity, IV: cellular localization of 125 l- and 131 l-labelled flagella in lymph nodes. Aust. J. Exp. Biol. 42, 311–330. Nossal, G.J.V., Abbot, A., Mitchell, J. and Lummus, Z. (1968). Antigens in immunity, XV: ultrastructural features of antigen capture in primary and secondary lymphoid follicles. J. Exp. Med. 127, 277–290. Pulendran, B., Kannourakis, G., Nouri, S., Smith, K.G.C. and Nossal, G.J.V. (1995). Soluble antigen can cause enhanced apoptosis of germinal-centre B cells. Nature 375, 331–334. Steinman, R.M. (1991). The dendritic cell and its role in immunogenicity. Annu. Rev. Immunol. 9, 271–296. Tew, J.G., Thorbecke, G.J. and Steinman, R.M. (1982). Dendritic cells in the immune response: characteristics and recommended nomenclature. J. Reticuloendothel. Soc. 31, 371–380. Tew, J.G., Kosco, M.H., Burton, G.F. and Szakal, A.K. (1990). Follicular dendritic cells as accessory cells, Immunol. Rev. 117, 185–211.

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Preface to the Second Edition The most beautiful experience we can have is the mysterious. It is the fundamental emotion which stands at the cradle of true art and true science. Albert Einstein, The World As I See It.

The substantial mystery surrounding antigen processing and presentation, initiation of the immune response and how the effector phase of the immune response vectored onto the dendritic cell (DC) over the last five years provided the impetus for our first edition. Much of the interest in DCs was made manifest in the increasing number of references and meetings devoted to this cell. We recall that there was a belief amongst outside advisors that a book dedicated to a single cell making for a paltry audience with limited scope would have equally limited value. This view, with the advantage of hindsight, was clearly wrong. We discovered, as our book was coming into print (quite rapidly from the time of initiation), that the field continued to grow with questions and answers leading inexorably to the next set of questions, apparently growing logarithmically. In addition, there was an appreciable interest in having much of the detailed information organized in one volume in a way that allowed credible exploration for the experienced dendritophile as well as the tyro. For those interested in these things, at the time of this edition, there have been 10,448 citations concerning DC in the literature since this cell was first described by Steinman and Cohn (1973) now almost 30 years ago; a prodigious literature to embrace and know – all on one cell! We have attempted to integrate many of these references by era (year) at the end of this volume, as an innovation for our readers and we hope that it will be of service. You can clearly see who are the authors of the DC citation classics by the number of chapters referencing them at the end of each citation! We have also introduced multiple new chapters and had several started anew to keep the volume contemporary and to reflect the new areas of biology and clinical application. So what have we learned in the last three years? The answer is obviously quite a bit, both in the clinical arena into which DCs have been thrust, and in the area of basic science where gene discovery has uncovered a veritable treasure trove of new molecular targets limited to or preferentially in DCs (Hartgers et al., 2000). The notion of lymphoid and myeloid DCs and the disparity between mouse and human biology are finally being resolved, although not yet to everyone’s satisfaction. The critical role of the DC in self and transplantation tolerance (Kurts et al., 2001; Thomson and Lu, 1999) and their association with a pathogen’s ability to identify and target this cell, both as a means of immunosuppression as well as a means to shuttle virus or other microbes to distant sites (Servet-Delprat et al., xviii

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2000) represent evolving areas. We have attempted to capture the current art, but additional information presents apace. The use of recently-identified tumor antigens to be presented by DC to cells of the immune system captured our interest and that of others. Moreover the number of clinical trials using these cells is now mature enough to support publication and presentation at national and international meetings. Now that we understand how tumor antigens interact with cells of the immune system, issues related to the source of antigen, adjuvanticity of DCs, and how tumors suppress DC function and maturation have become the central issues (Mayordomo et al., 1995; Lotze and Jaffe, 1998). Finally, it probably is a given, but we should recognize that in the absence of other immune cells, the DC plays at best a bit part. Likewise, we cannot limn the shape of cellular or humoral immunity without reflecting on the critical role of DC in shaping and selecting our immunity. Perhaps we should argue that virtually all treatises on modern B or T cell biology, without reflection on the critical role of DCs, are at best incomplete. And, at the end of this preface, we need to thank those individuals who have made our lives possible to lead meaningfully in the academic environment, providing support in every way they could to enable the preparation of this second edition. We would like to dedicate it to our administrative assistants – Maria Bond, Kathy Rakow, Margaret Corson, Joyce Caperilla, Shelly Conklin, as well as the staff of Academic Press, Lillian Leung, Jacqueline Read, and Tessa Picknett. In addition, Bridget Colvin’s talent uncovered some remarkably ‘DCesque’ literary quotations and to Thomas Lotze for assistance with the compilation of the references at the end. If we have been at all successful, it has been because of their fine efforts. We thank them for their patience and prudence at all stages of this book’s current evolution. Michael T. Lotze, GlaxoSmithKline Pharmaceuticals, King of Prussia, PA Angus W. Thomson, University of Pittsburgh, Pittsburgh, PA

REFERENCES Steinman, R.M. and Cohn, Z.A. (1973). J. Exp. Med. 137, 1142–1162. Hartgers F.C., Vissers J.L., Looman M.W et al. (2000). Eur J. Immunol. 30(12), 3585–3590. Kurts C., Cannarile M., Klebba I. et al. (2001). J. Immunol. 166(3), 1439–1442 Servet-Delprat C., Vidalain P.O., Azocar O. et al. (2000). J. Virol. 74(9), 4387–93 Mayordomo J.I., Zorina T., Storkus W.J. et al. (1995). Nature - Medicine. 1(12), 1297–1302. Lotze M.T. and Jaffe R. (1998). In: Lotze M.T. and Thomson A.W. (eds) Dendritic Cells. Academic Press, London pp. 325–338. Recigno, M., Urbana, M., Valzasina, B. et al. (2001). Nature Immunology 2, 361–367. Thomson A.W. and Lu L. (1999). Immunol. Today 20, 27–32

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The editors (Dr Lotze at right, Dr Thomson at left) pictured with the first edition of the book.

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Preface to the First Edition Fighting in the forefront of the Greeks, the Athenians crushed at Marathon the might of the gold bearing Medes. Simonides, c. 556–468 B.C.

This volume represents a special effort to bring togther in one place the information that would allow a scientifically oriented clinician, or even nonimmunologically oriented scientist, to appreciate the important role of this previously obscure cell. The notion of having an entire book dedicated to a single cell, albeit one as important as the dendritic cell (DC), is one which could be met with disapprobation in some quarters. It does, however, reflect the importance which the authors (who completed their tasks admirably) and the editors have placed on the extraordinary important role this cell has in dictating the initiation and persistence of the adaptive immune response. Indeed, all aspects of the ‘fight’ during the acute natural immune response but in particular the chronic infammatory immune response associated with cancer, chronic infections diseases, autoimmunity, and transplantation are importantly related to DC biology. While the first report of DC by Ralph Steinman and Zanvil Cohn, 25 years ago, posited an important role for this cell in immune regulation, this volume’s role is not soley celebratory (but it does have those elements!). It has thus been appreciated for the last 25 years that DC are specialized antigen-presenting cells (APC) with a unique ability to prime effective immune responses. This may give them a special importance in several human disease states known to have an immunological basis. While great strides were made in the understanding of the role of specific T cells and antibodies in mediating allergy, autoimmunity, graft rejection, infectious disease, and tumor immunity over this period, it was also clear that these effector cells and molecules represented the end stage of an immune response, the outcome of which was probably determined at its very initiation by the type of APC, the nature and state of the antigen, and the cytokine conditions under which the antigen was first presented. As is usually the case in science, testing of ideas must await development of new techniques and reagents. Several opportunities presented themselves in the 1970s with discovery and cloning of the first few cytokines, especially interferon-α and IL-2. This allowed for the first time use of immunologically important molecules themselves as means to manipulate immune responses or to support the growth and expansion of T cells and NK cells in vitro. This in turn led to the identification and characterization of the T cell receptor for antigen and examination of disease-specific T cell responses. T cells that mediate graft rejection, T cells specific for tumor antigens, and T cells specific for autoantigens were studied for their biology and function, as well as being used as reagents to fully define disease-specific antigens and the genes that encode them. The important role of cytokines, as well as the number of indentifiable cytokines, continued to rise xxi

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throughout that same period, now with possibly as many as 20–21 so-called ‘interleukins’ in addition to the colony-stimulating factors and interferons. Their pivotal role in directing the immune response was unveiled. An important outcome from this work has been the understanding of the polarity of the immune response at the T cell level, TH1 vs TH2 and Tc1 vs Tc2. The helper T cell response that develops as predominately TH1 type (in the presence of and producing TH1-type cytokines, IL-2, IFN-γ, GM-CSF) supports the development of cytotoxic T cells (CTL), often of the Tc1 type. Alternatively, the TH2-type response (developed in the presence of and producing the TH2-type cytokines, IL-4, IL-10) supports the development of humoral immunity and Tc2-type CTL. In systems where this can be tested in vivo, this dichotomy translates into a life or death, or disease or no disease situation. During this same period, understanding of immunoglobulin gene rearrangements and the advent of monoclonal antibody technology truly revolutionized all of the biological sciences, perhaps one of the greatest gifts that immunologists have yet bestowed on their nonimmunological colleagues. So how is this complex adaptive system initiated and maintained? We now believe that the DC plays an important role in initiating and maintaining immune reactivity. The sequential steps in the choreography from its birth in the bone marrow, maturation there or in the thymus or secondary lymphatic tissues, traversion of the initial endothelial barriers into virtually all tissues, sensitivity to inflammatory initiators and tissue damage, injury or ‘danger’, and transformation into a ‘mature cell’ capable of migrating across the lymphatic or postcapillary venules to enter secondary lymphoid sites, interacting with the resident T cells and B cells to rapidly screen and select immunological suitors make it one of the most versatile of dance partners. Not only does it have the potential to come into contact with all cells in all tissues, its athletic potential as the ‘track star of immunology’ makes it a marathon contender of the first order. And, just like the marathon runner who crossed the Plains of Marathon, after delivering its message in the lynph node or spleen, this cell dies, to be born again, Athena-like, in the bone marrow every day. The last five years have been particularly dizzying with application of cytokines now to the culture and maintenance of DC. Just as the advances in T cell and B Cell biology required means to grow and maintain these cells, the use of GM-CSF and IL-4 or TNF has made their study and application in preclinical and clinical disease models feasible. Prior studies, restricted by the relatively limiting numbers of these cells from any one site in tissue or blood, have now been extended by extraordinary new knowledge available from advances in these relatively simple and straightforward culture techniques. This new knowledge has emphasized the importance of the very early events in the priming of the immune response and turned full attention to the DC. As we prepare these comments, some of the important issues in DC migration and recruitment via specialized chemokines including MIP-3 and their receptors CCR6 and CCR7, as well as the preferred source of antigenic material for DC (apoplotic cells and bodies) are just coming to light. These and other recent insights will require some time to become integrated into the growing corpus of information and raise important additional questions about these centrally important cells. This volume celebrates them and, when it enters the second edition, hopefully many of the issues raised here will have been successfully addressed and new ones raised. We appreciate the careful assistance of our publishers from Academic Press, Tessa Picknett, Duncan Fatz, Lilian Leung and Emma White for their belief in the importance of this project, their patience, and persistence. To our families and in particular to Joan and Robyn, our spouses, acknowledgment for their gifts of time and support. To our students and colleagues who make the intellectual challenges and the great social role of science a pleasure and vocation, our gratitude and hopes for the future. Michael T. Lotze Angus W. Thomson

REFERENCE Steinman, R.M. and Cohn, Z.A. (1973). J. Exp. Med. 137 (5), 1142–1162.

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1 The development of dendritic cells from hematopoietic precursors Li Wu1 and Anne Galy 2 1

The Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia 2

Karmanos Cancer Institute, Wayne State University, Detroit, Michigan, USA

Experience is not what happens to a man. It is what a man does with what happens to him. Aldous Huxley

INTRODUCTION

DCs and DC populations found in other lymphoid organs. DCs have powerful functions in the immune system. They can capture and process antigens, then present the antigenic peptides and activate specific T cells (Steinman, 1991; Banchereau and Steinman, 1998). Variations among the tissue distribution of DCs and differences in their phenotype and function indicate the existence of heterogeneous populations of DCs (Hart, 1997). DCs were originally considered to be of myeloid origin and closely related to monocytes, macrophages and granulocytes. However, recent studies suggested that DCs can be generated along distinct developmental pathways and can originate from precursors of different hematopoietic lineages, with at least two DC lineages being identified so far, namely the conventional myeloid-related DC and the newly defined lymphoid-related DC lineages. Because various populations of DCs in mice (Kronin et al., 1996; Maldonado-Lopez

Blood cells are diverse types of cells that provide highly specialized functions such as tissue oxygenation, tissue repair, blood clotting or immune responses. The continuous demand for the supply of these various types of blood cells is provided by the proficient, yet tightly regulated development of hematopoietic progenitor cells. Cell fate specification of these uncommitted multipotential cells is therefore an important aspect of hematopoietic cell development, but one that remains incompletely understood. The study of the dendritic cell (DC) lineage specification has provided interesting insights into this area. DCs constitute a system of hematopoietic cells that are rare but ubiquitously distributed. Several DC types with different biological features have been identified in different tissues, including Langerhans cells (LCs) in the epidermis, interstitial DCs in various tissues, thymic Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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et al., 1999; Pulendran et al., 1999) or humans (Caux et al., 1997) are able to induce distinct types of immune responses, it becomes important to determine the role of their origin in determining functional heterogeneity. An important question is how this heterogeneity arises at the developmental level.

EARLY DEVELOPMENTAL DECISION CHECKPOINTS IN THE HEMATOPOIETIC DEVELOPMENT OF DCs The early developmental steps of DC formation from hematopoietic progenitor cells are not uniform and involve different types of progenitor cells, different developmental pathways and different signals. Understanding these early events is facilitated by the identification of early developmental checkpoints in the hematopoietic development of DCs.

Early hematopoietic progenitors The existence and identification of lineagerestricted progenitor cells has been helpful in our understanding of hematopoietic cell fate specification. Multipotent yet lineage-restricted progenitor cells identified and characterized in mice and in humans can be distinguished from the most primitive hematopoietic stem cells (HSCs) based on differences in cell surface phenotype and the capacity and durability of multilineage engraftment. In the murine thymus an early lymphoid-restricted precursor population termed the ‘low CD4 precursor’ has been identified. This precursor population does not express hematopoietic lineage markers (Lin), but expresses low levels of CD4 and Thy-1 and high levels of the hematopoietic progenitor cell markers c-kit and Sca-1 (Wu et al., 1991a). These precursors, although isolated from thymus, are not yet committed to the T-cell lineage and are able to produce T cells, B cells, natural killer (NK) cells and DCs (Wu et al., 1991b; Ardavin et al., 1993). However, they have no myeloid and erythroid differentiation potential (Wu et al., 1991b).

In murine bone marrow (BM), clonogenic common lymphoid and common myeloid progenitors have also been identified recently (Kondo et al., 1997; Akashi et al., 2000). IL-7Rα expression is a main marker to distinguish these two progenitors. The common lymphoid progenitors (CLPs) are Lin, IL7Rα, c-kitlo and Sca-1lo. Such cells can generate all lymphoid cells at clonal level and some DCs (Wu et al., unpublished), but not detectable myeloid or erythroid cells (Kondo et al., 1997). The common myeloid progenitors (CMPs) are Lin, IL-7Rα, c-kit, Sca-1, CD34, FcγRlo (Akashi et al., 2000). These cells can give rise to precursors for megakaryocytes/erythrocytes (MEPs) and precursors for granulocytes/macrophages (GMPs) (Akashi et al., 2000). CMPs also produce DCs (Traver et al., 2000) The human equivalent of mouse CMPs has not yet been described. However, progenitor cells with features similar to those of the CLPs in mouse have been identified in human. The human BM progenitor cells expressing CD34, CD45RA, CD10 and IL-7Rα, but no lineage-associated markers, have differentiation potential restricted to the production of lymphocytes and DCs, but not of myeloid cells and erythrocytes (Galy et al., 1995b; Ryan et al., 1997). This CLP arises from a myeloid/lymphoid-restricted progenitor cell with limited erythroid differentiation potential that is contained in the CD34CD45RA cell population (Galy et al., 1995a). Thus, hematopoietic cell fate specification occurs incrementally. The existenceofprogenitorcellsthatcangiverisetoseveral lineages but not to all hematopoietic lineages represents possible developmental checkpoints of hematopoietic differentiation.

Developmental relationships of myeloid lineage and DCs It is generally assumed that DCs have a ‘myeloid‘ origin because they arise from hematopoietic progenitor cells with myeloid differentiation potential and they can be produced from monocytes, a typical myeloid cell. Monocytes can generate immunostimulatory DCs. This differentiation process occurs without proliferation

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and is induced at a high frequency in culture by granulocyte–macrophage colony-stimulating factor (GM-CSF) and IL-4 (Romani et al., 1994; Sallusto and Lanzavecchia, 1994; Zhou and Tedder, 1996). In spite of this relationship, DCs can also develop independently from monocytes in ‘myeloid’ cell growth conditions. In mice, distinct pathways giving rise to granulocytes, monocytes/macrophages and DCs from a blood or BM precursor negative for MHC class II have been described (Inaba et al., 1993). DCs can be generated along with phagocytic myeloid cells from cells within a single colony in semisolid medium cultures. GM-CSF, but not granulocyte (G)-CSF or macrophage (M)-CSF, is required for DC development in such systems (Inaba et al., 1993). DCs generated under such conditions express MHC class II, display the characteristic morphology of DCs and are potent stimulators of resting T cells. They also have the capacity to home to T-cell regions in draining lymph nodes (Inaba et al., 1993). In the human system, it is also possible to grow pure colonies of DCs (DC-CFU) from BM in the presence of GM-CSF, TNFα and stem cell factor (SCF). These DC-CFU are distinct from mixed DC-myeloid CFU, therefore indicating that myeloid cells and DCs can have distinct clonogenic precursors at some point in their development (Young et al., 1995). The study of Langerhans cell (LC) production also indicates the existence of early developmental options within the DC lineage. By careful analysis of the in vitro differentiation of CD34 progenitor cells in culture, it is possible to recognize the existence of separate precursors of LCs that can be distinguished by their phenotype, by involvement of a recognizable monocyte stage and by their requirement for TGFβ (Caux et al., 1996; Geissmann et al., 1998; Jaksits et al., 1999). Among CD34 cells, the expression of low levels of the IL-3Rα chain (Olweus et al., 1997) or the expression of the cutaneous lymphocyteassociated antigen CLA (Strunk et al., 1997) defines progenitor cells able to give rise to LCs in culture in the presence of GM-CSF and TNFα. Two developmental pathways have been identified for the production of LCs and DCs from

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CD34 progenitor cells. One gives rise to HLADRbright cells with LC morphology, phenotype and function, via a CD14CD1a intermediate, after 4–5 days in culture with GM-CSF, TNFα and SCF (Caux et al., 1996). Another differentiation pathway of CD34+ cells was identified in these culture conditions that gives rise to a CD14+CD1a bipotential intermediate cell. This intermediate cell produces non-LC DCs but can be induced in the presence of TGFβ to differentiate into LCs (Jaksit et al., 1999). Alternatively, this intermediate cell can differentiate along a macrophage pathway when recultured with M-CSF (Caux et al., 1996; Szabolcs et al., 1996). Thus, the divergence in developmental pathways leading to the production of LCs and non-LC DCs has one origin within the small population of CD34+ progenitor cells. Both mouse and human studies indicate that there is a close lineage relationship between myeloid cells and DCs, and that some aspects of DC cell fate specification occur at the progenitor cell level.

Relationship between development of lymphocytes and some DCs Further insight into DC lineage specification has been obtained in studies of lymphoid progenitor cell subsets. Mouse thymic DCs and a subpopulation of DCs in spleen and lymph nodes express several markers of lymphoid cells, such as CD8αα, CD2, BP-1 and CD25 (Vremec et al., 1992; Wu et al., 1995). This was the first suggestion of a relationship between these DCs and lymphoid cells. Indeed, when the intrathymic lymphoid restricted precursor, the ‘low CD4 precursor’ population (Wu et al., 1991a, 1991b), was transferred intrathymically, thymic CD8α DCs were generated and when injected intravenously, both thymic DCs and the splenic CD8α DC population were generated (Ardavin et al., 1993; Wu et al., 1995, 1996). Unlike the bone marrow precursors, which produced both CD8α and CD8α DC populations in mouse spleen, the intrathymic precursor population can only generate the CD8α DCs (Wu et al., 1996). This suggests that the CD8α DC population represents a lymphoid-related DC lineage.

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In vitro studies showed that when CD8α or CD8α DC subsets were placed in short-term cultures to allow their further differentiation, they did not differentiate into one or the other (Vremec and Shortman, 1997 and unpublished). This again supports the theory that the CD8α and CD8α DC subsets represent separate DC lineages rather than DCs at different developmental stages of the same lineage. Further studies on developmental potential of the precursor populations downstream from the earliest ‘low CD4 precursors’ in T-cell development, namely the CD3CD4CD8 triple negative (TN) thymocyte precursor populations, revealed that the early TN precursor population, the ‘pro-T’ cell population, also retained the potential to form DCs (Wu et al., 1996). These pro-T cells resemble the ‘low CD4 precursors’ because their T-cell receptor (TCR) β gene is in germline state, they have the potential to produce T cells and DCs, but have lost the potential to form B cells and NK cells. This suggests a close relationship between T cells and DC development in the thymus. In contrast, the later TN precursor population, the pre-T cell, which has rearranged TCR β genes, is no longer able to produce any lineages other than T cells (Wu et al., 1996; Lucas et al., 1998). Therefore, it appears that full commitment to T cell lineage occurs at the stage of TCR β gene rearrangement. Interestingly, DCs can also be generated in cultures from the intrathymic ‘low CD4 precursors’ or from pro-T cells in the presence of a set of cytokines which was different from the ones normally used for generating myeloid-derived DCs (Saunders et al., 1996; Lucas et al., 1998). The main difference from the myeloid-derived DC cultures was the lack of requirement for myeloid cell growth factor GM-CSF to stimulate proliferation or DC differentiation. The cytokines required for DC generation from the thymic precursors include TNFα, IL-1, IL-3, IL-7, SCF, Flt-3L and CD40L. DCs could be generated from single low CD4 precursor cells in these cultures with a cloning efficiency of about 70%. Thus, the majority of the thymic lymphoid precursors are able to produce DCs (Saunders et al., 1996).

Interestingly, a recent report by Bjorck and Kincade (1998) showed that mouse bone marrow CD19 pro-B cells could also develop into DCs with T-cell stimulatory properties when cultured under conditions similar to those used for DC production from the thymic lymphoid precursors. This further illustrates the close relationship between some DCs and committed lymphoid progenitor cells and shows a potential link between DCs and the B lineage. However, it is not known whether DC differentiation from B-cell precursors occurs in vivo. Similarly, in the human system, relationships between DCs and lymphoid progenitor cells have been found. Hematopoietic progenitor cells expressing CD45RA, the high-molecularweight isoform of CD45, display a greater level of commitment for differentiation into lymphocytes (T, B and NK cells) than in the HSCs population (Galy et al., 1995a) and also seem to be more committed toward LCs than most progenitor cells as they contain CLA cells, the precursors for LCs (Strunk et al., 1997). In bone marrow, the CD34CD45RA progenitor cells are distinct from primitive HSCs phenotypically and functionally as they produce lymphocytes and myeloid cells (granulocytes and monocytes) but they are markedly depleted of erythroid progenitor cells, indicating the loss of some developmental options compared with HSCs (Craig et al., 1994; Galy et al., 1995a). This population of CD34CD45RA cells contains a common lymphoid progenitor (CLP) expressing CD10 but no lineage-associated markers (Lin), such as CD19, CD2, CD14, CD15, CD56 and glycophorin A. Such CLPs represent approximately 5% of progenitor cells in adult bone marrow. Their differentiation potential is restricted to the production of lymphocytes and DCs as they cannot produce myeloid cells (granulocytes or monocytes), erythrocytes, mast cells or platelets in spite of stimulation with multiple growth factors (Galy et al., 1995b). The ability of these CLPs to produce T cells rapidly (although not durably) and their overall differentiation potential into lymphocytes and DCs suggest that such cells could leave the bone marrow to colonize the thymus (Ardavin et al.,

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1993; Márquez et al., 1995; Res et al., 1996). Indeed, CD34LinCD10 cells are found in the human thymus and can be isolated from steadystate circulating blood to produce DCs in vitro (A. Galy, unpublished observations). It is not yet clear whether such CLPs are able to give rise to all of the types of thymic DCs which include interdigitating cells as well as plasmacytoid T-cell DCs (Res et al., 1999). Yet, the existence of this small progenitor cell population which contains clones of bipotential progenitors of lymphocytes and DCs strongly and directly argues for a tight developmental link between lymphocytes and DCs. Interestingly, the origin of DCs may be less of a determinant of their phenotype or function than previously thought. This is evidenced by recent studies showing that CD8α DCs as well as CD8α DCs can be produced from CMPs (Traver et al., 2000 and Wu et al. manuscript submitted) suggesting that the expression of CD8α does not delineate the ultimate lineage origin of DCs. An important area is to define what signals control the development of DCs.

SIGNALS REGULATING THE HEMATOPOIETIC DEVELOPMENT OF DCs Functions of the transcription factor Ikaros family in DC hematopoiesis Several signaling pathways, in which Ikaros is differentially involved, regulate DC development in vivo and in vitro. The Ikaros gene family was first implicated in DC hematopoiesis by studies in mutant mice. Ikaros is abundantly expressed in lymphoid tissues and encodes for a family of Kruppel-like zinc finger DNA-binding proteins. Potential Ikaros-binding sequences have been identified in many T cell- and B cellassociated genes such as the promoter and enhancer regions of CD3γ, δ and ε, the TCR α and β genes, the CD4 silencer, in the NFκB sites of the IL-2Rα, interferon β and MHC class II genes, in the HIV-LTR, in the LYF element of the TdT promoter, the EBF sites of the Igα promo-

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ter, and in the promoters of granzyme B, B29, TNFR p75 and BP-1 (Wargnier et al., 1995; Babichuk et al., 1996; Molnar et al., 1996; Santee and Owen-Schaub, 1996; Thompson et al., 1996). Mice homozygous for an Ikaros null allele lack B and NK lymphoid cell development and display specific alterations in T-cell development and a strong reduction in numbers of DCs in lymphoid organs (Georgopoulos et al., 1994; Wu et al., 1997). Deletions of the DNA encoding the Ikaros DNA-binding domain from the mouse germline that generate an Ikaros mutation with dominant negative properties (DN/) cause more serious lymphoid and DC defects (Wu et al., 1997). Proteins produced by the dominant negative locus interact and interfere with proteins produced by the wild-type Ikaros locus or with other family members, and compromise their activity (Sun et al., 1996; Morgan et al., 1997; Kelley et al., 1998). Hematopoietic defects in DN/ animals include a severe block in lymphopoiesis and a general depletion of DCs in lymphoid organs although monocytes and skin LCs are abundantly present, suggesting that several signaling pathways, in which Ikaros is differentially involved, regulate DC development in vivo. The human equivalent of Ikaros is highly homologous to its murine counterpart with almost complete identity in the DNA-binding region and protein interaction domains (Molnar et al., 1996). Ikaros mRNA is detectable in human CD34 cells (Galy et al., 1998) and murine dominant negative proteins interfere with the normal function of Ikaros proteins in human cells. One dominant negative Ikaros protein, Ik7, is the product of a gene targeting deletion of exons 3 and 4 which causes a strong reduction in the DNA-binding ability of heterocomplexes formed between Ik7 and other members of the Ikaros family of proteins through their C-terminal zinc finger modules (Sun et al., 1996). When the dominant negative Ik7 protein was overexpressed in human hematopoietic cells that were cultured in conditions promoting the development of

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lymphoid progenitor cells into DCs but not allowing differentiation of monocytes into DCs, the production of these lymphoid-related DCs was severely blocked by Ik7 but the formation of monocytes/macrophages was not (Galy et al., 2000). In contrast, DCs were produced when Ik7-expressing progenitor cells were cultured in conditions enabling DCs to be produced from monocytes. Thus, distinct signals that differentially implicate members of the Ikaros family of proteins control the formation of DCs. One of the molecular mechanisms by which the DN Ik7 protein blocks DC hematopoiesis appears to be downregulation of the Flt-3 receptor mRNA as shown in human progenitor cells (Galy et al., 2000). The downregulation of the murine homologue Flk-2 has also been described in progenitor cells of mutant DN/ mice (Nichogiannopoulou et al., 1999). This tyrosine kinase receptor, which is expressed in primitive hematopoietic cells and in mature cells such as lymphocytes, monocytes and DCs, plays a role both in multipotent stem cells and in lymphoid differentiation (Mackarehtschian et al., 1995). Its ligand (FL) exerts proliferative effects on progenitor cells in vitro in synergy with other cytokines. Importantly, FL has a dramatic effect in vivo on the growth of DCs (Maraskovsky, 1996; Lyman, 1998), thus the disruption of Flt-3/Flk-2-mediated signals in hematopoietic progenitor cells is likely to impair DC formation. The role of Ikaros proteins is not yet fully understood but there is evidence that they have functions in addition to their role as direct transcriptional activators. In lymphocytes, Ikaros proteins are localized in discrete heterochromatin regions (Brown et al., 1997). In these higher order chromatin structures Ikaros proteins associate with other protein complexes that are involved in chromatin remodeling and histone deacetylation (Kim et al., 1999) and control cell cycle transition and DNA replication in lymphocytes (Avitahl et al., 1999). Further identification of genes and proteins regulated by Ikaros proteins is likely to bring a better understanding of the mechanisms that control

hematopoietic cell development in general and DC development in particular.

Function of transcription factor RelB in DC development The family of NFκB/Rel transcription factors plays a pivotal role in the regulation of immunological processes, as demonstrated by the phenotypes of mice with gene mutations in different members of this family (Burkly et al., 1995; Weih et al., 1995; Franzoso et al., 1997, 1998). RelB, a member of this transcription factor family, is found to be expressed in lymphoid tissues. High levels of RelB protein expression are found in the nucleus of interdigitating DCs (Carrasco et al., 1993; Feuillard et al., 1996). Increase in protein expression and translocation of RelB protein from cytoplasm to nucleus are correlated with DC activation, maturation and function (Pettit et al., 1997; Clark et al., 1999; Ammon et al., 2000; Neumann et al., 2000). This suggests that RelB plays a role in DC development and function. Indeed, RelB-deficient mice display a lack of thymic DCs and impaired splenic antigen-presenting cell function and cellular immunity (Burkly et al., 1995; Weih et al., 1995). More detailed studies on these mutant mice revealed that the subset of splenic mature DCs lacking CD8α expression, the putative myeloid-related DCs, was severely reduced in these mice, whereas the CD8α DCs, the putative lymphoid-related DCs, were not directly affected by the mutation of RelB (Wu et al., 1998). Thus, the development of CD8α DCs, is dependent on RelB function. Although the molecular mechanism is not yet clear, these observations demonstrate a differential requirement for RelB in the development of different DC types, and again provide strong evidence supporting the concept that different DC subsets require different molecular signals for their development. It is interesting to note that although Ikaros family members and RelB are required for the development of either lymphoid- or myeloidrelated DC lineages, the skin Langerhans cells were present in both gene knockout mice. This

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indicates that the development of Langerhans cells, at least to the stage of their residence in the epidermis, does not require either Ikaros or RelB. This observation again raises the question of the lineage origin of Langerhans cells. This issue currently remains controversial. Further aspects of LC development will be discussed in subsequent chapters of this book.

2000). This suggested an indirect effect of PU.1 mutation on thymic DC development. The overall results from these studies indicate that PU.1 is essential for the development of myeloidderived DCs but not for lymphoid-related DCs.

Function of transcription factor PU.1 in DC development

The production of each of the different lineages of blood cells, including that of DCs, involves the differentiation of HSCs into intermediate cells with increasingly narrow differentiation potential (Plate 1.1). The identification of common lymphoid progenitor cells (Galy et al., 1995b; Kondo et al., 1997) and more recently of common myeloid progenitor cells in the mouse (Akashi et al., 2000) shows that hematopoietic cell fate specification occurs incrementally rather than all at once. Thus, common molecular programs are used for the development of groups of cells in related lineages. In that respect it is clear that some DCs and lymphocytes can develop very closely and share requirements for common molecules such as Ikaros proteins. However, the existence of other DCs that originate from myeloid precursors or monocytes, suggests that separate lineages of DCs exist having either a lymphoid-related origin or a myeloid origin. Undoubtedly, specific signals that regulate the development of each of these lineages of hematopoietic cells must specify important characteristics in the DC progeny. The purpose of DC heterogeneity at this level might be that the existence of separate lineages of DCs is justified by the need for specific and nonredundant functions. In favor of this argument, it has been recently shown that distinct mouse DC subsets have different influences on the type of immune response generated in vivo (Maldonado-Lopez et al., 1999; Pulendran et al., 1999). This appears to be the case in humans as well. Human plasmacytoid DCs which have been postulated to represent a type of lymphoidrelated DCs (Res et al., 1999) exhibit unique immune functions compared with monocytederived DCs (Rissoan et al., 1999). The immune function of human lymphoid-related DCs is

The transcription factor PU.1, a member of the ets family of DNA-binding proteins, is restricted in expression to hematopoietic cells (Klemsz et al., 1990; Hromas et al., 1993) and can regulate the expression of many myeloid and lymphoid genes (Tenen et al., 1997; Lloberas et al., 1999). The importance of PU.1 in the development of multiple hematopoietic cell lineages was previously revealed by studying mice with PU.1 gene targeted mutations. PU.1 is required for the development of both B lymphoid cells and macrophages (Scott et al., 1994, 1997; McKercher et al., 1996; Anderson et al., 1998; Spain et al., 1999). The importance of PU.1 in DC development has also been reported recently. PU.1 was shown to be expressed by mouse bone marrow-derived DCs and by human blood monocyte-derived DC (Anderson et al., 2000). The PU.1 mutant mice lack thymic DCs and mature macrophages in fetal or neonatal mutant thymus. Cells with a phenotype of lymphoid-related DCs (CD8αDEC-205) could be detected, whereas the myeloid-related CD8αDEC-205 DC subset was absent in these mice (Anderson et al., 2000; Guerriero et al., 2000). The PU.1-deficient hematopoietic progenitors lack the capacity of generating DCs in culture under conditions supporting the development of myeloid DCs (Anderson et al., 2000; Guerriero et al., 2000). Although thymic DC development was not detectable by histological staining for CD11c cells in the early embryonic thymus, when thymic lobes were placed into organ culture for 10 days, equal levels of CD11c and DEC-205 staining could be detected in both PU.1/ and wild-type thymus (Guerriero et al.,

CONCLUSION

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still unclear but it seems that DCs derived from CLPs provide T-cell stimulation signals distinct from monocyte-derived DCs (Wesa and Galy, 2001). Alternatively, it is also possible that separate lineages of DCs exist as a safeguard to provide redundant formation of such important immune cells, or to permit more versatile regulatory mechanisms as the different cells develop in distinct microenvironments. Further work is needed to understand the relative importance of these DC pathways.

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Galy, A.H.M., Cen, D., Travis, M. et al. (1995a). Blood 85, 2770–2778. Galy, A., Travis, M., Cen, D. and Chen, B. (1995b). Immunity 3, 459–473. Galy, A., Morel, F., Hill, B. and Chen, B.P. (1998). J. Immunother. 21, 132–141. Galy, A., Christopherson, I., Ferlazzo, G. et al. (2000). Blood 95, 128–137. Geissmann, F., Prost, C., Monnet, J.P. et al. (1998). J. Exp. Med. 187, 961–966. Georgopoulos, K., Bigby, M., Wang, J.H. et al. (1994). Cell 79, 143–156. Guerriero, A., Langmuir, P.B., Spain, L.M. and Scott, E.W. (2000). Blood 95, 879–885. Hart, D.N. (1997). Blood 90, 3245–3287. Hromas, R., Orazi, A., Neiman, R.S. et al. (1993). Blood 82, 2998–3004. Inaba, K., Inaba, M., Deguchi, M. et al. (1993). Proc. Natl Acad. Sci. USA 90, 3038–3042. Jaksits, S., Kriehuber, E., Charbonnier, A.S. et al. (1999). J. Immunol. 163, 4869–4877. Kelley, C.M., Ikeda, T., Koipally, J. et al. (1998). Curr. Biol. 8, 508–515. Kim, J., Sif, S., Jones, B. et al. (1999). Immunity 10, 345–355. Klemsz, M.J., McKercher, S.R., Celada, A. et al. (1990). Cell 61, 113–124. Kondo, M., Weissman, I.L. and Akashi, K. (1997). Cell 91, 661–672. Kronin, V., Winkel, K., Suss, G. et al. (1996). J. Immunol. 157, 3819–3827. Lloberas, J., Soler, C. and Celada, A. (1999). Immunol. Today 20, 184–189. Lucas, K., Vremec, D., Wu, L. and Shortman, K. (1998). Dev. Comp. Immunol. 22, 339–349. Lyman, S.D., and Jacobsen, S.E. (1998). Blood 91, 1101–1134. Mackarehtschian, K., Hardin, J.D., Moore, K.A. et al. (1995). Immunity 3, 147–161. Maldonado-Lopez, R., De Smedt, T., Michel, P. et al. (1999). J. Exp. Med. 189, 587–592. Maraskovsky, E., Brasel, K., Teepe, M. et al. (1996). J. Exp. Med. 184, 1953–1962. Márquez, C., Trigueros, C., Fernandez, E. and Toribio, M.L. (1995). J. Exp. Med. 181, 475–483. McKercher, S.R., Torbett, B.E., Anderson, K.L. et al. (1996). EMBO J. 15, 5647–5658. Molnar, A., Wu, P., Largespada, D.A. et al. (1996). J. Immunol. 156, 585–592. Morgan, B., Sun, L., Avitahl, N. et al. (1997). EMBO J. 16, 2004–2013. Neumann, M., Fries, H., Scheicher, C. et al. (2000). Blood 95, 277–285. Nichogiannopoulou, A., Trevisan, M., Neben, S. et al. (1999). J. Exp. Med. 190, 1201–1214. Olweus, J., BitMansour, A., Warnke, R. et al. (1997). Proc. Natl Acad. Sci. USA 94, 12551–12556.

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Pettit, A.R., Quinn, C., MacDonald, K.P. et al. (1997). J. Immunol. 159, 3681–3691. Pulendran, B., Smith, J.L., Caspary, G. et al. (1999). Proc. Natl Acad. Sci. USA 96, 1036–1041. Res, P., Martinez-Caceres, E., Cristina Jaleco, A. et al. (1996). Blood 87, 5196–5206. Res, P.C., Couwenberg, F., Vyth-Dreese, F.A. and Spits, H. (1999). Blood 94, 2647–2657. Rissoan, M.C., Soumelis, V., Kadowaki, N. et al. (1999). Science 283, 1183–1186. Romani, N., Gruner, S., Brang, D. et al. (1994). J. Exp. Med. 180, 83–93. Ryan, D.H., Nuccie, B.L., Ritterman, I. et al. (1997). Blood 89, 929–940. Sallusto, F. and Lanzavecchia, A. (1994). J. Exp. Med. 179, 1109–1118. Santee, S.M. and Owen-Schaub, L.B. (1996). J. Biol. Chem. 271, 21151–21159. Saunders, D., Lucas, K., Ismaili, J. et al. (1996). J. Exp. Med. 184, 2185–2196. Scott, E.W., Simon, M.C., Anastasi, J. and Singh, H. (1994). Science 265, 1573–1577. Scott, E.W., Fisher, R.C., Olson, M.C. et al. (1997). Immunity 6, 437–447. Spain, L. M., Guerriero, A., Kunjibettu, S. and Scott, E.W. (1999). J. Immunol. 163, 2681–2687. Steinman, R.M. (1991). Annu. Rev. Immunol. 9, 271–296. Strunk, D., Egger, C., Leitner, G. et al, (1997). J. Exp. Med. 185, 1131–1136. Sun, L., Liu, A. and Georgopoulos, K. (1996). EMBO J. 15, 5358–5369. Szabolcs, P., Avigan, D., Gezelter, S. et al. (1996). Blood 87, 4520–4530.

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Tenen, D.G., Hromas, R., Licht, J.D. and Zhang, D.E. (1997). Blood 90, 489–519. Thompson, A.A., Wood, W.J., Jr., Gilly, M.J. et al. (1996). Blood 87, 666–673. Traver, D., Akashi, K., Manz, M. et al. Science 290, 2152–2154. Vremec, D. and Shortman, K. (1997). J. Immunol. 159, 565–573. Vremec, D., Zorbas, M., Scollay, R. et al. (1992). J. Exp. Med. 176, 47–58. Wargnier, A., Legros-Maida, S., Bosselut, R. et al. (1995). Proc. Natl Acad. Sci. USA 92, 6930–6934. Weih, F., Carrasco, D., Durham, S.K. et al. (1995). Cell 80, 331–340. Wesa, A. and Galy, A. (2001). Cell Immunol. 208, In press. Wu, L., Scollay, R., Egerton, M. et al. (1991a). Nature 349, 71–74. Wu, L., Antica, M., Johnson, G.R. et al. (1991b). J. Exp. Med. 174, 1617–1627. Wu, L., Vremec, D., Ardavin, C. et al. (1995). Eur. J. Immunol. 25, 418–425. Wu, L., Li, C.-L. and Shortman, K. (1996). J. Exp. Med. 184, 903–911. Wu, L., Nichogiannopoulou, A., Shortman, K. and Georgopoulos, K. (1997). Immunity 7, 483–492. Wu, L., D’Amico, A., Winkel, K. D. et al. (1998). Immunity 9, 839–847. Young, J.W., Szabolcs, P. and Moore, M.A.S. (1995). J. Exp. Med. 182, 1111–1120. Zhou, L.J. and Tedder, T.F. (1996). Proc. Natl Acad. Sci. USA 93, 2588–2592.

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PLATE 1.1 Hypothetical schema for the specification of hematopoietic cell fate and critical requirement for transcription factors in DC hematopoiesis. Hematopoietic stem cells (HSC) differentiate into multipotential progenitor cells (MP) able to give rise to megakaryocytes (Meg), erythrocytes (E), granulocytes (G), monocytes (M), lymphocytes (L) and cells of the DC system. The existence of lineage-restricted progenitor cells has been demonstrated and segregates cells of the lymphoid lineage or cells of the myeloid lineage into separate developmental pathways. DCs appear to be myeloid- or lymphoid-related as they derive in close association with either lineage. The transcription factor Ikaros is essential for the formation of all lymphocytes and for some DCs, and is probably critical at least at the stage of common lymphoid/DC precursor cells. The transcription factor RelB is expressed at high levels in putative myeloid-derived DCs and is essential for their formation but is not required for the development of putative lymphoid-related DCs. The transcription factor PU-1 is essential for the formation of B cells, monocyte/ macrophages and is required only for the formation of putative myeloid-related DCs. Thus specific molecular events provide distinct cues in developing hematopoietic cells that control the formation of separate populations of DCs.

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C

H

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A

P

T

E

R

2 Dendritic cells in the thymus Hergen Spits Division of Immunology, Netherlands Cancer Institute, Amsterdam, The Netherlands

What does not destroy me, makes me strong. Friedrich Nietzche

INTRODUCTION

are removed. Induction of negative selection requires interaction of bone marrow-derived antigen-presenting cells with immature T cells. This process takes place mostly in the medulla, although in some experimental models negative selection has been shown to occur in the cortex (Surh and Sprent, 1994; Sprent and Webb, 1995). Since DCs are located in the medulla, it was suggested that DCs mediate negative selection in vivo. This was directly shown by Brocker and collaborators. They have generated a transgenic mouse model in which gene expression was targeted to DC using the CD11c promoter. I-E expression on thymic DCs was sufficient to negatively select I-E-reactive CD4 T cells, and to a less complete extent, CD8 T cells (Brocker et al., 1997). In contrast, when DCs expressed I-E in a class II-deficient background, positive selection of CD4 T cells could not be observed. Thus negative, but not positive, selection events can be induced by DCs in vivo. The other non-T cells found in the thymus are NK and B cells and macrophages. Macrophages

T cells mostly develop in the thymus. This organ delivers mature T cells to the periphery at a high rate during birth and early in life, and continues to do so later in life although at steadily declining rates. To be able to deliver the mature T cells, the thymus is continually populated with uncommitted precursor cells derived from bloodborne, bone marrow-derived progenitor cells. Within the thymus these progenitor cells go through a series of differentiation, expansion and maturation stages (Spits et al., 1995, 1998; Shortman and Wu, 1996; Shortman et al., 1998). Both hematopoietic and nonhematopoietic cells are essential for proper T-cell development. Dendritic cells (DCs) are found along with macrophages and B cells within the thymus. A considerable body of evidence indicates that DCs mediate negative selection (Sprent and Webb, 1995; Brocker, 1999). This is a process whereby developing T cells that express a T-cell receptor (TCR) with specificity for self antigens Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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remove dying cells in the thymus, including the negatively selected T cells as well as T cells that do not pass the positive selection step (Surh and Sprent, 1994). There is no evidence for a role of NK and B cells in thymic T-cell development, as some mouse strains lack B or NK cells but have normal T-cell numbers. DCs are highly efficient antigen-presenting cells that play a key role in inducing and regulating efficient immune responses (Banchereau and Steinman, 1998). They collect foreign antigens at the interface with the external environment and move to the lymph nodes where the antigenloaded DCs induce a T-cell response. It is undesirable that DCs that carry foreign antigens would be able to migrate into the thymus, as this would result in deletion of T cells recognizing foreign antigens. Internal development of DCs from precursors that enter the thymus, combined with a limited lifespan of these cells would prevent such an unwanted deletion of T cells specific for foreign substances. Indeed, as discussed below, there is now considerable evidence for a thymic development of DCs from

TABLE 2.1

immigrated precursor cells both from studies in mice as well as in humans.

EARLY STAGES IN T CELL AND DC DEVELOPMENT IN MICE AND HUMANS The earliest stages of T cell and DC development in mice and humans have now been mapped by cell surface phenotype as well as with functional assays (Table 2.1) (Spits et al., 1995; Shortman and Wu, 1996). The developmental pathways in these two organisms are comparable with regard to the developmental potential of precursor cell subsets and the sequence of TCR gene rearrangements. However, widely different sets of cell surface antigens are used to identify particular developmental stages. The earliest precursors present in the adult mouse thymus express low levels of CD4 and are denoted as CD4lo precursors (Wu et al., 1991a, 1991b). They also express c-kit ligand, the receptor for stem cell factor (SCF) (Matsuzaki et al.,

Early stages in human and mouse thymocyte development Mouse

Human

Phenotype

Hematopoietic activity

Phenotype

Hematopoietic activity

CD44 CD25 CD4 c-kit  NK1.1

T, B, NK, DC (myeloid)

CD34 CD38dim CD33 CD1a CD5

T, B, NK, DC, myeloid

CD44 CD25 CD4 c-kit  NK1.1

T, NK, DC

CD34 CD38 CD1a CD5

T, NK, DC

Stage II

CD44 c-kit  CD25 NK1.1

T (NK, DC)

CD34 CD1a CD4 CD8

T (NK)

Stage III

CD44lo/ c-kit lo/ CD25

T

CD1a CD4 CD8α/

T

Stage I

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1993), high levels of CD44 and lack CD25. The CD4lo precursors have their TCR genes in the germline configuration and have the capacity to develop into several cell types including T, B, NK cells and DCs upon injection into irradiated recipients (Ardavin et al., 1993b; Matsuzaki et al., 1993). Myeloid cells are not found in the periphery of irradiated mice injected with CD4lo thymocytes or with c-kit thymic precursors (Matsuzaki et al., 1993). Together, these data strongly suggest that CD4lo precursors have a very poor if any myeloid-developing potential. However, CD4lo precursors can readily develop into DCs in vitro in media containing IL-3 (Saunders et al., 1996). Cloning efficiencies of 70% were observed. The downstream ckitCD44CD25 population lacking CD4 or CD8 has also the capacity to develop into T, B, NK cells and DCs. These cells are functionally indistinguishable from the CD4lo population, but develop with more rapid kinetics. An important question is whether the c-kitCD44CD25 population contains bipotential or multipotential precursors or consists of a mixture of precursors committed to a particular lineage. Durum and coworkers established clones of CD44CD25 thymic precursors in IL-7 and SCF (Lee et al., 1999). They demonstrated that such clones could develop into both T and NK cells in a fetal thymic organ culture (FTOC), indicating that the adult thymus does contain bipotential T/NK precursors. More recently, Ikawa et al. (1999) used an elegant singlecell FTOC system to confirm that CD44CD25 thymocytes contain bipotential T/NK precursors that are unable to develop into B cells. Unfortunately in none of these studies was the DC precursor activity determined, leaving open the question as to whether and how many of these T/NK precursors have the capacity to develop into DCs. The stage that follows the CD44CD25 stage expresses CD25. Wu et al. (1996) reported that these cells have DC precursor activity, but the frequency of DC precursors is 3-fold less than that of CD4lo cells. Another group reported that this population also contains NK precursors (Moore and Zlotnik, 1995). No single-cell cloning experiments have been

15

performed and it is therefore unknown whether single CD44CD25 cells can give rise to T cells and DCs but not to NK cells. Thus, whether T cells and DCs are more closely related to each other than to NK cells is an unresolved issue. In the human thymus the earliest precursors can be found in a population that expresses CD34, high levels of CD7 and CD44 and lacks CD1a (Galy et al., 1993; Schmitt et al., 1993; Márquez et al., 1995). In the fetal thymus more primitive precursor cells (weeks 16–18 of gestation) have been found within this population (Sánchez et al., 1993; Res et al., 1996). These primitive CD34 cells express relatively low levels of CD38, are CD5lo and express CD33. In thymus samples from young children, there are CD34CD33 early precursor cells but CD34 cells that lack CD5 are almost undetectable. As in the mouse, it has not been firmly established that the thymus contains tripotential precursors that are able to develop into T, NK cells and DCs. We have performed single-cell analyses of fetal CD34CD38lo precursor cells, that are also CD5lo and CD33 in medium that contain either cytokines favoring DC development (GM-CSF and TNFα) or NK cell development (IL-2, IL-7, GM-CSF and SCF) (Res et al., 1996). We found that the sum of NK cells and DC cloning efficiencies exceeded 100, indicating that this population contains bipotential NK/DC clones. Unfortunately, no assays exist to test the T-cell developmental potential of single precursor cells in the human system. Sánchez et al. (1994) used an indirect approach to show the existence of pre-T/NK clones within the CD34CD5lo population (which is also CD38lo). These investigators sorted single CD34CD5lo thymocytes and cultured these cells for 5 days in a mixture of IL-2, IL-7 and SCF. The best growing single cellderived cultures (containing up to 25 cells) were split; one half was cultured further in IL-2, IL-7 and SCF and the rest of the cultures were pooled and tested in a FTOC. Virtually all the cytokineassisted single cell-derived cultures developed into NK cells (Sánchez et al., 1994). In addition, T cells were found to develop in the FTOC (Sánchez et al., 1994). This experiment proved

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the existence of bipotential T/NK precursors. T and NK cells are much more similar to each other than to mature DCs. Intuitively, a developmental scheme in which a tripotential T/NK/DC precursor precedes a bipotential T/NK precursor appears, most plausible. Indeed, investigating the kinetics of T-cell development of fetal liver progenitor cells in an FTOC system, Plum et al. (1999) observed that development of DCs preceded that of NK cells. They concluded, therefore, that DCs branch off before the T/NK split. It is, however, important to note that so far there is no direct proof for the existence of a tripotential T/NK/DC precursor. An alternative model has been proposed by Toribio and collaborators (Márquez et al., 1998). They have investigated the developmental relationship of T, NK cells and DCs using postnatal rather than fetal thymus. CD34CD33 precursors were cultured for 3 days in a cytokine mixture of IL-7, GM-CSF, IL-1 and SCF and subjected to single-cell cloning in the same medium with or without IL-2. This experiment yielded NK clones in medium with IL-2 and DC clones in medium without IL-2. The sum of the cloning efficiencies was more than 100, indicating the existence of bipotential NK/DC precursor cells in the cultured CD34CD33 thymocytes. These cultured CD34CD33 cells had lost T-cell characteristics such as CD5 and CD7. Márquez et al. (1998) also identified a precursor cell with the phenotype of cultured CD34CD33 cells in situ. Based on these findings, the authors proposed that NK cells and DCs develop from a common, bipotential precursor that has lost T-cell precursor activity. Verification of this interesting model requires testing the T-cell precursor activities of cultured CD34CD33 and freshly isolated CD34loCD33CD44hi cells. Summarizing the data in both human and mouse: in the thymus of both species T, NK cells and DC developmental activities have been found in the primitive precursor compartment. The existence of a bipotential T/NK precursor has been directly demonstrated and there is indirect evidence for a common DC/NK precursor in the human thymus. The presence of

tripotential T/NK/DC precursors in the thymus has not been directly demonstrated.

CHARACTERISTICS OF THYMIC DCs IN THE HUMAN THYMUS As discussed in the preceding section, CD34CD1a precursors can differentiate into DCs upon culture in GM-CSF, TNFα, IL-7 and SCF. These in vitro generated DCs express CD1a and several myeloid markers such as CD13 and CD33 (Márquez et al., 1995, 1998; Res et al., 1996). However, it has been reported that CD1a is not typically present on human thymic DCs in situ (Lafontaine et al., 1992). This observation raises the question whether the DCs generated in vitro with GM-CSF-containing media are the same as the thymic DCs in situ. Therefore, we performed an extensive characterization of DClike cells freshly isolated from thymic tissue (Res et al., 1999). Previously it was shown that human thymic cells resembling DCs express CD4 and are negative for CD8α. Phenotypic analysis of the CD4CD8α thymocytes revealed two populations: one that express CD1a and lacks CD45RA and another one that expresses CD45RA and lacks CD1a (Res et al., 1999). The CD1aCD45RA cells are committed T-cell precursors, as these cells have their TCR genes rearranged and are able to differentiate into T cells in an FTOC but not to NK cells (Blom et al., 1999). The CD1aCD4 cells however, were, unable to differentiate into T cells (Res et al., 1999). The CD1aCD4 cells express high levels of CD123 (IL-3Rα). In the presence of IL-3 or IL-3 and CD40L they differentiate into cells with dendritic extensions that express high levels of HLA-DR and CD54 (ICAM-1). Moreover, the cells acquire CD80 and CD83 and upregulate CD86 upon culture in IL-3 plus CD40L. The CD1aCD3CD4 thymocytes are therefore DC precursors (pDC). We employed the high level of expression of CD123 to analyze the localization of the pDC

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in the thymus. It was found that these cells are present mainly in the thymic medulla. Furthermore immunohistology revealed that the CD123hi cells express CD4, CD45RA and lack CD1a, consistent with the phenotype determined with the fluorescence-activated cell sorter (FACS) analysis. We also found mature DCs in the thymus as indicated by the presence of CD83 cells in the medulla. These mature DCs are distinct, however, from the CD123hi cells, as a confocal laser scan analysis of thymic sections demonstrated that the great majority of the CD123hi cells did not coexpress CD83 (Res et al., 1999). The CD83 cells coexpressed CD86 as expected. Interestingly, the phenotype of thymic CD3CD4 pDCs is similar to that of CD3CD4 cell types found in the tonsil and peripheral blood of adults (Grouard et al., 1997; Rissoan et al., 1999). They are all CD1a, CD11c and CD45RA and share expression of CD123 (Table 2.2). It seems likely, therefore, that the CD3CD4 cell populations in thymus and periphery represent the same pDC type. There are differences with respect to expression of the lymphoid markers CD2, CD5 and CD7 (Grouard et al., 1997; Res et al., 1999) The latter antigen is expressed on the thymic cells but not on those in peripheral blood and tonsil. CD2 and CD5 are TABLE 2.2 Phenotype of CD3CD4CD8 human thymic subpopulations and DC precursors from tonsil and peripheral blood

17

present on all thymic cells but only on subsets of peripheral blood and tonsillar CD3CD4 cells. These differences in expression of CD2, CD5 and CD7 could be caused by the different microenvironments. It is, however, also possible that thymic and peripheral pDCs represent different subsets. Liu and collaborators reported that the mature DCs, generated from CD123hi pDCs in IL-3 and CD40L, stimulated naïve T cells to produce TH2 cytokines (IL-4 and IL5) (Rissoan et al., 1999). In contrast, DCs generated from monocytes with GM-CSF and IL-4 stimulated naïve T cells to become TH1 cells that fail to produce IL-4 and IL-5. Based on these functional differences Liu proposed to name the CD123hi pDCs as pDC2 and monocytes as pDC1 (Rissoan et al., 1999). Another group characterized a CD3CD4 cell type that expresses the antigen ILT-3, an antigen related to NK inhibitory receptors (Cella et al., 1999). Despite the fact that these cells closely resembled the pDC2 described by Liu and collaborators, Cella et al. reported that they generated TH0 cells from naïve T cells (Cella et al., 1999). A discussion of the possible reasons for the different results obtained by these two groups falls outside the scope of this chapter. Both groups reported that the pDC2 have a function in their own right. They produce exceptional high levels of type 1 interferons (IFNα and β) upon stimulation with virus (HSV-1 and influenzavirus) (Cella et al., 1999; Siegal et al., 1999). The observation that a cell population highly enriched for thymic CD11c pDC also produces IFNα following stimulation with HSV-1 (B. Blom and Y.-J. Liu, personal communication) is consistent with the notion that pDC2 and the thymic pDC are similar.

CD4 immature single-positive pre-T cells

Thymic pDC

pDC2 (tonsil, PBMC)

CD1a CD45RA

 

 

 

CD13 CD14 CD40 CD80 (B7.1) CD86 (B7.2) IL-3Rα

     

     

     

DEVELOPMENTAL ORIGIN OF CD1aCD3CD4 THYMIC pDCs

CD2 CD5 CD7 pTαRNA

   

   

   

As discussed above, in the mouse it has been demonstrated that thymic DCs originate from thymic precursors and are more closely related to lymphocytes than to myeloid cells. By analogy

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with the mouse the thymic CD123hi pDC could be of lymphoid origin. Several characteristics of these cells are consistent with this notion. In the first place, the thymic pDC express ‘lymphoid’ markers as CD2, CD5 and CD7 and lack typical ‘myeloid’ markers, such as CD11c, CD13 and CD33. Moreover, DCs generated from these pDCs lack CD13 and CD33 and have only a low level of expression of CD11c. In contrast, these antigens are abundantly expressed on DCs generated from monocytes which can be considered to be myeloid-derived DCs. More importantly, the thymic CD123hi pDCs expressed transcripts of pTα. This polypeptide forms, together with the TCRβ chain, the pre-T-cell receptor (preTCR). This receptor selects developing T cells that have undergone productive, in-frame, TCRβ gene rearrangements, as these cells will receive a proliferation and differentiation signal through the pre-TCR (Fehling et al., 1995; von Boehmer and Fehling, 1997; von Boehmer et al., 1998). Pre-T cells that fail to rearrange their TCRβ genes productively are unable to form a pre-TCR complex and they will not receive a signal and die. Because of the essential role of the pTα in T-cell development, this polypeptide was considered T cell-specific. Our data indicate that this is not the case (Res et al., 1999). Nonetheless, the fact that pTα transcripts are present in thymic DC and T-cell precursors but have not been detected in NK or B-cell precursors does suggest a common developmental origin of thymic DCs and T cells. The function of the pTα in thymic pDCs is not yet known. To verify whether thymic CD34 precursor cells are capable of developing into CD123hi DC precursors, it was important to develop a differentiation assay. We have found that the murine stromal cell line S17 not only supports development of human precursor cells into B cells (Rawlings et al., 1995; Jaleco et al., 1999) but also into CD123hi DC precursors (Spits et al., 2000). Using this assay we have shown that CD34CD1a thymocytes are capable of developing into CD123hiCD45RA cells upon culture with S17 cells, whereas CD34CD1a cells are unable to develop into pDCs. These data indicate that in principle the thymic pDCs can

develop within the organ. To investigate whether the thymic microenvironment can support development of pDC precursors, we injected carboxyfluorescein diacetate succinimidyl ester (CFSE)-labeled CD34CD38 CD123 fetal liver precursors into a human thymus graft of transplanted SCID mice. One week after injection, CFSE cells expressing high levels of CD123 were clearly detectable (H. Spits et al., manuscript in preparation). These cells coexpressed CD4. Our data together indicate that CD123hi thymic pDCs can develop from primitive precursors introduced in a ‘natural’ thymic microenvironment.

ARE THERE TWO TYPES OF DC IN THE THYMUS? Ardavin and colleagues have shown that the murinethymuscontainstwotypesofDC-likecells (Ardavin et al., 1993a, 1997). These two DC types can be distinguished on the basis of expression of MHC class II antigens: one expresses high levels and the other low levels of MHC class II antigens. Both DC types express CD8α and another DC marker, DEC-205, that is not expressed on CD8α splenic DCs. There are also morphological differences between these subsets: MHC class IIlo cells are round while the MHC class IIhi cells are irregular-shaped veiled cells. Upon intrathymic injection of CD4lo precursors these two subsets appear sequentially, the MHC class IIlo subset appearing earlier than the MHC class IIhi subset. It is therefore possible that these two subsets have a precursor/progeny relationship. It would be of interest to determine whether the mouse MHC class IIlo DC population is equivalent to the human CD123hi pDCs which also have a relatively low expression of MHC class II antigens. Recently, Rodewald et al. (1999) reported the presence of CD11cloCD45 DCs which express Mac-1 (CD11b) in the thymus of wild-type mice and of mice deficient for the γc and c-kit. As Mac-1 is a myeloid marker, the data of Rodewald suggest that the murine thymus may contain myeloid-related DCs in addition to lymphoidrelated DCs.

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REFERENCES

Human CD34CD1a thymic precursors also have the capacity to develop into two types of DC, suggesting that different DC subsets can exist in human thymus. Upon culture in cytokine mixtures, DCs are generated that express high levels of CD13 and CD33 (Márquez et al., 1995, 1998). Many of these cells also express CD1a. When the CD34CD1a cells are co-cultured with the S17 stromal cells, some of the cells develop into CD123hi DC precursors that develop into mature DC following culture with IL-3 and CD40L (H. Spits et al., manuscript in preparation). These two DC types are clearly different with respect to phenotype and requirements for their in vitro development. The presence of CD123hi pDCs in the medulla has been firmly established. However, it remains to be investigated whether mature thymic DCs are generated through the ‘GM-CSF’ pathway or via the CD123hi pDC stage or whether there are two types of DC in the thymus. A careful examination of the mature DC compartment in the human thymus will be necessary to reveal a possible heterogeneity in thymic DCs. Taken together, there is evidence both in human and mouse systems for the presence of different subsets of DCs in the thymus. It is now well established that thymic DCs can mediate negative selection. If indeed the thymus contains more that one DC type, it would be of interest to know whether they both mediate negative selection. One may speculate about other functions that DCs might have within the thymus. It might be possible that a certain DC subset or its precursor is involved in formation of the thymic organ. It has been proposed that a fetal CD3CD4 DC precursor is necessary for the formation of Peyer’s patches and mesenterial lymph nodes in the mouse (Mebius et al., 1997). Another possible function is the maintenance of of single positive thymocyte survival after they have been positively selected and before they exit from the thymus. Note: A recent report by the Shortman group presented conclusive evidence for the existence of two DC populations in the human thymus (Vandenabeele et al., 2001).

ACKNOWLEDGEMENTS I thank Drs Pieter Res, Bianca Blom, Christel Uittenbogaart, Florry Vyth and Kees Wijer and Franka Couwenberg, Trees Dellemijn and Arie Voordouw for their contributions to the thymic DC project.

REFERENCES Ardavin, C. (1997). Immunol. Today 18, 350–361. Ardavin, C., Wu, L., Ferrero, I. and Shortman, K. (1993a). Immunol. Lett. 38, 19–25. Ardavin, C., Wu, L. and Shortman, K. (1993b). Nature 362, 761–763. Banchereau, J. and Steinman, R.M. (1998). Nature 392, 245–252. Blom, B., Verschuren, M.C., Heemskerk, M.H. et al. (1999). Blood 93, 3033–3043. Brocker, T. (1999). J. Leukoc. Biol. 66, 331–335. Brocker, T., Riedinger, M. and Karjalainen, K. (1997). J. Exp. Med. 185, 541–550. Cella, M., Jarrossay, D., Facchetti, F. et al. (1999). Nature Med. 5, 919–923. Fehling, H.J., Krotkova, A., Saint Ruf, C. and von Boehmer, H. (1995). Nature 375, 795–798. Galy, A., Barcena, A., Verma, S. and Spits, H. (1993). J. Exp. Med. 178, 391–402. Grouard, G., Rissoan, M.C., Filgueira, L., Durand, I., Banchereau, J. and Liu, Y.J. (1997). J. Exp. Med. 185, 1101–1111. Ikawa, T., Kawamoto, H., Fujimoto, S. and Katsura, Y. (1999). J. Exp. Med. 190, 1617–1626. Jaleco, A.C., Stegmann, A.P., Heemskerk, M.H. et al. (1999). Blood 94, 2637–2646. Lafontaine, M., Landry, D. and Montplaisir, S. (1992). Cell. Immunol. 142, 238–251. Lee, C.K., Kim, K., Geiman, T.M., Murphy, W.J., Muegge, K. and Durum, S.K. (1999). Cell. Immunol. 191, 139–144. Márquez, C., Trigueros, C., Fernández, E. and Toribio, M.L. (1995). J. Exp. Med. 181, 475–483. Márquez, C., Trigueros, C., Franco, J.M. et al. (1998). Blood 91, 2760–2771 Matsuzaki, Y., Gyotuku, J-I., Ogawa, M. et al. (1993). J. Exp. Med. 178, 1283–1292. Mebius, R.E., Rennert, P. and Weissman, I.L. (1997). Immunity 7, 493–504. Moore, T.A. and Zlotnik, A. (1995). Blood 86, 1850–1860. Plum, J., De Smedt, M., Verhasselt, B. et al. (1999). J. Immunol. 162, 60–68. Rawlings, D.J., Quan, S.G., Kato, R.M. and Witte, O.N. (1995). Proc. Natl Acad. Sci. USA 92, 1570–1574.

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Res, P., Martínez Cáceres, E., Jaleco, A.C., Noteboom, E., Weijer, K. and Spits, H. (1996). Blood 87, 5196–5206. Res, P.C., Couwenberg, F., Vyth-Dreese, F.A. and Spits, H. (1999). Blood 94, 2647–2657. Rissoan, M.C., Soumelis, V., Kadowaki, N. et al. (1999). Science 283, 1183–1186. Rodewald, H.R., Brocker, T. and Haller, C. (1999). Proc. Natl Acad. Sci. USA 96, 15068–15073. Sánchez, M.-J., Spits, H., Lanier, L. and Philips, J.H. (1993). J. Exp. Med. 178, 1857–1866. Sánchez, M.-J., Muench, M.O., Roncarolo, M.G., Lanier, L. and Phillips, J.H. (1994). J. Exp. Med. 180, 569–576. Saunders, D., Lucas, K., Ismaili, J. et al. (1996). J. Exp. Med. 184, 2185–2196. Schmitt, C., Ktorza, S., Sarun, S., Blanc, C., de Jong, R. and Debré, P. (1993). Blood 82, 3675–3685. Shortman, K. and Wu, L. (1996). Early T lymphocyte progenitors. Annu. Rev. Immunol. 14, 29–47. Shortman, K., Vremec, D., Corcoran, L.M., Georgopoulos, K., Lucas, K. and Wu, L. (1998). Immunol. Rev. 165, 39–46.

Siegal, F.P., Kadowaki, N., Shodell, M. et al. (1999). Science 284, 1835–1837. Spits, H., Lanier, L. and Phillips, J.H. (1995). Blood 85, 2654–2670. Spits, H., Blom, B., Jaleco, A.C. et al. (1998). Immunol. Rev. 165, 75–86. Spits, H. Couwenberg, F., Bakker, A.Q. et al. (2000). J. Exp. Med. 192, 1775–1784. Sprent, J. and Webb, S.R. (1995). Curr. Opin. Immunol. 7, 196–205. Surh, C.D. and Sprent, J. (1994). Nature 372, 100–103. Vandenabeele, S., Hochrein, H., Mavaddat, N. et al. (2001). Blood 97, 1733–1741. von Boehmer, H. and Fehling, H.J. (1997). Annu. Rev. Immunol. 15, 433–452. von Boehmer, H., Aifantis, I., Azogui, O. et al. (1998). Immunol. Rev. 165, 111–119. Wu, L., Antica, M., Johnson, G.R., Scollay, R. and Shortman, K. (1991a). J. Exp. Med. 174, 1617–1627. Wu, L., Scollay, R., Egerton, M., Pearse, M., Spangrude, G.J. and Shortman, K. (1991b). Nature 349, 71–74. Wu, L., Li, C.L. and Shortman, K. (1996). J. Exp. Med. 184, 903–911.

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3 T-cell activation and polarization by dendritic cells Yong-Jun Liu, Vassili Soumelis and Nori Kadowaki DNAX Research Institute, Palo Alto, California, USA

If a man loses pace with his companions, perhaps it is because he hears a different drummer. Let him step to the music in which he hears, however measured, or far away. Henry David Thoreau

INTRODUCTION

product rapidly induce DCs to produce large amounts of IL-12 suggests that DCs may not need a third cell type to polarize activated T cells toward TH1 effectors (Plate 3.2) (Macatonia et al., 1995; Cella et al., 1996; Koch et al., 1996; Sousa et al., 1997). The questions are: (1) Are there distinct types of DCs? (2) Do distinct types of DCs induce different types of immune responses, such as TH1 versus TH2 or immunity versus tolerance? (3) How do pathogen-induced innate immunity or cytokine microenvironment determine the functions of DCs? (4) Is there an IL-4-independent mechanism for the induction of TH2 differentiation? (5) What are the ideal DCs for tumor therapy or for treatment of autoimmune diseases and graft-versus-host diseases (GVHD)?

Dendritic cells (DCs) are professional antigenpresenting cells capable of activating naïve T helper (TH) cells (Cella et al., 1997; Banchereau and Steinman, 1998; Reise e Sousa et al., 1999). Differentiation of activated TH cells into IFNγproducing effector TH1 cells or IL-4, IL-5 and IL-10-producing effector TH2 cells depends respectively on cytokines such as IL-12 or IL-4, possibly produced by a third cell type (Macatonia et al., 1993; Abbas et al., 1996; Hilkens et al., 1997; O’Garra, 1998; Reis e Sousa et al., 1999). IL-12 produced by activated macrophages was believed to be critical for TH1 differentiation (Plate 3.1) (Macatonia et al., 1993; Hilkens et al., 1997; O’Garra, 1998). Because IL-4 is a TH2 prototype cytokine, the original source of IL-4 required for TH2 differentiation has been controversial (Abbas et al., 1996; O’Garra, 1998). During early cognate DC–T cell interaction, activated T cells rapidly express the T-cell activation antigen, CD40 ligand. The finding that CD40 ligand and microbial Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

DC LINEAGE AND FUNCTION IN MICE The concept of tolerogenic dendritic cells (DCs) came from experiments in mice showing that

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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thymic DCs mediated clonal deletion of emerging autoreactive T cells within the thymus (Brocker et al., 1998). The finding that lymphoid precursors give rise to both T cells and CD8CD11b DCs within the thymus suggests the existence of a lymphoid pathway (Ardavin et al., 1993), in addition to a well-established myeloid DC pathway (Inaba et al., 1993; Randolph et al., 1999; Schreurs et al., 1999). In mouse spleen, both CD8CD11b “myeloid” DCs and CD8CD11b “lymphoid DCs” were identified. The finding that “lymphoid” DCs express higher levels of self peptide–MHC class II complex (Inaba et al., 1997) and Fas-binding protein (Suss and Shortman, 1996) suggests that “lymphoid” DCs may be tolerogenic for T cells, in contrast to immunogenic “myeloid” DCs. This hypothesis was not supported, however, by studies showing that CD8CD11b “lymphoid” DCs produce a high level of IL-12 and induce potent TH1 response to foreign antigens (Pulendran et al., 1997, 1999; Sousa et al., 1997; Maldonado-Lopez et al., 1999; Smith and de St Groth, 1999).

EVIDENCE OF HUMAN MYELOID AND LYMPHOID DCs The most direct evidence that humans may also have different DCs comes from the finding that human blood contains two distinct types of dendritic cell precursors. Monocytes (pre-DC1) were shown to differentiate into immature myeloid DCs (im-DC1) after 7–10 days of culture with GM-CSF and IL-4 (Romani et al., 1994; Sallusto and Lanzavecchia, 1994). A second type of DC precursors (pre-DC2), previously known as plasmacytoid T cells (Lennert et al., 1975) or plasmacytoid monocytes (Facchetti et al., 1988) were shown to differentiate into im-DC2s after 3–6 days of culture with IL-3 or monocyte conditional medium (O’Doherty et al., 1994; Grouard et al., 1997; Olweus et al., 1997; Strobl et al., 1998; Kohrgruber et al., 1999; Res et al., 1999; Rissoan et al., 1999; Robinson et al., 1999; Sorg et al., 1999). Unlike CD11c immature DCs from

blood and tonsils, both pre-DC1s and pre-DC2s were unable to differentiate into DCs in the absence of cytokines. Pre-DC1s and pre-DC2s display many different features: (1) Pre-DC1s but not pre-DC2s express MHC class-like molecules CD1a, b, c and d and mannose receptors, suggesting that pre-DC1s and pre-DC2s may recognize different types of antigens (Kadowaki et al., 2000). (2) While pre-DC1s depend on GM-CSF to differentiate into im-DC1s, pre-DC2s mainly depend on IL-3 to differentiate into im-DC2s. This correlates with high GM-CSFRα and low IL-3Rα on pre-DC1s and low GM-CSFRα and high IL-3Rα on pre-DC2s (Grouard et al., 1997; Olwens et al., 1997; Rissoan et al., 1999). (3) While IL-4 is required for pre-DC1s to differentiate into imDC1s, IL-4 kills pre-DC2s when cultured with IL3, indicating different regulations for the two DC differentiation pathways (Rissoan et al., 1999). (4) In contrast to pre-DC1s, which express many myeloid antigens, such as CD11b, CD11c, CD13, CD33 and mannose receptors, and are able to differentiate into macrophages when cultured with M-CSF, pre-DC2s express low or no myeloid antigens and are unable to differentiate into macrophages (O’Doherty et al., 1994; Grouard et al., 1997; Olweus et al., 1997; Strobl et al., 1998; Kohrgruber et al., 1999; Res et al., 1999; Rissoan et al., 1999; Robinson et al., 1999; Sorg et al., 1999). (5) Pre-DC2s express high levels of pre-Tcell receptor α chain (pre-Tα), suggesting that pre-DC2s may be differentiated from lymphoid precursors (Res et al., 1999).

CD40L-ACTIVATED DC1s AND DC2s RESPECTIVELY INDUCE TH1 VERSUS TH2 DIFFERENTIATION CD40L not only triggers DC maturation (Caux et al., 1994; Peguet-Navarro et al., 1995), but also stimulates DCs to produce cytokines (Macatonia et al., 1995; Cella et al., 1996; Koch et al., 1996; Sousa et al., 1997). While im-DC1s produce a large amount of IL-12p75 (over 500 pg/mL), imDC2s produce a much lower amount of IL-12p75

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(less than 50 pg/mL) during 24 hours of CD40L activation (Rissoan et al., 1999). This was confirmed by quantitative RT-PCR analyses showing that im-DC1s expressed 20 times more IL-12p40 mRNA (10 000–40 000 fg/50 ng DNA) than do im-DC2s (500–600 fg/50 ng cDNA) after CD40L activation. In addition, CD40L-activated imDC1s but not im-DC2s produced significant amounts of IL-1α, IL-1β, IL-6 and IL-10. Both CD40L-activated im-DC1s and im-DC2s produce comparable amounts of the chemokine IL-8. Neither DC1s nor DC2s produce IL-4 mRNA or protein. Both CD40L-activated DC1s and DC2s induce 35 30 25 20 15 10 5 0

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strong proliferation of naïve CD4 T helper cells. To examine the function of DC1s and DC2s in T helper cell differentiation, CD4CD45ROCD45RA naïve T cells were cultured for 6 days with: (1) 24-hour CD40Lactivated DC1s; (2) 24-hour CD40L-activated DC2s or (3) anti-CD3 and anti-CD28 coated on a culture plate. After 6 days of priming, T cells were re-stimulated with anti-CD3 and anti-CD28 coated on a culture plate for either 5 hours to examine intracellular cytokine expression using flow cytometry or 24 hours to examine cytokines secreted into the culture supernatants using ELISA. CD40L-activated DC1s induced allogeneic naïve CD4 T cells to produce a large amount of IFNγ (over 30 000 pg/mL) and little IL-4, IL-5 and IL-10. By contrast, CD40L-activated DC2s induced CD4 naïve T cells to produce a much lower level of IFNγ (less than 500 pg/mL), but significant levels of IL-4 (over 200 pg/mL), IL-5 (over 600 pg/mL) and IL-10 (over 800 pg/mL) (Figures 3.3 and 3.4). These results suggest that: (1) DCs not only activate naïve T cells, but also directly polarize activated T cells toward TH1 and TH2 differentiation; (2) while CD40L-activated DC1s induce TH1 differentiation, CD40L-activated DC2s preferentially induce TH2 differentiation. The fact that DC1s induced TH1 differentiation was only partially inhibited by anti-IL-12 antibody suggests that either IFNγ or other undefined factors might be involved. The findings that DC2s do not produce IL-4 and that anti-IL-4 does not block DC2-induced TH2 differentiation suggests that DC2s may express an undefined TH2 differentiation factor. Trans-well culture experiments suggest that DC2-induced TH2 differentiation requires direct DC–T cell contact. In addition, exogenous IL-12 did not block DC2induced generation of IL-4-producing cells.

THE EFFECTS OF PATHOGENS AND CYTOKINES ON DC1 EFFECTOR FUNCTION ImDC1s derived from monocytes after 5–7 days of culture with GM-CSF and IL-4 have the

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FIGURE 3.4 Intracellular cytokine expression. (a) DC1–T cell co-cultures with control goat IgG antibodies and goat antibody to IL-12. (b) DC2–T cell co-culture with medium, goat Ig, goat antibody to IL-4 and IL-12. Double staining with anti-IL4 plus anti-IFNγ or anti-IL-10 plus IFNγ was performed. 104 cells were analyzed and the percentage of each T-cell population is indicated in the plots. ORIGIN AND MOLECULAR BIOLOGY OF DENDRITIC CELLS

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potential to induce both TH1 and TH2 differentiation (Hilkens et al., 1997). Two groups of signals were shown to stimulate imDC1s to induce TH1 differentiation: (1) LPS-derived from gram-negative bacteria, gram-positive bacteria particle SAC (Kalinski et al., 1999), unmethylated bacterial CpG-containing oligonucleoside (Jakob et al., 1998, 1999; Sparwasser et al., 1998; Hartmann et al., 1999) and double-stranded viral RNA (Verdijk et al., 1999); (2) T-cell signals such as CD40L (Macatonia et al., 1995; Cella et al., 1996; Koch et al., 1996; Sousa et al., 1997) and IFNγ (Kalinski et al., 1999). Many signals were shown to stimulate imDC1s to induce TH2 differentiation or to inhibit TH1 differentiation: (1) anti-inflammatory molecules such as IL-10, TGFβ, PGE2 and steroid (King et al., 1998; Kalinski et al., 1999) and (2) OX40 ligand expressed on T cells (Delespesse et al., 1999; Akiba et al., 2000). IFNα/β may directly stimulate TH1 differentiation by activation of STAT4 (Rogge et al., 1998). However, IFNα/β may also inhibit TH1 differentiation through downregulating IL-12 production by DCs (McRae et al., 1998, 2000; Bartholome et al., 1999a, 1999b). The clinical application of IFNβ in treating multiple sclerosis suggests that IFNα/β may have anti-inflammatory functions (Bartholome et al., 1999b). Some studies found that CD40L strongly stimulates imDC1s to produce IL-12 and induce TH1 differentiation (Macatonia et al., 1995, Cella et al., 1996; Koch et al., 1996; Sousa et al., 1997; Rissoan et al., 1999). Other studies found that CD40L is insufficient to stimulate imDC1s to produce IL-12 to induce TH1, and that exogenous IL-12 was required for CD40L activated imDC1s to induce TH1 differentiation (Hilkens et al., 1997). Two technical differences may need to be considered: (1) soluble CD40L and CD40Ltransfected L cells may have different potency in stimulating imDC1s to produce IL-12; (2) restimulation of primed T cells with anti-CD3 plus anti-CD28 or with PMA and ionomycine may have different results.

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THE EFFECTS OF PATHOGENS AND CYTOKINES ON DC2 EFFECTOR FUNCTION Recent studies have finally established that preDC2s in human blood are identical to natural IFNα–producing cells (IPCs), which produce enormous amounts of IFNα in response to viruses (Cella et al., 1999; Siegal et al., 1999). Thus, IPCs/pre-DC2s may represent a crucial effector cell type in antiviral innate immunity. Interestingly, unlike other effector cell types in the innate immune system such as neutrophils, macrophages and mast cells, which immediately die after performing their innate immune function, pre-DC2/IPCs undergo differentiation into DCs after producing IFNα/β in response to human herpes simplex virsues (HSV) (Kadowaki et al., 2000). This differentiation is mediated by the effect of autocrine cytokines IFNα/β and TNFα. Thus, pre-DC2/IPCs may differentiate into DCs by two pathways: (1) IL-3 and (2) viruses (HSV, by triggering endogenous IFNα/β and TNFα). Interestingly, unlike IL-3-induced DC2s that preferentially induces TH2, HSVinduced DC2s induce T cells to produce large amount of IFNγ and IL-10 (Kadowaki et al., 2000). Upon invasion of certain parasites or allergens, IL-3 produced by activated mast cells (Plant et al., 1989) may cause IPCs to differentiate into TH2-inducing DCs, which may contribute to the establishment of T cell-mediated allergic responses. On the other hand, HSV-DCs induce naïve CD4 T cells to produce IFNγ and IL-10, different from a classical TH1 or TH2 type. The existence of a T-cell population producing IFNα and IL-10 during intracellular infection with viruses (Sarawar and Doherty, 1994), bacteria (Gerosa et al., 1999) and parasites (Lee et al., 1999) suggests that IFNγ- and IL-10producing T cells may play an important role in immune responses to intracellular pathogens. In fact, it has been suggested that IL-10 protects the host against detrimental effects of excessive cellular immune responses elicited during acute infection (Gazzinelli et al., 1996). The finding

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that IL-3-induced DCs and HSV-induced DCs induce distinct types of T-cell differentiation illustrates the critical role of innate immunity in determining the type of adaptive immune responses, depending on the nature of pathogens.

THE EFFECT OF TISSUE MICROENVIRONMENTS ON DC EFFECTORS CD11c DCs isolated from mouse mucosa such as Peyer’s patches or respiratory tracts preferentially induce TH2 differentiation (Stumbles et al., 1998; Iwasaki and Kelsall, 1999; Khanna et al., 2000). By contrast, CD11c DCs isolated from mouse spleen preferentially induce TH1 differentiation. DCs derived from bone marrow precursors or from liver precursors also displayed different effector function in polarizing T helper cells. While liver-derived DCs produced high IL-10 and induced allogeneic T cells to undergo TH2 differentiation, bone marrow-derived DCs produced low IL-10 and induced allogeneic T cells to undergo TH1 differentiation during primary MLR in cultures or in allogeneic recipient mice after DC transfer (Khanna et al., 2000). The functional differences among different tissue DCs may result from differences in tissue cytokine microenvironments as well as in the lineage origin of different tissue DCs.

CONCLUSIONS AND FUTURE PERSPECTIVES Dendritic cells play two critical roles in T cellmediated immune responses: (1) they activate naïve T helper cells; and (2) they polarize activated T helper cells toward TH1 or TH2 effector cells. During early cognate DC–naïve T cell interaction, T cells rapidly express CD40L, which strongly stimulates DCs to upregulate MHC class I and II, CD80 and CD86, and to express/secrete T cell-polarizing molecules/ cytokines. While CD40L-activated DC1s produce large amounts of IL-12 and cause TH1 differentiation, CD40L-activated DC2s produce a low

level of IL-12 and mainly cause TH2 differentiation in an IL-4-independent fashion. T helper cell differentiation may depend on both the types of antigen-presenting cells and microenvironment signals/pathogens that act on imDCs. From our studies, CD40L-activated DC1s may be the most efficient natural adjuvant in vaccination against tumors and infectious pathogens, because DC1s probably induce strong inflammatory TH1 responses as well as strong CTL responses in vivo. By contrast, CD40L-activated DC2s may have the potential to be used in treating TH1-mediated autoimmune diseases and GVHD, because DC2s may induce anti-inflammatory TH2 immune responses in vivo.

ACKNOWLEDGEMENTS I would like to thank the members of my lab, B. Blom, H. Kanzler, M. Gilliet, S. Ho and S. Antonenko, for their contributions and critical reading of the manuscript. J. Katheiser for excellent editorial assistance.

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PLATE 3.1 Previous model regarding T helper cell differentiation. DCs activate naïve T helper cells due to their high expression of MHC class II and costimulatory molecules. Differentiation of activated T helper cells into TH1 or TH2 depends on, respectively, IL-12 produced by activated macrophages or IL-4 produced by a third cell type such as NKT cells or eosinophils.

PLATE 3.2 Reciprocal interaction between DCs and T helper cells. T helper cells rapidly express CD40 ligand after activation by DCs. CD40 ligand not only induces T helper cells to express high MHC class II and co-stimulatory molecules such as CD28, but also stimulates T helper cells to secrete T cell polarizing cytokines.

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4 Follicular dendritic cells: molecules associated with function Marie H. Kosco-Vilbois Serono Pharmaceutical Research Institute, Plan-les-Ouates, Switzerland

Whatever you are, be a good one. Abraham Lincoln

INTRODUCTION

using a tissue-specific promoter, has led to the development of de novo FDC networks in the murine pancreas (Kratz et al., 1996). The hurdle of isolating large numbers of primary cells as well as the absence of a clearly defined precursor continues to restrict our capacity to fully characterize the molecular and biological nature of FDCs. Indeed, following the identification of molecules responsible for deriving DCs ex vivo by the laboratories of J. Banchereau, R. Steinman, A. Lanzavecchia and others, a plethora of studies have become possible that significantly contributed to our understanding of events associated with T-cell priming. However, although this situation remains to be resolved for FDCs, investigators have endeavored to provide new information concerning the cytokines, chemokines and other novel molecules relevant to FDC function. This review will highlight some of these recent insights.

Follicular dendritic cells (FDCs), although sharing similar aspects of morphology, are quite different from the T-cell-priming dendritic cell (DC) described elsewhere throughout this book. FDCs are present within primary B-cell follicles, appearing in murine secondary lymphoid tissues as early as 10 days after birth (M. KoscoVilbois, unpublished observation). In contrast, cells expressing markers of FDCs such as CD21 and Ki-M4p have been observed in human fetal tissue at 21 weeks of gestation (Kasajima et al., 1997). FDCs form an extensive network in germinal centers of secondary lymphoid tissues as well as germinal centers developing ectopically. Examples of the latter have been observed using biopsies of arthritic joints, Hashimoto’s thyroid, Chlamydia infected eye and Helicobacter pyloriinfected gastrointestinal tissue. Furthermore, overexpressing the cytokine lymphotoxin (LT),

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ROLE OF TNF, LT AND THEIR RECEPTORS IN FORMATION OF FDC NETWORKS TNF exists in membrane as well as soluble homotrimeric forms, the latter being produced as a result of the activity of the TNFα-converting enzyme (TASE) (Black et al., 1997; Moss et al., 1997; Fu and Chaplin, 1999). In addition, LTα exists as a secreted homotrimer. These cytokines interact with the receptors TNFRI (p55) made up of a 55-kDa protein or the TNFRII (p75) consisting of a 75-kDa protein. Both receptors are capable of initiating signal transduction although they result in unique downstream events (Le Hir et al., 1996). The membraneassociated form of LT exists as an LTα1β2 heterotrimer and signals through the lymphotoxin β receptor (LTβR) (Crowe et al., 1994). Another form of LT exists as a LTα2β1 heterotrimer, but the biological relevance of this molecule remains to be determined. Analyzing the phenotype of mice targeted to delete one or more of these gene products provided the first evidence of a growth factor necessary for FDC formation. Mice deficient in either TNFα, LTα or LTβ lack or have severely underdeveloped FDC networks (Matsumoto et al., 1996; Pasparakis et al., 1996; Alimzhanov et al., 1997; Koni et al., 1997). In addition, mice lacking expression of the TNFRI or LTβR are also unable to produce FDC networks (Le Hir et al., 1996; Futterer et al., 1998). Using these mice in adoptive transfer studies has revealed further details. For example, several authors, including those investigating the transfer of compound LTβ/ plus B-cell deficient (BCR/) or TNF/ plus BCR/ bone marrow mixtures into LTβ/ hosts, have demonstrated a key role for LTα-, LTβ- and TNF-expressing B cells for FDC development (Fu et al., 1998; Gonzalez et al., 1998; Endres et al., 1999). Furthermore, similar chimeric studies with combinations of wild-type bone marrow cells transferred into deficient mice and vice versa, it was established that TNFRI and LTβR were required on nonhematopoietic cells in order to form an FDC network (Futterer et al., 1998; Tkachuk et al., 1998; Endres et al., 1999).

These conclusions were reinforced by Mackay and Browning, who demonstrated the collapse then disappearance of the FDC networks in adult mice following injection of a soluble form of the LTβR (Mackay and Browning, 1998). Taken together, it appears that FDC formation is coordinated by TNF, LTα3 and LTα2β1 via TNFR1. However, LTβR also plays a role, but may prove subtly different in mechanism (Futterer et al., 1998).

PLASTICITY OF THE FDC NETWORK Exploring further the mechanics of this microenvironment, termination of the LTβR-Ig treatment permitted the re-establishment of FDCs (Mackay and Browning, 1998). In addition, using the above mentioned adoptive transfer models, it was noted that when LTα/ bone marrow was used to reconstitute wild-type mice, the FDC network involuted and disappeared (Fu and Chaplin, 1999). Furthermore, this feature has been documented in patients undergoing antiretroviral treatment for the HIV-1 infection. The FDC network, previously crippled by viral load and lymphocyte malfunction, regenerated even at advanced stages of HIV-1 disease (Zhang et al., 1999). These observations underscore the potential susceptibility of the FDC compartment to modifications in morphology and consequently function following certain therapeutic regimes or pathogenic insults.

POSSIBLE CLUES TO THE ORIGIN OF FDCs In the first edition of this book, published in 1999, Tew and colleagues reviewed the literature for evidence favoring non-bone marrow versus extranodal versus primary lymphoid organ origin of the FDC (Tew et al., 1999). Their conclusion was that the exact nature of the cell type giving rise to FDCs remains an enigma, and this statement remains true at the time of composing this second edition. Using gene-targeted

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mice described above, as well as mice deficient in the complement receptors CR1 and CR2 (Ahearn et al., 1996; Croix et al., 1996), numerous investigators have endeavored to shed light on the nature of the precursor cell type (Matsumoto et al., 1997). In most studies, the FDC network remains of host origin and is not reconstituted if the receptor is deficient in the donor cells derived from bone marrow. However, it is evident that other cell types within follicles may contribute to proper architectural development. Stromal cells exist both in the B-cell follicle as well as T-cell areas and may be influential in directing the migration of lymphoid and nonlymphoid cells to their ‘proper’ microenvironment (Ngo et al., 1999). Indeed, plasma cells in TNFRI-deficient mice lack a signal to move out of the white pulp of the spleen and into extrafollicular locations (Tkachuk et al., 1998). As such, they remain in the periarteriolar lymphoid sheaths (PALS) and complete their differentiation atopically. For this reason, as well as the observation that FDCs can arise from primary lymphoid tissues when used as a source to establish networks in severe combined immunodeficiency (SCID) mice (Kapasi et al., 1998), caution must be maintained in reaching any conclusion concerning an origin. Certainly, the nature of this elusive precursor is complex in that only in situations where lymphocytes have never entered a secondary lymphoid tissue (i.e. SCID mice) can the precursor cell be provided by the donor population (Kapasi et al., 1998). Studies of scrapie replication in lymphoid tissues have also reinforced the concept of a precursor population established in adult secondary lymphoid organs. Using prion protein (PrP)-deficient mice crossed onto the SCID background, FDC networks developing posttransfer of wild-type bone marrow maintained their PrP-negative nature (Brown et al., 1999). Along these lines, an interesting experiment to conduct would involve the use of mice expressing LT under the control of the insulin promoter (Kratz et al., 1996). The pancreas normally does not support an FDC network. Establishing bone marrow chimeras in which the donor cells were distinguishable from host, turn on the insulin

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promoter-driven LT gene in an inducible manner, and determine whether the FDCs that develop in the pancreas are of host versus donor origin. If FDCs are indeed derived from a circulating source, this may be the model to test potential precursor candidate populations. Currently, experiments are underway to isolate a population of FDC-M1- or FDC-M2-binding cells in the spleen of TNFRI/ mice. These cells do not appear in the spleens of RAG or SCID mice and, therefore, may represent a precursor population that can establish itself when lymphocytes are present. Until a cell type can be isolated, adoptively transferred and give rise to a FDC network, conclusions on this issue should remain open.

EARLY EVENTS INITIATED IN GERMINAL CENTER REACTIONS Evidence from various sources has shown that the initial events leading to a high-affinity antibody response occur in the periphery of primary follicles of lymph nodes and the outer PALS of the spleen. Using B and T cells isolated from B-cell receptor (BCR)–hen egg lysozyme and T-cell receptor–ovabumin transgenic mice, respectively, Jenkins and colleagues have elegantly demonstrated the location of the antigenspecific lymphocytes in the lymph node during the first days following an antigenic insult (Garside et al., 1998). Furthermore, using a soluble form of the mannose receptor, a subset of DCs was shown to appear in these regions where antigen-specific B and T cells accumulate (Berney et al., 1999). These studies complement the earlier illustration by Kelsoe and colleagues of these events in the spleen (Jacob et al., 1991; Jacob and Kelsoe, 1992). The next step of a humoral response would be to attract these activated B cells into the FDC network and establish a germinal center response. We have observed that already on day 4 postantigen injection, B cells in cell cycle form peanut agglutinin-binding clusters within the processes of FDCs (Kosco-Vilbois et al., 1996).

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CXCR5 (previously known at Burkitt’s lymphoma receptor-1, BLR1) appears, at least in the spleen, to be the receptor responsible for this movement (Forster et al., 1996). The attraction is most likely mediated by the chemokine B lymphocyte chemoattractant (BLC)/B cellattracting chemokine1 (BCA-1) (Gunn et al., 1998; Legler et al., 1998) which is produced by FDCs as well as follicular stromal cells (Ngo et al., 1999). Future investigations should reveal the role of additional chemokines, especially for the lymph nodes. Knowing the nature of the FDC precursor as well as revealing conditions to mature it in vitro would certainly provide significant insight into this area.

MOLECULES PRESENT ON FDCs INDUCING THE FORMATION OF CLASSICAL GERMINAL CENTERS Antigen retained on FDC networks in the form of immune complexes has long been established to provide signals to germinal center B cells (Kosco-Vilbois et al., 1993). Previously, we have observed at the ultrastructural level that germinal center B cells will associate and internalize antigen trapped on the immune complexcoated bodies (iccosomes) produced by morphological changes in the processes of FDCs (Szakal et al., 1988). During this process, the germinal center B cells are induced to become efficient antigen-presenting cells (Kosco et al., 1988; Kosco-Vilbois et al., 1993), an essential step for eliciting necessary T cell help. Furthermore, crosslinking the BCR with the FDC-retained antigen induces release of the chemokines macrophage inflammatory protein1β (MIP-1β) and MIP-1α, which induce the chemotaxis of CD4CD45RO T cells (Krzysiek et al., 1999). Either anti-MIP-1β or anti-CCR5 antibodies will block this migration. These results demonstrate a mechanism for T-cell recruitment into and specific activation within the germinal centers following B-cell activation by antigen on FDCs. Recently, Li and colleagues have described a novel molecule expressed by FDC that

stimulates germinal center B-cell growth (Li et al., 2000). Generating the antibody 8D6 that recognizes a novel 282-amino-acid protein with homology to the LDL-R, these investigators were able to demonstrate that FDC–B cell interactions provide signaling for cell cycle progression. Furthermore, this antigen appears to play a role in the development of certain lymphomas as proliferation of the cell line L3055 is also inhibited in the presence of this antibody and the FDC-like cell line HK. Another molecule produced by FDCs and important for proper expansion of the germinal center reaction is IL-6. We have previously shown using IL-6-deficient mice that germinal centers remain small and antigen-specific IgG2a and IgG2b titers are diminished in comparison with wild-type mice (Kopf et al., 1998). We have also documented that FDCs and not the germinal center B or T cells produce IL-6. After searching extensively for a mechanism, it appears that in the absence of IL-6, local (i.e. in the follicles) production of complement 3 (C3) molecules is impaired. Macrophages in the germinal center have been subsequently identified as responsible for the production of C3 (Fischer et al., 1998). Hence, it appears that IL-6 produced by FDCs regulates molecules produced by other cell types in order to maintain stepwise progression of the high-affinity humoral responses.

IL-13Rα1 EXPRESSION BY FDCs IL-13 is a TH2-type cytokine that exerts biological effects similar to those produced by IL-4 (Zurawski et al., 1993; Minty et al., 1993; Zurawski and de Vries, 1994; Chomarat and Bancherau, 1997). The receptors for IL-13 consist of the IL-4Rα chain coupled to either the IL13Rα1 or IL-13Rα2 molecules (Caput et al., 1996; Aman et al., 1996; Hilton et al., 1996; Donaldson et al., 1998). Recently, using a monoclonal antibody generated in our laboratory, C41, that recognizes the extracellular portion of the murine (m)IL-13Rα1 molecule, we have observed reactivity both with FDCs and germinal center B

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REFERENCES

cells (Poudrier et al., 2000). Furthermore, using a soluble form of the mIL-13Rα1 protein, we encountered several unexpected results (Poudrier et al., 1999). Soluble mIL-13Rα1 promoted proliferation and antibody production of germinal center B cells. However, while IgG2a and IgG2b levels were augmented, IgG1 titers were surprisingly unaltered. Furthermore, this activity was not inhibited by an anti-IL-4Rα monoclonal antibody yet was dependent on IL-6 as the effect of mIL-13Rα1 was absent using germinal center cells isolated from IL-6-deficient mice. Taken together, our results suggest that a soluble form of the mIL-13Rα1 may signal through the IL-13R complex or a yet to be defined counter-structure. Germinal center B cells may be the source of soluble mIL-13Rα1 providing a signal to FDCs for IL-6 production. We are currently testing this hypothesis as well as screening for a novel component of this complex. Similar to the 8D6 antigen (Li et al., 2000), IL-13 appears to play a role in Hodgkin’s and nonHodgkin’s lymphomas (Billard et al., 1997; Kapp et al., 1999). As the Reed–Sternberg cell may derive from germinal center B cells (Kapp et al., 1999), greater understanding of IL-13 receptor biology during FDC–B cell interactions may provide novel insights for future therapeutic strategies.

PERSPECTIVES FDCs are central players in humoral immunity. The recent studies involving members of the TNF and TNF receptor family have provided an important clue to cytokines that drive FDC network formation. These various reagents and mouse lines will continue to provide significant tools allowing the discovery of other features of FDCs. Furthermore, numerous investigators are obtaining evidence that FDCs express molecules other than immune complex receptors which are pivotal in driving high-affinity antibody responses. While working with FDCs in vitro remains limited, the field is slowly accumulating new insights into the nature of FDC biology. Hopefully, the nature of its precursor is not too distant a discovery.

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Kapasi, Z.F., Qin, D., Kerr, W.G. et al. (1998). J. Immunol. 160, 1078–1084. Kapp, U., Yeh, W.C., Patterson, B. et al. (1999). J. Exp. Med. 189, 1939–1946. Kasajima, N., Maeda, K., Liu, D. et al. (1997). Dendritic Cells 7, 53–58. Koni, P.A., Sacca, R., Lawton, P., Browning, J.L., Ruddle, N.H. and Flavell, R.A. (1997). Immunity 6, 491–500. Kopf, M., Herren, S., Wiles, M.V., Pepys, M.B. and Kosco-Vilbois, M.H. (1998). J. Exp. Med. 188, 1895–1906. Kosco, M.H., Szakal, A.K. and Tew, J.G. (1988). J. Immunol. 140, 354–360. Kosco-Vilbois, M.H., Gray, D., Scheidegger, D. and Julius, M. (1993). J. Exp. Med. 178, 2055–2066. Kosco-Vilbois, M.H., Zentgraf, H., Gerdes, J. and Bonnefoy, J.Y. (1996). Immunol. Today 18, 225–230. Kratz, A., Campos-Neto, A., Hanson, M.S. and Ruddle, N.H. (1996). J. Exp. Med. 183, 1461–1472. Krzysiek, R., Lefevre, E.A., Zou, W. et al. (1999). J. Immunol. 162, 4455–4463. Le Hir, M., Bluethmann, H., Kosco-Vilbois, M.H. et al. (1996). J. Exp. Med. 183, 2367–2372. Legler, D.F., Loetscher, M., Roos, R.S., Clark-Lewis, I., Baggiolini, M. and Moser, B. (1998). J. Exp. Med. 187, 655–660. Li, L., Zhang, X., Kovacic, S. et al. (2000). J. Exp. Med. 191, 1077–1084. Mackay, F. and Browning, J.L. (1998). Nature 395, 26–27.

Matsumoto, M., Lo, S.F., Carruthers, C.J.L. et al.(1996). Nature 382, 462–466. Matsumoto, M., Fu, Y.X., Molina, H. et al. (1997). J. Exp. Med. 186, 1997–2004. Minty, A., Chalon, P., Derocq, J.M. et al. (1993). Nature 362, 248–250. Moss, M.L., Jin, S.L., Milla, M.E. et al. (1997). Nature 385, 733–736. Ngo, V.N., Korner, H., Gunn, M.D. et al. (1999). J. Exp. Med. 189, 403–412. Pasparakis, M., Alexopoulou, L., Episkopou, V. and Kollias, G. (1996). J. Exp. Med. 184, 1397–1411. Poudrier, J., Graber, P., Herren, S. et al. (1999). J. Immunol. 163, 1153–1161. Poudrier, J., Graber, P., Herren, S. et al. (2000). Eur. J. Immunol. Submitted. Szakal, A.K., Kosco, M.H. and Tew, J.G. (1988). J. Immunol. 140, 341–353. Tew, J.G., Kapasi, Z., Qin, D., Wu, J., Halley, S.T. and Szakal, A.K. (1999). In: Lotze, M.T. and Thomson, A.W. (eds) Dendritic Cells. London: Academic Press p. 51. Tkachuk, M., Bolliger, S., Ryffel, B. et al. (1998). J. Exp. Med. 187, 469–477. Zhang, Z.Q., Schuler, T., Cavert, W. et al. (1999). Proc. Natl Acad. Sci. USA 96, 5169–5172. Zurawski, G. and de Vries, J.E. (1994). Immunol. Today 15, 19–26. Zurawski, S.M., Vega, F.J., Huyghe, B. and Zurawski, G. (1993). EMBO J. 12, 2663–2670.

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5 Langerhans cells Dieter Maurer and Georg Stingl Department of Dermatology, University of Vienna Medical School, Vienna, Austria

À la condition que la vaccination soit faite strictement dans la peau, à une dose susceptible de regresser, et que l’épreuve soit faite après que la tumeur intracutanée ait complètement disparu, on réalise chez la souris une immunité anti-sarcomateuse solide et spécifique. A. Besredka and L. Gross, Ann. Inst. Pasteur 55, 491–499, 1935

INTRODUCTION

(original quotation: ‘Epithelzellen können sie nach ihrer Gestalt nicht sein; es handelt sich somit um die Frage: bindegewebig oder nervös?’), their reactivity with gold salts made him believe that ‘his cells’ represented sensory nerve endings (Langerhans, 1868). Today, we know that LCs are dendritic leukocytes which reside mainly within stratified squamous epithelia and constitute approximately 2–4% of all epidermal cells (see Schuler, 1991 for review). In the epidermis, they are usually located at a suprabasal position and attach to neighboring keratinocytes via an E-cadherin- and Ca2dependent mechanism (Tang et al., 1993). LCs cannot be easily identified on routine H&E sections. Their visualization and quantification in situ therefore requires the use of appropriate histochemical and/or immunolabeling techniques. Using adenosine triphosphatase (ATPase) histochemistry, Chen et al. (1985) found the LC density within the human epidermis to range from ~200/mm2 (palms, soles) to ~970/mm2 (face, neck). By electron

An intact immune system is required for all higher organisms to detect and destroy invading microorganisms (viruses, bacteria, fungi and parasites) and to eliminate cells that undergo malignant transformation. Anatomically, the first barrier to microbiological invasion is the skin, an organ that for many years was considered only a passive barrier against this invasion. Over the last two decades, however, concepts of a previously unrecognized role for skin have unfolded, a role in which dendritic leukocytes of the epidermis (i.e. Langerhans cells; LCs) and dermis (i.e. dermal dendritic cells; DDCs), initiate immune responses that protect the integrity of this organ. In 1868, the medical student Paul Langerhans, driven by his interest in the anatomy of skin nerves, discovered in the suprabasal region of the epidermis a population of dendritically shaped cells which now bear his name. While he was uncertain about their histogenetic nature Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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microscopy, LCs exhibit unique trilaminar cytoplasmic structures (Birbeck granules, BGs) that allow their identification (reviewed in Schuler, 1991; Figure 5.1). Using this organelle as a marker, it became clear that histiocytosis X represents a LC neoplasm (Basset and Nezelof, 1966), which is now referred to as Langerhans cell histiocytosis. In adults, the disease is mostly localized to skin and bone and usually runs a rather benign and protracted course. In children, it may also affect multiple internal organs and thus, represents a life-threatening condition (reviewed in Caputo, 1999). Langerhans cells are a mobile cell population with a relatively slow turnover. Epidermal residence is only one step in their life cycle. They originate from bone marrow precursors which, upon circulation in the peripheral blood, populate the skin. Their immigration into the skin is most prominent during late gestation (Foster and Holbrook, 1989); in adulthood, their putative precursors appear in the peripheral blood after severe epidermal injury (Gothelf et al., 1988). There also exist substantial numbers of dendritic leukocytes in the mammalian dermis. While some of them represent LCs on their way into or out of the epidermis, most of these

dermal dendrocytes are phenotypically slightly different from LCs and are generally referred to as dermal dendritic cells (DDCs) (Cerio et al., 1989). DDCs are located primarily in the perivascular areas of the superficial plexus. They have a folded nucleus and a highly ruffled, irregular surface. By electron microscopy, their cytoplasm is relatively dark, contains the organelles needed for an active cellular metabolism but is devoid of BGs. Phenotype and function of LCs/DDCs are critically influenced by their cellular and molecular microenvironment. Keratinocytes (KCs) are the major symbionts of LCs. They are capable of producing and secreting various mediators of the inflammatory reaction and of the immune response such as eicosanoids, cytokines as well as neuropeptides, e.g. proopiomelanocortin (POMC). KC-derived cytokines include interleukin (IL)-1, IL-6, IL-7, IL-8 and a steadily growing number of chemokines, IL-10, IL-12, IL-15, IL-18, granulocyte/macrophage colonystimulating factor (GM-CSF), M-CSF, tumor necrosis factor (TNF) α as well as some of the factors regulating the growth of epithelial and/ or mesenchymal cells, e.g., transforming growth factors (TGF) α and β, platelet-derived growth factor (PDGF), basic fibroblast growth factor

FIGURE 5.1 Electron micrograph of a Langerhans cell within the human epidermis. Depending on the plane of sectioning, Birbeck granules appear as either rod-shaped (arrowheads) or tennis racket-shaped (stars) structures (original magnification  40 000). ORIGIN AND MOLECULAR BIOLOGY OF DENDRITIC CELLS

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(bFGF), as well as vascular endothelial growth factor (VEGF) (reviewed in Luger et al., 1996; Weninger et al., 1996). POMC is synthesized by KCs as a large prohormone and is cleaved posttranslationally in active peptide hormones such as α-, β- and γ-MSH (Schauer et al., 1994). With the notable exception of IL-1, IL-7, TGFβ, macrophage inflammatory protein (MIP)-3α, CTACK, and stromal cell-derived factor (SDF)1α (Morales et al., 1999; Kilgus et al., 1993, Charbonnier et al., 1999), most biological response modifiers of KC origin are not expressed constitutively but only after perturbation of the epidermal homeostasis, for example, by hypoxia, chemical (e.g. haptens, drugs) and physical (ultraviolet radiation (UV ), cell dissociation) injury as well as microbial invasion (Enk et al., 1991; Kilgus et al., 1993; Luger and Schwarz, 1995; Harder et al., 1997; Cumberbatch et al., 1999). KCs are not the only cutaneous source of LC-/DDC-modulating mediators. Leukocytes (e.g. T cells, mast cells) as well as nonleukocytes (nerves, fibroblasts, endothelial cells) are also involved in this process, and certain pathogens (e.g. bacteria, viruses) can directly act on cutaneous DCs. Upon an antigenic encounter, LCs/DDCs can leave the cutaneous compartment and migrate to peripheral lymphoid organs where they initiate T-cell responses. Having accomplished this task, they initiate events which finally result in their own demise (reviewed in Schuler, 1991) (see Plate 5.2).

PHENOTYPIC AND FUNCTIONAL FEATURES OF LCs AND DDCs Although the cardinal features of human and rodent LCs (Birbeck granules, MHC class II expression) are the same, there exist certain phenotypic differences between the species. In this section, we will describe the characteristics of human epidermal LCs, unless otherwise stated. Resident LCs display nonspecific esterase and ATPase activity, and are the only cells within the normal epidermis to express

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CD1a, Fc-IgG receptors type II (FcRγII, CD32), Fc-IgE receptors type I (FcεRI), C3bi receptors (CD11b–CD18), and major histocompatibility complex (MHC)-encoded class II antigens (Table 5.1). Similar to other DCs in nonlymphoid tissues, LCs in situ have only a limited potency for stimulating naïve resting T cells but are quite efficient in antigen capture (Romani et al., 1989). The mechanism(s) by which LCs accomplish this task (is) are not fully understood. They are capable of engulfing large particles (e.g. latex beads (Wolff and Konrad, 1972), Leishmania major amastigotes (Blank et al., 1993) but, on the other hand, lack classical antigen uptake receptors such as the mannose receptor (Mommaas et al., 1999). In analogy to that which has been shown for peripheral blood DCs (Maurer et al., 1996), one can assume that IgE bound to FcεRI on LC surfaces can serve as an allergen-focusing molecule which, upon ligation by allergen, would facilitate processing and presentation of the latter. Investigators from Schering-Plough, Dardilly, France have produced the mAb DCGM4. It reacts selectively with LCs staining both the cell surface and the cytoplasm (Valladeau et al., 1999). By immunoprecipitation, DCGM4 identifies a 40 kDa molecule, termed Langerin. It is a type II Ca-dependent lectin displaying mannose-binding specificity. While sparing MHC class II cytoplasmic compartments, Langerin co-localizes with Birbeck granules. When incubated with freshly isolated LCs, the anti-Langerin mAb is internalized into Birbeck granules. Together with the observation that transfection of Langerin cDNA into fibroblasts results in Birbeck granule formation, one can hypothesize that Langerin is an antigen-capturing molecule which channels the antigen into Birbeck granules and, thus, possibly provides access to a nonclassical, nonMHC class IIdependent antigen-processing pathway (Valladeau et al., 2000). Upon receipt of certain stimuli [e.g. cytokines such as GM-CSF, IL-1, TNFα (Heufler et al., 1988), chemical haptens (Aiba and Katz, 1990), bacterial DNA and CpG-containing oligodeoxynucleotides

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LANGERHANS CELLS

Phenotype of resident versus cytokine-activated human epidermal Langerhans cells

Property

Resident LCs

Cytokine-activated LCs

Morphology Birbeck granules Cytoplasmic veils

 

/ 

Enzyme profiles Adenosine triphosphatase Nonspecific esterase

 

/ 

Antigenic profiles MHC class I MHC class II CD1a CD1b CD1c CD3-TCR CD4 CD8 CD14 CD15 CD19 CD20 CD45 CD45RA CD45RB CD45RO FcRI (CD64) FcRII (CD32) FcRIII (CD16) FcRI Langerin

                    

      ? /             /

 or   or /

? (4, 6) ? or (CD11c)

  /  

   /  or 

 /  /     /  ?

          

Adhesion molecules b 1 integrins b 2 integrins CD44 pan v7/v8 v4–v6, v9 E-cadherin CLA (cutan. lymphocyte-assoc. ag) Costimulatory molecules/Activation markers CD24 CD40 CD50 (ICAM-3) CD54 (ICAM-1) CD58 (LFA-3) CD69 CD80 (B7-1) CD83 CD86 (B7-2) CD98 CD101

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PHENOTYPE AND FUNCTIONAL FEATURES OF LC S AND DDC S

Property Cytokine receptors/Receptors for chemotactic factors GM-CSFRa (CD116) GM-CSFRb (CD131) M-CSFR (CD115) TNFR II (75kD) (CD120b) IL-1RI (CD121a) IL-1RII (CD121b) IL-2Ra (CD25) IL-2Rb (CD122) IFN-R (CD119) CCR1 CCR2 CCR3 CCR5 CCR6 CCR7 CXCR1 CXCR2 CXCR3 CXCR4/Fusin CXCR5 C5aR (CD88)

(Jakob et al., 1999)], LCs undergo a process of activation usually termed LC maturation (Schuler and Steinman, 1985). This process is characterized by: (1) neosynthesis and then efficient peptide loading of MHC molecules followed by the transport of stable peptide– MHC complexes to and prolonged display at the surface; (2) by downregulation of antigen uptake moieties (e.g. FcεRI, Birbeck granules); and (3) by a massive upregulation of costimulatory molecules of T cell activation (e.g. CD24, CD40, CD54, CD80, CD83, CD86; see Table 5.1) and certain cytokines such as IL-1β (Schreiber et al., 1992), IL-6 (Schreiber et al., 1992), IL-12 (Kang et al., 1996), IL-15 (Blauvelt et al., 1996) and IL-18 (Stoll et al., 1998). As a consequence, LCs acquire the capacity of stimulating naïve resting T cells for helper function and cytotoxicity. Antigens which induce primary immune responses when presented by LCs include alloantigens (Aberer et al., 1982; Inaba et al., 1986), soluble protein antigens and haptens (Hauser and Katz, 1988; Hauser, 1990) as well as tumor antigens (Grabbe et al., 1991, Celluzi and Falo, 1997) and microorganisms (Konecny et al., 1999). Phenotypic features of DDCs include the

Resident LCs

Cytokine-activated LCs

 /         /    /    /  /

   / /              /  

expression of MHC class I and II molecules, CD45, CD1c and, to a lesser extent, CD1a antigens, CD11c, CD40, CD54, CD80, the myeloid marker CD33, CD36 and the subunit A of the clotting proenzyme factor XIII (factor XIIIa). Notably, these cells either lack or express only negligible amounts of CD14 and, thus, do not qualify as macrophages. Their immunostimulatory capacity is similar to that of other mature DCs and far exceeds that of dermal macrophages (Lenz et al., 1993; Meunier et al., 1993; Nestle et al., 1993). While LC-driven immune responses to protein antigens are usually but not always (Ulanova et al., 1999) restricted by MHC class I and II antigens, nonpeptide bacterial antigens are usually presented in the context of CD1 molecules (Sieling et al., 1995). Microenvironmental signals to LCs and DDCs donotnecessarilyresultintheiractivationbutcan have quite opposite effects. It has been known for a long time that UV irradiation of epidermal cells leads to an impairment of the immunostimulatory properties of LCs (Stingl et al., 1981; Aberer et al., 1982) and, in fact, endows these cells with tolerogenic properties (Cruz et al., 1989). Evidence exists that an UV-induced increase in

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IL-10 (Niizeki and Streilein, 1997) and perhaps also IL-6 (Nishimura et al., 1999) synthesis by KCs is responsible for the latter phenomenon. The immunoinhibitory effect of α-MSH (Grabbe et al., 1996) and calcitonin gene-related peptide (CGRP; Hosoi et al., 1993; Carucci et al., 2000) may have a similar molecular basis (Bhardway et al., 1996; Torii et al., 1997). In the past few years, it has become clear that TGFβ1 profoundly affects LC development (see further below) and function. According to Geissmann et al. (1999), this cytokine prevents the final maturation of monocyte-derived LCs in response to TNFα, IL-1 and LPS with respect to MHC class II, CD80, CD86 and CD83 expression, loss of FITC-dextran uptake, production of IL-12, and antigen presentation. Therefore, due to the effects of (KC-derived) TGFβ1, mild inflammatory stimuli introduced or originating in the skin may not be sufficient to induce full maturation of LCs, thus avoiding potentially harmful immune responses. Increasing evidence exists that, in many cancer patients, the lack of protective tumor immunity is partly due to an impairment or loss of the immunostimulatory function of LCs/DCs (Enk et al., 1997). Cancer cell-derived immunoinhibitory cytokines (e.g. IL-10) as well as direct negative effects of certain carcinogens (Woods et al., 2000) are responsible for this phenomenon. In contrast to the immunoactivating properties of bacterial DNA and CpG-containing oligodeoxynucleotides (Jakob et al., 1999), viral infections (e.g. HIV) can often result in profound immunosuppression. LCs/DCs can be the target of HIV/SIV infection (Tschachler et al., 1987; Miller and Hu, 1999) and, as a consequence, become impaired in their immunostimulatory capacity (Blauvelt et al., 1995). Measles virus (MV) can infect LCs/DCs and, if so, interferes with their CD40L-dependent terminal maturation and thus, prevents CD8 T-cell activation (Servet-Delprat et al., 2000). This event may well contribute to the severe immunosuppression not infrequently seen in MV-infected children. Summarizing these findings, it appears that cutaneous DCs can subserve a dual function in skin-derived immune responses. Under

conditions which favor their terminal maturation, LCs/DCs will provide the skin with unique immunosurveillance mechanisms, i.e. the capacity to generate protective responses against exogenous and endogenous pathogens. In this context, it should be emphasized that cutaneous genetic immunization with naked DNA results in the transfection of and/or uptake of DNAencoded proteins by skin DCs which migrate to draining lymph nodes and efficiently elicit antigen-specific, cytotoxic and helper T-cell responses (Condon et al., 1996; Casares et al., 1997). On the other hand, stimuli which interfere with the acquisition of maturation-related immunostimulatory molecules and/or endow LCs/DDCs with tolerizing/anergizing properties (e.g. IL-10; Enk et al., 1993, 1994; Steinbrink et al., 1997) would not only render the skin more vulnerable towards potentially harmful invaders but also prevent exaggerated tissue responses to innocuous moieties, e.g. autoantigens and allergens. A third consequence of antigen uptake by cutaneous DCs could be an immunological null event. A ‘gatekeeper’ function of cutaneous DCs could be an important facet of the skin’s barrier function, but its actual occurrence has yet to be experimentally demonstrated. An understanding of the mechanisms that regulate the molecular signals to and from cutaneous DCs should give us a clue as to the role of the skin immune system in the maintenance of the host’s homeostasis and integrity and provide the basis for reestablishing it when perturbed. This knowledge will also have a major impact on vaccine design leading to a new generation of immunogens which, depending on the host’s (patho)physiological situation, will evoke a desired response in either a prophylactic or therapeutic setting.

DEVELOPMENT OF LCs FROM CD34 HEMATOPOIETIC PRECURSORS A major breakthrough in the understanding of LC development came from the observation that

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DEVELOPMENT OF LC s FROM CD 34  HEMATOPOIETIC PRECURSORS

the exposure of CD34 hematopoietic progenitor cells (HPCs) to GM-CSF and TNFα gives rise to a progeny of CD1a, E-cadherin, BGcontaining cells with immunostimmulatory properties strikingly resembling those of LCs isolated from human skin (Caux et al., 1992; Strunk et al., 1996) (Plate 5.2). Additional factors promoting the development of LCs/DCs from CD34 progenitors have been identified. The DCpromoting effect of GM-CSF can be replaced by IL-3, a finding that is consistent with the observations that the skin of GM-CSF/ mice harbors LCs and that IL-3 and GM-CSF receptors display similarities in structure and function (Caux et al., 1996a). Moreover, other stimuli, i.e. stem cell factor (SCF) and Flt3 ligand (FL), amplify the DC differentiation pathways initiated by GM-CSF and TNFα without any apparent selectivity for LC or nonLC-DC development (Siena et al., 1995; Szabolcs et al., 1995) (Plate 5.2). Many efforts have been undertaken to identify the LC progenitors at their various stages of maturation/differentiation. Strunk et al. provided evidence that, already at the CD34 HPC stage, cells exist which are committed to the LC lineage. An apparently useful marker to identify these cells is the cutaneous leukocyte homing antigen (CLA) which is abundantly expressed by LC precursors rather than by cells giving rise to nonLC-DCs (Strunk et al., 1997) (Plate 5.2). It has still to be determined whether and when this fucosylated PSLG-1 moiety (Fuhlbrigge et al., 1997) with affinity for vascular E- and P-selectin can mediate skin homing of LCs/LC progenitors. In GM-CSF- and TNFα-supplemented liquid cultures, LC precursors acquire CD1a expression around days 4–6 (Caux et al., 1996b). Evidence exists that a LC precursor of similar maturational stage circulates in the peripheral blood. Ito et al. identified and isolated a CD1a CD11c DC precursor from the peripheral blood that starts to express E-cadherin and Langerin and to display Birbeck granules within 1 day of culture in the presence of GM-CSF, IL-4, and TGFβ1 (Ito et al., 1999). In vitro-generated CD1a LC precursors, upon further culture in GM-CSF- and TNFαsupplemented medium until days 12–14,

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develop into DCs displaying all features found in and on epidermal LCs. It appears that the CD1a precursor cells are strictly committed to LC differentiation since they undergo rapid cell death when cultured in the sole presence of macrophage development-promoting factors. Simultaneously with the appearance of CD1a LC precursors, CD14CD1a cells emerge in the cultures. The lineage commitment of these cells is much less restricted than that of the CD1a precursors. They give rise to a monocyte/ macrophage progeny when exposed to M-CSF and develop into nonLC ‘DDC-like’ DCs in the presence of GM-CSF and TNFα (Caux et al., 1996b) (Plate 5.2). Phenotypically, these DDCs are characterized by the expression of factor XIIIa, CD1a, CD68, CD11b, CD36 and the lack of E-cadherin and Birbeck granules and, in contrast to LCs, are macropinocytotic and capable of producing IL-10 and of inducing differentiation in pre-activated B cells (Caux et al., 1997). Strobl et al. showed that TGFβ1 is a critical factor for LC generation in GM-CSF- and TNFα-supplemented serum-free stem cell cultures (Strobl et al., 1996, 1997). Recent work demonstrates that the LC-promoting action of TGFβ1 occurs through the induction of progenitor cell proliferation and of CD14 precursor differentiation. In the presence of TGFβ1, CD14 progenitors differentiate into DCs that display overlapping phenotype and functions with CD1a precursor-derived LCs (Caux et al., 1999; Jaksits et al., 1999). Thus, it appears that a major LC-promoting function of TGFβ1 is to redirect an early monopoietic precursor population into the LC differentiation pathway. Two independent studies conclude that TGFβ1 is also important for the development of DDC-type DCs (Caux et al., 1999; Jaksits et al., 1999). The most likely explanation is that a subset of GM-CSFand TNFα-stimulated CD14 precursors at day 6 has already lost its full LC differentiation potential but can still be aided by TGFβ1 stimulation to give rise to DDC-like cells. It also appears that CD14 precursors of advanced monocytic differentiation stage as well as monocytes require IL-4 plus GM-CSF rather than TNFα plus GM-CSF to enter the DC differentiation pathway.

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IL-4, when combined with GM-CSF, also induces TGFβ1-independent preferential development of nonLC-DCs from CD34 HPCs. Similarly, the GM-CSF- and IL-4-mediated development of DCs from monocytes (mdDCs) is TGFβ1 independent. Whether mdDCs, when generated in the presence of TGFβ1, display all features of LCs is a matter of debate (Geissmann et al., 1998, 1999; Jaksits et al., 1999). Collectively, it appears that TGFβ1 and IL-4 promote the development of true LCs and of nonLC-DCs, respectively. In vivo data strongly support a major role of TGFβ1 in LC development (Plate 5.2). TGFβ1/ mice lack epidermal LCs while showing normal bone marrow morphology and proper differentiation of the major myeloid cell lineages (Borkowski et al., 1996). Since bone marrow from TGFβ/ mice when transplanted into irradiated wild-type mice gives rise to a progeny of epidermal LCs, it appears that paracrine TGFβ1 stimulation of LC precursors is sufficient for LC differentiation in vivo (Borkowski et al., 1997). Furthermore, similar numbers of bone marrowderived LCs were found in skin grafts from TGFβ1/ and wild-type mice after transplantation onto nude mice. These findings, together with the observation of normal LC numbers in mice that lack TGFβ type II receptor expression in keratinocytes, suggest that the LC deficiency in TGFβ/ mice reflects a requirement of LCs or LC precursors for TGFβ1 and is obviously not due to a modulation of the cutaneous environment. It is not yet clear which cell types serve as the biologically relevant source of TGFβ1 in LC differentiation. As suggested by cell transfer and transplantation studies, radiation-resistant host cells other than keratinocytes are important in this regard (Borkowski et al., 1997). The exact relationship between LCs and other members of the DC family is not entirely understood. Although prevailing opinion holds that DCs residing within the T-cell zones of skindraining lymph nodes originate, at least partly, from a pool of antigen-laden LCs, it was somewhat surprising to learn that mice lacking the transcription factors rel B and Ikaros harbor essentially no DCs within their lymphoid organs, but contain a phenotypically normal-appearing

LC population (Georgopoulos et al., 1994; Burkly et al., 1995; Wu et al., 1997). Some evidence supports the idea that LCs could be members of the CD8 lymphoid rather than CD8 myeloid lineage of DCs. As shown by Anjuere et al. (2000), LCs start to express CD8 upon migration to draining lymph nodes or following ligation of CD40. However, in strong support for a myeloid derivation of LCs stands the recent observation that Notch1-deficient mice harbor normal epidermal LCs (as well as thymic DCs) while T-cell development is arrested at a very early stage, i.e. before the expression of T-cell lineage markers (Radtke et al., 2000).

RECEPTORS AND CYTOKINES PRESUMABLY INVOLVED IN THE SELECTIVE TISSUE-HOMING PROPERTIES OF LCs/LC PRECURSORS The mechanisms responsible for the rather selective homing of LCs/LC precursors to tissues covered by squamous epithelium are still not entirely understood. Several groups of investigators have tried to delineate the migratory potential of LCs along their differentiation from CD34 HPCs and have focused on the impact of the chemokine (CK) and CK receptor system. Chemokines represent a family of small basic chemotactic proteins that mediate their effect by binding to seven transmembrane-spanning G protein-coupled receptors on target cells. Currently, CKs are subdivided into four subfamilies according to the position of the first cysteine pair (CXC, CC), the lack of the second and the fourth cysteine (C), or the presence of three spacing amino acids in the first cysteine tandem (CX3C; for review see Baggiolini, 1998; Rossi and Zlotnik, 2000; Zlotnik and Yoshie, 2000). To date, five and 10 receptors have been identified for CXC and CC chemokines, respectively. By in vitro migration and CK receptor expression studies, we detected a functional dichotomy between the LC and the nonLCDC lineage already at the precursor level

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(Charbonnier et al., 1999). CD34 precursorderived CD1a LC precursors at day 6 express only one CC chemokine receptor, CCR6, in a prominent fashion and migrate to its ligand, macrophage inflammatory protein (MIP)-3α in a selective manner, i.e. they do not, or only very poorly respond to other CC chemokines. In contrast, day 6 CD14 nonLC-DC precursors migrate to many different chemokines. They respond to the CCR1 and CCR5 ligands MIP-1α and RANTES as well as to the CCR2 ligand monocyte chemotactic protein (MCP) -1, -2, -3, -4, and are less MIP-3α responsive than their CD1a counterparts (Charbonnier et al., 1999; Dieu-Nosjean et al., 2000). This migration profile resembles that of monocyte-derived (md)DCs; unlike CD34 stem cell-derived LCs (Greaves et al., 1997; Sozzani et al., 1998; Charbonnier et al., 1999; Dieu-Nosjean et al., 2000), mdDCs do not express CCR6 constitutively and display migratory responsiveness to MCP-1, 2, 4, RANTES, MIP-1α/β, MIP-5/HCC2, macrophage-derived chemokine (MDC), the CXC chemokines IL-8 and SDF-1α (Sozzani et al., 1995, 1997; Xu et al., 1996; Godiska et al., 1997; Delgado et al., 1998; Lin et al., 1998; Rubbert et al., 1998; Sallusto et al., 1998). Linked to its ability to induce LC features in mdDCs, TGFβ upregulates CCR6 expression and thereby induces MIP-3α reactivity in these cells (Sato K. et al., 2000). Are MIP-3α and CCR6 of critical importance for LC trafficking in vivo? The expression of MIP3α by keratinocytes and by other epithelia (Charbonnier et al., 1999; Tanaka et al., 1999, Iwasaki and Kelsall, 2000) along with the expression of CCR6 by LCs in situ (Charbonnier et al., 1999; Dieu-Nosjean et al., 2000) and by certain DC populations in Peyer’s patches (PPs) (Cook et al., 2000; Iwasaki and Kelsall, 2000) suggests that MIP-3α is responsible for the epitheliotropism of various DC types. In support of this assumption stands the observation that, unlike wild-type mice, CCR6/ mice fail to display CD11b DCs in the subepithelial dome of their PPs and, perhaps as a consequence, have a severe defect in the homeostasis and functionality of their gut-associated lymphoid tissues (Cook et al., 2000). Although some MIP-3α mRNA

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and protein is produced by nonstimulated keratinocytes ex vivo (Charbonnier et al., 1999; Dieu-Nosjean et al., 2000) and in vitro (DieuNosjean et al., 2000; Homey et al., 2000b; Kriehuber and Maurer, unpublished observations), it has not been resolved as to whether the observed MIP-3α expression level suffices for and whether additional factors are involved in the recruitment of LCs into an intraepidermal location. Another candidate chemokine constitutively produced by keratinocytes is the cutaneous T cell-attracting chemokine (CTACK; Morales et al., 1999). CCR10 binds CTACK and is expressed by endothelial cells, fibroblasts, melanocytes, T cells and LCs, but not in either CD34 hemopoietic progenitor-derived or monocyte-derived DCs (Homey et al., 2000a; Jarmin et al., 2000). Recently, it was shown that MIP-3α mRNA and protein expression is upregulated in psoriatic skin lesions. This appears to be associated with an inflammation-related increase in LC turnover rather than an enhanced epidermal LC density (Dieu-Nosjean et al., 2000). We also have evidence that MIP-3α can be expressed at sites of leukocyte entry into and exit from the dermal tissue, i.e. by endothelia of postcapillary venules and lymphatic vessels. Thus, the orchestrated MIP-3α expression by endothelial and epithelial cells could be central to the regulation of LC entry in and LC emigration from the skin and, thus, be of importance for the regulation of LC turnover under both normal and inflammatory conditions. Although these data suggest a specific role for MIP-3α in LC/DC recruitment to the skin and the small intestine, one should not forget that this CK is also expressed in other tissues such as fetal lung, liver and pancreatic islets (Power et al., 1997; Rossi et al., 1997). Perhaps of major importance is the observation that tumors of epithelial origin produce MIP-3α (Bell et al., 1999). That tumor cell-derived MIP3α is functionally relevant is suggested by an accumulation of immature, Langerin LC precursors within MIP-3α-producing adenocarcinomas of the breast. It is currently not known whether the migration of immature LCs into the tumor is followed by augmented tumoricidal immune responses or, alternatively,

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by the DCs’ demise within the tumor and, thus, the persistence of immunologic ignorance for tumor antigens. Unexpectedly, systemic treatment of mice with IL-12 leads to a significant increase in epidermal LC numbers while mature dermal DCs accumulate in FL-treated animals. The lack of FL-mediated DC accumulation in IL-12/ mice supports an important role of IL-12 for LC/DC trafficking, although the mechanism involved remains elusive (Esche et al., 1999).

EMIGRATION AND MATURATION OF LCs Evidence exists that perturbation of the cutaneous microenvironment leads to phenotypic and functional changes in the LC population

which are similar to those seen in epidermal single cell cultures (Figure 5.3). A few hours after skin transplantation, LCs begin to enlarge and to exhibit increased amounts of surface-bound MHC class II molecules. Subsequently, a marked reduction in the number of epidermal LCs occurs concomitantly with the appearance of strongly Ia cells in the dermis of the transplants (Larsen et al., 1990). Other investigators found that 24 hours after application of a contact sensitizer LCs appear larger than normal, exhibit more intense anti-Ia staining, and are severalfold more potent in their T-cell stimulatory capacity than LCs from nontreated or vehicletreated animals (Aiba and Katz, 1990). Furthermore, it was possible to identify antigen-bearing LCs/DCs in draining lymph nodes after the application of contact sensitizers (SilberbergSinakin et al., 1976; Macatonia et al., 1987) and

FIGURE 5.3 Phenotypic changes and emigration of epidermal Langerhans cells after perturbation of the cutaneous microenvironment. Cryostat sections of freshly excised or ex vivo cultured (48 hour 96 hour) human split-thickness skin were reacted with antibodies against CD1a, FcεR or CD83 and then processed for peroxidase immunolabeling. Before culture, LCs are CD1a, FcεRI, CD83 and reside mainly at a suprabasal position within the epidermis. In skin organ culture, LCs progressively lose surface-bound CD1a and FcεRI, begin to express CD83, and move downward (original magnification  40). ORIGIN AND MOLECULAR BIOLOGY OF DENDRITIC CELLS

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to demonstrate antigen-specific T-cell activation by these cells. Thus, it appears likely that LC function involves two components, i.e. antigen uptake and processing by ‘immature’ cutaneous LCs and, in the regional lymphoid organs, actual antigen presentation by mature skin-derived LCs. Recent studies have identified a role for epidermal cell-derived cytokines in regulating LC migration from the skin into draining lymph nodes. Antibodies to TNFα and IL-1β prevent the early migration of LCs from the epidermis, the accumulation of DCs in lymph nodes, and the development of optimal contact sensitization (Cumberbatch and Kimber, 1995; Cumberbatch et al., 1997). In keeping with these observations is the finding that the intradermal injection of TNFα or IL-1β stimulates the migration of LCs out of the epidermis and the accumulation of DCs in draining lymph nodes (Rambukkana et al., 1996). The effect of TNFα is apparently mediated by the 75 kDa TNFR II as LC migration in TNFR I gene targeted knock out mice is unchanged and still sensitive to TNFα neutralization (Wang et al., 1996). The further sequence of events occurring in LC emigration include the loosening of the E-cadherindependent attachment of LCs to neighboring KCs which can be, at least partly, explained by a LC maturation-related downregulation of this molecule (Schwarzenberger and Udey, 1996). Other yet poorly investigated conditions may include changes in the ligand binding activity of LC-expressed E-cadherin and/or the modification of the E-cadherin anchoring to cytoskeletal elements perhaps resulting in enhanced mobility of this moiety. It is of interest that the intradermal injection of unmethylated cytosineguanosine (CpG) oligonucleotides (Jakob et al., 1999; Ban et al., 2000), the topical application of the synthetic immune response modifier imiquimod (Suzuki et al., 2000), and infection with certain live viruses (Johnston et al., 2000) promotes LC maturation and migration. All these strategies could therefore be useful for the development of antimicrobial and anti cancer vaccines. A possible involvement of T cells and/or

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T cell-associated molecules in LC migration/ maturation is suggested by the observations that SCID and RAG/ mice as well as CD40L/ mice have reduced LC numbers in lymph nodes draining hapten-sensitized skin (Shreedhar et al., 1999; Moodycliffe et al., 2000). The findings in SCID and RAG/ mice, however, were not confirmed in a subsequent study (De Creus et al., 2000) and, thus, await further clarification. The migratory LC defect in CD40L/ mice could be overcome by administration of TNFα or antiCD40 antibodies (Moodycliffe et al., 2000). This indicates that LCs somehow and somewhere encounter CD40L on their way to the draining node and that TNFR signals can compensate for the necessity of CD40–CD40L interactions. Recently, two additional LC surface receptors were implicated as being essential for LC emigration to occur. One is CD44, a hyaluronic acid receptor putatively involved in the tissue homing of leukocytes and certain cancer cells. Antibody blocking studies suggest that an N-terminal epitope of CD44 is involved in LC emigration while the differentiation-related expression of the CD44 splice variant v6 allows for LC binding to T cell rather than to B-cell areas of lymph nodes (Weiss et al., 1997). It will be important to determine which skin- and lymph node-bound ligands of CD44 and of its different splice variants are responsible for the observed phenomena. The other LC-bound receptor structure involved in tissue emigration is a heterodimer built up by the integrin chains α6/β1 or α6/β4 (Price et al., 1997). Importantly, the prototype cell expressing the latter moiety is the KC. These cells express this receptor in the hemidesmosome where it mediates KC attachment to laminin, a major constituent of the basement membrane. It is conceivable that in vivo stimulated LCs loosen their KC-binding sites, and use α6-containing integrin receptors to specifically recognize basement membrane components. The antigenic stimulus itself, stimulation-induced KC products, or the receptor-mediated interaction with extracellular matrix proteins may then induce LCs to secrete proteolytic activity, e.g. type IV collagenase (matrix metalloproteinase (MMP)-9)

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(Kobayashi, 1997, Kobayashi et al., 1999), allowing them to penetrate the basement membrane and to pave their route through the dense dermal network into the lymphatic system. It is still unclear whether LCs enter lymphatic vessels in an active, perhaps directed fashion or are subject to passive transport by the constitutive flow of tissue fluid. If the first scenario applies in vivo, the identification of factors governing DC trafficking may allow us to devise strategies aiming at preventing DC-mediated allosensitization in transplantation settings. However, a definitive answer will require a detailed analysis of the functional potential of lymphatic vessel-derived endothelial cells which is currently not available. Recent observations favor an active recruitment of LCs into lymphatics and suggest a role of CCR2 ligands in this regard. Sato et al. reported that LCs from CCR2/ mice are markedly impaired in their ability to reach regional lymph nodes while the epidermal LC density in these mice is similar to that of littermate controls (Sato N. et al., 2000). In studies on skin explants, the authors observed that MHC class II-positive cells in CCR2/ mice accumulate in the dermis in a cord-like configuration and, thus, are possibly trapped within or around dermal lymph vessels. It remains to be seen whether the emigration defect in these mice primarily involves LCs or, perhaps, other MHC class II-positive skin cells present in the murine dermis. Following this line of thought, it is well established that monocytes/macrophages and monocyte-related DCs express CCR2 and respond to MCPs while LCs, at least those generated from human hemopoietic precursors or isolated from human skin, express no, or only trace amounts of CCR2 and fail to migrate in response to CCR2 ligands (Charbonnier et al., 1999; Dieu-Nosjean et al., 2000). While only certain immature DC subsets express CCR6, all mature DC types analyzed so far express CCR7 and migrate in response to its ligands MIP-3β and secondary lymphoid organ chemokine (SLC), chemokines that are expressed by yet poorly defined cells in T cellrich areas of secondary lymphoid tissue and by

high endothelial venules (HEVs; Greaves et al., 1997; Lin et al., 1998; Sozzani et al., 1998; Yanagihara et al., 1998; Charbonnier et al., 1999; Sallusto and Lanzavecchia, 1999). Besides its presence on mature DCs, CCR7 is expressed by naïve T cells and by a small subset of circulating, lymph node-seeking ‘central memory’ T lymphocytes (Sallusto et al., 1999). These expression patterns suggest that CCR7–CCR7 ligand interactions are of central importance spatially in bringing together mature DCs and naïve T cells. Indeed, severely reduced numbers and altered morphology of LCs/DCs in nodes draining contact sensitizer-exposed skin were seen in CCR7/ mice and in mice that do not constitutively express SLC in lymphoid tissues (i.e. mice that carry the paucity in lymphoid tissue (plt) mutation) (Förster et al., 1999; Gunn et al., 1999). While the lack of mobilized DCs in draining nodes of CCR7/ mice and plt mice cannot be attributed to reduced epidermal LC numbers or to impaired LC entry into the dermis, it appears that CCR7 signaling is important for LC entry into lymphatic channels and/or for their proper localization/ retention within T-cell zones. In line with an important role of CCR7 and SLC during the sensitization phase is the observation that hapten-specific CHS responses can be prevented when mice are treated with anti-SLC antibodies at the time of sensitization but not when they receive antibodies prior to challenge (Engeman et al., 2000). LC upregulate chemokine expression and secretion during their migration from contactsensitized skin towards T-cell zones of draining nodes. Among the plethora of factors probably elaborated by lymph node-bound mature LCs, the CCR4 ligand MDC was found to be of particular importance for T-cell attraction towards the antigen-laden LCs (Tang and Cyster, 1999). The observation that CCR4 is not expressed by naïve T cells but is rapidly upregulated upon antigen-specific activation of these cells suggests that MDC is a factor that helps to stabilize LC–T cell contacts once initiated and/ or that LCs in lymph nodes preferentially encounter T cells that were activated by

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REFERENCES

resident lymph node APCs presenting antigen delivered via the afferent lymph. A candidate cell type that could fulfill this task is the plasmacytoid DC present in T-cell zones that has the indigenous ability to induce TH2-biased immune responses (Grouard et al., 1997; Kohrgruber et al., 1999; Rissoan et al., 1999). In this scenario, the appearance or nonappearance of IL-12-producing LCs could decisively control whether a TH1- or TH2-biased T-cell response follows a cutaneous encounter of antigen.

FUTURE DIRECTIONS As mentioned above, LCs share many phenotypic and functional features with other members of the DC family. Accordingly, conventional read-out systems for DC functionality have largely failed to detect LC-restricted functions. As a result, the in vivo role of LCs relative to that of other, e.g. dermal DCs is far from being entirely understood. Further progress in this field will certainly benefit from the observations that LCs display certain unique features which include their peculiar antigen-uptake receptor and chemokine receptor repertoire as well as their ability to form intimate, receptor-mediated contacts with epithelia. It is thus expected that thorough investigations focusing on antigen processing and presentation pathways as well as on LC migration to and communication with epithelia will reveal profound insights into the true biologic role of these cells both in homeostasis and in inflammatory and neoplastic disease states. The new tools that already exist and/or are being currently established, e.g. the availability of genes that are selectively expressed in LCs and the cloning of their promotors, will in the future allow the investigation of LC biology in a fundamental manner. Among many possible experimental approaches, the generation of conditional LC knock-out animals as well as of animals that express genes of interest in LCs selectively will be instrumental in uncovering LC functions that are of relevance in vivo.

ACKNOWLEDGEMENTS This work was supported, in part, by a grant from Novartis Ltd, Basel, Switzerland and by the ICP Programme of the Austrian Ministry for Research. We thank S. Wichlas, M.D. and E. Csinády, M.D. for technical assistance.

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PLATE 5.2 Schematic representation of ontogenetic, migration and maturation pathways of cutaneous dendritic leukocytes. α6/β1,4, α6/β1,4 integrins; BM, bone marrow; CLA, cutaneous leukocyte antigen; CD, cluster of differentiation; CTACK, cutaneous T cell-attracting chemokine; DC, dendritic cell; DDC, dermal DC-type DC; FL, Flt-3 ligand; GM-CSF, granulocyte/macrophage colony-stimulating factor; HPC, hemopoietic progenitor cell; IL-1β, interleukin 1 beta; IL-4, interleukin 4; LC, Langerhans cell; MCP, monocyte chemotactic protein; M-CSF, macrophage colony-stimulating factor; MIP-3α, macrophage inflammatory protein 3α; MIP-3β, macrophage inflammatory protein 3β; MMP-9, matrix metalloproteinase 9; PB, peripheral blood; SCF, stem cell factor; SLC, secondary lymphoid organ chemokine; TGF-β1, transforming growth factor β1; TNF-α, tumor necrosis factor alpha.

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6 Dendritic cell-related immunoregulation: signals and mediators Pawel Kalinski 1, Michael T. Lotze 2 and Martien L. Kapsenberg 3 1

University of Pittsburgh, Pittsburgh, Pennsylvania, USA GlaxoSmithKline Pharmaceuticals, King of Prussia, PA, USA 3 University of Amsterdam, Amsterdam, The Netherlands

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missus dominicus (Latin: ‘envoy of the lord’), plural MISSI DOMINICI, officials sent by some Frankish kings and emperors to supervise provincial administration. Used sporadically by Merovingian and early Carolingian rulers, the missi became a normal part of the administrative machinery under Charlemagne (reigned 768–814). (. . .). They had full investigatory powers and were to rectify all error and injustice. Missi administered the oath of allegiance exacted from all freemen on the occasion of a new sovereign, informed local communities of the content of imperial decrees, and reported back on local conditions and needs. © 1999–2000 Britannica.com and Encyclopædia Britannica, Inc.

INTRODUCTION

selection of effector mechanisms that are most effective against the particular pathogen and are most suitable for the affected tissue. Migrating DCs provide naïve T cells with information about the antigenic structure of the pathogen (signal 1: antigen-specific stimulation; ensuring the specificity of response) and on its immunogenic potential (signal 2: co-stimulation; governing the magnitude of the response). Accumulating evidence indicates that in addition to providing signal 1 and signal 2, migrating DCs can also carry information about the characters of both the invader and the infected tissue and affect the initial TH1/TH2 pattern of the

Dendritic cells (DCs) initiate immune responses by providing the lymph node-based naïve T and B lymphocytes with pathogen-related information from the affected tissues. The effectiveness of the immune response depends on the information about the identity and pathogenicity of its potential target, and about the character of the pathogen. The availability of such information allows the pathogens to be targeted selectively and avoids potentially harmful responses to innocuous substances and nonpathogenic microorganisms. It also allows the Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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immune response, thereby contributing to the selection of the most appropriate effector mechanisms (signal 3: polarization). While the mechanisms used by DCs to regulate the antigenic specificity of immune response are addressed elsewhere, the current chapter focuses on the mechanisms allowing DCs to provide co-stimulatory and polarizing signals and to regulate efficiently the magnitude and character of the immune response.

REGULATION OF THE MAGNITUDE OF THE IMMUNE RESPONSE: DCs AS CARRIERS OF SIGNAL 1 AND SIGNAL 2 Activation of immature DCs in peripheral tissues results in an increase in their immunostimulatory capacity and migration to the draining lymph nodes where they initiate responses of antigen specific T helper (TH ) cells, CD8 CTLs and B cells, and act as a source of activating signals for NK cells (Banchereau and Steinman, 1998). In addition, locally activated DCs recruit additional immune cells to the site of pathogen entry and to the draining lymph nodes, orchestrating cooperation between different components of the immune system. The ability of DCs to induce an immune response depends on their effectiveness as the carriers of pathogen-related antigenic peptides and on their ability to provide T cells with costimulatory signals. Antigenic peptide–MHC complexes present on the DC surface provide TCR ligands for antigen-specific T cells and inform about the identity of the invader (signal 1). High expression of co-stimulatory molecules on the DC surface reflects the ability of the pathogen to activate DCs. DCs can detect pathogenic invasion either directly, due to their ability to recognize certain classes of pathogens (Medzhitov and Janeway, 1998), or indirectly, due to their ability to react to the pathogeninduced nonspecific inflammatory response of affected tissues, and to the pathogen-caused damage to local cells that release ‘danger

signals’, e.g. heat-shock proteins (Matzinger, 1998). Each of these mechanisms allows DCs to translate the pathogenicity of the invader into high expression of co-stimulatory molecules on their surface, and to provide T cells with signal 2, or co-stimulation, informing about the need to initiate specific immune response (Plate 6.1). The combination of signal 1 and signal 2 results in priming naïve T cells, which start to proliferate and acquire effector functions. By shifting the expression pattern of chemokine receptors and homing molecules, T cells acquire the ability to enter the appropriate tissues to provide other immune cells with helper signals or to directly kill the pathogen or the affected cells. There is increasing evidence that, in addition to their role in priming naïve T cells, DCs are critically important for the effector phase of the T-cell response, promoting the survival of committed T cells in the tissues and supporting their functions (Lambrecht et al., 1998). DCs are also critically important for B-cell responses. They directly enhance the proliferation of CD40-activated naïve and memory B cells and their differentiation into antibodysecreting plasma cells (Caux et al., 1994a; Dubois et al., 1998, 1999; Johansson et al., 2000), also participating in isotype class switching (Johansson et al., 2000). Similarly, interaction with DCs is important for induction of cytotoxic activity of NK cells (Fernandez et al., 1999). Interaction of DCs with at least some of these subsets of immune cells is bidirectional. This feedback is most evident in the case of T helper cells, the major regulatory lymphocyte population. The interaction of T helper cells with DCs brings about not only activation of T helper cells but also rescues DCs from apoptosis, and elevates the stimulatory activity of DCs and their cytokine production (see below). Although CTLs were perceived as pure recipients of DC-related activating signals, it was recently proposed that in the absence of CD40-mediated signals, constituting the major DC-activating pathway, CTLs can also contribute to the activation of DCs, bypassing the need for T helper cells (Ruedl et al., 1999). This observation may explain the

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relative T helper cell independence of some antiviral responses.

REGULATION OF THE CHARACTER OF THE IMMUNE RESPONSE: DCs AS CARRIERS OF ‘SIGNAL 3’ In addition to the specificity and magnitude of the immune response, a third element defining its effectiveness is the character of the effector mechanisms used, or the class of the immune response. One can distinguish two principal components of immune response: (1) cellular immunity mediated by inflammatory cells capable of killing the pathogen within infected cells and (2) humoral immunity, mediated by antibodies produced by antigen-specific B lymphocytes. Humoral immunity is largely ineffective against the pathogens localized inside cells. Antibodies released into the circulation, intracellular fluids and mucosal compartments are, however, extremely powerful against pathogens localized outside the cell. They effectively control infections with most bacteria and prevent the transmission of viruses between cells. The balance between these two components of immunity needs to be appropriately matched to a particular type of pathogen and to a character of the infected tissue. Inappropriate match results in chronic disease, either due to uncontrolled spread of the pathogen or due to autoimmune damage to host tissues. CD4 T helper cells are essential in regulating the balance between cell-mediated and humoral immunity. So called type 1 T helper (TH1) cells, the main producers of such effector molecules as IFNγ and TNFβ (lymphotoxin), are directly cytotoxic against infected or transformed cells. Perhaps even more importantly, they provide helper signals to cytotoxic CD8 T lymphocytes, and nonspecific killer cells, including NK cells and macrophages. Type 2 T helper (TH2) cells, produce B-cell-stimulatory factors, including IL-4 and IL-5, and promote the proliferation, survival, cytokine production and the Ig class switch in B cells (Mosmann and Sad, 1996). In

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addition to these two archetypal populations of effector/memory T helper cells, and an intermediate population of unpolarized TH0 cells, two additional phenotypes have been defined more recently: TGFβ-producing TH3 cells, implicated in the regulation of mucosal immune responses (TGFβ is a potent inducer of IgA) (Weiner, 1997), and the IL-10-producing TR1 cells (Groux et al., 1997), with a predominantly suppressor function, possibly important for the protection against autoimmunity. The development of strongly polarized ‘clearcut’ TH1 and TH2 phenotypes can usually only be observed after repetitive antigenic challenges, e.g. during chronic infections. However, an initial bias towards TH1 or TH2 cytokine production is already present at the moment of the first division of lymph-node based naïve T helper cells, as early as day 3 following exposure to immunogen, being dependent on the character of the immunogen and the type of infected tissue (Sangster et al., 1997; Toellner et al., 1998). Rapid and concomitant delivery of information about the antigenic structure and local pathogenicity of the infectious agent, but also about its character and the character of affected tissue, suggests that migrating DCs deliver to the lymph nodes not only the antigenic peptides (signal 1) and co-stimulatory factors (signal 2) but also the polarizing signals defining the character of the subsequent immune response (signal 3) (see Figure 6.1). In this respect, rat lung- and gutassociated DCs were found to be preferential inducers of the TH2 responses, when compared with spleen-derived DCs (Stumbles et al., 1998; Iwasaki and Kelsall, 1999), while numerous pathogens are known to differentially regulate the TH1/TH2-promoting function of myeloid (reviewed by Kalinski et al., 1999b) and lymphoid (Kadowaki et al., 2000) DCs. Just as there are a large number of factors involved in the delivery of co-stimulatory signal 2, the delivery of differential polarizing signals by DCs depends on several secreted and membrane-bound factors. While IL-12 appears to be the dominant TH1-skewing factor during intracellular infections with mycobacteria and intracellular parasites, IFNα (also capable of

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signalling via STAT4 in human cells) may well be the key factor promoting the development of TH1-biased cells during viral infections (Kadowaki et al., 2000). This second mechanism may counteract the ability of many viruses to suppress IL-12 production by DCs. In addition to IFNα and IL-12 and other soluble DC products, several particular surface-bound molecules (B7.1, B7.2, ICAM-1, OX40L, 4-1BBL, T1/ST2L) and the density of antigen/MHC complexes displayed on the DC surface all affect the TH1/TH2 profile of the immune response. The role of the individual membrane-bound soluble immunoregulatory factors in regulating the magnitude and character of immune response by DCs (Table 6.1) will be addressed next.

DC MEMBRANE-BOUND MODULATORY FACTORS B7 family members as the ligands for CD28, CTLA-4, and ICOS Expression of B7.1 (CD80) and B7.2 (CD86), two related molecules belonging to the Ig superfamily, appears crucial to the ability of the APCs to activate T cells (reviewed by Lenschow et al., 1996). Ligation of CD28 by these molecules provides an essential co-stimulatory signal for resting T cells (Damle et al., 1988) and allows for the induction of cytolytic activity in NK cells (Damle et al., 1988; Azuma et al., 1992). B7–CD28 interaction co-stimulates the induction of IL-2, an essential autocrine T-cell growth factor, that rescues TCR-triggered cells from activationinduced cell death and allows for their proliferation and acquisition of effector functions (Lenschow et al., 1996). Apart from induction of IL-2, CD28 ligation has a distinct IL-2-independent positive effect upon the proliferation of CD8 T cells and the development of their cytotoxic potential (Guerder et al., 1995). While proliferation of CD8 CTLs is just as dependent on B7–CD28 interactions as the proliferation of TH cells, their primary effector functions, cytotoxicity and IFNγ production (Van Gool et al., 1993; Guerder et al., 1995; Krummel et al., 1999)

are relatively CD28 independent. CD28 ligation is required for the de novo generation of CTLs within the naïve T-cell subset and for the induction of cytotoxicity in resting memory T cells. Although the CD28–B7 interaction is not required for cytotoxic effector cell function (Azuma et al., 1993; van Gool et al., 1993; Krummel et al., 1999), the CD28-related expansion of antigen-specific CTLs does contribute to the overall intensity of the CTL-mediated killing (Krummel et al., 1999). Similarly, B7-dependent signals support the acquisition of cytolytic activity by NK cells (Azuma et al., 1992; Chambers et al., 1996) but it is unclear to what extent this mechanism is involved in the activation of NK cells by the DCs (Fernandez et al., 1999). In addition to being essential for the development of cellular immunity, CD28-mediated signaling is required for induction of IL-4producing capacity in T helper cells (Seder et al., 1994; King et al., 1995; Kalinski et al., 1995), critical for their function as helper cells for B cells. It is also important for the induction of high levels of CXCR5, the chemokine receptor that mediates entry of T helper cells into germinal centers to support the germinal center reaction (Walker et al., 1999). Although memory/effector T helper cells are considered to be less dependent on the B7–CD28-mediated costimulation than their naïve counterparts, they also depend on this signaling pathway for optimal proliferation and cytokine production (van Neerven et al., 1998). In contrast to CD28, another ligand for both B7.1 and B7.2, CTLA-4 plays a distinct T-cell inhibitory function. In contrast to CD28, which is constitutively expressed on resting T cells, CTLA-4 only appears on the surface of T cells late during their activation and is implicated in suppressing their activity (Chambers et al., 1996; Oosterwegel et al., 1999). It binds both members of the B7 family with higher affinity than CD28 and is believed to protect against uncontrolled activation of the T-cell system and resultant autoimmunity (Walunas et al., 1994). Apart from regulating the magnitude of the immune response, B7.1 and B7.2 may also have a role in determining the class of the response,

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TABLE 6.1

Major functions and major target cells of DC-produced co-stimulatory and polarizing factors

DC, Mf

IL-1Ra IL-18

() TH1, NK

IL-6

CTL, B, DC, Mf

IL-12

TH, CTL, NK, B

IL-15

CTL, TH, NK, B, NK-T

IFNγ

CTL, NK, DC, Mf

TH1(h)

IL-10

TH (, m), CTL (/), DC (), Mf

TH2, TR1

B7 family

B7.1 (CD80) B7.2 (CD86) B7h/B7RP-1/ GL50* B7-H1*

CTL, TH, NK CTL, TH, NK TH(m)

TH1 (?) TH2 (?) TH2/TR1

TNF family

TNFα

DC, Mf, CTLs (?)

OX40L

TH

Hematopoietin/ activin family

Interferon family

Polarization

Comments

TH1

Upregulated by IL-18. Produced mostly by mLCs, rare in other DCs. Enhances DC stimulatory function Downregulated by IL-4/IL-13, HIV infection Interferon γ−inducing factor. Acts synergistically with IL-12. Pro-IL-18, like IL-1, is cleaved by caspase 1 following activation of APCs. Enhances production of IFNγ, IL-1, GM-CSF and IL-13; ↑ in mature DCs

TH1

TH(h)

TH2

Made by most DCs. Its proposed TH2-driving function was not confirmed in IL-6 knockout mouse Important survival factor for CTLs, TH1 and DCs. In humans, it is made predominantly by immature DCs Several IL-2-like activities. In contrast to IL-2, IL-15 promotes survival of T cells, rather than their effector functions. Made by many cells, including DCs, but not by T-cells Induced by viral infection. Promotes survival of T and DC and DC maturation. Essential factor for TH1 induction by human plasmacytoid IL-3Rα DCs Direct action on mouse T helper cells; in humans mostly via APCs. Antitumor activity in some models: recruitment of neutrophils and monocytes. ↑ angiogenesis. Production elevated in mucosal DCs and PGE2-exposed DCs Co-stimulatory and antiapoptotic via CD28; suppressive via CTLA-4 Ligand for ICOS. Selective inducer of IL-10, co-stimulator of CD4 cell proliferation Ligand unknown but different from CD28, CTLA-4 or ICOS Induces DC activation/maturation. Produced by most human DCs, rarely murine DCs Also on B cells, microglia, endothelial cells. Ligation enhances cytokine production by DCs. Although OX40L favors TH2 responses, it is also important for the induction of TH1 cytokines continued

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IL-1

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IL-1 Family

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Family/group

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Major functions and major target cells of DC-produced co-stimulatory and polarizing factors (continued ) Co-stimulation

Polarization

Comments

TNF Family (continued )

4-1BBL

CTL, less important for TH cells

TH1 (?, see comments)

CD40

TH, CTL (/), B, NK DC (enhancing their survival and function)

Expressed on B cells and Mf (binding of 4-1BBL stimulates both these cell types). In the absence of CD28 costimulation, 4-1BBL does not promote TH1 responses but acts as a selective IL-4 inducer Also on B cells. Ligation promotes production of IL-12 and other cytokines by DCs High on mature DCs and osteoclasts. Promotes DC survival and cytokine production, enhancing their stimulatory capacity. Ligand (TRANCE) on memory CTLs, T helper cells; inducible on naïve T cells Inducible on human CD11c DCs after treatment with IFNα or IFNγ (h); mediates direct killing of tumor cells. Involved in CMV-related immunosuppression. Also inducible on CTLs and T helper cells Constitutive on CD8α, inducible on CD8α mouse DCs. Can induce CD4 T cell apoptosis

RANK (TRANCE-L)

Other

Chemokines

TRAIL

() upon susceptible targets

FasL LIGHT

() upon susceptible targets T cells

BAFF

B cells

CD70

CTL

ICAM-1/2/3

TH1

LFA-3

CTL (less important) for TH cells), NK, B CTL, TH

T1/ST2L

CTL, TH

TH2 (m, ?)

see the text

Restricted to immature DCs. Downregulated after LPSor CD40Lo-induced activation/maturation of DCs Also made by activated T cells. B-cell co-stimulator lacking antiapoptotic function On mDC CD11 subpopulation of lymph node-derived cells Ligand for LFA-1 (CD11a/CD18). Important in early adhesion. Crucial for effector functions Ligand for CD2. Proposed to set the threshold level for TCR-dependent CTL activation T1/ST2 is an orphan receptor related to type-1 IL-1R Several of the DC produced chemokines affect the polarization of the immune response

(m), only in mouse; (h), only in human; (), negative impact; (/), context-dependent impact; (?), contrasting reports available; (*), unconfirmed expression on DCs.

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by modulating the cytokine profile of newly recruited naïve T helper cells. Triggering of CD28 primes naïve T helper cells for elevated production of the TH2 cytokines IL-4, IL-5 and IL-13 upon subsequent restimulation, and is less important for the ability to produce the TH1 cytokine IFNγ (King et al., 1995; Kalinski et al., 1995; Rulifson et al., 1997). The ability to promote production of TH2 cytokines may be particularly pronounced in the case of B7.2 (Freeman et al., 1995; Kuchroo et al., 1995), opening the possibility that the relative expression of B7.1 and B7.2 on a particular APC population may contribute to its TH1 or TH2-driving function. However, since B7.1 and B7.2 molecules were shown to be similar in importance in the induction of TH2 cytokines by DCs (Salomon and Bluestone, 1998), it remains to be established whether their differential expression indeed plays a physiological role in the polarization of the immune response. Expression of B7.1 and B7.2 depends on the stage of maturation of the DCs and on their source. Mouse and human Langerhans cells in healthy skin, analyzed in situ or directly after isolation, are either negative or show only marginal expression of either molecule. The expression of both of these molecules is rapidly upregulated in culture (Larsen et al., 1992, 1994; Symington et al., 1993; Razi-Wolf et al., 1994; Yokozeki et al., 1996). Similar requirement for the culture-associated induction/upregulation of both molecules were reported for mouse and rat DCs isolated from spleen, lung or kidney (Larsen et al., 1992, 1994; Stumbles et al., 1998). Human resting peripheral blood DCs are negative or show only marginal B7.1 and B7.2 expression on their surface but upregulate expression on culture (O’Doherty et al., 1993; McLellan et al., 1995). The requirement for activation of DCs restricts the expression of these critical costimulatory molecules to the situation of pathogen invasion, eliminating the risk of untoward expression of signal 2 in otherwise healthy tissues. In vitro-generated mouse and human DCs express significant amounts of B7.1 and B7.2, possibly due to the culture-associated activa-

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tion, but require an additional activation step for the optimal expression of these molecules (Caux et al., 1994b; Sallusto and Lanzavecchia, 1994). Increased B7.1/B7.2 expression accompanies the DC maturation induced by microbial products, factors released from dying cells, the mediators of nonspecific tissue response to pathogens such as GM-CSF, IL-1β, TNFα or type I interferons, or induced by CD40L-bearing T helper cells. Recently, two additional members of the B7 family were identified in humans and mice that do not interact with CD28 or CTLA-4. Murine B7h/B7RP-1/GL50 (Swallow et al., 1999; Yoshinaga et al., 1999; Ling et al., 2000) is a ligand for ICOS, (activation-induced costimulator), the CD28-related molecule originally identified on activated human T cells (Hutlof et al., 1999). B7h–ICOS interaction efficiently induces T-cell proliferation, but in contrast to B7–CD28 interaction, is poorly effective in the induction of IL-2 and IFNγ, cytokines associated with cellular immunity. Instead, it promotes IL-10 production, suggesting a role in a shift from cellular to humoral immunity. The role of co-stimulation via B7h–ICOS interaction appears to be restricted to CD4 T-cell responses, with limited or no impact on CTLs (Kopf et al., 2000). B7-H1 is a human B7 family-related molecule which also induces T-cell proliferation and preferential IL-10 production (Dong et al., 1999). Its’ T cell counter-receptor remains unknown but appears to be different from CD28, CTLA-4 or ICOS (Dong et al., 1999). The expression of both of these new members of B7 family have been demonstrated on macrophages, B cells and T cells. It remains to be established if they are also expressed on DCs.

ICAM-1 (CD54), -2, -3 as the ligands for LFA-1 (CD11a/CD18) LFA-1 interaction with ICAM-1, -2 and -3 represents another co-stimulatory event occurring early during DC–T helper cell interaction. LFA-1 acts as an adhesion molecule, facilitating optimal TCR-mediated signaling and B7–CD28mediated co-stimulation, but also has a direct

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co-stimulatory effect, as demonstrated in experiments utilizing a model of CD3 mAbmediated stimulation of T cells (Dubey et al., 1995). LFA-1 was originally described as CTLspecific accessory molecule, critically important in CTL-mediated cytotoxicity (Davignon et al., 1981; Sanchez-Madrid et al., 1982). In accordance, LFA-1-deficient mice show impaired activity of peripheral cytotoxic T cells and reduced antitumor immunity (Schmits et al., 1996). Although LFA-1-mediated co-stimulation appears more important for direct co-stimulation of CD8 cells, compared with CD4 T helper cells (Deeths and Mescher, 1999), LFA-1-dependent co-stimulation appears to be an important factor in the development of TH1 cells that support cellular-type immunity. ICAM-1–LFA-1 interaction was also shown to be important for the activation of NK cell activity and the resulting cytotoxicity. In addition, ICAM-1 and LFA-1 appear to be involved in the interaction of DCs with B lymphocytes (Kushnir et al., 1998). Several studies suggested the involvement of the ICAM–LFA-1 interaction in the differential regulation of the class of the immune response by the differential impact on the production of TH1 and TH2 cytokines. Ligation of LFA-1 by ICAM (ICAM-1, ICAM-2 and ICAM-3) molecules in primary mouse T-cell cultures inhibits IL-4 and IL-10 production, whereas it has little effect on IL-2 and IFNγ, and consequently results in a TH1 cytokine bias in effector T helper cells. Absence of ICAM, or blocking this costimulatory pathway, results in strongly TH2biased cytokine responses of effector T helper cells (Salomon and Bluestone, 1998; Luksch et al., 1999). The analogous ability to selectively support the production of TH1-type cytokines was recently demonstrated in human T helper cells (Jenks and Miller, 2000). The expression of ICAM-1 is low in immature DCs and increases upon maturation (Teunissen et al., 1990; Sallusto and Lanzavecchia, 1994). The levels of ICAM-1 expression in mature DCs reflect conditions of DC maturation. For instance, human monocyte-derived DCs matured in the presence of IFNγ show high

ICAM-1 expression, whereas the PGE2-matured DCs express low levels of this molecule (reviewed by Kalinski et al., 1999b; Vieira and Kapsenberg, unpublished results).

LFA-3 (CD58)–CD2 interaction LFA-3 was also originally identified as one of the factors required for CTL function (Krensky et al., 1984; Shaw et al., 1986). In CD8 cells, the stimulation via LFA-3 is mainly required for the induction of their effector functions, cytotoxicity and IFN production, while being less important for their IL-2 production and proliferation (Sanchez-Madrid et al., 1982; Parra et al., 1997; Le Guiner et al., 1998). The CD2–LFA-3 interaction activates T cells in the absence of TCR triggering, which may be important for setting the threshold level for CTL activation (Le Guiner et al., 1998; Bachmann et al., 1999a). LFA-3–CD2 interaction is important for the initial clustering and resultant proliferation of T cells interacting with human tonsillar, thymic, blood-isolated and monocyte-derived DCs (King and Katz, 1989; Landry et al., 1990; Young et al., 1992). It also protects T cells from apoptosis (Daniel et al., 1999). Although LFA-3 is expressed in substantial amounts on freshly isolated blood DCs (Freudenthal and Steinman, 1990), tissue-type DCs, such as Langerhans cells express only low levels of LFA-3 and need an activation step for its significant expression (Teunissen et al., 1990).

T1/ST2L–T1/ST2 T1/ST2 is an orphan receptor with homology to the IL-1 receptor family, and is preferentially expressed on mouse TH2 cells. T1/ST2 ligation is important in the TH2 effector function. T1/ST2blocking antibody or a recombinant fusion protein suppresses TH2 cytokine production in vitro and in vivo and attenuates eosinophilic inflammation, but fails to inhibit TH1-mediated inflammation (Lohning et al., 1998; Coyle et al., 1999). Similarly, eosinophil infiltration and IL-5 and IgE production is inhibited in the absence of T1/ST2 (Coyle et al., 1999). Independently,

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T1/ST2 was found to be indispensable in T1/ST2-deficient mice in a primary granuloma model induced with Schistosoma eggs (Townsend et al., 2000). The role of T1/ST2 as a TH2-driving factor became controversial recently, since the T1/ST2 deficiency has no effect on the development of TH2 cells in response to IL-4 and during infection with the helminthic parasite Nippostrongylus brasiliensis (Hoshino et al., 1999). To date, no data are available on the role of T1/ST2 in human T-cell polarization. The expression of a putative ligand of murine and human T1/ST2 was demonstrated in several cell types (Gayle et al., 1996). Unpublished data from the authors suggest that T1/ST2 ligand is expressed by human monocyte-derived DCs.

Membrane-bound TNF–TNFR family members TNF–TNF receptor family members appear to be an important group of factors regulating the magnitude and character of the immune response in human and murine models. The interactions of OX40L with OX40, CD40–CD40L, 4-1BBL–4-1BB, RANK–TRANCE, TRAIL–DR4/ DR5, LIGHT–LIGHTL, BAFF–BAFF-R and Fas– FasL were shown to be important for the ability of DCs to deliver co-stimulatory and polarizing signals to T cells. The presence of additional members of this family on DCs and their physiologic role still need to be demonstrated (Kwon et al., 1999). OX40 Ligand Among the members of the TNF–TNF receptor family, the role of OX40–OX40L interaction in the ability of DCs to provide co-stimulatory and polarizing signals is probably the best studied so far. OX40 (CD134) was initially identified as a surface antigen on activated rat CD4 T cells (Paterson et al., 1987). Its ligation augmented T-cell proliferation, especially at late stages of the immune response. Several years later, its analogues, showing high interspecies homology, were also identified in mouse and in human (Calderhead et al., 1993; Latza et al., 1994, Baum

59

et al., 1994). OX40 is expressed on activated T cells, while the expression of its ligand is found on B cells, T cells, microglia, endothelial cells, as well as DCs (Calderhead et al., 1993; Baum et al., 1994; Godfrey et al., 1994; Stuber et al., 1995; Imura et al., 1996; Ohshima et al., 1997; Weinberg et al., 1999). Crosslinking of OX40 costimulates the proliferation and the production of cytokines by T cells (Baum et al., 1994; Godfrey et al., 1994; Flynn et al., 1998; Gramaglia et al., 1998; Ohshima et al., 1998; Kaleeba et al., 1998), affecting the intensity of the immune response. Interestingly, apart from serving as a source of co-stimulatory signals for T cells, OX40–OX40L interaction also activates DCs, enhancing their maturation and their production of cytokines (Ohshima et al., 1997). Ligation of OX40 was also proposed to contribute to the preferential production of TH2 cytokines (Baum et al., 1994). Co-stimulation of OX40 by OX40L-transfected cell line induces high IL-4 expression in mouse naïve T helper cells in vitro (Flynn et al., 1998). Co-stimulation via OX40 also enhances IL-4 expression during priming of human naïve T helper cells in vitro and promotes the development of TH2 cells (Ohshima et al., 1998). A recent murine experimental leishmaniasis study demonstrated the critical role for OX40L in the development of functional TH2 cell responses in vivo (Akiba et al., 2000). OX40–OX40L interaction is also essential for the ability of activated T helper cells to enter germinal centers, provide help to B cells, promote germinal center formation and support immunoglobulin secretion (Stuber et al., 1995; Stuber and Strober, 1996; Flynn et al., 1998; Walker et al., 1999; Brocker et al., 1999). While the above studies demonstrate the key role of the OX40–OX40L interaction in the development of TH2-driven humoral immunity, another set of studies with OX40- or OX40Ldeficient mice demonstrate the importance of this signaling pathway for optimal induction of TH1-type responses (Chen et al., 1999; Kopf et al., 1999). The role of OX40–OX40L interaction in the TH1-driven inflammatory-type responses was demonstrated by showing its requirement for the induction of encephalitogenic activity in

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murine experimental allergic encephalomyelitis (EAE) models (Kaleeba et al., 1998; Weinberg et al., 1999). Elimination of OX40-positive cells or blocking this molecule ameliorates ongoing EAE (Weinberg et al., 1996). Blocking OX40–OX40L interaction also prevents induction of inflammatory bowel disease in several mouse models (Higgins et al., 1999). The same pathway is also crucial for the induction of CTL activity and contact hypersensitivity reaction to dinitrofluorobenzene (Chen et al., 1999). Since OX40L-deficient mice show relatively well-preserved production of IL-2, with concomitant deficiency of both IFNγ and IL-4, this suggests that OX40-dependent co-stimulation may be particularly important for the development of T helper cell effector functions, rather than for the expansion of Ag-specific cells (Chen et al., 1999). OX40L is not expressed on resting spleenderived mouse DCs but is induced following CD40-dependent stimulation (Chen et al., 1999). It is also strongly upregulated in CD40Lstimulated human monocyte-derived and blood-isolated DCs, although a small fraction of blood-derived DCs constitutively express this molecule (Ohshima et al., 1997). The heterogeneous expression of OX40L within the population of peripheral blood DCs raises the possibility that OX40L may have a differential regulatory role in T helper cell polarization. Indeed, similar to IL-12, the level of OX40L expression on mature DCs depends on the conditions of DC maturation.The ability of helminthic egg proteins to induce elevated OX40L expression on maturing DCs suggests its regulatory role in TH2 cell-mediated protection against helminths (de Jong and Kapsenberg, unpublished results). 4-1BB Ligand 4-1BB (CD137/ILA: receptor induced by lymphocyte activation), another member of the TNF family, is a co-stimulatory molecule expressed on activated T cells (Kwon and Weissman, 1989). 4-1BB ligand (4-1BBL) is an inducible molecule present on several APC types, including B cells, macrophages and DCs (Pollock et al., 1994; DeBenedette et al., 1997). Crosslinking of

4-1BB provides a TRAF2-NIK-mediated, CD28independent co-stimulatory signal for both CD4 and CD8 T cells (Saoulli et al., 1998), inducing their proliferation and activation of effector functions (reviewed by Vinay and Kwon, 1998; Watts and DeBenedette, 1999). The co-stimulation via 4-1BB is especially effective in the presence of high-intensity TCRmediated signal (Saoulli et al., 1998). In this case, 4-1BB is as potent as CD28 in co-stimulating IL2 production. 4-1BB synergizes with the CD28 pathway and with ICAM-1 in the induction of CD4 T-cell activation in the presence of limiting amounts of antigen (Kim et al., 1998; Gramaglia et al., 2000). Despite the ability of 4-1BB to act as an IL-2 inducer, at least a part of its costimulatory activity for CD8 cells appears to be IL-2 independent (Shuford et al., 1997). In addition, 4-1BB signaling also exerts an antiapoptotic effect, allowing for prolonged immune responses (Hurtado et al., 1997; Takahashi et al., 1999). 41BB stimulation, similar to LFA-1-mediated signaling, appears especially important for CD8 Tcell responses, augmenting their proliferation, IFNγ production, cytotoxicity and survival, while being less critical for CD4 T helper cells (Shuford et al., 1997; Tan et al., 1999; Takahashi et al., 1999). The importance of 4-1BB in tumor rejection, graft-versus-host disease, transplant rejection, and antiviral immunity (Shuford et al., 1997; Tan et al., 1999, 2000; DeBenedette et al., 1999) underlines the essential role of 4-1BB in cell-mediated immunity. 4-1BB was proposed to play a role in the differential regulation of the class of immune response, as a factor enhancing the production of IFNγ, with concomitant suppression of TH2-type cytokines (Kim et al., 1998). This TH1-promoting function of 4-1BB depends critically on the availability of CD28 co-stimulation. In its absence, 4-1BB co-stimulation selectively induces the production of IL-4 and IL-2 but not IFNγ (Chu et al., 1997). The above described differential regulation of TH1 and TH2 cytokines was not confirmed, however, in two recent studies addressing the role of 4-1BB in T helper cell responses (Gramaglia et al., 2000; Akiba et al., 2000). 4-1BB–4-1BBL interaction appears to be

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bidirectional. Ligation of 4-1BBL itself provides co-stimulation of anti-IgM-stimulated B-cell responses and induces the monocyte production of proinflammatory cytokines, TNFα, IL-6, IL-8, and upregulation of the ICAM-1 expression, while decreasing the production of IL-10 by monocytes (Pollock et al., 1994; Langstein et al., 1998). It was proposed that the triggering of 4-1BBL on human peripheral blood monocytes can promote their proliferation (Langstein et al., 1999). Expression of 4-1BBL on resting DCs is limited and requires their activation (DeBenedette et al., 1997). The role of 4-1BBL–4-1BB interaction in DC function still needs an in-depth analysis. CD40 CD40–CD40L (CD154) interaction (reviewed by van Kooten and Banchereau, 2000) provides an important maturation signal for DCs, induces the production of cytokines, including IL-1β, TNFα, IL-6, IL-8 and IL-12, prevents DC apoptosis and allows the effective stimulation of CTLs (Bjorck et al., 1997; Ridge et al., 1998). In addition, CD40 present on the surface of DCs provides a co-stimulatory factor for other immune cells. Direct CD40L-mediated signals are important for the proliferation of murine CD4 T helper cells and, to a lesser extent, for the activation of CD8 CTLs (Cayabyab et al., 1994). B cellexpressed CD40L is also implicated in the ability of B cells to receive survival signals from DCs (Wykes and MacPherson, 2000). Interestingly, human blood-derived DCs can also express functional CD40L. Its expression is enhanced by ligation of CD40 (Pinchuk et al., 1996), which may have additional roles in DC–B cell interaction. Although ligation of CD40L present on human NK cells stimulates this cell type, activating their cytolytic machinery (Carbone et al., 1997), other pathways appear to be more critical for the ability of DCs to activate NK cells (Carbone et al., 1999; Fernandez et al., 1999). RANK (TRANCE ligand) TRANCE (TNF-related activation-induced cytokine, or osteoprotegerin ligand (OPG-L), or

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RANK-L, or osteoclast differentiation factor (ODF)) is an inducible member of the TNF family expressed by T cells following TCR ligation (Anderson et al., 1997; Wong et al., 1997). TRANCE mRNA is constitutively expressed in memory, but not naïve T cells. The expression of TRANCE mRNA and the surface protein are rapidly upregulated in CD4 and CD8 T cells following TCR triggering, especially in the presence of CD28-mediated co-stimulation of CD4 cells (Josien et al., 1999). Interaction of TRANCE with its receptor can provide a back-up mechanism for the generation of functional virusspecific T-cell responses in the absence of CD40–CD40L interaction, as demonstrated in knockout animals (Bachmann et al., 1999b). The receptor for TRANCE, RANK (receptor activator of NFκB, osteoprotegerin (OPG)) is a member of the TNFR superfamily. It is expressed in high amounts on osteoclasts and mature DCs (Anderson et al., 1997) but only in low amounts on macrophages and B cells. Its expression is weak on resting DCs but is strongly upregulated by the culture of spleen-derived DCs and by CD40L stimulation (Anderson et al., 1997; Wong et al., 1997; Yun et al., 1998; Josien et al., 1999). RANK provides DCs with an antiapoptotic signal, resulting in the prolongation of DC survival during interaction with T cells, leading to increased T-cell stimulation (Anderson et al., 1997; Wong et al., 1997). In this respect, RANKmediated signals synergize with CD40- and TNFα-mediated signals (Josien et al., 1999). Engagement of RANK also stimulates DCs to produce cytokines, including IL-1, IL-6, IL-15 and IL-12 (Josien et al., 1999; Bachmann et al., 1999b). TRANCE-deficient animals (that have preserved spleen and Peyer’s patches but not lymph nodes) show a deficiency in T- and B-cell differentiation but they do not display overt abnormalities of DCs (Kong et al., 1999). TRAIL TRAIL (TNF-related apoptosis inducing ligand, of Apo-2 ligand), another TNF family member, signals via two apoptosis-inducing receptors, while an additional TRAIL receptor has an

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apparent decoy function (Wiley et al., 1995; Pitti et al., 1996; Sheridan et al., 1997). TRAIL is expressed by activated CD4 and CD8 T cells as a surface molecule or released in vesicles following cell activation (Kayagaki et al., 1999; Martinez-Lorenzo et al., 1999; Kaplan et al., 2000). Human DCs can express TRAIL following their exposure to IFNγ and IFNα, which results in their ability to kill the receptor-positive tumor cell lines (Griffith et al., 1999; Fanger et al., 1999). This ability appears to be restricted to CD11c DCs. IL-3Rα plasmacytoid DCs do not display this function. In addition, DCs infected with measles virus (MV) express TRAIL, which may play a role in MV-associated immunosuppression (Vidalain et al., 2000). Fas ligand FasL is expressed on a CD8α subpopulation (putative lymphoid DC subset) of mouse spleen DCs and can be induced on CD8α DCs. It is implicated in the ability of CD8α DCs to induce apoptosis of Fas-positive CD4 cells (Suss and Shortman, 1996). FasL can be also expressed by bone marrow-derived GM-CSF and IL-4cultured DCs. Although such FasL-positive cells could kill Fas-sensitive Jurkat cells, their concomitant expression of high levels of B7.2 acted as a source of signals preventing healthy T cells from apoptosis (Lu et al., 1997), suggesting that the balance between the level of expression of co-stimulatory molecules and FasL can dictate the function of DCs as inducers or inhibitors of immunity.

Interestingly, in contrast to other molecules of this group, the expression of LIGHT is restricted to immature DCs and is shut down following LPS- or CD40L-induced maturation, the precise significance of which is unknown. BAFF A recent report (Schneider et al., 1999) demonstrated that BAFF (B cell-activating factor belonging to the TNF family), another novel TNF family member previously known to be expressed by activated T cells, is also produced by human monocyte-derived DCs. Expression of BAFF may contribute to the ability of DCs to promote the proliferation of B lymphocytes. BAFF activation induces somewhat different response of B cells than CD40 crosslinking. The proliferation-enhancing effect of BAFF is more strictly dependent on the presence of anti-IgM stimulation and, in contrast to CD40 ligation, it does not have an antiapoptotic effect. CD70 CD27, a membrane glycoprotein similar to CD40, OX40 and 4-1BB, is present on most human and mouse T cells. Anti-CD27 antibodies augment antigen receptor-mediated T-cell proliferation (Camerini et al., 1991; Gravestein et al., 1993). CD70, the ligand for CD27, is expressed on a subpopulation of mouse lymph node-isolated CD11c DCs, but the physiological role of DC-expressed CD70 (as well as that of the related molecule CD30) remains elusive (Akiba et al., 2000).

LIGHT LIGHT (TNFSF14, herpesvirus entry mediator (HVEM) ligand), is another member of the TNF receptor family, with the ability to co-stimulate proliferation of HVEM or lymphotoxin-βexpressing T cells, but also with a proapoptotic function against carcinoma cells (Harrop et al., 1998). LIGHT produced by human monocytederived immature DCs was recently shown to be important for induction of T-cell proliferation in response to alloantigen (Tamada et al., 2000).

DENDRITIC CELL-PRODUCED CYTOKINES TNFα TNFα (cachectin), first described as a factor with direct cytotoxic activity following intratumoral administration (Carswell et al., 1975), is a pleiotropic proinflammatory cytokine with

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multiple immunomodulatory functions (Beutler and Cerami, 1989). Its targets include DCs that are susceptible to TNF-induced activation, maturation and migration to lymph nodes (Cumberbatch and Kimber, 1992; Sallusto and Lanzavecchia, 1994). TNFα production was documented in many types of DCs, including human Langerhans cells, bone marrow- and monocyte-derived DCs (Larrick et al., 1989; Nickoloff et al., 1991; Caux et al., 1994b; Verhasselt et al., 1997). Murine DCs, however, produce no or only marginal amounts of this cytokine (Oxholm et al., 1991; Enk and Katz, 1992; Shreiber et al., 1992; Sprecher and Becker, 1992). In contrast to the several membranebound members of TNF family, discussed above, the importance of TNFα itself for DC costimulatory and polarizing functions has not been extensively studied.

IL-1 Interleukin 1, a prototypic proinflammatory cytokine, is produced as IL-1α or IL-1β, both binding the same set of receptors with similar affinities (Dinarello, 1996). IL-1 was identified as an important factor for the initiation of antigenspecific responses by resting DCs, allowing for their initial activation and resultant ability to activate T cells (Koide et al., 1987). Dendritic cells themselves are relatively poor producers of this factor, both in human and mouse (Koide et al., 1987; Vakkila et al., 1990; Zhou and Tedder, 1995), with the possible exception of mouse LCs, which produce significant amounts of IL-1β, especially after culture (Heufler et al., 1992). Human DCs can also produce another member of the IL-1 family, IL-1 receptor antagonist (IL-1Ra) (Lore et al., 1999).

IL-6 IL-6 (IFNβ2, BSF-2, HGF) is a pleiotropic cytokine, the functions of which include induction of acute-phase proteins, co-stimulation of proliferation and antibody production by B cells, participation in the development of CTL responses, and enhancement of the survival of

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naïve T cells (Van Snick, 1990; Kishimoto et al., 1992; Teague et al., 2000). IL-6 may also act as a suppressor factor, as it inhibits the production of proinflammatory cytokines: IL-1β, TNFα, GMCSF and IFNγ, while it induces the production of soluble TNFαR and IL-1Ra (Opal and DePalo, 2000). Although IL-6 was shown to shift the balance of cytokine production in mouse in vitro activated T helper cells towards TH2, IL-6-deficient mouse do not demonstrate a TH1 bias (Kopf et al., 1998). In our own experience, IL-6 does not mediate any TH1- or TH2-biasing effect upon human CD4 T helper cells from peripheral blood. In contrast to IL-1, IL-6 is produced by most populations of DCs tested. Mouse LCs, lymph node-isolated DCs, as well as DCs generated from bone marrow cultures are good producers of this cytokine (Hope et al., 1995; Cumberbatch et al., 1996; Josien et al., 1999). IL-6 gene expression is also detected in freshly isolated human blood DCs, as well as monocyte-, and CD34 progenitor-derived DC populations (Zhou and Tedder, 1995; Verhasselt et al., 1997; de Saint-Vis et al., 1998). IL-6 is abundantly produced by CD40L-stimulated immature DCs and the ability to produce this factor is further enhanced in fully mature cells. In contrast, bacterial products, such as LPS and SAC, induce the production of IL-6 only in immature CD83 cells, but not in mature DCs (Verhasselt et al., 1997; Kalinski et al., 1999a).

IL-10 Interleukin 10 is another pleiotropic immunomodulatory factor with functions at different levels of immune response. Although IL-10 can play an active role in cellular immunity, as a factor facilitating the generation and survival of CTLs (Yang et al., 1995; Santin et al., 2000), its immunosuppressive, anti-inflammatory, and TH2-biasing functions predominate. IL-10 modulates cellular immunity acting directly on T cells (especially in mouse), but also affects T-cell responses indirectly, by inhibiting migration, expression of co-stimulatory molecules and

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cytokine and chemokine production by DCs and other APC populations (reviewed by Howard and O’Garra, 1992; Opal and DePalo, 2000). As expected from such an array of inhibitory functions of IL-10, the IL-10 knockout mice spontaneously develop chronic inflammatory enteritis similar to human inflammatory bowel disease. They are also markedly more susceptible to endotoxin-induced shock. Major sources of IL-10 are stromal cells, monocytes and macrophages, as well as B cells and T cells. In the mouse, IL-10 is produced effectively by TH2 cells but not TH1 cells. In humans IL-10 production is regulated independently from the TH1/TH2 commitment of T helper cells. More recently, within CD4 cell populations a unique TR1 cell subset was identified that produces mainly IL-10 and plays a role in preventing uncontrolled inflammatory responses and autoimmunity (Groux et al., 1997). Both human and murine DCs are generally poor producers of IL-10 with several significant exceptions. DCs freshly isolated from murine bronchi and from Peyer’s patches are good IL-10 producers, which is associated with their TH2driving function (Stumbles et al., 1998; Iwasaki and Kelsall, 1999). A similar TH2-driving function, associated with high IL-10-producing capacity and suppressed IL-12 production, can be induced in human monocyte-derived DCs by their pretreatment at early stages of maturation with prostaglandin (PG)E2 (Kalinski et al., 1997), a factor present in mucosal tissues at high concentrations. High IL-10-producing capacity was also found in the subpopulation of human CD34 progenitor cell-derived DCs developing via the CD14 intermediate stage (de Saint-Vis et al., 1998). This last DC population produces IL-10 in response to CD40 ligation, while PGE2modified DCs produce high levels of IL-10 following stimulation with bacterial products, SAC and LPS, but not CD40L.

IL-12 Interleukin 12, a key factor in the development and maintenance of cellular immunity, provides

essential signals for the activation, functional activity and survival of NK cells, cytotoxic T lymphocytes and TH1 cells (reviewed by Trinchieri, 1998; Gately et al., 1998). IL-12 is also important for B cells (Dubois et al., 1998), explaining perhaps the relevance of the paradoxical ability of TH2 cells and IL-4 to support the production of bioactive IL-12 in DCs (Kalinski et al., 2000). Another recent study suggests that autocrine IL-12 production contributes to the activation of mouse DCs following CD40 triggering (Bianchi et al., 1999). IL-12 is perhaps the best-defined T helper cell-polarizing factor, affecting the character of immune response to different classes of pathogens. IL-12- or STAT-4-deficient mice show a broad array of defects associated with impaired TH1 responses, but surprisingly wellpreserved antiviral responses (Schijns et al., 1998). The few identified individuals lacking a functional IL-12R have recurrent infections with salmonella and mycobacteria, but not with viruses, stressing the important role of IL-12 in the defense against pathogens located in intracellular vesicles (Altare et al., 1998; de Jong et al., 1998), while being apparently less important for the induction of cellular responses to viruses. Most DC populations are capable of producing IL-12 following activation by a variety of stimuli, including infection with pathogens or exposure to their products. IL-12 production is also induced by T helper cells after ligation of CD40 by CD40L present on activated T helper cells. Efficient induction of bioactive forms of IL-12 (IL-12p70) by T helper cells depends on co-stimulatory signals provided by T helper cellproduced IFNγ (Hilkens et al., 1997; Snijders et al., 1998) or IL-4 (Hochrein et al., 2000; Kalinski et al., 2000). Until recently, DCs were regarded as a strictly TH1-driving APC type, because of their ability to produce IL-12. However, several recent in vitro and in vivo studies show that DCs are good inducers of both TH1 and TH2 cytokine responses. An important factor in this respect is that IL-12 production in DCs strongly depends on the stimulus type and the conditions of stimulation of DCs. While IFNγ and IL-4 are potent

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enhancers of bioactive IL-12 production (although IL-4 suppresses the secretion of the inactive p40 homodimer), many other factors, including IL-10, corticosteroids and PGE2, suppress IL-12 production by DCs (Moser et al., 1995; Buelens et al., 1997; Vieira et al., 1998, 2000; Kalinski et al., 1998). IL-12-modulating factors may act directly on DCs interacting with T helper cells. Such direct effects may be especially important in affecting the polarization of T helper cells during secondary responses. Probably more important for the primary responses is the fact that the levels of IL-12 production in mature DCs are strongly influenced by the environmental conditions at the site of induction of DC maturation (reviewed by Kalinski et al., 1999b). IL-12-producing ability of DCs can also be affected by pathogens. Induction of DC maturation by helminthic proteins result in mature DCs lacking the ability to produce. This in turn allows for the development of helminth-specific TH2 cells (de Jong and Kapsenberg, unpublished results).

on their long-term survival. IL-15 knockout mice lack functional NK cells and have severely reduced numbers of spleen and lymph node memory CD44hi CD8 T cells, thymic NK-T cells, as well as the thymus-independent population of intraepithelial lymphocytes in the gut, and show a reduced ability to control vaccinia infection (Kennedy et al., 2000). IL-15 is produced by human epidermal Langerhans cells and dermal DCs (Blauvelt et al., 1996; Castagnoli et al., 1999), human CD83 mature monocyte-derived DCs, CD34-progenitor derived DCs (Blauvelt et al., 1996; Jonuleit et al., 1997; de Saint-Vis et al., 1998), and mature BMderived mouse DCs (Josien et al., 1999). While many papers reported spontaneous production of IL-15 by DCs, at least two signals, CD40L and TRANCE, are known to upregulate or induce its production (Josien et al., 1999; Kuniyoshi et al., 1999). CD40L-induced IL-15 production was shown to be important for the effectiveness of DCs in inducing CTL activity (Kuniyoshi et al., 1999).

IL-15

IL-18

IL-15 (Grabstein et al., 1994; Burton et al., 1994) binds to a common IL-2Rγ and IL-2Rβ chains and has multiple IL-2-like activities, although it does not have sequence homology to IL-2. IL-15 induces the proliferation of CD4 and CD8 T cells, and promotes the generation of antigenspecific CTLs and the development of cytotoxic activity in NK cells. In addition, IL-15 costimulates proliferation and antibody production by B cells and induces proliferation of mast cells (reviewed by Kennedy and Park, 1996; Waldman and Tagaya, 1999). Recently, it has been shown that IL-15 is especially important for the survival of intraepithelial T cells, rather than for the induction of their effector functions (Lai et al., 1999), while another report demonstrated a key role of IL-15 in sustaining the persistence and slow division rate of memory CD8 T cells in the mouse (Ku et al., 2000). This activity of IL-15 was in apparent contrast to IL-2, which is an inducer of short-term activation of CD8 cells, but exerts a negative impact

IL-18 (Interferon γ-inducing factor; IGIF), is a potent enhancer of IFNγ production in T cells and an enhancer of NK-cell activity. Although it does not induce TH1 differentiation by itself, it enhances the expression of the IL-12 receptor, and synergizes with IL-12 in the induction IFNγ production in primed effector cells (Okamura et al., 1998; Akira, 2000; Chang et al., 2000). Like IL-1β, IL-18 is synthesized as an inactive precursor and requires further processing by the IL-1βconverting enzyme (ICE)/caspase 1 to acquire biological activity and to be secreted (Dinarello, 1999). Bioactive IL-18 is produced by mouse LCs and bone marrow-derived DCs (Stoll et al., 1998). Expression of IL-18 mRNA and secretion of IL-18 protein has been demonstrated in several populations of human DCs, including CD34 progenitor- and monocyte-derived DCs (Stoll et al., 1998; de Saint-Vis et al., 1998; Bohle et al., 1999; Gardella et al., 1999; Demeure et al., 2000). Like IL-12, IL-18 is induced by CD40-triggering and by

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stimulation of CpG oligonucleotides. In contrast to IL-12 production, which is reduced in mature DCs (Kalinski et al., 1999a; Demeure et al., 2000), IL-18-producing capacity increases in mature DCs (Demeure et al., 2000).

IFNα IFNα has multiple immunomodulatory and antiviral actions. Its antiviral activity depends both on a direct inhibition of virus replication as well as the activation of NK cells and CTLs. It also promotes CTL survival (Tough et al., 1996; Biron, 1998; Marrack et al., 1998). IFNα is involved in virus-induced DC activation, promoting DC survival, and participating in the induction of DC maturation (Luft et al., 1998; Cella et al., 1999; Radvanyi et al., 1999). IFNα may contribute to TH1 development by its direct ability to activate STAT4 in human T helper cells (Brinkmann et al., 1993; Frucht et al., 2000; Kadowaki et al., 2000), as well by upregulation of the IL-12Rβ chain (Rogge et al., 1997). However, at the same time IFNα inhibits the production of IL-12 (McRae et al., 1998). IFNα production is induced in murine DC cell lines, human peripheral-blood isolated and monocyte-derived DCs by enveloped viruses, including HSV or influenza virus (Eloranta et al., 1997; Milone and Fitzgerald-Bocarsly, 1998; Cella et al., 1999). IFNα is produced in especially high amounts by human CD11cIL-3R plasmacytoid DCs (Siegal et al., 1999), allowing this cell type to induce IFNγ-dominated TH1-like responses upon viral infection (Kadowaki et al., 2000).

Chemokines Activated DCs produce a wide array of chemokines that attract DCs themselves, CD4 and CD8 T cells, NK cells, and other immune cells, such as neutrophils and macrophages. This allows for recruitment of additional immune cells to the site of ongoing inflammation and facilitates their interaction with DCs. Several chemokines are produced spontaneously by freshly isolated DCs. Murine

Langerhans cells (LCs), spleen DCs, and bone marrow-derived DCs produce macrophage inflammatory protein 1γ (MIP-1γ) (Mohamadzadeh et al., 1996) a CC chemokine attracting both CD4 and CD8 T cells. Responsiveness to this factor does not require prior activation of T cells, making it a good candidate as a primary factor inducing local accumulation of potentially antigen-reactive cells. Human DCs spontaneously produce macrophage-derived chemokine(MDC)/stimulatedTcellchemotacticprotein (STCP-1) (Godiska et al., 1997; Chang et al., 1997). MDC, a ligand for CCR4 (Imai et al., 1998), is weakly chemoattractive for freshly activated T lymphocytes, but much stronger for long-term activated T cells. It also recruits DC and NK cells, but not monocytes, neutrophils, eosinophils, nor resting T lymphocytes. The selective activity of this molecule upon activated T cells makes it a likely candidate in the trafficking of activated/effector T lymphocytes to inflammatory sites. STCP-1 and some other DC-derived chemokines, TARC (thymus and activation-regulated chemokine) and MCP-4 (Hashimoto et al., 1999; Imai et al., 1999; Lieberam and Forster, 1999) preferentially recruit T cells displaying TH2-type cytokine profiles (Andrew et al., 1998; Imai et al., 1999), which predominate in chronic inflammation. The production of MDC/STCP-1 is enhanced by IL-4, the TH2 cytokine, which may result in a positive feedback loop promoting TH2-dominated chronic disease state. ABCD-1 (Schaniel et al., 1998) represents the murine orthologue of human MDC/STCP-1. A related DC molecule, ABCD-2, preferentially attracting murine TH2-type cells (Schaniel et al., 1999). In case of the above discussed molecules it remains to be confirmed whether DCs indeed produce these factor constitutively, or whether their production is just triggered by DC isolation. The same question applies to SDF-1 (stroma cellderived factor), a CXC chemokine produced by human skin CD1a dendritic cells (Pablos et al., 1999). While the above factors are likely to act mainly in peripheral tissues, another CC chemokine, DC-CK1 (Adema et al., 1997)/PARC

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(pulmonary and activation-upregulated chemokine) (Hieshima et al., 1997) preferentially attracts naïve CD45RA T cells. DC-CK1 production is specific for DCs, particularly the DCs present in germinal centers and T-cell areas of secondary lymphoid organs, consistent with its role during the initiation of primary immune responses. ELC/MIP-3β, constitutively produced by mouse DCs within the T-cell zone of secondary lymphoid tissues, has a similarly selective activity upon CCR7 (EBI-1)-expressing naïve T cells (Ngo et al., 1998). ELC strongly attracts both CCR7-expressing naïve CD4 and CD8 T cells, in addition to activated B cells. Thymic DCs constitute a source of TECK (thymusexpressed chemokine), a CC chemokine recruiting macrophages, DCs and thymocytes, and a factor involved in T-cell development (Vicari et al., 1997). Despite the ability of DCs to produce numerous cytokines in the apparent absence of any pathogen-related signals, the production of the majority of chemokines requires activation of DCs and their resulting maturation. Mature DCs are good producers of MIP-1α, MCP-1, IL-8 and RANTES (Zhou and Tedder, 1995; Caux et al., 1994b; Sallusto et al., 1999). Constitutive production of MDC by mouse LCs is rapidly upregulated following their local activation in the skin and accompanies the process of their maturation and migration to the lymph nodes (Tang and Cyster, 1999). The same was reported for mouse ex vivo matured spleen- and epidermis-isolated DCs as well as for monocyte-derived DCs in humans (Sallusto et al., 1999; Kanazawa et al., 1999). This feature is common for several other ‘constitutively’ expressed chemokines, such as MDC, PARC (Sallusto et al., 1999), TARC (Lieberam and Forster, 1999) and fractalkine (Kanazawa et al., 1999). The expression of membranebound fractalkine in human Langerhans cells and CD83 positive DCs in lymph nodes is also upregulated during activation of DCs, particularly via CD40 ligation (Papadopoulos et al., 1999). The requirement for DC activation is even more strict in the case of MIP-1α, MIP-1β and IL-8 production, which are absent in immature DCs but are rapidly induced after stimulation of

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DCs with LPS, TNFα or CD40 ligand (Sallusto et al., 1999). While the above factors were induced early in monocyte-derived DCs and expressed only transiently after activation, RANTES and MCP-1 were produced in a sustained fashion (Sallusto et al., 1999). Since DCs are one of the earliest infiltrating cells recruited to the site of ongoing inflammatory responses (McWilliam et al., 1996), their ability to attract multiple other components of the immune system is compatible with the role of DCs in amplification of local immune responses in the peripheral tissues. The ability of mature, lymph node type DCs to attract naïve and memory T and B cells suggests a role for DCs in the structural organization of lymphoid tissue.

MODULATION OF IMMUNOSTIMULATORY AND POLARIZING FUNCTIONS OF DCs BY TISSUES AND PATHOGENS Immature DCs residing in peripheral tissues can be activated by pathogens either directly or indirectly, resulting in a series of events collectively referred to as the maturation of DCs. Phagocytosis of bacteria, direct infection with some viruses, as well as contact with several bacterial toxins and components of bacterial wall, bacterial DNA, CpG oligonucleotides, etc., can all directly activate resting DCs, resulting in the initiation of the antigen-specific immune response and the elimination of the pathogen. However, many other pathogens have developed means of turning DCs into allies, either by inactivating infected DCs and rendering them tolerogenic (Gabrilovich et al., 1994; Grosjean et al., 1997; Salio et al., 1999) or by inducing the production of immunosupressive factors (Ruebusch et al., 1986; Farrel and Kirkpatrick, 1987; Vellupilai and Harn, 1994). The ability of some pathogens to control the functions of directly infected DCs highlights the importance of additional, indirect mechanisms of DC activation for effective induction of immuneresponses.Onesuchindirectmechanism

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may rely on the mediators of nonspecific tissue responses to pathogens, including GM-CSF, IL1β, TNFα or type 1 interferons, which can induce the maturation of resting DCs. Perhaps even more importantly, DCs can also be activated by the factors released from necrotic cells, such as the heat-shock protein (HSP)-60 (Ohashi et al., 2000) or HSP-70 (Todryk et al., 1999; Moroi et al., 2000). Such a direct translation of the cell damage into a source of DC-activating signals may serve as an additional, fail-safe alarm mechanism, or a ‘danger signal’ for the immune system (Matzinger, 1998). Interestingly, these different classes of exo- and endogenous maturation-inducing factors may utilize the same receptors (e.g.TLR-4 recognizes both HSP-60 and LPS (Ohashi et al., 2000)). The ability of these distinct types of signals to induce activation of DCs may reflect competition between the hostadapting capacity of pathogens and the adaptive ability of the immune system. The same competition may have led to the heterogeneityofmechanismsthatallowDCstoadapt their TH1/TH2-inducing capacity to the type of pathogen and to the type of the infected tissue. Direct infection with some viruses and bacteria and contact with many pathogen-related factors includingnucleicacidsandcomponentsofbacterial walls effectively induce either IL-12 or IFNα production. Both of these mediators induce STAT4 activation in human T helper cells, resulting in the induction of TH1-biased responses. However, many other intra-cellular pathogens have developed means to prevent DCs from producing factors like IL-12 and from inducing cellular immunity. Such intra-cellular parasites as Leishmania major or Babesia microti fail to induce IL-12 production, inducing instead the production of IL-12-inhibitory factors(Ruebusch et al., 1986; Farrel and Kirkpatrick, 1987). Also several viruses, including rhinoviruses (Stockl et al., 1999), human immunodeficiency virus (HIV), or measles virus (Yooetal.,1996;Karpetal.,1996)can impair IL-12 production. These examples underline the dangers associated with a direct interaction of antigen-presenting cells with pathogens and illustrate the benefits of additional indirect waysofjudgingthecharacteroftheinvader.

One possibility is the indirect modulation of the DC polarizing function by distinct mediators of pathogen-induced nonspecific tissue response. DCs matured under the influence of TNFα and IL-1β or LPS show intermediate IL-12-producing ability and induce the development of the TH0-like T helper cells. The additional presence of IFNγ during DC maturation results in effector DCs with remarkably high IL12-producing ability and TH1-inducing capacity (Vieira et al., 2000). In contrast, IL-10, glucocorticoids or PGE2 induce a TH2-inducing DC subtype, deficient in IL-12 production (reviewed by Kalinski et al., 1999b). Such environmental polarization of DCs may contribute to the deficient IL-12-producing ability of DCs isolated from mucosal tissues. Freshly isolated bronchial DCs and DCs from Peyer’s patches show impaired ability to produce IL-12 and are efficient inducers of TH2-type cytokines (Stumbles et al., 1998; Iwasaki and Kelsall, 1999). The different abilities of DCs matured in different environments may allow efficient adaptation of the effector immune mechanisms not only to the type of the pathogen, but also to distinct tissues. Such DC-mediated early polarizing signal, reflecting the modulation of DC functions by different tissues can explain the early development of different classes of immune response in a single lymph node, depending on whether the immunization occurred via a mucosal route or via the skin (Sangster et al., 1997). Differences in the TH1/TH2-inducing function of DCs may also result from intrinsic differences between distinct lineages of these cells. Mouse CD8αCD11b DCs were shown, at least in vitro, to favor the development of TH1-biased responses, as compared with their myeloid counterparts (Maldonado-Lopez et al., 1999; Pulendran et al., 1999). In contrast, human myeloid DCs are superior TH1 inducers compared with the DC-like CD11c IL-3Rα plasmacytoid cells that are deficient in IL-12-producing capacity and were proposed to be the ‘obligatory’ inducers of TH2-type responses (Rissoan et al., 1999). However, the observation that murine Peyer’s patch DCs, showing a

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REFERENCES

pronounced TH2-driving capacity, and predominantly TH1-inducing spleen DCs, have similar frequencies of CD11b and CD11b cells (Iwasaki and Kelsall, 1999) argues against a major role for the lineage differences in the above respect, suggesting the dominant involvement of the environment-guided functional adaptation of DC functions. While the ability of DCs to adapt to pathogen- and tissue-imposed requirements became first evident in case of myeloid DCs, lymphoid DCs also appear to have similar functional plasticity. It was recently shown that human IL-3Rα plasmacytoid DCs, originally appearing to be the ‘default’ inducers of TH2 responses, produce large amounts of IFNα in response to viral infection, resulting in induction of IFNγ-dominated responses (Kadowaki et al., 2000). Interestingly, the induction of IFNγ by human plasmocytoid DCs is accompanied by induction of substantial amounts of IL-10, a factor involved in the induction of CTL function of CTLs. The above observations indicate that a functional modulation of DCs may be a more general way of regulating the character of the immune response, than a selective presentation of a pathogen by DCs of different lineages. The ability of DCs to acquire pathogen- and tissue-related information is strictly regulated during the course of DC maturation. Newly activated DCs transiently increase the intensity of their antigen uptake and terminate this process after several hours, increasing instead the stability of their surface MHC/peptide complexes (Cella et al., 1997). This confers relative selectivity of antigen uptake, restricting it to the site and the moment of pathogen entry, and allows the pathogen-related antigenic signals to be retained for a prolonged period of time. Similarly, mature DCs acquire resistance to many of the environmental factors that can impair the stimulatory capacity of immature DCs. The same timeframe is relevant for the ability of DCs to pick up tissue-related polarizing signals (Vieira et al., 2000). These features allow for consolidation of distinct pathogen-related signals within a single migrating cell (Plate 6.1) and reduce the risk of their distortion, making

DCs particularly well-equipped to transmit the pathogen-related information from the affected peripheral tissues to the lymph nodes. Environmental signals present during the priming of naïve T helper cells drive their development into functionally distinct effector/ memory TH1 and TH2 cell subsets. Such polarized effector cells, showing only limited differences in surface phenotype, nonetheless being functionally stable and relatively resistant to repolarization, orchestrate the effector phase of the immune response (Mosmann and Sad, 1996). The ability of immature/sentinel-type DCs to receive the tissue- and pathogen-derived clues and to develop into type 1 and type 2 polarized effector/mature DC subsets appears to follow the same paradigm (Plate 6.2), providing the immune system with the plasticity required to fight different pathogens effectively, and adapting the character of response to particular tissues of the host.

ACKNOWLEDGEMENTS The authors thank Kathy Rakow for secretarial support. Grant support: NCI-1RO1CA73816-01 to Walter Storkus, MTL; NIH NCI-1PO1CA73743-01 to MTL, Olivera Finn; PAR-97-PI/Project 4 to MTL, Cara Wilson; and NCI 1RO1CA8201601A29 to MTL.

REFERENCES Adema, G.J., Hartgers, F., Verstraten, R. et al. (1997). Nature 387, 713–717. Akiba, H., Miyahira, Y., Atsuta, M. et al. (2000). J. Exp. Med. 191, 375–380. Akira, S. (2000). Curr. Opin. Immunol. 12, 59–63. Altare, F., Durandy, A., Lammas, D. et al. (1998). Science 280, 1432–1435. Anderson, D.M., Maraskovsky, E., Billingsley, W.L. et al. (1997). Nature 390, 175–179. Andrew, D.P., Chang, M.S., McNinch, J. et al. (1998). J. Immunol. 161, 5027–5038. Azuma, M., Cayabyab, M., Buck, D., Phillips, J.H. and Lanier, L.L. (1992). J. Immunol. 149, 1115–1123.

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PLATE 6.1 Dendritic cells as carriers of pathogen-related information. Immature ‘sentinel-type’ DCs in the tissues (left) sample their environment in search of antigens. They respond with activation and maturation when they encounter pathogenic microorganisms. Pathogens can activate DCs either directly (red arrow) or indirectly. Indirect DC activation can be caused either by the mediators of pathogen-induced inflammatory response of affected tissues (orange arrows) or by the factors released from dying cells, such as heat-shock proteins (yellow arrows). Apart from a direct or indirect impact of pathogens, the activation state of DCs is also affected by tissueproduced inhibitory factors, which may elevate the threshold for DC activation, especially at such immunoprivileged sites as the eye or in mucosal tissues. As a result of their activation, DCs increase their surface expression of co-stimulatory molecules and migrate to the lymph nodes, where they provide naïve T helper cells with so-called signal 1 (triggering of TCR by antigenic peptides bound to MHC molecules) and signal 2 (co-stimulation; mediated by surface-bound and soluble DC-related factors). Pathogen-related molecules and tissue-derived factors, either produced constitutively or induced by the pathogen, can also modulate the expression of polarizing factors by DCs (signal 3). These DC-transmitted signals inform naïve T helper cells about the identity, pathogenicity and the nature of the invader, and about the character of the affected tissue, allowing the induction of a highly specific immune response and the selective mobilization of the most appropriate effector mechanisms.

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PLATE 6.2 Polarized effector subsets of dendritic cells. Differences in the ability to induce TH1 or TH1 responses can be observed between immature DC populations in distinct peripheral tissues. Such heterogeneity may result from early polarization of DC precursors developing in different environments and, possibly, from intrinsic lineage-specific polarization of DC functions. Incoming pathogens can further modulate the TH1/TH2-inducing capacity of immature DCs; either directly or by distinct pathogen-induced inflammatory factors. In analogy to committed memory/effector T helper cells, which are relatively stable and resistant to repolarization, maturing DCs acquire significant resistance to polarizing factors, resulting in the development of polarized effector DC1 (TH1-inducing) and DC2 (TH2-inducing) subsets. *The factors involved in the induction of DC1 phenotype and/or mediating the TH1-inducing function of human IL-3α plasmacytoid DCs. **The factors inducing the TH2-promoting function of DCs but suppressing their immunostimulatory potential.

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7 Isolation, culture and propagation of dendritic cells Weiping Zou, Jozef Borvak, Florentina Marches, Shuang Wei, Tatyana Isaeva and Tyler J. Curiel Baylor Institute for Immunology Research, Dallas, Texas, USA

Man’s mind, once stretched by a new idea, never regains its original dimensions. Oliver Wendell Holmes

INTRODUCTION AND GENERAL CONSIDERATIONS

company. Different manufacturers may use different units to measure cytokine potency. Care should be taken to determine the correct concentrations of cytokines from different vendors. This chapter will focus on mouse and human DCs, but it should be noted that DCs have also been cultured from rhesus macaques (Messmer et al., 2000; O’Doherty et al., 1997; Stahl-Hennig et al., 1999), chimpanzees (Barratt-Boyes et al., 1996, 1997, 1998), rats (Stumbles et al., 1998), pigs (West et al., 1999), horses (Hammond et al., 1999; Siedek et al., 1999) and cows (Hope et al., 2000). Surprisingly, culture of DCs from dogs or cats has not been reported to our knowledge, although that will surely change soon. DC phenotype and function may be affected by the precursor cells from which the DCs were derived, mobilization factors, factors used to effect differentiation or maturation and the anatomic location from which the DC was recovered. Differential effects are well known, but have not been fully evaluated.

The discovery of effective culture conditions in which large numbers of DCs could be produced in vitro revolutionized their study. The historical background related to those early discoveries is discussed in detail elsewhere. This chapter will focus on in vitro techniques to cultivate DCs. Mobilization of peripherally circulating DC progenitor cells (Siena et al., 1995; Ratta et al., 1998; Raje et al., 1999; Rondelli et al., 1999) and distinct DC subsets using agents such as G-CSF or Flt-3L (Maraskovsky et al., 1996; Pulendran et al., 1999a, 1999b) are discussed briefly here, but are covered more fully in Chapter 10. Mouse DC cell lines for in vitro culture are discussed in Chapter 12. All reagents described are recombinant and commercially available unless otherwise specified. R&D Systems (Minneapolis, MN, USA) produces a variety of high-quality mouse and human cytokines. We have no financial interest in this

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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MOUSE DENDRITIC CELLS General principles Most published reports of mouse DCs use cells derived from Balb/c or C57Bl/6 mice. However, there is no reason not to use mice of other genetic backgrounds for certain applications. Consideration should be given to the experimental needs and available reagents and cell lines. In general, mice 8–12 weeks old yield sufficient numbers of DCs, precursor or progenitor cells, and organs of sufficient size to be easily manipulated. Mice younger than 6 weeks may yield fewer cells or be too small to manipulate easily, and mice older than 12 weeks may be more expensive with no additional gain in cell yields. Consideration should be given to the effects of anesthetizing or euthanatizing agents. We use CO2 asphyxiation followed by cervical dislocation for minimal perturbation. DC isolation from tissue usually involves a digestion procedure followed by DC enrichment (buoyant density or depletion of lineage cells) and a final sorting procedure. The final cell population may range from 70 to 100% pure depending on the efforts expended. Collagenase contaminated with trypsin-like activity may reduce the expression of certain DC surface molecules. Trypsinlike activity in collagenase may be determined as described (Vremec and Shortman, 1997). Nycomed publishes an excellent technical manual on density gradient materials and their properties (distributed by Accurate Chemical, 800-645-6264; www.accuratechemical.com). Characteristics of cell surface marker expression may change depending on whether DCs are studied freshly isolated or after in vitro culture (Vremec and Shortman, 1997) and after treatments such as trypsinization (Anjuere et al., 1999). Finally, the composition of myeloid versus lymphoid DCs varies both from organ to organ within mice, and from mouse strain to mouse strain within the same organ. These differences are detailed in Chapters 2 and 8 and section IV. DCs of apparently the same phenotype may function differently depending

on their anatomic location (Iwasaki and Kelsall, 1999). Age- and sex-related differences in DC composition are not as well defined. Some investigators use specific pathogen-free mice to avoid pathogen-specific effects.

Bone marrow-derived DCs The technique described by Inaba and associates (1992) has been modified over the years. Using sterile dissecting equipment, fascia and muscle of the thigh are parted and the femur and tibia removed. Muscle remnants are removed with sterile gauze and the bones sterilized with ethanol. Bone ends are cut with scissors, and marrow is flushed out with medium. Debris is reduced by passage through a wire mesh and cells are pelleted by centrifugation. Red blood cells are lysed in 17 mM Tris–HC1 pH 7.2 plus 144 mM NH4Cl for 10 minutes on ice. Care must be taken not to allow this lysis step to progress much beyond 10 minutes, or viability and yield of DCs may be compromised. T and B lymphocytes and class II cells are depleted using antibody-coated magnetic beads. Further purification can be achieved by fluorescence-activated cell sorter (FACS) sort on CD11b and CD11c, or by using CD11c-coated paramagnetic beads. Cells are plated at 3 to 4 million/well in 3 mL of medium supplemented with murine GM-CSF in sixwell plates overnight. Granulocyte depletion with specific antibodies, and intermittent removal of granulocytes from culture may be necessary if a final sort is not accomplished, as GM-CSF is a powerful stimulus to granulopoiesis. Cells can be cultured in RPMI-1640 supplemented with 10% heat-inactivated fetal calf serum, 10 mM HEPES buffer, 2 mM glutamine, 50 µM 2-mercaptoethanol and antibiotics. Iscove’s modified Dulbecco’s minimal essential medium may be substituted for RPMI-1640. Clusters of loosely adherent DCs may be attached to spreading, adherent macrophages. Note that adherent cells through day 4 will become DCs, whereas after 5 or 6 days, with further differentiation, the DCs become nonadherent. Typical DC yields are approximately 5  106/mouse after 6–7 days of culture, if both

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femurs and tibias are used. Maturation of DCs is effected by LPS or TNFα treatment. CD34 cells obtained from mouse bone marrow will differentiate into DCs when cultured with GM-CSF plus Flt-3L (Vecchi et al., 1999). CD34 cells may be obtained from mouse marrow by depletion of lineage, nonadherent cells followed by sorting.These cells will differentiate into CD11bCD11c and CD11b/dullCD11c subsets upon culture with GM-CSF,TNFα and SCF (Zhang et al., 1998) which may be sorted by FACS on day 6 forindependentculture.Cellsproliferatesuchthat the cultures are split on day 4, and fresh cytokines are added every 3 or 4 days. Addition of TGFβ to these three cytokines preferentially drives precursors down a Langerhans DC differentiation pathway (Zhang et al., 1999). Contaminating macrophages and granulocytes are greatly reduced using CD34 sorted cells. Addition of SCF or Flt-3L to bone marrow cultures will increase the yield of DCs ultimately derived, especially when used in combination. High-quality DCs will also be derived in the absence of these growth factors, but yields will be lower.

Spleen-derived DCs Short-term cultures Spleen cells may be dissociated mechanically by teasing with sterile forceps (Winzler et al., 1997). In addition, following mechanical disruption, spleen matter may be further processed by digestion with collagenase and DNAse (Leenen et al., 1998). Spleens may also be mechanically disrupted by other means including gentle compression between etched glass microscope slides. Enzymatic digestion of spleen tissue is reported not to alter the FACS phenotype, DC yield, or functional properties of the differentiated cells (Leenen et al., 1998). Debris is partially clarified by passage through a wire mesh and the cells are pelleted by centrifugation. Red blood cells are lysed as described above. Recovered cells are collected and depleted of lineage cells as described above, followed by culture in GM-CSF plus IL-4. Spleen- and bone marrow-derived DCs from

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culture of precursor cells in GM-CSF plus IL-4 were equivalent in induction of an allogeneic MLR, but bone marrow derived DCs were better than spleen derived DCs in processing and presentation of exogenous antigen (Garrigan et al., 1996). However, the precise method of generating bone marrow precursor cells was not specified. Long-term cultures Spleen cells are plated at 500 000/mL in medium supplemented with 10–20 ng/mL mouse GMCSF supplemented with 30% filtered supernatant from the NIH 3T3 cell line. The cytokines and supernatant are replaced every 3 or 4 days. Nonadherent and loosely adherent DCs are passaged after about 2 weeks, and adherent cells are discarded. These cells are reported to maintain an immature phenotype and remain functional for up to 1 year (Winzler et al., 1997). These DCs are class II and express CD40, CD80 and CD86. Up to 20% are class IIbright suggesting a mature phenotype. Maturation of the remaining cells can be effected with LPS, IL-1β or TNFα. TNFα-matured cells have reduced capacity to capture and present the soluble antigen ovalbumin, and secrete significantly more bioactive IL12 than immature DCs, as expected. Direct isolation Spleen cells contain variable amounts of CD4CD8αDEC-205loCD11bhi and CD4CD8α DEC-205hiCD11blo DCs that can be sorted to high purity for immediate use or cultured in GM-CSF overnight prior to use (MaldonadoLopez et al., 1999). Flt-3L or GM-CSF treatment of animals significantly increases the number of both types of DCs, which may then be sorted from spleen for immediate use (Maraskovsky et al., 1996; Pulendran et al., 1999b). A third major spleen DC subset which is CD4CD8αDEC-205loCD11bhi was recently reported (Vremec et al., 2000). Details of these studies and the immunologic consequences of these distinct DC subsets are presented in Chapter 27.

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Peyer’s patch, thymus, lymph node and lung-derived DCs Because of the small size of organs, material from syngeneic animals may be pooled for analysis. Organs are cut into small fragments and digested with collagenase and DNAse I (Vremec and Shortman, 1997). At the end of digestion, EDTA is added for 5 additional minutes with agitation to disrupt DC–T cell aggregates and undigested fragments are removed by passage through a steel sieve. Addition of 5 µg/mL DNAse may help to prevent material from clumping. Recovered cells are washed and the low-density fraction collected over iso-osmotic Nycodenz (Pharma, Oslo, Norway). T and B lymphocytes, granulocytes, macrophages and other cells are removed using antibody-coated Dynabeads (Lake Success, NY, USA). Shortman and colleagues (Vremec and Shortman, 1997) no longer add anti-CD25 oranti-CD11bantibodiesto avoid removal of variable populations of DCs expressing these markers. DCs are maintained in medium containing buffered EDTA and fetal calf serum for final analyses. Recent technical advances in FACS staining and sorting have helped to define two major thymic and three splenic DCsubsets and to characterize their phenotypes (Vremec et al., 2000). Resident lung DCs are collected by lavaging and perfusing lungs of killed animals to remove lung macrophages and blood. Removed tissue is removed and digested with collagenase and DNAse and freed of large debris by passage through a metal mesh. Low-density cells are collected over discontinuous Percoll. Cells are then separated according to adherence properties (Pollard and Lipscomb, 1990). FACS sorting or bead selection may replace adherence purification. Peyer’s patch DCs are isolated by treatment of tissue with dithiothreitol, HEPES and EDTA to remove epithelial cells, digested with collagenase and DNAse, and CD11c cells are positively selected from single-cell suspensions using beads. FACS sorting CD11cB220 cells following bead selection allows for high purity (98–100%) recovery of DCs away from CD11cB220 B cells (Iwasaki and Kelsall, 1999).

Skin-derived DCs The skin is a significant source of both Langerhans DCs (isolated from epidermal material) and interstitial DCs (isolated from dermis). Sterilely prepared ears are split in half with forceps and incubated in 0.5% trypsin which separates epidermal and dermal tissue planes. Epidermal cells (containing Langerhans DCs) are obtained by filtering trypsin-treated material through steel mesh, followed by washing and collection of the low-density fraction over Optiprep (Life tech/GibcoBRL, Rockville, USA). Langerhans DCs will account for 2–10% of the starting material, and preparations will be approximately 20–30% pure. Additional density gradient centrifugations can be used to increase purity (Schuler and Steinman, 1985). Note that trypsin treatment will reduce DEC-205 expression, but expression will recover after 1 day of in vitro culture. An alternative procedure is to prepare epidermal and dermal sheets as just described and culture them for 24 hours in 24-well plates in medium containing no cytokines, or containing mouse GM-CSF, which induces migration of both DCs and keratinocytes out of the ear and into the tissue culture medium. Nonadherent DCs may be carefully removed from the mostly adherent keratinocytes to yield Langerhans DCs from epidermis (60–80% purity) and interstitial DCs from dermis (30–40% purity), by collection of the low-density fraction as described above (Lenz et al., 1993). This method allows for parallel isolation and analysis of Langerhans DCs and interstitial DCs, but the Langerhans DCs here may not be exactly equivalent to the Langerhans DCs freshly isolated from ears owing to the 24-hour incubation in GM-CSF and to selection of migratory cells.

Monocyte-derived DCs Monocyte-derived DCs (MDDCs) are the most commonly used type of human DC, but are only rarely prepared from mice largely because the yield is small, and there are other easier methods available to produce mouse DCs.

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Nonetheless, DCs may be derived from circulating mouse monocytes. Blood is obtained by cardiac puncture. PBMCs are separated by density gradient centrifugation (Lympholyte, Amersham Pharmacia, Biotech, Uppsala, Sweden) and plated at 8–10 million cells/well in a six-well plate. After 2 hours of incubation, nonadherent cells are gently rinsed away. The

remaining adherent cells are then cultured further in 20 ng/mL murine GM-CSF plus 20 ng/mL murine IL-4. Fresh cytokines are added on days 3 and 6 and nonadherent DCs are collected on day 7 or 8. The yield of mouse MDDCs is approximately 1  105/mouse compared with 7  106/mouse when bone marrow is used. The significantly lower yield reflects the

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40%

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DC/ well

FIGURE 7.1 Properties of mouse peritoneal macrophage-derived DCs (φDCs). (a) FACS demonstrates typical CD11bCD11c cells, which are also CD8α (not shown). A portion are class IIdim/, which greatly increases following maturation. (b) These φDCs elicit a significant allogeneic mixed lymphocyte reaction which is mediated by the CD11c cells in this mixed population. CD11c cells were purified using magnetic beads. These are representative data from over 20 experiments with similar results. DENDRITIC CELL BIOLOGY

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low frequency (3–6%) of circulating monocytes in mice, and the poorer proliferative capacity of these circulating monocytes compared with bone marrow cells. Significant morphologic, phenotypic and functional differences between these MDDCs and marrow-derived DCs were not demonstrated. Mouse MDDCs effected significant antitumor immunity in vivo (Schreurs et al., 1999).

cells in these cultures effected a weak allogeneic MLR. Thioglycollate-elicited peritoneal macrophages do not differentiate into DCs with GMCSF alone or in combination with TNFα as described (Rezzani et al., 1999). A detailed description of these cells is in progress (T. Isaeva et al., manuscript in preparation).

FETAL LIVER DCs Peritoneal cell-derived DCs Resident cells The mouse peritoneal cavity harbors a small resident population of myeloid cells that are generally regarded as being macrophages. Peritoneal contents are recovered by gentle lavage. TNFα cooperates with GM-CSF to differentiate these resident peritoneal cells into DCs that are efficient in activating an allogeneic mixedlymphocytereaction(MLR),butareslightly less efficient than control spleen-derived DCs in this regard. Thioglycollate-derived peritoneal macrophages do not differentiate into DCs under these conditions (Rezzani et al., 1999).

Mature mouse DCs will differentiate from fetal liver progenitor cells using a stromal cell feeder layer, GM-CSF, Flt-3L and SCF to induce differentiation, followed by GM-CSF plus TNFα to induce maturation (Zhang et al., 2000).

Tumor-derived DCs DCs isolated from mouse myeloma (Dembic et al., 2000) or GM-CSF and CD40L-transduced colon carcinoma cells (Chiodoni et al., 1999) will activate tumor-specific T cells without exogenous antigen loading. Parallel results in humans have not been reported to date.

Cell lines Elicited macrophages Macrophages are elicited by inoculation into the peritoneum of 0.5 mL of 3% thioglycollate. Macrophages are collected 4 or 5 days later, adhered to plastic plates and cultured with 20 ng/mL murine GM-CSF plus 20 ng/mL murine IL-4 (R&D Systems) (optimized doses, based on extensive preliminary experiments). Nonadherent cells with DC morphology are apparent after about 4 days of culture. After 8 days of culture, nonadherent cells are CD11bCD11cCD8αDRhi by FACS, consistent with a murine myeloid DC phenotype (Figure 7.1a). These macrophage-derived DCs elicit a significant allogeneic MLR (Figure 7.1b) and present a superantigen efficiently to autologous spleen T cells. Addition of TNFα did not enhance differentiation of these DCs as was observed for human macrophages, and was toxic to cultures when used at greater than 2.5 ng/mL. Adherent

Several murine DC lines are available, some of which have many features typical of primary DCs. There are few reports of immortalized mouse tumor cell lines that will differentiate into DCs under the influence of cytokines.

HUMAN DENDRITIC CELLS Monocyte-derived DCs Classical The best-studied human DCs are those derived from monocytes, which may be obtained by plastic adherence of Ficoll-Hypaque (Amersham Pharmacia Biotech) (Sallusto and Lanzavecchia, 1994; Bender et al., 1996; Kiertscher and Roth, 1996; Chapuis et al., 1997;) or Lymphoprep (Life tech/GibcoBRL) (Romani et al., 1996) purified

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PBMCs which may be further purified over a metrizamide (Zhou and Tedder, 1996) or Percoll (Sallusto and Lanzavecchia, 1994) density gradient column, by elutriation (Czerniecki et al., 1997) or by more complicated sorting strategies (Ito et al., 1999). Various methods to reduce contaminating lymphocytes, platelets and other cells include plastic adherence, sorting, paramagnetic beads, neuraminidase-treated sheep red blood cells and other techniques. However, significant differences in DCs differentiated from these monocytes collected and purified in these distinct ways have not generally been noted. PBMCs can be obtained by phlebotomy or leukopheresis. Whole, heparinized blood may be kept at room temperature in a plastic syringe overnight and MDDCs can be differentiated from these cells the following day. The yield and quality of these MDDCs varies, although goodquality MDDCs can be obtained. In our experience, leukopheresis cells held overnight produce poor-quality MDDCs. Ten million PBMCs are adhered to plastic sixwell plates, or approximately 500 000 purified monocytes in 3 mL of medium per well are cultured in medium containing GM-CSF and IL-4. For cytokine-mobilized samples, adjustments in the numbers of PBMCs per well will be necessary to reflect the altered proportions of mononuclear cells. Cells will start to detach within 2 days, and may form small clusters. Fetal calf serum may contain trace amounts of endotoxin, TGFβ or other factors (Strobl et al., 1997). Some investigators thus use serum-free medium or autologous plasma instead of fetal calf serum. To produce immature MDDCs, monocytes are cultured with GM-CSF plus IL-4 for 5–7 days. A good culture will contain 95–99% CD1aCD14CD83lo/ cells. If cells are cultured much beyond 8 days, they will undergo spontaneous maturation with upregulation of CD83 and loss of ability to effect pinocytosis of antigen. Reported amounts of GM-CSF and IL-4 used to differentiate MDDCs vary considerably. We use much less IL-4 (5 ng/mL, R&D Systems) than generally reported to induce efficient MDDC differentiation (Zou et al., 2000). Cultures at day 7 should contain 5–10%

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CD14 and CD83 cells. If higher percentages are found (or if CD1a expression is low), endotoxin or Mycoplasma contamination, poor cytokines or poor monocytes are possibilities. Maturation can be effected by an additional 2–2.5 days of culture with 1 µg/mL Escherichia coli LPS. Other investigators report using much lower doses (10 ng/mL) to effect maturation. DCs may also be matured by a variety of other agents, well-described elsewhere. Interferonα is reported to mature MDDCs (Luft et al., 1998), a finding disputed by others (Cella et al., 1999b). Although not extensively studied, significant differences in phenotype and function may exist in MDDCs matured with different factors (Cella et al., 1999b) and for varying lengths of time. In addition to GM-CSF plus IL-4, other culture conditions will effect monocyte to MDDC differentiation. For example, adherent PBMCs cultured with Flt-3L plus IL-4 will differentiate into CD1a DCs that effect a significant allogeneic MLR. Addition of TNFα or CD40L-expressing cells enhances differentiation and maturation. GM-CSF alone or with TNFα will not effect MDDC differentiation (Brossart et al., 1998). It is not clear that specific culture systems are advantageous over others, aside from cost considerations. Langerhans DCs Circulating monocytes will differentiate upon culture with GM-CSF, IL-4 and TGFβ into cells with many features of immature Langerhans DCs (Geissmann et al., 1998). Addition of LPS or TNFα plus IL-1β induces maturation, although CD40 ligation is more effective in this regard (Geissmann et al., 1999). These monocytederived Langerhans DCs effect a significant allogeneic MLR comparable to MDDCs. MDDCs differentiated with GM-CSF plus IL-4 also differentiate into Langerhans DCs following addition of TGFβ (Geissmann et al., 1998). A small subpopulation of cells will express langerin (Valladeau et al., 1999), whose detection may be a reasonable substitute for Birbeck granule detection.

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Macrophage-derived DCs

variable CD1a, and effect an allogeneic MLR, but significantly less efficiently than nonadherent cells. We call these macrophage-derived DCs ‘ΦDCs’ (phi-DCs). These ΦDCs secrete up to 10-fold less IL-12 than MDDCs following LPS activation, yet induce up to 9-fold more T cell interferon γ through β-chemokines (Zou et al., 2000). ΦDCs secrete up to 10-fold more β-chemokines than MDDCs, express significantly more antigen/ apoptotic body capture receptors and effect up to 3-fold more pinocytosis, although the immunologic consequences are largely unknown at present. A detailed analysis of differences between macrophage- and monocytederived DCs has been performed (Zou et al., 2000).

Macrophages differentiated in vitro from monocytes Ten million PBMCs are seeded into wells of a six well plate, adhered for 2 hours, and nonadherent cells are then gently washed away. Adherent cells are cultured in 25 ng/mL M-CSF for 4 or 5 days to differentiate them into macrophages. M-CSF is then removed and the medium is replaced with 100 ng/mL GM-CSF plus 5 ng/mL IL-4. Nonadherent cells in M-CSF should be returned to culture with adherent cells. After 2–4 days, a detached population will appear, which are the nonadherent macrophage-derived DCs. These cells are optimal around day 5–7 of culture in GM-CSF plus IL-4. Over half of CD1a cells will coexpress CD14 (Figure 7.2a). Maturation with E. coli LPS significantly increases CD1a expression, reduces CD14 expression (Figure 7.2b), and induces CD83. Approximately 50–70% of these macrophages will become nonadherent DCs under these conditions. Adherent cells express

(a)

Tumor-associated macrophages in malignant ascites Malignant ascites is a rich source of tumorassociated macrophages that will differentiate

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FIGURE 7.2 FACS analysis of ΦDCs derived from M-CSF in vitro macrophages. (a) Immature ΦDCs are up to 99% CD1a, but the majority are also dually CD14. (b) CD14 expression is significantly reduced following maturation with E. coli LPS. ΦDCs also express CD4, CD11c, CD25, CD36, CD40, CD54, CD68, CD80, CD86, CD95, αvβ3 and αvβ5 and upregulate CD83 upon maturation (not shown). Data are representative of over 10 experiments with similar results. DENDRITIC CELL BIOLOGY

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into DCs upon culture with GM-CSF plus IL-4. These ΦDCs exhibit some phenotypic and functional characteristics of ΦDCs derived from M-CSF macrophages (Figure 7.3). Cell composition of ascites varies greatly. In malignant ascites from ovarian carcinoma, there is a preponderance of macrophages (40–95% of cells). We plate these at 2 million cells/well in six well plates. Ascites from chronic hepatitis C on the other hand may contain predominantly lymphocytes. We thus seed more cells per well, depending on the percentage of CD14 cells present. These ΦDCs are surprisingly efficient stimulators of an allogeneic MLR in most cases. They present superantigen and tetanus toxoid to autologous T cells, and activate tumor-associated T cells. A sample of ascites may contain up to 600 million macrophages, which is sufficient for adoptive immunotherapy using ΦDCs. We thus propose that ascites macrophages represent a significant pool of easily accessible DC precursors that merit consideration for use in antitumor immunity protocols. A detailed study of these tumor-associated macrophage-derived (a)

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ΦDCs has been performed (Borvak, et al., manuscript in preparation). Other tissue macrophages We have differentiated ΦDCs from macrophages in ascites of patients with cirrhosis (including due to chronic hepatitis C infection) and pulmonary alveolar macrophages by culture in GM-CSF plus IL-4. Detailed studies of these ΦDCs are in progress in our laboratory.

Peripherally circulating progenitor cells General principles Progenitor cells are distinctly different from precursor cells in that the former proliferate. Thus, final DC yield can be increased by expanding the progenitor pool prior to terminal DC differentiation. Some protocols include an initial expansion period (usually with Flt-3L, SCF or both, in combination with other (b)

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1000 3000 10,000 Number of dendritic cells/ well

FIGURE 7.3 Macrophages from malignant ascites are DC precursor cells. Macrophages from ascites of a patient with ovarian carcinoma were cultured in GM-CSF plus IL-4 for 6 days. (a) These ΦDCs are CD1aDR and have other FACS features similar to M-CSF macrophage-derived ΦDCs (see the legend to Figure 7.2). (b) Ascites macrophage-derived ΦDCs elicit a significant mixed lymphocyte reaction using naïve, CD4CD45RA T cells as the responder cells. Macrophages induced 2200 counts per minute in this experiment. Data are representative of over 20 experiments with similar results. DENDRITIC CELL BIOLOGY

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cytokines) to boost DC progenitor cell numbers, followed by a differentiation step (which usually includes GM-CSF plus TNFα). Serum-free culture generally requires more cytokines to effect differentiation than serum-replete culture. More primitive precursor cells may require more cytokines to effect differentiation. DC progenitor cells tend to differentiate down at least two distinct DC pathways that can be separated for independent culture and terminal differentiation. CD34 cells Steady-state cells CD34 cells are usually collected by leukopheresis as insufficient CD34 cells may be obtained from phlebotomy. Nonmobilized leukopheresis may yield up to 109 mononuclear cells, affording approximately 5 million CD34 cells. Mononuclear cells are collected by FicollHypaque density gradient centrifugation and the resultant mononuclear cell population is enriched for CD34 cells by depletion of lineage cells. CD34 cells can then be positively selected with paramagnetic beads, by FACS sort or over an affinity column. Becton-Dickinson (San Jose, CA, USA) makes a useful kit for FACS detection of circulating DCs and precursors, and Coulter (Miami, FL, USA) will launch their product soon. Growth factor-mobilized cells Mobilizing agents such as G-CSF greatly enhance the yield of CD34 circulating hematopoietic progenitor cells in cytopheresis specimens. CD34 cells obtained from nonmobilized or mobilized mononuclear cells are purified as just described and cultured in GMCSF plus TNFα. After 5–9 days distinct CD1a and CD14 precursor cells can be sorted and cultured for 4–7 more days in GM-CSF plus TNFα to effect terminal differentiation. CD34 cells may also be cultivated without separation of precursor cells to afford a mixed DC population. As CD34 cells proliferate, cultures are split around day 5, at which time additional cytokines

are also added. Significant differences in CD34 cells obtained after cytokine mobilization compared to steady state have not been described, but may exist. Herbst et al. (1996) used chemotherapy plus G-CSF to mobilize peripheral blood CD34 cells which were positively selected on a CellPro Ceprate column (no longer commercially available). Cells grown in IL-3, IL-6 and SCF for 8 days, followed by GM-CSF plus IL-4 thereafter expanded 100-fold after 21 days of culture, but retained an immature Langerhans DC phenotype. In earlier studies, this group demonstrated that a different cytokine cocktail supported CD34 cell proliferation and Langerhans DC differentiation, but these cells matured and further differentiated into interstitial DCs (Mackensen et al., 1995). Twice as many DCs may be derived from CD34 cells in G-CSF mobilized peripheral blood compared with bone marrow. The DCs differentiated from these mobilized CD34 cells are qualitatively similar to those in bone marrow when differentiated with GM-CSF, TNFα, SCF and Flt-3L (Ratta et al., 1998). Continuous flow conditions (Cellmax, CELLCO, Germantown, MD, USA) may slightly improve the function (allostimulation) of DCs differentiated from CD34 cells using GM-CSF, TNFα, SCF, Flt-3L and TGFβ under serum-free conditions, although nonflow (static) culture produced the largest yield of DCs (Soligo et al., 1998). The timing of introduction of IL-4 into cultures may also help to boost yields of DCs (Rosenzwajg et al., 1998). SCF and Flt-3L are equivalent in increasing CD34 cell proliferation, but used together provide an additional augmentation of DC yield (Rosenzwajg et al., 1998). CD34 cells may be divided into CD34CLA and CD34CLA precursor cells. The latter differentiate into Langerhans DCs upon culture with GM-CSF plus TNFα (Strunk et al., 1997). GM-CSF mobilized CD34 cells in cancer patients differentiate into DCs when cultured with GM-CSF plus TNFα, but are less efficient in activating an allogeneic mixed lymphocyte reaction than MDDCs (Triozzi and Aldrich, 1997).

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DC precursors defined by CD11c or ILT3/1 expression Mononuclear cells from cytopheresis are purified by Ficoll-Hypaque density gradient centrifugation and depleted of lineage cells (in some cases after an additional elutriation step) using a fluorochrome-labeled antibody cocktail and beads, followed by staining for DR and CD11c. Two distinct populations of linDRCD11c (CD11c) and linDRCD11c (CD11c) cells are readily sorted by FACS. These cells have features of DCs including expression of CD80 and CD86. The CD11c cells are CD45RO, express the GM-CSF receptor and differentiate into DCs (myeloid DCs) under the influence of GM-CSF plus IL-4. CD11c DCs on the other hand are CD45RA, express high IL-3α receptor but very low GM-CSF receptor, and differentiate into lymphoid DCs and mature in response to IL-3 plus CD40L (Grouard et al., 1997; Rissoan et al., 1999), or IL-3 plus TNFα (Kohrgruber et al., 1999). IL-4 alone or in combination with IL-10 kills CD11c cells (Kohrgruber et al., 1999; Rissoan et al., 1999), but toxicity can be overcome by pretreatment with CD40L or interferon γ (Rissoan et al., 1999) or after several days of in vitro culture with IL-3 and with GMCSF (Kohrgruber et al., 1999). A single cytopheresis of unmobilized peripheral blood can yield 1–8 million CD11c cells and about onefifth as many CD11c cells. Mobilization with growth factors will increase these numbers and may alter the ratios of cells recovered. Lin PBMCs presumably containing both CD11c and CD11c cells yield predominantly CD11c cells after 2 days of culture in monocyteconditioned medium, whereas CD11c cells predominate following culture without cytokines (O’Doherty et al., 1993). Both CD11c and CD11c precursor cells elicit a significant allogeneic MLR when freshly isolated, but CD11c cells are generally better in this regard. CD11c precursors can be matured by culture in TNFα, thereby enhancing MLR induction. Regarding antigen loading, CD11c cells are regarded as being poor in phagocytosis and pinocytosis when freshly isolated. Most

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groups (Grouard et al., 1997; Rissoan et al., 1999) report that this capacity does not improve with culture. However, there may be a window of maximal antigen capture capacity acquired during in vitro culture (Kohrgruber et al., 1999). These features should be taken into account in specific experimental conditions. Lymphoid DCs rarely survive more than 5 days under these conditions (Ito et al., 1999; Kohrgruber et al., 1999; Rissoan et al., 1999). Improved methods to culture these rare, fastidious cells are in progress in several laboratories. Effective means to induce significant cell expansion of either DC precursor have not been reported. Details of these two DC precursor cells are reported in Chapters 1, 3 and 6. CD11c circulating DC precursor cells may be divided into CD1aCD11c and CD1aCD11c cells. The former differentiate into Langerhans DCs with GM-CSF, IL-4 and TGFβ (Ito et al., 1999). Alternatively, DC precursor cells can be classified based on differential expression of immunoglobulin-like transcript (ILT)3/1 expression in lineage circulating cells (Cella et al., 1999a). LinILT3ILT1 cells appear to correspond to the CD11c myeloid DC precursor cells described above, whereas LinILT3ILT1 cells appear to be essentially equivalent to the CD11c lymphoid DC precursor subset. A hallmark of both ILT3ILT1 and CD11c subsets is their striking production of type I interferons following viral infection (Cella et al., 1999a; Rissoan et al., 1999), which we use to confirm functional identity in vitro. G-CSF preferentially mobilizes CD11c DC precursor cells, although it also mobilizes CD11c DC precursors, whereas Flt-3L mobilizes large numbers of both in humans (Pulendran et al., 1999a). The effects of cyclophosphamide, GM-CSF, SCF, and other agents on mobilizing specific subsets in human DCs has not been reported in detail as of this writing, but are under investigation by several groups. Adherent cells Populations of PBMCs have been defined following overnight adherence in the absence of

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cytokines. Tightly adherent cells are CD14bright macrophages. Loosely adherent cells are further depleted of T and B lymphocytes and CD14 and CD16 cells, followed by sorting to remove CD45RO cells. This yields two populations of circulating DCs based on light scatter characteristics in FACS that may correspond to lymphoid (low side scatter) and myeloid (high side scatter) DC precursors (Kiertscher and Roth, 1996). These freshly isolated DC populations effect a significant allogeneic MLR.

functionally similar, including those obtained from patients with multiple myeloma (Raje et al., 1999). Comparison of adherent bone marrow-derived DCs to bone marrow CD34 cell-derived DCs has not been reported to our knowledge. Care must be taken in this technique to aspirate only a few milliliters of bone marrow per aspirate, or samples may become contaminated with blood cells that also adhere and differentiate into DCs.

Cord blood DCs CD2 cells CD2 monocytes may represent circulating myeloid DCs, and effect an efficient allogeneic MLR without culture in cytokines (Crawford et al., 1999).

Bone marrow-derived DCs CD34 cells CD34 cells in bone marrow are positively selected using paramagnetic beads following density gradient centrifugation of mononuclear cells (Young et al., 1995). GM-CSF plus TNFα cooperate to induce DC differentiation, and addition of SCF further increases the number of resultant DCs, similar to cord blood CD34 cells (Caux et al., 1992, 1996). Addition of Flt-3L to SCF or substitution for it may also be used to augment DC yield. These DCs effect a significant allogeneic MLR. CD34 bone marrow cells in HIV-infected and cancer-bearing patients can be derived in similar fashion. Adherent cells Bone marrow aspirates are subjected to FicollHypaque density gradient centrifugation, and the 2-hour adherent cells are then cultured in GM-CSF plus IL-4 to yield DCs. This technique has been used successfully in tumor-bearing patients, including those with marrow involvement. In a comparative study, adherent bone marrow cells and adherent PBMCs cultured in GM-CSF plus IL-4 were phenotypically and

CD34 cells Mononuclear cells are obtained by FicollHypaque density centrifugation, and contain many more CD34 cells than normal adult blood. CD34 cells can be obtained through positive selection using Miltenyi beads (Caux et al., 1996) or Dyna Beads followed by Detach-a-Bead treatment (Arrighi et al., 1999) with equivalent yields, or can be obtained by FACS. Beads are faster and less expensive; FACS yields higher purity. Cells are cultured in GM-CSF plus TNFα. After 12 days of culture, a CD1a cell population with features of immature DCs is evident (Caux et al., 1992). Three distinct cell populations are evident after 5 or 6 days of culture: CD1aCD14 cells, CD1aCD14 cells and CD1aCD14 cells (Caux et al., 1996). Furthermore, addition of SCF to GM-CSF and TNFα at culture initiation increases the yield of differentiated DCs (Szabolcs et al., 1995). Thus, cord blood CD34 cells may be cultured in GM-CSF, TNFα and SCF all together to effect simultaneous proliferation of progenitors and differentiation of precursors from them. After 5 or 6 days, these cells can be FACS sorted to obtain the CD1aCD14 and CD1aCD14 cells, which are then seeded at 1 to 2  105/mL and cultured 6 or 7 days further in GM-CSF plus TNFα. SCF is not added at this point, as these cells no longer proliferate significantly (Caux et al., 1996). Langerhans DCs are derived from the CD1aCD14 population. Interstitial DCs resembling monocyte-derived DCs are derived from the CD1aCD14 population. Langerhans DCs express dendrites by

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around day 9 of culture, whereas CD14-derived DCs lag a few days behind and may not exhibit significant dendrites until around day 12 (Caux et al., 1996). CD1aCD14 cells do not differentiate into DCs. Both types of DCs produced in this fashion express typical myeloid DC T cell co-stimulatory molecules including CD40, CD80, CD86 and DR, and are potent inducers of an allogeneic MLR (Caux et al., 1996). CD14-derived DCs express M-CSF receptor and will differentiate into macrophages upon culture with M-CSF, whereas CD1a-derived Langerhans DCs will not differentiate into macrophages under these conditions and die (Caux et al., 1996). Specific functional differences and detailed descriptions of the differences in expression of surface molecules between these two DC subsets are discussed in Chapters 1 and 8. Culture of CD34 cells under serum-free conditions in vitro (Bio Whittaker, Walkersville, MD, USA) with GM-CSF, TNFα, SCF and TGFβ induces significant differentiation of CD1a DCs (Strobl et al., 1996). Addition of Flt-3L almost doubles the percentage of CD1a DCs and quadruples their absolute numbers after 10 days of culture compared with GM-CSF, TNFα, SCF and TGFβ without Flt-3L (Strobl et al., 1997). The DCs differentiated in the presence of Flt-3L have a normal morphologic appearance and Langerhans DC FACS profile, and elicit a significant allogeneic MLR. SCF and Flt-3L are comparable in enhancing DC output in combination with GM-CSF, TNFα and TGFβ, yet the two together effected an additive augmentation of total DC output. Study of fewer cytokines in serum-replete conditions, and analysis of DCs differentiating from CD1a versus CD14 precursors was not performed here. TGFβ in these cultures is reported to favor differentiation of Langerhans DCs, but Flt-3L alone does not compensate for TGFβ in inducing DC differentiation or suppressing differentiation of other myeloid cells in serum-free conditions. It is likely that exogenous TGFβ replaces serum TGFβ in these effects. Such culture conditions, while complicated and expensive, may be relevant to groups developing adoptive

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immunotherapy protocols wherein cells grown ex vivo will later be infused back into human patients. CD34 cord blood cells can be induced to proliferate with thrombopoietin plus Flt-3L in human or fetal calf serum-supplemented medium to increase cell numbers over 16 000fold in 4 weeks. Addition of SCF boosts cell numbers an additional 5-fold after 4 weeks. Two populations of CD34CD1aCD14 and CD34 CD1aCD14 cells are derived. These cells can then be induced to differentiate into DCs with no further proliferation by culture in GM-CSF plus IL-4. The CD34CD1aCD14 cells also differentiate into DCs upon culture with GMCSF plus TNFα, followed by GM-CSF plus IL-4 after proceeding through a CD14 intermediate. The CD34CD1aCD14 cells did not differentiate into DCs upon culture with GM-CSF plus TNFα. Thus, these precursor cells are not equivalent to those described by Caux et al. (1996) or Rosenzwajg et al. (1996), but may be the same as those described by Herbst et al. (1996). The proliferating CD34cell-derived cell population can also be frozen, thawed later and induced to differentiate into DCs. As in other long-term proliferating DC precursor cultures, it is necessary to split cells weekly to maintain an optimal growth density of around 1 to 2  105/mL (Arrighi et al., 1999). G-CSF mobilized CD34 cells purified by CD40 expression after overnight incubation in TNFα differentiate into a CD34CD40 population with high potential to differentiate into DCs with GM-CSF, TNFα, Flt-3L and SCF. The CD34CD40 population is poorer at differentiating into stimulatory DCs, and is proposed to be of potential use in allograft tolerance (Rondelli et al., 1999). CD11c cells DRlinCD11c cells with features of lymphoid DCs have been demonstrated in cord blood cells, where they comprise about 0.3% of the total cell population (Sorg et al., 1999). CD34negative selection is also used to purify these cells. These cells are linCD4CD11cCD45RA,

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and express IL-3α receptor but not CD80, CD83 or CD86 and are comparable to similar cells circulating in peripheral blood. CD11c precursor cells are much rarer. Effects of culture of these cells in cytokines has not yet been reported.

DCs From fetal tissue Liver Chimeric organ culture is produced by allowing human fetal hepatocytes to migrate into a fragment of SCID mouse thymus, allowing study of thymic DC development. Human fetal livers are mechanically disrupted and purified over a Lymphoprep density centrifugation gradient. Lineage and CD38 cells are depleted using Dynabeads. Resultant CD34CD38/dimlin cells are purified from these cells by FACS. Thymus lobes are obtained from gestational age day 14 or 15 fetal SCID mice. Human hepatocytes are incubated with fetal mouse thymus in a hanging drop culture in Terasaki plates to allow hepatocytes to migrate into the organ culture. Thymus lobes are then placed in organ culture and cells are recovered at later time points by mechanical disruption of the thymus. CD4DR human cells defined as DCs are apparent between 4 and 11 days of culture, with numbers plateauing around day 12 at approximately 500–1500 cells/lobe. Day 11 DCs effect a significant allogeneic MLR and have FACS features of DCs (Plum et al., 1999). A potential advantage of this system over using DCs sorted from human thymus is that in this organ culture, the DCs may be more or less synchronized and all at the same developmental stage at the same time. In sorted human thymus cells, on the other hand, all DC developmental stages are likely to be present. However, the extremely small yield of cells from this technique remains problematic. Thymus An alternative system uses human thymus instead of fetal liver as the source of progenitor

cells to seed the thymus. In this system, postnatal thymus is collected from children undergoing heart surgery, or from aborted 16- to 18week gestational age fetuses. Thymus is minced and passed through a steel mesh to disrupt tissue, and large aggregates are discarded. Lineage cells are depleted using paramagnetic beads and CD34CD38/dim cells are collected by FACS. CD34CD38/dim cells differentiate into CD4CD40DRhi lin cells after culture in GMCSF, TNFα and SCF. Some cells also express CD1a. Insufficient cells were available for functional analyses (Res et al., 1996). Using similar strategies, a CD1aCD3 CD4CD8 precursor cell can be obtained from postnatal human thymus that will differentiate into a DC with lymphoid DC characteristics (CD1aCD45RADRhi and expressing IL-3α receptor chain) upon culture with IL-3 and CD40L-expressing mouse L cells (Res et al., 1999). Unlike some other thymic progenitor/ precursor cells, these CD1aCD3CD4CD8 cells are unable to differentiate into T lymphocytes or natural killer cells.

Lymph node, tonsil and spleen-derived DCs Lymph nodes or tonsils are cut into small pieces, digested with collagenase and DNAse and cells are collected, pooled and subjected to density gradient centrifugation over Percoll (Grouard et al., 1996, 1997). Low-density cells are depleted of lineage cells using paramagnetic beads. Other cells including CD34, CD11c and CD1a cells can also be removed in this fashion. Lineage cells are stained with FITC-conjugated antibodies, DR is labeled with Tricolor and CD11c with phosphatidylethanolamine (PE). Populations of DRCD11clin and DRCD11clin DC precursor cells can be obtained as described for isolation from peripheral blood. The CD11c population expresses myeloid markers including CD13 and CD33 and is CD45RAdimCD45ROhi, whereas the CD11c population is negative for myeloid markers and is CD45RAhiCD45ROdim. Germinal center DCs in lymph nodes are

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CD1aCD3CD4CD11cCD20 and may be discriminated from interdigitating DCs by higher CD40 and lower CD4 intensity in the former (Grouard et al., 1996). Lymphoid DC precursor cells in tonsils are CD3CD4CD11cDRlin cells that can be cultured with IL-3 plus CD40Lexpressing recombinant mouse cells for up to 6 days to effect differentiation (Grouard et al., 1997). These tonsil CD11c DC precursor cells are likely equivalent to circulating CD11c DC precursor cells (Kohrgruber et al., 1999; Rissoan et al., 1999). Although there is no theoretical reason not to study DCs and DC precursor cells in human spleens, this organ has received little study simply because of the inconvenience in getting the organ. DC isolation from human gutassociated lymph tissue can be accomplished as has been described for mice (Iwasaki and Kelsall, 1999).

DCs from skin Although Langerhans DCs may be derived from CD34 precursor cells and from monocytes, they were originally described in skin. Human skin for such studies can be obtained following surgeries (Lenz et al., 1993). Skin is split cut with a keratome, and trypsinized to separate dermal and epidermal layers, and to disaggregate epidermal tissue into a single cell suspension. The epidermis (containing Langerhans DCs) may then be detached from the dermis with forceps and pooled with other epidermal sheets in medium supplemented with 20% fetal calf serum to stop trypsin action. Final disruption of the epidermal sheets into a single cell suspension is accomplished by vigorous pipetting, and the cells are then washed three times. This cell suspension contains a mixture of Langerhans DCs, keratinocytes and other cells. To enrich for Langerhans DCs, cells are suspended at 4  106/mL in medium and applied to a Lymphoprep density gradient. The interface is collected after centrifugation and washed. Keratinocytes are depleted by adherence on collagen-coated glass plates and a repeat density centrifugation step for 20 minutes yields a cell suspension containing 70–95% Langerhans DCs.

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DCs from tumors Acute and chronic leukemias Peripherally circulating CD34 leukemic blasts from both myeloid and lymphoid leukemias will differentiate into DCs in vitro under appropriate conditions (Cignetti et al., 1999). Leukemic blasts were obtained from PBMCs of a patient with M2 acute myelogenous leukemia and used without further manipulation, as they were 95% CD34 at collection. Cells were cultured with GM-CSF, IL-4 and TNFα. Medium with fresh cytokines was replaced every 5–7 days. After 20 days of culture, approximately one-third of the cells had upregulated CD80, CD86, CD1a and CD40, and expressed typical myeloid DC morphology. These cells effected an allogeneic MLR, but poorly. The leukemic origin of these cells was confirmed by detection of the leukemic monosomy 7 abnormality. Interestingly, the majority of these DCs continued to express CD34. DCs were also derived from acute B-cell leukemic blasts by culture of these cells in soluble, recombinant CD40L plus IL-4. The resultant ‘DCs’ upregulated CD80, CD86 and CD83, and downregulated the B-cell marker CD19 slightly from 82% to 70%, but did not express CD1a. They induced an allogeneic MLR slightly better than the untreated leukemic blasts, but absolute T cell activation was poor. The leukemic origin of these cells was confirmed by detection of the leukemic bcr-abl translocation protein or aberrantly expressed surface molecules. As DCs theoretically will activate T cells better than leukemic cells, DCs produced from leukemic cells expressing a tumor-specific antigen might make useful reagents for anticancer immunotherapy protocols. Further work will be required to determine the utility of these cells. Whether or not one wishes to classify them as DCs, the cytokine-induced upregulation of T-cell co-stimulatory molecules on leukemic blasts may nonetheless have therapeutic utility, although not all leukemic cells upregulate co-stimulatory molecules under these conditions (Mutis et al., 1998; Cignetti et al., 1999).

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Adherent PBMCs or CD34 cells from patients with chronic myelogenous leukemia will differentiate into DCs which express the Philadelphia chromosome (Heinzinger et al., 1999) and stimulate tumor-specific immunity (Choudhury et al., 1997; Eibl et al., 1997). Myeloid precursor cells in acute myelogenous leukemias also differentiate into DCs, some of which express tumor-specific genetic defects (Choudhury et al., 1999). Progenitor cells in chronic myelogenous leukemia differentiate into DC-like cells with calcium ionophores (Engels et al., 1999).

lymphoid DCs will be presented elsewhere (Zou et al., manuscript submitted). Solid tumors As of this writing, there were numerous histologic descriptions of DCs in tumors, including detection of Langerhans DCs, myeloid DCs and lymphoid DCs, but no reports regarding the functional capacity of these tumor-associated DCs in humans (Chiodoni et al., 1999).

Cytokine-independent DC differentiation

Malignant ascites We have demonstrated that cells in ascites of humans with ovarian carcinoma are 40–95% CD14 macrophages. These macrophages differentiate into DCs when cultured with GM-CSF plus IL-4. A typical ascites fluid sample of 4 L can yield 300–800 million macrophages. In addition, malignant ascites may be rich in lymphoid DCs that are CD4CD11clin, express IL-3α receptor and have the morphologic appearance of peripheral blood lymphoid DCs by light microscopic examination of Giemsa-stained cytocentrifuge specimens. To obtain these lymphoid DCs, ascites cells are purified over a FicollHypaque density centrifugation gradient. CD14 and CD3 cells in the pellet are depleted using Miltenyi beads, and the CD4CD11c cells are obtained by FACS. Culture with 10 ng/mL IL-3 may enhance differentiation. Yields of lymphoid DCs are approximately 450 000–900 000 after FACS sorting, which is a figure comparable to what can be obtained following cytopheresis of peripheral blood. These lymphoid DCs are functional in elicitation of an allogeneic MLR and secretion of interferon α following viral infection. Lymphoid DCs were not detected in significant numbers in ascites of patients with cirrhosis or chronic hepatitis. Thus, only a subset of patients with ascites will have sufficient numbers of lymphoid DCs for in vitro study. Given the ease and safety of collection, and the quantity of cells obtained, ascites lymphoid DCs represent a significant new source of these cells for in vitro studies. Details of these tumor-associated

By CD40–CD40L interactions Adherent PBMCs are cultured with irradiated, recombinant, mouse L cells expressing human CD40L and analyzed after 7 days of culture. CD40L expressed on the surface of mouse L cells induces significant upregulation of DR, CD80, CD86 and CD54 that is comparable to that effected by GM-CSF plus IL-4. However, about half the cells remain CD14, which is significantly greater than with GM-CSF plus IL-4. CD40 is also upregulated, but not to the extent seen with GM-CSF plus IL-4. Addition of IL-4 to CD40L effects further upregulation of CD40 and decreased CD14 expression. CD1a expression is also significantly increased by CD40L, although a substantial percentage of cells (up to 80%) remain CD1a negative. DCs differentiated by exposure to CD40L alone effect an allogeneic MLR that is comparable to that effected by DCs differentiated with GM-CSF plus IL-4. Others have been unable to replicate this finding. The difference may be explained by different starting cell populations, experimental conditions or contaminating cells in adherent PBMCs (FloresRomo et al., 1997; Brossart et al., 1998). CD34 cells cultured with irradiated CD40Lexpressing mouse L cells differentiate into DCs as demonstrated by expression of CD80, CD86 and DR, but cells do not express CD1a or CD40 (as do CD34 cells cultured in GM-CSF plus TNFα), and also exhibited other differences in surface molecule expression. Thirty to forty per

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cent of CD34 cells differentiate into DCs under these conditions (more resembling interstitial than Langerhans DCs). Function was proven by significant elicitation of an allogeneic MLR (Flores-Romo et al., 1997). A mouse anti-CD40 agonist antibody induces monocytes to differentiate into DCs, in a process augmented by addition of GM-CSF plus IL-4 (Zhou et al., 1999). Calcium ionophores Addition of the calcium ionophores A23187 or ionomycin to elutriated monocytes induces de novo CD83 expression, upregulation of CD40, CD80, CD86, class I and DR and downregulation of CD14 expression over 20–72 hours. Calcium ionophore-treated cells are morphologically distinct from cytokine-induced DCs in increased vacuole formation and increased endosomal and other ultrastructural features. Calcium ionophore-treated monocytes activate a significant allogeneic MLR and activate both CD4 and CD8 T lymphocytes when assayed after 40 hours of culture. The elicitation of an allogeneic MLR is more efficient than control MDDCs cultured in GM-CSF plus IL-4 (Czerniecki et al., 1997). However, the assay was performed after these DCs had also been in culture for 40 hours, which was insufficient time for complete cytokine-driven differentiation. Thus, the efficiency of elicitation of an allogeneic MLR by calcium ionophore-derived DCs compared with that by cytokine-derived DCs is not clear from these data. A similar technique was applied to the myeloid cell line HL-60 and CD34 bone marrow cells. Calcium ionophore-treated cells elicit a significant MLR in HL-60 cells that is further boosted by addition of GM-CSF plus interferon γ to A23187. Nonetheless, classical MDDCs are more efficient in induction of an allogeneic MLR. Bone marrow CD34 cells were cultured with SCF, GM-CSF and TNFα for 6 days to induce expansion and differentiation. Cells were then washed and cultured in GM-CSF, TNFα and A23187. After 2 or 3 days these cells upregulated T-cell co-stimulatory molecules and effected a

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modest increment in an allogeneic MLR compared with control CD34 cells (Koski et al., 1999). CD33 precursor cells in human chronic myelogenous leukemia acquire DC properties following treatment with A23187 (Engels et al., 1999). Reverse transmigration across endothelium Monocytes contacting an endothelial surface in the reverse transmigration direction will differentiate into DCs, particularly after undergoing phagocytosis (Randolph et al., 1998). This special case of monocyte to DC differentiation is discussed in detail in Chapter 21. Miscellaneous culture conditions Thyroid hormones enhance monocyte to DC differentiation possibly by induction of autocrine GM-CSF and TNFα (Mooij et al., 1994; Delemarre et al., 1995). Vitamin D (Singh et al., 1999; Penna and Adorini, 2000), steroids (Singh et al., 1999), prostaglandin E2 (Kalinski et al., 1998), bacterial LPS (Palucka et al., 1999) and other factors (Gabrilovich et al., 1998; MenetrierCaux et al., 1998) inhibit DC differentiation or maturation in vitro. Retinoic acid may play a role in DC differentiation (Kreutz et al., 1998).

Special considerations DCs grown for adoptive immunotherapy in humans Many groups are studying the immuneenhancing effects of human DCs grown ex vivo, which are loaded with antigen (with or without maturation) and then adoptively transfused back into human subjects. These types of trials require special attention to the nature of the cell collection, culture conditions, differentiation agents, maturation agents and antigens used. In the United States, the Food and Drug Agency closely scrutinizes these trials and must approve them before they can be implemented. Cells must be grown in specialized facilities called

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Good Laboratory Practice facilities. These are loosely defined and not as stringently controlled as the Good Manufacturing Process facilities used to manufacture drugs and biologics by pharmaceutical and biotechnology companies. Culturing DCs from human immunodeficiency virus (HIV)-infected individuals Monocytes and bone marrow CD34 cells from HIV-infected patients will differentiate into DCs when cultured with GM-CSF plus IL-4 (Triozzi and Aldrich, 1997; Chougnet et al., 1999). These cells will be infected with variable amounts of live HIV, depending on the clinical status and the antiretroviral treatment status of the patient at the time the cells are collected. As there is always the potential for infectious virus to be present in cultures of these cells, even if the patient has no detectable circulating virus, it is prudent to consider cultivating these cells under BL2 or higher level laboratory containment. G-CSF has been used successfully and safely to mobilize CD34 cells in HIV-infected individuals (Slobod et al., 1996). On currently available, highly active antiretroviral therapies, the monocyte-derived DCs from HIV-infected individuals express normal levels of T-cell co-stimulatory molecules and DC differentiation markers and effect a significant allogeneic MLR that is comparable to that effected by MDDCs from HIV-seronegative individuals. IL-12, IL-10 and β-chemokine production following bacterial or CD40L activation is also comparable to that seen in uninfected subjects (Chougnet et al., 1999). Of interest, circulating DCs in HIV-infected individuals may express naturally processed HIV peptides when examined in vitro (Fernandez et al., 1998). Cell lines Several investigators have reported differentiating DCs or DC-like cells from permanent human cell lines such as K562, HL-60 or U937 cells with conflicting results. The relationship of these DCs to any of the DCs discussed above has not yet been clearly established.

ACKNOWLEDGEMENTS We acknowledge the tremendous efforts of our colleagues, from whose work we have extracted summaries. We regret not mentioning the work of others owing to space limitations. This work was supported by NIH grants AI39379 and AI44322, the Baylor Endowment, and Golfers against Cancer (to T.J.C.). Thanks to Elizabeth Kraus for expert assistance with flow cytometry and to Dr. Ruth Berggren for critical review of the manuscript.

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8 Phenotypic characterization of dendritic cells Derek N.J. Hart, Kelli MacDonald, Slavica Vuckovic and Georgina J. Clark Mater Medical Research Institute, Mater Misericordiae Hospitals, South Brisbane, Queensland, Australia

We think in generalities, but we live in details. Alfred North Whitehead brane phenotype identifies DCs as CD45 leukocytes that express high levels of MHC class II molecules in the absence of markers associated with other leukocyte lineages including CD3, CD15, CD16, CD56, CD19, CD20 and CD14. These so-called lineage-negative (lin) populations of leukocytes are referred to as DC preparations, but the details with regard to cell preparation, the monoclonal antibodies (mAb) used, and source of serum additions, etc. may have profound influences on the characteristics of the resulting DC populations. At least one population of DCs develops from committed myeloid precursors derived from the pluripotent stem cell. The description of a lymphoid precursor-derived DC certainly provoked much thought and needs to be reconciled with the phenotypic definition of several subpopulations of DCs. The production of dendritic-like cells from in vitro culture protocols has improved experimental productivity but has also increased the complexity of the DC populations, which need to be considered both

INTRODUCTION Definition of DCs Dendritic cells (DCs) are a population of leukocytes, that have specialist antigen-presenting cell (APC) functions. As such, they link the innate and adaptive immune systems. Defined originally on morphological grounds, DCs are now better defined by functional criteria as described in earlier chapters. These criteria include the ability (1) to take up, process and present antigen (Ag), (2) to migrate selectively through tissues, and (3) to interact with, stimulate and direct primary T- and B-lymphocyte responses (Hart, 1997). However, more recently the ability to subclassify DCs has raised the possibility of different functional subsets. Thus once again, the phenotype (particularly cell membrane antigen phenotype) of DCs assumes critical importance for understanding DC biology, and certainly for exploiting them safely in clinical interventions. The basic surface memDendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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phenotypically and functionally. In rewriting this chapter we have felt compelled to provide an interpretation of the currently identified DC populations and their putative differentiation prior to describing the surface markers used to identify them and relating this information to DC function.

Differentiation pathway It is possible to define a number of phenotypically distinguishable populations related to the stage of DC differentiation. The DC originates in the bone marrow (BM) and migrates into the blood as an immature population before forming an extensive network of interstitial DCs in most nonlymphoid organs except the brain, parts of the eye and testes (Hart, 1997). Migration of these interstitial DCs into the afferent lymphatic system and then to the T-lymphocyte areas of the lymphoid organs designated as the interdigitating DCs (IDCs), is induced by a variety of stimuli, including inflammatory mediators, microbial products and other ‘danger’ signals. The phenotype of the cell changes throughout the differentiation and migration corresponding both to the location and functional state of each particular DC population. Upon this basic differentiation pathway one can layer some potential subclassifications or extensions, which may need more experimental justification to become widely accepted. First, it is possible there is early separation of an epithelial-associated versus nonepithelialassociated surveillance DC population. This makes potential biological sense. The possibility that blood monocytes (Mo) as opposed to DCs traffic to tissues and differentiate into monocyte-derived DCs (Mo-DCs) is becoming accepted. This at first sight makes less biological sense as a primary APC, and we have proposed that it may reflect a secondary boost pathway for developing an amplified APC population (Vuckovic et al. 1999). An alternative view is that certain phenotypically defined subsets of blood monocytes are destined to differentiate into tissue DCs. The so-called lymphoid DC population is a real conundrum. Mouse data appear

compelling (Maraskovsky et al., 1996), but in humans the subpopulations generate confusion and the term has been applied to different populations, none of which match the properties of mouse DCs. The possibility that some of these populations represent relatively fixed stages of differentiation merits consideration. A summary of these possibilities has been published previously (Hart, 1997). A more stylized version, acknowledging the uncertainty of the ‘lymphoid’ lineage, allowing individual antigen expression to be classified was developed for the 7th International Leucocyte Differentiation Antigen Workshop (Plate 8.1).

Species issues Langerhans cells (LCs) were originally identified in human skin on the basis of their morphology and ATPase staining. The morphological, cytochemical and functional description of DCs following isolation from mouse spleen was a major step forward (Steinman and Cohn, 1973). Subsequently similar cells were identified in other organs of rats, pigs, rabbits, sheep and humans (Drexhage et al., 1979; Hart, 1997). However, it was difficult to define DCs further due to the lack of specific markers. Additional complications arose from the differential expression of some markers between species making the criteria used to identify DCs in one species not always relevant in others. For example, the expression of MHC class II is restricted to APCs in the mouse but is expressed by APCs, kidney tubules and activated T lymphocytes in humans. Certain critical reagents, e.g. CD14 and CD1 mAb, are not yet readily available for studying these molecules in mice. Moreover mice have only one CD1 molecule (CD1d) compared to several in humans. At a practical level, most mouse studies use a complex mixture of spleen DCs (again the preparative method is crucial), and human studies generally use one form or another of blood DCs. Now cultured mouse BMDCs are used commonly and these likewise are very different populations. The inability to match DC subpopulations between mouse and human (and other species) is a major challenge.

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CELL SURFACE ANTIGENS ASSOCIATED WITH DCs The production of monoclonal antibodies that recognize DC-specific markers has proved difficult. No lineage-defining reagents have been produced as yet but a series of ‘first generation’ mAb are proving to be useful reagents.

Markers commonly used to define mouse and rat DCs Three mAbs recognize relatively DC-specific epitopes on mouse DCs. Marginal zone spleen DCs can be isolated by the 33D1 rat mAb (Nussenzweig et al., 1982), although the details on the structure of this antigen are unavailable. A second rat mAb, NLDC-145, (Kraal et al., 1986) recognizes the DEC-205 antigen, expressed by DCs in the T-lymphocyte areas, including the splenic white pulp (and not the marginal zone DCs or LCs) and by thymic cortical epithelium and activated macrophage (Mφ). A cDNA clone has been isolated encoding the mouse DEC-205 antigen (Jiang et al., 1995), suggesting that the molecule is related to the Mφ mannose receptor. The third mAb, the rat N418 mAb that binds a CD11c (β2-integrin family) epitope (Metlay et al., 1990) is expressed in high density on mouse DCs but is also found on other leukocytes including Mφ. CD11b reagents have been used more recently to define myeloid DCs. A mouse mAb, OX62, has been produced that recognizes an integrin expressed predominantly on rat DCs (Brenan and Puklavec, 1992). Similarly, the OX41 mAb, which recognizes a member of the SIRP family, has been useful in identifying rat DC subpopulations (Liu et al., 1998).

Markers used to define human DC populations The complex task of defining a human DC phenotype is dependent on the tissue of origin, method of preparation and state of activation. The use of certain well-defined markers to analyse linHLA-DR populations has proved fruitful.

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Reagents which detect HLA class II products define human blood DC populations as the density of MHC class II expression varies with cell activation which alone defines human blood DC subpopulations. Members of the CD1 family of molecules, which have structural similarity with MHC class I molecules, are expressed by cortical thymocytes and differentially by DC populations. There are five discrete gene products CD1a, b, c, d and e. LCs express CD1a and variable amounts of CD1c (Porcelli and Modlin, 1999). Dermal or migrating LCs are reported to express CD1b (Richters et al., 1996). All three members of the CD1 gene family, CD1a, CD1b and CD1c, are probably expressed by the IDCs draining the skin (Cattoretti et al., 1987). Blood and tonsil DCs do not express CD1a. Both the absence and expression of CD1c has been noted on blood DCs and this remains controversial (Hart and McKenzie, 1988; Xu et al., 1992; Egner et al., 1993a) and may depend on reagents used. A more recent analysis of human blood DCs suggests that CD1a is not on linHLADR blood DCs but that CD16 defines a subset (Ito et al., 1999). The integrin CD11c is present on a subset of blood DCs and other tissue DC subsets (O’Doherty et al., 1993; Thomas and Lipsky, 1994). As it is also present on Mo and Mφ, its use is best confined to defining subpopulations. The three first-generation mAbs to human DCs, CD83, CMRF-44 and CMRF-56, recognize antigens primarily expressed on activated or cultured DCs. The CD83 mAb, HB15a, was raised against transfectants expressing the HB15 cDNA. It stains cultured human blood DCs, LCs and some IDCs in the lymph node as well as showing reactivity with activated B lymphocytes (Zhou and Tedder, 1995). Analysis of the cDNA indicated that this antigen is a member of the Ig gene superfamily (Zhou et al., 1992). The molecule appears to make a functional contribution to DC–T lymphocyte interactions (D. Munster, unpublished) and it is released from the cell surface in vitro (Hock et al., in press). Reagents to the mouse homologue are expected shortly. The CMRF-44 mAb binds an antigen

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expressed in high density on cultured or differentiated/ activated DCs (Hock et al., 1994; Fearnley et al., 1997). Tissue staining and functional analysis suggest that the CMRF-44 antigen is associated with an active stimulatory DC population. The antigen, which has not been characterized biochemically as yet, is expressed at low levels by B lymphocytes, and by Mo and Mφ following treatment with high dose IFNγ (above physiological levels). The CMRF-56 mAb identifies another distinct differentiation/activation marker on cultured DCs (Hock et al., 1999). Efforts to characterize the 95-kDa antigen are under way. Another mAb, CMRF-58, appears to define yet another DC differentiation/activation antigen. This reagent defines three populations of cultured blood DCs on the basis of their expression of an antigen that has not been identified as yet. A neo epitope of C9, described on human blood DCs is recognized by the mAb X-11, but this reagent has had limited use (Wurzner et al., 1991). The Lag mAb identified a cytoplasmic molecule present within LCs (Strunk et al., 1997). This molecule has been re-identified as part of a molecular screening programme and re-named langerin (Valladeau et al., 1999). It is a C type lectin and appears to be expressed selectively by the epithelial DC differentiation pathway. Functionally, the molecule appears to be involved in cell membrane invagination, Birbeck granule formation and possibly cytoskeletal association (Valladeau et al., 2000). Other cytoplasmic markers, e.g. CD68 associated with lysosomal granules, have been described on DCs (Hart and McKenzie, 1988; Prickett et al., 1988). More recently DC–LAMP has been identified as an antigen associated with mature DC lysosomal granules, detected by cytoplasmic staining (de Saint-Vis et al., 1998). Several other molecules of the lectin family have been described on human DCs. Type I (multidomain) molecules include the macrophage mannose receptor and DEC-205, thought to be associated with antigen uptake. The type II (single domain) lectins include some regulating molecules (discussed later) and also the

molecule DC-SIGN. This molecule first identified as placental gp120-binding lectin was reidentified as an ICAM-3-binding molecule present on Mo-DCs (Geijtenbeek et al., 2000a, 2000b). It may have limited expression on other DCs but because of its HIV p120-binding properties it is the subject of widespread attention.

THE SURFACE PHENOTYPE OF DIFFERENT DC POPULATIONS Although the ability to define human DC subpopulationshasimprovedsignificantly,thisremainsa complex area as cross comparison of populations and data requires considerable insight. Predictably, the direction of DC differentiation and other leukocytes is driven by transcription factors, and as predicted, their investigation is also becoming a fruitful area of research.

Bone marrow precursors A subpopulation of human CD34 BM cells are immunostimulatory and may represent a DC precursor (Egner et al., 1993c). These cells still appear to have the capacity to differentiate into both the lymphoid and myeloid lineage. The predominant population of CD34, CD33 cells present in human BM appear to be the precursor population for the myeloid DCs. The freshly isolated precursor appears to be CD14 but a CD14 precursor may evolve during in vitro culture. An alternative CD34 precursor is thought to differentiate along an independent pathway to provide LagCD1 epithelial-associated LCs (Strunk et al., 1996). Certainly data on progenitor populations capable of generating DCs is becoming available from mouse knockout models. One model indicates that Pax5-deficient animals can generate myeloid DCs from dedifferentiated ‘committed B lymphoid progenitors’ (Nutt et al., 1999), whilst CD19 pro-B cells can differentiate into DCs when cultured with IL-1β, IL-3, IL-7 and TNFα (Bjorck and Kincade, 1998). Deletion of the Ikaros genes prevents myeloid but not lymphoid

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reveals very few lin HLA-DR/ DCs (Figure 8.2a). The presence of a linHLA-DR population (mainly basophils but potentially including other rare cell populations) needs to be remembered. Finally, as illustrated in Figure 8.2a the presence of a lin/ shoulder is inevitable after immunodepletion, and additional populations may be included if the gate for analysis or sorting is set well into the lin/ region. Two subsets of blood DCs have been identified: CD11cCD123/lo and CD11cCD123 (Olweus et al., 1997), but the situation is already more complicated. Analysis of the human linHLA-DR population for CD11c and CD123

differentiation in the mouse (Wu et al., 1997). Human studies involving Ikaros mutants also disrupted myeloid DC production (Galy et al., 2000).

Blood DCs Fresh blood DCs The heterogeneity of the preparations of lin blood DCs has always been evident (Egner et al., 1993a; O’Doherty et al., 1994) but has been somewhat underappreciated. An analysis of PBMCs for standard lineage (CD3, CD14, CD11b, CD16 and CD19) markers and HLA-DR

(a) PBMC

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≤1%

102 CD3; CD14; CD11b; CD16; CD19

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SORT PURIFY

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CD11c

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CD123

Detection and isolation of human blood DCs. PBMCs were isolated from healthy donors by Ficoll density gradient centrifugation. (a) Two-colour flow cytometry with HLA-DR and a lineage mAb cocktail (CD3, CD14, CD11b, CD16 and CD19) demonstrate linHLA-DR DCs comprising 1% of PBMCs. PBMCs were enriched for DCs by depletion of lin cells by immunomagnetic beads, followed by purification of lin cells by FACS sorting. (b) Three colour flow cytometry with HLA-DR, CD11c and CD123 reveals three distinct populations among linHLADR DCs. FIGURE 8.2

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in fact provides clear evidence for three blood subpopulations: (1) CD11cCD123/loHLADR, (2) CD11cCD123HLA-DR, and (3) CD11cCD123HLA-DR (Figure 8.2b). The HLA-DRCD11cCD123 population is also CD34 and probably represents circulating BM cells. They are allostimulatory and may represent a precursor population in the blood (Egner et al., 1993a, 1993b). More recently, Schmitz and colleagues have produced further mAbs that identify antigens on these blood DC subsets. BDCA-2 and BDCA-4 bind the CD11cCD123 population whilst BDCA-3 stains the CD11cCD123 myeloid DC subset (Dzionek et al., 2000). Previouslyacquireddatasuggestthatfreshlyisolated DCs express a variety of adhesion molecules, few co-stimulatory molecules (Freudenthal and Steinman 1990; Hart and Prickett 1993; McLellan et al., 1995, 1996) and some Fc receptors (Davis et al., 1988; Fanger et al., 1996). Freshly isolated DCs express the crossreacting DC-24 epitope defined by IgM CD24 mAb but not the CD24 protein (Williams et al., 1996). The CD13CD33CD14dim DC precursor population isolated directly from PBMCs differentiate, after culture, into cells with dendritic processes, a more mature phenotype and the ability to stimulate T lymphocytes in an MLR or to induce T-lymphocyte responses to soluble antigens such as tetanus toxoid (Thomas et al., 1993; Thomas and Lipsky, 1994). The CD2HLADRlin population of PBMCs is capable of processing and presenting nominal antigen to T lymphocytes (Takamizawa et al., 1997). A novel CD1 reagent does, as published, identify precursors of LCs within the CD11c subset of blood DCs (Ito et al., 1999). A re-analysis of the expression of these molecules on the three subpopulations described above is underway in our laboratory as part of our commitment to the 7th International Leukocyte Differentiation AntigenWorkshop. Fresh monocyte subsets Two recent publications have suggested that CD2 monocytes and CD34 monocytes acquire

DC-like characteristics after short culture (Ferrero et al., 1998; Crawford et al., 1999). The CD11c subset of Mo was described as a subset committed to Mφ differentiation. It is absent from patients with chronic myelomonocytic leukaemia, consistent with a Mo precursor (Vuckovic et al., 1999). Recent functional data suggest that these cells may develop immunostimulating capacity. Thus they may complicate the shoulder of lin cells described above, and decisions here relate fundamentally to the ability to sort CD14 populations with high purity. At present the identity of this subset must remain an open question. The nature of the population of CD2CD14 monocytes as a DC-like population also presents a conundrum (Crawford et al., 1999). Further analysis of this population and consideration of its relationship with the CD2CD14 DCs is again essential. Cultured blood DCs DCs used to be purified most readily by virtue of their low density compared with other PBMC populations after a brief period of tissue culture. This characteristic allows the use of a variety of gradient media to enrich DCs from other contaminating cells. BSA, metrizamide, Nycodenz and Percoll gradients have all been used successfully, although their osmotic effects may induce phenotypic or functional changes in the purified cells. Morphologically, these cultured cells have a prominent cytoplasm and some dendritic processes; phenotypically there is upregulation of surface antigen associated with DC activation. This allows purification by positive selection using the CMRF-44, CMRF-56 or CD83 reagents (Fearnley et al., 1997). There is also upregulation of a number of critical functional cell surface antigens. The CD33, CD13 antigens (Thomas et al., 1993; Fearnley et al., 1997) and the novel DC-24 carbohydrate epitope (Williams et al., 1996) are downregulated on this population. Recently, we have adopted an alternative approach to investigate antigen upregulation on cultured blood lin cells in short-term

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culture. The use of IL-3 and GM-CSF to culture the cells prior to phenotypic analysis enables three distinct populations to be detected in a majority of individuals (it should be stressed that there is significant individual variation in blood DC subpopulations in all individuals). HLA-DR and CD11c analysis defines the populations well, as does CMRF-58 and CD11c (Plate 8.3a, b). Thus, again three populations were noted with distinct morphological features: (1) HLA-DRCD11cCMRF58, (2) HLA-DRCD11cCMRF-58 and (3) HLA-DRCD11cCMRF-58 (Vuckovic, submitted) (Plate 8.3b,c). These data provide evidence to support the possibility that the subsets may reflect rigorously controlled stages of differentiation, perhaps even including recirculating (memory?) DCs.

Tissue (nonlymphoid) DCs – the interstitial DCs The blood delivers DCs to the tissues. The resident tissue-associated DCs can be described as surveillance cells. It is possible that this population is boosted during an inflammatory episode. These (nonlymphoid) tissue DCs can be separated into two phenotypically distinct populations, which may have specific properties relevant to their site. Superficial epithelial DC populations DC populations have been identified in the superficial epithelial tissues of the skin, the gut, the urogenital tract and the respiratory tract (Hart, 1997). The skin epidermal LCs are identified as the MHC class IICD1aCD14 cell in the epidermis and the presence of Birbeck granules within them can be demonstrated by electron microscopy (Romani and Schuler, 1992). In situ, some cells express very low levels of the CMRF-44, CD83, CD40 and CD86 antigens but not the CD80 antigen. Isolated LCs express these antigens in greater density (except CD80), which may reflect upregulation during isolation (McLellan et al. 1998). There is selective expression of certain β1-integrins (VLA subfamily α1-α9,

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αv). The expression of E-cadherin on LCs appears to play a key role in maintaining their contact with epithelial cells – downregulation is associated with mobilization (Borkowski et al., 1994; Blauvelt et al., 1995); LCs stain strongly for langerin (Valladeau et al., 1999, 2000). Rare CD1a DCs are identified in the epithelium of the gut but more extensive numbers are found in the lamina propria. CD1a cells have recently been identified in the urothelium of bladder, ureter and kidney (Troy et al., 1998b). The respiratory mucosa hosts an extensive population of DCs. Again, few are located within the epithelial surface but an extensive network of DCs is localized beneath the basement membrane (McWilliam et al., 1995, 1996). The phenotype of these cells reflects the influence of inflammatory and other mediators on DC activation. Nonepithelial DC populations This population of cells includes the dermal DCs, (Davis et al., 1988) and DCs in the heart, kidney, lung and liver (Hart and Fabre, 1981; Austyn et al., 1994) and perhaps deep epithelial glandular structures. Dermal DCs do not have prominent Birbeck granules and are MHC class IICD1aCD36Lag (Romani et al., 1989). Staining for factor XIIIa also identifies a dermal cell with dendritic morphology (Cerio et al., 1989). Further phenotypic analysis of these populations is required to distinguish these DCs from CD1a LCs, which migrate through the dermis. Langerin provides an important discriminating marker (Strunk et al., 1997; Valladeau et al., 1999). In situ, only a subpopulation of the dermal DCs express the CMRF-44 and CD83 antigens (McLellan et al., 1998) but these upregulate rapidly on isolated cells. Isolated mouse dermal DCs are reported to be N418 and NLDC-145 (Austyn et al., 1994). In situ phenotypic analysis of human liver DCs distinguished these from Mφ (Prickett et al., 1988). Interstitial DCs have been isolated from mouse heart and kidney, (Austyn et al., 1994) rat liver, (Hart and Fabre, 1981) and the rat iris (Steptoe et al., 1995). These cells express low-density MHC

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molecules (which upregulate in culture) and lack significant expression of the 33D1, N418 and DEC-205 antigens.

Lymphoid tissue-derived DCs – Interdigitating DCs DCs migrate from the tissues, either spontaneously or as a result of inflammatory ‘danger’ signals traffic, in the afferent lymph to the draining lymph nodes. The resulting populations appear to be phenotypically complex as judged by an analysis of isolated cells. This may reflect differing states of activation, different locations, B-lymphocyte (follicular) versus T-lymphocyte (interfollicular) areas. Again, species differences may be important. In humans, lin DCs associated with the T-lymphoid areas of tonsil are CD13, CD33, CMRF44 and CD83. A proportion of MHC class II DCs in these areas are activated, expressing CMRF-44, CD83, CD11b and co-stimulatory molecules (Summers et al., in press). These activated DCs are relB-positive and also express the cytoskeletal marker p55. A population of CMRF44 tonsil IDCs phenotypically resembles the CD4CD11c ‘plasmacytoid T lymphocytes’ described by Grouard et al. (1996). It is likely that these cells represent the same population as the HLA-DRlinCD123hi cells described by Olweus et al. (1997) and may represent a DC lineage distinct from the LC pathway. No major functional difference in the properties of the CMRF-44 and CMRF-44 IDC populations has been revealed as yet. A third population, found predominantly in the germinal centres, can be distinguished from the follicular DCs by the absence of CD21 but expression of CD4, CD11c, CD11b, CD13, CD33 and CD45 (Grouard et al., 1997). More recently, a more extensive phenotypic analysis of tonsil DC was undertaken by us on both isolated cells and tonsil sections. This allowed five phenotypic subpopulations to be defined: (1) HLA-DRmodCD11c CDw123, (2) HLA-DRhiCD11cCMRF-44hi, (3) HLA-DRmodCD11cCD13CMRF-44lo, (4) HLADRmodCD11cCDw123, (5) HLA-DRmodCD11c

germinal centre DCs (Summers et al., in press). Perhaps the most important functional conclusion from this phenotypic data is to emphasize that the majority of DCs trafficking to lymph nodes have a nonactivated phenotype, perhaps as a result of active downregulation (McLellan et al., 1999). Phenotypically distinct populations of IDCs are also found in the mouse spleen. The mouse DC marker NLDC-145 reacts with cells in the inner periarteriolar lymphocyte sheath around the central arteriole and not in the follicular or marginal zone (Kraal et al., 1986). A second population identified by the 33D1 mAb is found in the periphery of the T-lymphocyte area and marginal zone (Metlay et al., 1990). This may be an activated cell. MUC-1 staining has been found on activated mouse spleen DCs and this DC population is also located in the marginal zone (McGuckin et al., 2001). Double staining to clarify their identity is required. New isolation procedures have isolated three populations based on CD8α and CD11c staining. The analysis of Flt-3L-stimulated mouse DCs identified up to four populations from spleen based on staining by CD11c and NLDC-145 mAb (Maraskovsky et al., 1996). Thus again the complexity suggests that stages of activation contribute to the phenotypic diversity. Deletion of RelB appears to arrest DC differentiation (migration and activation) at the LC stage and depletes lymphoid tissue of the myeloid (? activated) DC population (Burkly et al., 1995). A lymphoid progenitor gives rise to the mouse thymic DC (Fairchild and Austyn, 1990; Ardavin and Shortman, 1993). The cells are characterized by the expression of CD8α, CD44, HSA, CD11a and MHC class II molecules in the absence of CD3 and rearranged TCR (Vremec et al., 1992; Wu et al., 1995). Recent data suggest the CD8α mouse DC has a myeloid origin and CD8α is induced on mouse lines. A similar cell identified in peripheral lymphoid tissues has been proposed to have a tolerogenic function rather than the allostimulatory function associated with the myeloid DC. Again, this may require reinterpretation. The human CD34CD38dim thymic precursor

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reportedly differentiates, in the presence of GM-CSF and TNFα, into T lymphocytes, NK cells and DCs (Res et al., 1996). The DCs resulting from this culture are CD4, CD40, MHC class II, CD33 with a subset expressing CD1a. They do not express other lineage markers, including CD8. A second report observed in vitro a CD10 human lymphoid progenitor-derived DC (Galy et al., 1995) that gives rise to T and B lymphocytes, NK cells and DCs. More recent transcriptional factor studies have failed to delineate human lymphoid and myeloid subsets clearly (Galy et al., 2000).

In vitro cultured DC-like cells DCs have been generated from a number of sources by culture of cells in the presence of a variety of cytokines and media supplements (Inaba et al., 1992, 1993a, 1993b; Scheicher et al., 1992; Mayordomo et al., 1995, 1996; Celluzi et al., 1996; Gabrilovich et al., 1996; Zitvogel et al., 1996; Dillon et al., 1997). Again, there appear to be clear species differences. Cultured mouse cells Mouse MHC class II bone marrow cells cultured in the presence of GM-CSF develop into a complex cellular population. This includes a subpopulation of DCs expressing high levels of MHC molecules, CD44 and CD11b, medium levels of CD24, CD45 and CD8 and low levels of NLDC-145, 33D1, CD11c as well as the Mφ marker F4/80 and Fcγ receptors (Inaba et al., 1992). Current culture conditions select for an activated ‘myeloid’ cell. Recently, mouse MoDCs were reported to be similar to the cells generated from bone marrow (Schreurs et al., 1999). Cultured human cells Various cultured human cell preparations have been exploited to provide large numbers of DC-like cells for basic studies and therapeutic administration. The starting populations of cells have included the CD34 BM, cord blood and GM-CSF mobilized peripheral blood cells.

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Although there are definite similarities with the ‘gold standard’ – directly purified blood DC populations – there are also notable differences. Generally, these culture systems result in heterogeneous populations of cells, some of which undoubtedly have the true DC phenotype (accounting for the functional properties) but many do not. The induction of CD1a on monocytic cells may confound the results obtained with this marker and the langerin molecule may be a better alternative marker for LC-like progeny. Cells cultured from CD34 BM in the presence of GM-CSF with a variety of cytokines, including IL-4, IL-13, TNFα and SCF, generate cells which include a CD14HLA-DR subpopulation that coexpress many of the accessory molecules and are able to stimulate allogeneic T lymphocytes (Szabolcs et al., 1995; Young et al., 1995). These cultures include 10–15% CD1a cells. A more substantial commitment to differentiating epithelial type LCs results from including TGFβ in the culture (Strobl et al., 1996). The culture of CD34 cord blood cells with GM-CSF and TNFα generates a subpopulation of cells with some characteristics of DCs and LCs including expression of CD1a, HLA-DR, costimulatory molecules and, in some cells, Birbeck granules (Caux et al., 1992). Yields of 5–15% CD1a cells are reported. Further characterization has suggested that DCs are derived from the CD13 cells (Rosenzwajg et al., 1996). Similarly, the generation of CD1a, CLA LC precursor from mobilized peripheral blood CD34 cells cultured with GM-CSF has been reported (Strunk et al., 1996). Apparently higher yields of CD1a cells (35-55%) are obtained from this source. Mo-DC can be grown from either PBMC or CD14 Mo in the presence of a combination of cytokines, generally including GM-CSF and IL-4 (Sallusto and Lanzavecchia, 1994) or Moconditioned medium. The presence of IL-4 may induce the downregulation of CD14 (Lauener et al., 1990) and suppress Mo development (Jansen et al., 1989). The resulting populations are heterogeneous with respect to many markers, including CD1a. The CMRF-44CD14 cells

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isolated from such cultures indicated that the allostimulatory activity was present only in a subpopulation of the CD1a cells (Vuckovic et al., 1998). The CD83 antigen was upregulated on the CD1a cell population of Mo-DCs (Zhou and Tedder, 1996a) as is the RelB transcription factor (Akagawa et al., 1996). The latter study also distinguished commitment to either Mφ or DC differentiation by expression of the M-CSF receptor. Other studies have performed detailed analyses of Mo-DCs derived from culture of CD14 Mo with GM-CSF and IL-4 or TNFα (Pickl et al., 1996; Zhou and Tedder, 1996a). The resulting cells expressed CD1a/b/c, CD80 and CD5, further upregulated CD40, MHC class II, CD4, CD11b/c, CD43, CD45, CD54, CD58 and CD59 whilst downregulating CD14, CD15s, CD64 and CDw65. The populations again were mixed and contained significant amounts of mRNA for myeloperoxidase and lysozyme, suggesting that the resulting cells are different from DCs purified without in vitro culture. Thus these cultures contain potent allostimulatory cells which are probably true DCs, whilst at the same time containing a number of contaminating cell populations. Finally, the plasticity of these populations has been emphasized by elegant studies delineating monocytic differentiation versus DC differentiation dependent on growth factor exposure (Akagawa et al., 1996). Additional TGFβ induces an LC phenotype (Birbeck granules, Langerin, CD1) (Strobl et al., 1996; Valladeau et al., 1999).

DCs derived from diseased tissue Lin DCs isolated from reactive sites of autoimmune diseases (e.g. chronic arthritic joints in rheumatoid arthritis) have a somewhat unexpected phenotype in that they lack highdensity expression of the CD80/CD86 costimulator molecules (Summers et al., 1995, 1996). Otherwise the isolated cells are morphologically similar to activated DCs with high-density expression of CD40 and adhesion molecules. A small number of the cells express the CMRF-44 activation marker but not the CD83 marker (Summers et al., 1995). However, these markers

and the co-stimulator molecules are induced after culture in the absence of synovial fluid. The phenotype of these cells is almost certainly modified by the presence of T lymphocytes. Analysis of the DCs found in tumour tissue has also revealed striking new information. First, relatively few DCs are present in renal cell carcinoma and the majority of these have an unactivated phenotype (Troy et al., 1998b). Only a small proportion (5–15%) of tumour-associated DCs are CMRF-44 and few have upregulated the CD86 molecule. Similar results pertain to prostatic carcinoma (Troy et al., 1998a). Likewise, phenotypically immature DCs predominate in breast and bowel carcinoma and only a proportion are CD1a (Coventry et al., submitted). The analysis of DC phenotype and function in malignant disease is now a major activity.

FUNCTIONAL PHENOTYPE OF DCs Many molecules useful for surface phenotyping and purification of DC populations have unknown functions. DC populations can also be phenotyped using molecules associated with specific functions.

Molecules involved in antigen uptake, processing and presentation Receptors involved in the uptake of antigen can be divided into a number of classes based on their mode of action. These include the patternrecognition receptors, the complement (C) receptors and the Fc receptors. Pattern-recognition receptors The pattern-recognition receptors bind a range of molecules with structural patterns common to the surface of many microorganisms but absent from the surface of mammalian cells (Pearson, 1996). Such antigens tend to be complex carbohydrates that show some specificity in the terminal sugar residues. Mo-DCs express surface molecules that are thought to bind these

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antigens including the macrophage mannose receptor, (Kraal et al., 1986; Sallusto et al., 1995; Engering et al., 1997; Tan et al., 1997) and other lectin-like molecules, i.e. the NKRP1A molecule (Poggi et al., 1997), DC-SIGN (Geijtenbeek et al., 2000a), DCIR (Bates et al., 1999) and perhaps MLD1. Expression of these molecules on other DC populations remains to be established. Functional studies on mouse DCs suggest that LCs (Reis e Sousa et al., 1993) express mannose receptors. The DEC-205 antigen is related to the macrophage mannose receptor and probably acts as an antigen receptor (Jiang et al., 1995). It is present on all mouse DC populations but in greater density on the so-called lymphoid (? less activated) DCs. A cDNA clone encoding the human DEC-205 homologue has been isolated (Kato et al., 1998) and the production of mAb now allows the expression of this molecule to be analysed. DEC-205 is present on both CD11c and CD11c blood DC populations with marginally higher levels on the former. Expression on tissue DCs is very limited and it is proposed that upregulation must occur before this receptor can contribute to intracellular trafficking of antigen. Another family of molecules important in the innate immune response and recognition of pathogens include the Toll-like receptors (TLR) (Muzio and Mantovani, 2000; Muzio et al., 2000b). TLR4 plays a crucial role in LPS signalling and as such has been suggested to play an important role in dendritic cell activation. A recent systematic analysis of the expression of six members of the TLR family identified to date shows that TLR1–5 mRNA are expressed in mature Mo-DCs, although only TLR3 is expressed exclusively in these cells (Muzio et al., 2000a).

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tors) (Morelli et al., 1996). Tonsillar DCs do not express the CD21 (CR2) and CD35 (CR1) receptors (Hart and McKenzie, 1988). However, immature Mo-DCs have been found to phagocytose apoptotic cells via CD36 (Albert et al., 1998). Protection of blood DCs against C-mediated lysis may be via the expression of CD55, CD59 and CD46 (Hart and Fearnley, 1997). CD46 antigen is also expressed by cytokine-derived DCs (Fugier-Vivier et al., 1997; Grosjean et al., 1997). Fc receptors Fc receptors allow the specific uptake of opsonized antigen by various cells. Both CD32 and CD64 but generally not CD16 are expressed by fresh blood DCs early in their differentiation; however, careful cell preparation was required for this analysis. These two Fcγ receptors enable DCs to phagocytose ox red blood cells, although at a reduced rate (Fanger et al., 1996, 1997). A recent report has used a novel mAb M-DC8 to identify a blood DC population that expresses HLA-DR, CD33 and CD16 (Schakel et al., 1998). Human LCs express CD32, CD64 and both the high and low receptors for IgE (Bieber et al., 1992; Rieger et al., 1992). However the FcεRI complex on DC lacks the β chain required for activity (Maurer et al., 1996).

Migration and adhesion of DC populations The phenotype of DCs migrating through the tissues prior to interaction with T lymphocytes relates to adhesion molecule and chemokine receptor expression. Adhesion molecules

Complement (C) receptors Some C receptors have been described on DCs. Low levels of CD11b (Weber-Matthiesen and Sterry, 1990; Egner et al., 1993a; Thomas and Lipsky, 1994) and CD11c (Shibaki et al., 1995) are found on blood DCs and LCs. Dermal DCs and a subpopulation of LCs express CD88 (C5a recep-

DCs express a wide variety of adhesion molecules. The ligands for CD11a (LFA-1); CD54 (ICAM-1), CD50 (ICAM-2) and CD102 (ICAM-3) are all expressed on DCs but show differential regulation. CD54 is expressed at low density on blood DCs and LCs but is quickly upregulated by activation (Starling et al., 1995), whereas CD50 is

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expressed at high density showing little change in expression level with activation (Hart and Prickett, 1993). CD102 is expressed in highest density on DCs and may be the most important CD11a ligand involved in early DC–T lymphocyte adhesion (Starling et al., 1995). LCs also express high levels of CD102 (Vedel et al., 1992; Lee et al., 1993; Zambruno et al., 1995). An interesting hypothesis detailing DC-SIGN as the initial DC ligand to stabilize DC interactions with T lymphocytes is based on Mo-DC data (Geijtenbeek et al., 2000b). Clarification of the tissue distribution of this molecule is awaited. Ecadherin has a role in migration and is expressed by mouse (Borkowski et al., 1994) and human LCs, (Blauvelt et al., 1995) and blood DCs but downregulates from LCs as the cells migrate. The presence of E-cadherin on LCs may reflect their unique adhesive interaction with the squamous epithelium. Although selectins have yet to be described on DCs, LCs do express the E-selectin ligand or cutaneous lymphocyte-associated antigen (CLA) (Koszik et al., 1994), which may be involved in cell interaction during trafficking. Interaction between DCs and the connective tissue may be stabilized by isoforms of the CD44 molecule that is expressed in high density by DCs (Prickett et al., 1992). Detailed analysis of the expression of the isoforms has only been performed on Mo-DCs which express the V3, V6 and V9 isoforms (Sallusto and Lanzavecchia, 1994), whilst LCs and blood DCs upregulated V4, V5, V6 and V9 following exposure to antigen (Weiss et al., 1997, 1998). Other molecules that may play adhesive roles reportedly expressed on DCs include syndecan (CD138), the endothelial molecule endoglin (CD105) and neurothelin (CD147) (Hart and Fearnley, 1997). In addition, subpopulations of DCs express CD106 (V-CAM) (Norton et al., 1992) and CD31 (PECAM-1) is expressed on blood and tonsil DCs (Hancock and Atkins, 1984). Receptors for chemoattractants As DCs respond to chemokines they must express receptors for chemoattractant molecules that presumably influence their migra-

tion and antigen-presenting roles. Mo-DCs differentiated in the presence of GM-CSF and IL-13 express the CCR1, CCR2, CCR5, CXCR1, CXCR2 and CXCR4 receptors but not the CCR3, CCR4 or CXCR3 receptors (Sozzani et al., 1997). Lung DCs express a receptor for the CC chemokine MIP-3α (Power et al., 1997). However, the expression of a particular receptor does not always indicate that DCs demonstrate chemotaxis in response to all chemokines that bind that receptor. Sozzani et al. have shown that although these DC populations should bind IL-8 and MCP-1, they do not migrate in response (Sozzani et al., 1997), whilst others showed that CD34derived DCs did migrate in response to MCP-1 (Xu et al., 1996). Migration of Mo-DCs is found in response to formylpeptides, C5a, SDF-1. MCP-3, MCP-4, MIP-1α, MIP-1β, MIP-5, RANTES and platelet-activating factor (Sozzani et al., 1995). Low numbers of LCs localized at the basement membrane of the epidermis express the C5aR (CD88), the expression of which is increased following culture of purified LCs with GM-CSF. These cells were also shown to migrate in response to C5a (Morelli et al., 1996). Mo-DCs and blood DCs differ in their response to chemokines (J. Roake, J. McKenzie and S. Vuckovic, unpublished). It is the expression of CCR4 and CCR7 that has attracted the most credence. CCR4 is proposed to allow recruitment of DCs to inflammatory sites. Critical differences in the DC populations might be expected. Its downregulation followed by upregulation of CCR7 is thought to encourage migration to T-cell areas of lymph node (Sallusto et al., 1999).

Adhesion and signalling Direct adhesion between T lymphocytes and DCs is probably mediated by CD2/LFA-1/ICAM-3 on the T lymphocyte and LFA-3/LFA-1/ICAM-3 (Prickett et al., 1992) or the recently described ICAM-3 ligand DC-SIGN (Geijtenbeek et al., 2000b) on the DCs, and results in phenotypic changes due to upregulation of co-stimulatory and other functional molecules. Molecules that may play a role in signalling between DC and T lymphocytes following adherence and antigen

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recognition are only now beginning to be elucidated. One causing interest at present in mouse models is the interaction between Serrate 1, expressed by CD11c splenic DCs and Notch (Hoyne et al., 2000).

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macrophages (Wong et al., 1997). Ligation of RANK on DCs enhances DC survival and this signal can also substitute for signals through the CD40L (Anderson et al., 1997). Inhibition/regulation of DC function

Co-stimulatory molecules Following migration of DCs to the T-lymphocyte areas of the lymphoid tissue DCs acquire a phenotype relating to their stimulatory properties. CD40, which is only present in low amounts in blood DCs (McLellan et al., 1996) and LCs (Romani et al., 1989), is upregulated following stimulation of the DCs. It is present on the more mature tonsillar DC population (Hart and McKenzie, 1988). Likewise, CD80 and CD86 are not expressed by resting blood DCs but are rapidly upregulated following activation (Hart et al., 1993; Symington et al., 1993; McLellan et al., 1995). Expression of other members of the B7 family have not been studied in detail (Swallow et al., 1999). Reciprocal T-lymphocyte CD40L feedback after antigen recognition induces major CD40-mediated increases in DC CD80/ CD86 expression (McLellan et al., 1996). The Ig superfamily member SLAM (CDw150) has been identified on blood DCs, upregulates following activation and has a putative co-stimulatory role (Hart and Fearnley, 1997). Cytokines may also modulate DC phenotype and function. Type I IL-1 receptors are expressed by LCs and IL-1α and IL-1β clearly induce CD40 expression on human blood DCs (McLellan et al., 1996). Both type I and type II TNF receptors are expressed on human blood DCs (McKenzie et al., 1995), GM-CSF receptors has been demonstrated on human blood DCs (Zhou and Tedder, 1996b) and tonsil DCs (Hart and Calder, 1994). Other members of the TNFR family of molecules are expressed by DCs and play signalling roles in the interaction between DCs and T lymphocytes. OX40L is expressed by mature human DCs (Ohshima et al., 1997) and contributes to co-stimulatory activity. Similarly, RANK (or TRANCE receptor) is expressed on mature DCs but not by B or T lymphocytes, or

Although it was widely anticipated that DCs underwent immune effector-mediated death or perhaps regenerated cell death (apoptosis), the expression on DCs of cell surface molecules with inhibitory functions has been documented only relatively recently. The mouse CD8α lymphoid DC population found in the thymus and spleen express high levels of FasL (Suss and Shortman, 1996). The possibility that myeloid-derived DCs were regulated by Fas-mediated apoptosis after performing APC function is an intriguing one. Recent data suggest they do express Fas on their surface. They also express receptors for IL-10 and TGFβ, both cytokines known to down regulate DC function. Rat spleen DCs express the NKR-P1 marker, a molecule associated with NK cell function (Josien et al., 1997). Members of the immunoregulatory signalling family of molecules expressed by DCs include the Ig family members CMRF-35A and CMRF-35H ( Jackson et al., 1992; Green et al., 1998), ILT1/ILT2,3 (Cella et al., 1999), SIRPa/SIRPb (Brooke et al., 1998; Liu et al., 1998), LAIR (Meyaard et al., 1997; St. Louis et al., 1999), DORA (Bates et al., 1998) and the C-type lectin molecule DCIR (Bates et al., 1999). Inhibitory members of this family are able to deliver inhibitory signals through association with cellular phosphatases, whereas activatory members associate with adapter molecules that contain immunotyrosine-based activatory motifs (ITAM) that are able to activate tyrosine kinase activity. The CMRF-35 antigen is a molecule that has considerable similarity with the Fc receptor for polymeric IgA and IgM, although ligands for CMRF-35 have not been identified (Jackson et al., 1992). This antigen is expressed in different forms on human blood DCs and some, but not all, LCs and tonsil DCs. Furthermore, this molecule is not expressed by Mo-DCs possibly allowing for another means of purifying Mo-DCs.

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Transcripts for a novel molecule (ILT3) were cloned from EBVB-cell lines using RT-PCR and primers designed to amplify Ig superfamily members (Samaridis and Colonna, 1997). An antibody to this molecule stained monocytes, CD83+ DCs and Mo-DCs (Cella et al., 1997). A putative immunotyrosinebased inhibitory motif is found in the cytoplasmic region of ILT3 and the ILT3-specific antibody prevented Ca2 mobilization, indicative of an inhibitory signal. These molecules have allowed the further discrimination of the myeloid blood DC population, which have the CD11c, ILT3, ILT1, CD13, CD33 phenotype, whereas the blood DC2 plasmacytoid precursor has the CD11c, ILT3, ILT1, CD123, CD36 phenotype (Cella et al., 1999). Both of these subsets express CD4, CLA, E-cadherin, CD40, CD80 and CD86, but the linILT3CD123 subset also expressed CD62L and CXCR3. At a functional level, the CD11cILT3ILT1CD123CD36 subset produced large amounts of type I IFN after being stimulated by viral infection. The DCIR molecule, which is structurally related to the C type lectins, also contains a cytoplasmic inhibitory motif and is expressed by lin DCs, Mo-DCs and CD34-derived DCs of the myeloid phenotype and not those with a LC phenotype (Bates et al., 1999). Activation of these cells with LPS, CD40L or TNFα down regulates its expression. Another molecule that is downregulated by CD40L activation of DCs is the Ig superfamily member DORA (Bates et al., 1998). This molecule contains three ITIMs and is expressed by immature Mo-DCs, CD34-derived DCs, CD11c blood DCs and LCs, as well as by monocytes and granulocytes. Indeed, we would argue that the central (i.e. negative) regulation of DC function may become the next major area for investigation. Functional data suggest that human DCs not actively involved in productive antigenspecific responses are downregulated (McLellan et al., 1999). The expression of several immunoregulatory receptor family molecules on DCs suggests that these may contribute to this process.

Other molecules The production of mAbs recognizing DCspecific surface antigens has been frustrating and researchers have looked at alternative ways of characterizing these cells. A number of mAbs binding nonsurface-expressed molecules have proven useful and a phenotypic profile of these secreted molecules may further characterize DC populations. Secreted molecules: cytokines and chemokines Interaction with T lymphocytes and signalling to other cells certainly involves cytokines and chemokines, and DCs may be further phenotyped by their limited production of either. Populations of DCs may express small amounts of IL-1 (Caux et al., 1994; Hart and Calder, 1994; Zhou and Tedder, 1996b), TNFα (McKenzie et al., 1995; Zhou and Tedder, 1996b), lymphotoxin and GM-CSF (Hart and Calder, 1994). Activated DCs have been reported to produce IL-7 (Sorg et al., 1998) and IL-12 (Yawalkar et al., 1996), while Mo-DCs produce IL-15 (Jonuleit et al., 1997). CD83 DCs purified from blood as metrizamide low-density mononuclear cells were found to express mRNA for a variety of chemokines; MIP-1β, IL-8, low levels for MCP-1 and RANTES and, on activation, for MIP-1α (Zhou and Tedder, 1996b). Mo-DCs, differentiated in the presence of GM-CSF and IL-13, produce MCP-1, RANTES and MIP-1α and the recently described chemokine, macrophagederived chemokine, (MDC). MDC acts as a chemoattractant for monocytes, Mo-DCs and NK cells (Godiska et al., 1997). Expression of MDC by DCs purified without exposure to cytokines has not been demonstrated. A second chemokine expressed by Mo-derived DCs, DCCK1 is a chemoattractant for CD45RA T lymphocytes and in situ hybridization identified its expression in germinal centres and the T-lymphocyte areas of tonsil (Adema et al., 1997). DCCK1 expression appears to be induced by IL-4 but inhibited by GM-CSF, indicating that its expression will be markedly influenced by the cytokine combination used to differentiate the DC population.

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Enzyme reactivity is a useful phenotypic marker for distinguishing DCs from other cells, particularly from classic Mo/Mφ. The latter cell populations show myeloperoxidase activity (Steinman and Cohn, 1973; Witmer and Steinman, 1984; Buckley et al., 1987; Egner et al., 1993a), and low levels of 5-nucleotidase, dipeptidyl peptidase and cathepsin B activity (Knight et al., 1986; Romani and Schuler, 1992; Thomas et al., 1993), whereas DCs appear to lack these. DCs also express low levels or show a different intracellular distribution of other enzymes such as nonspecific esterase (Arkema et al., 1991; Liu et al., 1998), acid phosphatase or markers such as CD68, a lysosomal marker (Hart and McKenzie, 1988). The S100 mAb detects an intracellular isomer useful for histological studies on formalin-fixed sections (Takahashi et al., 1982), but because of its solubility, not in frozen section. Recent data suggest it is associated with activated DCs (Troy and Hart, 1997). DC-LAMP, a novel DC-associated lysosomal protein, provides an excellent marker of DC maturation (de Saint-Vis et al., 1998). Enzyme activity is also related to the antigen-processing and presentation function of DCs. A number of proteases, including cathepsin S and D (Sallusto et al., 1995) and protease inhibitors such as cystatin C (Pierre and Mellman, 1998a, 1998b), have been identified in Mo-DC populations.

regulated differently in DCs compared with other cells (e.g. B lymphocytes) in terms of basal activity, levels and kinetics. Transcription factors regulating the DC phenotype may provide a further level of phenotyping. However, it may be the pattern of expression of a variety of transcription factors and not the absolute presence or absence of any one factor that will provide the best distinction of DC from other cells. The NFκB transcription family member RelB shows some selective expression in mouse thymus, spleen and lymph node DCs (Carrasco et al., 1993) and mice with a deleted RelB gene show, amongst other phenotypes, a lack of mature functional DCs (Burkly et al., 1995). Human DCs grown from peripheral blood with GM-CSF express c-rel, relB, NFκBp65 and NFκBp50 (Granelli-Piperno et al., 1995) but lack the widely expressed transcription factor SP-1. RelB has been identified in Mo-DCs (Akagawa et al., 1996) and human tonsil DCs (Feuillard et al., 1996). Peripheral blood DCs purified following culture expresses RelB, which is absent in freshly purified blood DCs (Clark et al., 1999). In another study, the Oct-2 transcription factor was shown to be downregulated during differentiation of DCs but not during macrophage development (Neumann et al., 2000). The Ikaros gene product has been involved in early DC differentiation events. Mice with targeted Ikaros mutations lack DCs (Wu et al., 1997; Nichogiannopoulou et al., 1999). CD34 progenitor cells expressing a retrovirally transduced dominant negative isoform of Ikaros, Ik7, fail to generate DCs when cultured in conditions that would normally allow DC production (Galy et al., 1998, 2000). Curiously, Pax5 deletion in mice allows committed B lymphoid progenitors to dedifferentiate into DCs (Nutt et al., 1999). The isolation and study of promoters for DC genes is likely to identify interesting new transcription factors.

Transcription factors

Other cytoplasmic markers

The expression of many surface molecules, including CD80/CD86, MHC class II, is clearly

The specialized function of DCs in processing and presentation of antigens suggests that other

Cytoskeletal markers A 55-kDa actin bundling protein (p55, fascin) is expressed by the majority of human blood DCs, IDCs, cytokine-derived DCs but not other leukocyte lineages (Mosialos et al., 1996). Messenger RNA for a protein called restin associated with the intermediate filament cytoskeletal network of Reed–Sternberg cells (RSCs) in Hodgkin’s disease has been detected in human DC populations (Bilbe et al., 1992). Cytoplasmic markers

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intracellular enzymes or regulatory proteins may be useful in identifying this cell lineage. The techniques of differential display are fast describing new DC molecules and a number of groups have identified many transcripts expressed by DC populations. In general, these reports have studied the expression of molecules from in vitro-derived DCs (Marland et al., 1997; Hashimoto et al., 1999), however most reports are as abstracts at specialist meetings rather than published papers. Molecules identified in these experiments to date include transcripts for proteases and protease inhibitors including decysin (Mueller et al., 1997b) and serpin (Mueller et al., 1997a). However, the value of these studies in identifying phenotypic markers requires corresponding data showing expression of translated products.

APPLICATION OF PHENOTYPE Detailed knowledge of the phenotype of DCs is crucial to understanding the biology of the cells being studied. In addition, as many applications of DCs for clinical treatments are being investigated, it is crucial that the most appropriate cell population to be used for each is identified. Purification of homogeneous DC populations for particular uses is presently difficult for most populations except the activated CD83, CMRF44, HLA class II, CD14 human blood DC or N418bright, DEC-205bright mouse spleen DCs. The ability to positively or negatively select a population for a particular purpose requires the availability of the necessary reagents. Drug targeting or antigen loading of cells for therapeutic purposes may be more readily achieved using immature DCs that express antigen uptake receptors, rather than cells with a more mature phenotype. The homogeneity of a population may influence the outcome, particularly if DC populations that are able to tolerize (Matzinger and Guerder, 1989) are included in a tumour therapy protocol. The availability of the CMRF-44 mAb that reacts with blood DCs after a brief period of in

vitro culture led us to exploit the DC phenotype to develop the first routine blood test for human DCs. Using a method suitable for most diagnostic flow cytometers, reliable counts can be obtained from 5 mL EDTA blood samples. This has allowed DC counts to be monitored in several clinical situations and this has given basic information, stimulating more research. More importantly, this has allowed DC numbers to be tracked in patients to facilitate DC collection for immunotherapy programmes. Phenotypic characterization of DCs can also be applied directly to certain potential DC malignancies. Class I histiocytosis is a rare childhood condition that is defined by CD1 staining, suggesting an LC origin of this malignancy (Egeler et al., 1993). This is further supported in one case by positive staining with the CMRF-44 and CD83 mAb (D.N.J. Hart, unpublished). A DC origin for the malignant cells in Hodgkin’s disease has been suggested for a proportion of cases in which the malignant cell expresses a number of DC markers, including CD83 (Sorg et al., 1997) and p55 (Mosialos et al., 1996). At a clinical level mAb to various DC molecules might be exploited for DC purifications. The functions of molecules expressed specifically in DC populations is relevant to the ability to use DC in clinical strategies. Thus DEC205 or other receptors may be targets for antigen loading for tumour immunotherapy. CTLA-4 may target antigen effectively to DCs. The ability to specifically inhibit a particular enzyme or DNA-binding protein that is important to DC function may allow new immunosuppressive treatments that may lead to applications in clinical transplantation and autoimmune disease. An understanding of the function of the CD83, CMRF-44 and CMRF-56 antigens may likewise contribute to therapeutic interventions, particularly in transplantation.

THE FUTURE A DC lineage-specific marker has yet to be identified. The human genomic sequence and DC gene discovery programmes will soon reveal

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REFERENCES

they exist. For the moment, DCs are still characterized by combinations of phenotypic markers, morphology and functional criteria, but the first generation of mAbs are already helping. A greater understanding of these cells requires delineation of the fundamental question – is there a single DC lineage with many DC subpopulations determined by microenvironment and other stimuli or are there distinct lineages? The molecular dissection of the unique properties of DCs promises to be very informative and therapeutic!

ACKNOWLEDGEMENTS We would like to acknowledge the support of the NH&MRC of Australia, the Queensland Cancer Foundation and other team leaders in the Dendritic Cell Laboratory of the Mater Medical Research Institute – M. Kato, M. Wykes and J.A. Lopez.

Note Further phenotypic data on important new molecules expressed on various DC populations was collated at the 7th Leukocyte Differentiation Antigen Workshop (7th Leakocyte Differentiation Antigen Workshop; DC section summary, Hart, D.N.J., Clark, G.J., MacDonald, K., Kato, M., Vuckovic, S., Lopez, J.A., Wykes, M. and Munster, D. in press). The new CD classification for DC related molecules includes; CD85 - ILT/LIR family, CD205 - DEC205, CD206- Macrophage mannose R, CD207 - Langerin, CD208 - DC-LAMP, CD209 - DC-SIGN. Further phenotypic data clarifying the human blood DC populations is being submitted for publication; MacDonald, K.P.A. et al., further characterization of human blood dendritic cell subsets.

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PLATE 8.3 Phenotype and morphology of cultured blood lin DCs. Lin blood DCs were prepared as outlined in Figure 8.2, cultured for 12 hours in RPMI/10% FCS supplemented with GM-CSF (200 U/mL) and IL-3 (10 ng/mL) and stained with HLA-DR, CD11c and CMRF-58. (a) Linblood DCs included HLADRCD11c and HLA-DRCD11c- DCs. (b) CMRF-58 mAb defined three populations among linHLA-DR DCs with (c) distinct morphological features. PLATE 8.1 Leucocyte Differentiation Antigen Workshop diagram presented staining with CMRF-44 monoclonal antibody. The diagram represents the differentiation of DC precursors from the bone marrow into mature DCs. It attempts to emphasize the complexity of the subpopulations of DCs that are studied. The CMRF-44 mAb-stained B lymphocytes and monocytes in blood, blood CD11c DCs, in vitro maturated CD11c and CD11c blood DCs, immature and mature Mo-DCs, Mo-LCs, CD34-DCs, epidermal and dermal DCs, subsets of tonsil DCs and scattered cells in spleen.

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C

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9 Targeted knockouts affecting DCs David Lo1, Lian Fan2, Will Redmond2 and Christina R. Reilly2 1

Digital Gene Technologies Inc., La Jolla, CA, USA

2

The Scripps Research Institute, La Jolla, CA, USA

What we label scientific knowledge is ‘lots of fun. You get lots of correlations, but you don’t get the truth. Things are much more marvelous than the scientific method allows us to conceive.’ Barbara McClintock, in A Feeling for the Organism, by Evelyn Fox Keller Researchers have already cast much darkness on the subject, and if they continue their investigations we shall soon know nothing at all about it. Attributed to Mark Twain

INTRODUCTION

targeted disruptions of genes thought to be relevant to DC function. In this chapter we will briefly discuss some of the applications of this approach, and how our own studies have used combinations of transgenic and knockout models of DC development. While these approaches may bring new complications and considerations, they should provide some interesting new insights into the function of DCs and their role in the development of secondary lymphoid tissues.

Studying the biology of dendritic cells (DCs) can be complicated by their relative rarity within tissues, so a variety of approaches have been used to circumvent this limitation. For example, cultures of bone marrow in the presence of GMCSF and IL-4 can give rise to cells that look very much like bona fide DCs, both in terms of their cellular morphology and their ability to stimulate T-cell proliferation in vitro. However, these cultures do not reproduce all of the phenotypic features of DCs freshly isolated from tissue, and so it is reasonable to expect that some of the in vivo behavior of DCs might also be lacking in these cells. A more physiological approach to studying DCs might be to examine the phenotype and behavior of DCs in vivo in mice with Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

PARADOXES IN EXISTING MODELS OF DC SUBSETS AND BIOLOGY As the studies on DC functions have grown in intensity, a variety of schemas have developed to

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explain the diversity in DC phenotypes and functions. For example, Shortman and colleagues have identified two strikingly different DC phenotypes, and their studies suggest that these subsets have distinct functions in vivo. The ‘lymphoid’ DC was named for its apparent origin from stem cells that also give rise to thymocytes and its expression of the lymphoid markers CD8α and DEC-205 (Ardavin et al., 1993; Wu et al., 1996; Shortman et al., 1998). It has been suggested that these cells mediate thymic negative selection. In the periphery, they are located within the T-cell dependent regions of the spleen white pulp, comprising 15–20% of splenic CD11c DCs, and they may contribute to tolerance induction through the expression of FasL (Suss and Shortman, 1996). In contrast, the ‘myeloid’ DCs are negative for CD8α and DEC-205, and instead express the marker 33D1 (Crowley and Lo, 1999). These are thought to be the most potent stimulators of T cells, comprising about 80% of splenic CD11c DCs, but their anatomic location is in the marginal zone of the spleen white pulp, so their interactions with naïve T cells have not been as easy to describe. Interestingly, Moser has ascribed rather different functions to these two DC phenotypes (Maldonado-Lopez et al., 1999). Thus, the CD8α DEC-205 subset appears to be primarily responsible for the generation of TH1 effector Tcell responses. In contrast, the other major subset (the CD8α subset), appears to drive TH2 effector T-cell development. In studies on human DCs by Liu and colleagues, further variation has been described. A DC precursor expressing the lymphoid marker CD4 has been identified as the major source of type 1 interferons (Siegal et al., 1999). This cell gives rise to DCs that drive TH2 development, and so they have been designated DC2 (Rissoan et al., 1999). The DC2 appears to be derived from a plasmacytoid cell, and is distinguished from monocyte-derived DCs capable of driving TH1 responses (DC1). Curiously, the type 1 interferons produced by the DC2 precursors have been shown to be potent stimulators of TH1 development even in the absence of IFNγ (Rogge et al., 1997, 1998), so it would

appear that the same cell may be driving both TH1 and TH2 generation. By now it may be evident that these various schemas, while being relatively simple when considered individually, actually raise a number of complicating conflicts. First, the Shortman lymphoid DC, which is toleragenic, is phenotypically equivalent to the Moser CD8α subset driving TH1 responses instead. In both models, the localization of the myeloid CD8α subset to the marginal zone also raises questions about how they are to engage with T cells in the development of immune responses, since T cells are not commonly found in the marginal zone. The conflict with the studies on human DCs is even more striking, as human monocytederived DCs, presumably myeloid in origin, drive TH1 responses. This, of course, is the opposite from the situation in mouse where myeloid DCs (CD8α subset) drive TH2 responses.

THE CONTRIBUTIONS OF KNOCKOUTS TO DC STUDY: RelB AND FRIENDS One approach to the study of DC biology is to examine the in vivo functions of DCs in mice with defects in relevant genes. Previously, we had focused on DC defects in mice deficient in the NFκB transcription factor RelB (Crowley and Lo, 1999). The findings in RelB knockout mice illustrate the complexities in dealing with knockout phenotypes, as the plasticity in the immune system can drive partial compensation for targeted gene defects. In normal mice, RelB is normally constitutively expressed in only a few cells: DCs of lymphoid tissues (spleen, thymus, lymph nodes and Peyer’s patches) and a subset of thymic medullary epithelial cells (Carrasco et al., 1993; Burkly et al., 1995). In RelB knockout mice the thymic medullary epithelial cells are absent (Burkly et al., 1995; DeKoning et al., 1997), so in this case it may be inferred that the development of this cell type is absolutely dependent on RelB for its development and function. As for DCs, development is not so easily

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attributed to RelB expression. This is in part due to the fact that the working definition of a ‘dendritic cell’ can be ambiguous, depending on the investigators and the situation. Thus, cells presenting many phenotypic features of DCs are present throughout the body, but they are not quite equivalent to functional antigen-presenting DCs. One example is the skin Langerhans cell, which shows a dendritic morphology, expresses high levels of MHC class II antigens, and also expresses the co-stimulatory molecule B7-2 (Inaba et al., 1994). It might be most appropriate to consider this to be a DC precursor, as they are not functionally engaged with T lymphocytes while they reside in the skin, and only do so on activation and migration to the draining lymph node. Since the expression of RelB is only associated with DCs that have arrived in the draining lymph nodes, a deficiency in RelB would not be expected to alter the development of DC precursors such as Langerhans cells. Indeed, RelB knockout mice appear to have normal numbers of Langerhans cells in the skin, and DC precursors appear to be present in other tissues as well (Burkly et al., 1995). Since these precursors can develop to a fairly differentiated stage (e.g. resident tissue macrophage, airway epithelium DC, etc.) in the absence of RelB, deficiencies in RelB can have rather subtle and complex effects on the final stages in DC differentiation. Thus, as individual DC functions are assessed, we find that a number of defects can be identified that do not fit in neatly with defined characteristics of DC subsets. The first, most direct defect noted in RelB-deficient mice is the alteration in the phenotype of DCs isolated from mutant mice. While normal splenic DC populations show an approximate 80:20 distribution of CD8α to CD8α DCs (Crowley et al., 1999), RelB knockout mice only contained CD8α DCs (Wu et al., 1998; Crowley and Lo, 1999). Since RelB deficiency also causes a defect in regulation of inflammatory cytokines/chemokines in nonhematopoietic cells, causing an inflammatory syndrome (Xia et al., 1997; Lo et al., 1999), we also studied bone marrow chimeric mice using RelB knockout stem cells to reconstitute

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irradiated normal mice. In these chimeras, the inflammatory syndrome was absent, but the DC population was still limited to CD8α cells (Wu et al., 1998). While phenotypic analysis suggests that RelB knockout mice are profoundly deficient in ‘CD8α/myeloid/DC2’ DCs, retaining the ‘CD8α/lymphoid/DC1’ DCs, the functional defects in DC function are quite another matter. Our studies so far have examined two separate predictions from these phenotypic effects, and found some rather puzzling results. The CD8α subset, which is the dominant subset found in the thymus, is associated with negative selection in the thymus. Since the RelB knockouts appear to retain this subset, it might be expected that thymic negative selection in the T-cell repertoire would be intact. Surprisingly, negative selection is significantly impaired in mutant mice; the thymus fails to delete T cells reactive to endogenous retroviral superantigen/I-E complexes (Laufer et al., 1996), and peripheral T cells show strong autoreactive proliferative responses to normal syngeneic antigen-presenting cells (DeKoning et al., 1997). The export of T cells with significant autoreactivity may also be an important factor in the multiorgan inflammatory syndrome of RelB-mutant mice (Weih et al., 1995, 1996). Studies on peripheral T cells in mutant mice show that an unusually high proportion of T cells express inflammatory cytokines, especially TNF (DeKoning et al., 1997). Thus, it appears that RelB is important in providing thymic DCs with the ability to engage developing T cells during their selection, and delete the autoreactive population. Despite this important defect, the maturation of T cells is unimpaired, illustrating the principle that positive selection and maturation of T cells are processes carried out independently of negative selection. It has also been suggested that the CD8α subset drives TH1 responses, so the RelB knockout mice would be predicted to produce mainly TH1 responses. Unfortunately, a variety of studies on immune responses in RelB knockout mice fail to support that prediction. For example, delayed-type hypersensitivity (DTH) responses, which are mainly driven by TH1 cells,

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cannot be generated in RelB knockout mice (Weih et al., 1995). Effective immune responses to the parasite Toxoplasma gondii (Caamano et al., 1999) also require TH1 effectors to clear the parasite, and here too the RelB mice fail to generate the necessary IFNγ response. Cytokine production by T cells of RelB mice has been studied by intracytoplasmic staining of activated T cells. There was no absolute skewing in the profile of cytokines produced, since both IFNγ and IL-4 could be detected, but there appeared to be more cells producing IL-4 than IFNγ (DeKoning et al., 1997). Interestingly, there was an even higher proportion of cells expressing TNF, although the significance of this finding is unclear. Skewing of immune responses in vivo can also be assessed by the relative amounts of antibody isotypes; TH1 responses tend to drive the switch toward IgG2a, while TH2 responses drive switching toward IgG1 and IgE. In this case, we found that control mice immunized with influenza virus generated strong IgG2a antibody responses to the influenza hemagglutinin. By contrast, RelB knockout bone marrow chimeras generated lower antibody titers, consisting mainly of IgG1 (Burkly et al., 1995). Thus, by several criteria, RelB knockout mice actually may be slightly predisposed toward TH2 responses. The results with the RelB knockouts suggest an alternative model for the generation of DC subsets. The deficiency in RelB results in the specific loss of the CD8α subset, while the CD8α/lymphoid subset is retained. Yet the functional defects seen in RelB knockouts (defects in negative selection, defects in T cell priming toward TH1 effectors) had previously been associated with CD8α/lymphoid subset functions. It would seem that in an attempt to compensate for CD8α/lymphoid functional deficits, DC precursors that might have otherwise produced CD8α/myeloid cells are instead driven toward the CD8α/lymphoid phenotype. Despite this phenotypic shift, the DCs still fail to function in the face of RelB deficiency. This suggests that RelB regulates genes important to DC function, while other RelB-independent mechanisms may regulate expression of

markers such as CD8α or DEC-205. Indeed, expression of CD8α by T cells and expression of DEC-205 by thymic epithelium remains normal in RelB knockout mice. Perhaps even more importantly, this shift in DC phenotypes may also indicate a single lineage for both DC phenotypic subsets, since the functional defect in one subset clearly draws precursors away from the other subset. Viewed in this context, a variety of other targeted gene knockouts may be evaluated according to whether they represent partial defects in function or absolute defects in development. For example, Ikaros, a transcription factor associated with lymphoid cell development (Georgopoulos et al., 1997), can in some cases have overlapping functions with the related gene Aiolos (Morgan et al., 1997). Thus, in Ikaros knockout mice, significant defects in DC development have been described, and interestingly, the retained DC phenotype matches the CD8α subset (Wang et al., 1996; Wu et al., 1997). As with the RelB knockouts, we would interpret this surviving phenotype as a cellular attempt to compensate for the loss of Ikaros-dependent gene expression through partial compensation by Aiolos. In the case of an Ikaros dominant negative mutant, DC development is entirely lost, as the partial compensation by genes such as Aiolos is blocked by the dominant negative effect. A similar analysis can be applied to the role of PU.1 in DC development (Anderson et al., 2000). PU.1 is an ets family transcription factor expressed in multiple hematopoietic lineages, and it appears to regulate expression of many myeloid and lymphoid genes. Mice deficient in PU.1 show profound defects in myeloid lineage cells such as macrophages and neutrophils, but T-cell development in the thymus appears to be normal. As might be expected, studies on mutant mice in vivo and in vitro found that myeloid phenotype DCs are absent. Interestingly, lymphoid phenotype DCs were also absent from the thymus and spleen, even though development of the thymic medullary compartment appeared to be normal. In this case, defects in DC precursors (myeloid

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macrophages?) appeared to have dramatic effects on DC development, since sources of compensatory mechanisms are apparently not available. Indeed, the defects observed in this case add further support to the notion that both DC phenotypic subsets have a common progenitor stem cell, since lymphoid cells (thymic T cells) can develop in these knockouts, yet fail to provide progenitor cells for ‘lymphoid’ DCs.

LYMPHOTOXIN LIGANDS AND DC DEVELOPMENT Up to this point, we have been considering DC development as independent cells without regard to how other cells or tissues influence DC development. Yet DC clearly do not simply develop in place on their own, and in vitro studies have sometimes provided misleading indications as to the events driving DC development. For example, a common protocol for generating large numbers of DCs involves the addition of GM-CSF and IL-4 to bone marrow cultures (Inaba et al., 1992); as reliable as this technique may be, it is quite clear that mice deficient in these cytokines or their receptors do not have DC defects in vivo. Recent studies have begun to provide some interesting information on the importance of TNF/lymphotoxin ligand signals provided to DC precursors in vivo. Interestingly, these ligands turn out to be important in the development of lymphoid tissues as well (DeTogni et al., 1994; Ettinger et al., 1996; Koni et al., 1997; Alimzhanov et al., 1997; Futterer et al., 1998; Rennert et al., 1998; Wu et al., 1999). There are at least three TNF/LT ligands consisting of various trimeric combinations: TNFα3, LTα3 and LTα1β2, with a possible LTα2β1. Both the TNFα3 and LTα3 forms are bound by the receptors TNFRI and TNFRII, and the LTα1β2 form is bound by the LTβR. While knockouts in TNF and TNFR can affect germinal center development, LTα, LTβ and LTβR knockouts have much more dramatic effects on secondary lymphoid tissue development. Initial studies on various LT knockout mice found major effects on secondary lymphoid

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tissue development. In the case of LTα and LTβR knockouts (DeTogni et al., 1994; Koni et al., 1997; Futterer et al., 1998), mice entirely lack any peripheral lymph nodes or Peyer’s patches, and the spleen shows poor organization in the white pulp. Thus, the defect in lymphoid tissue development appears to be complete enough that we conclude that these signals are absolutely required for development and function. In the case of LTβ knockouts (Alimzhanov et al., 1997; Koni et al., 1997), there is a partial phenotype; occasional mice can be found with cervical or mesenteric lymph nodes. This may be due to partial compensation by related ligands. Are DCs similarly affected? Previously, TNF signaling had been associated with activation of DC precursors; in the case of LT ligands, studies are beginning to identify more basic effects on DC development. In preliminary studies, we have examined the DC subsets present in the spleen of LTβ knockout mice (Alimzhanov et al., 1997), and found that among the CD11c cells, both CD8α/DEC205 and CD8α subsets are present. However, in contrast to normal mice, the proportions of these subsets are slightly skewed toward the CD8α cells (unpublished results). Curiously, this is very different from the published observations on the LTα knockout mice, in which the overall numbers of DCs are drastically reduced (Wu et al., 1999). Reconstitution of LTα knockout mice with normal spleen cells was found to be sufficient to restore DC development (although DC subsets were not characterized). Are these strikingly different results compatible? As we have already discussed, the plasticity in the immune system can have a significant impact on the observed phenotype. In the case of LTα and LTβR knockouts, the defects appear to be so profound that there are no avenues to compensate, and the mice are always devoid of any secondary lymphoid tissue. By extension, if LTα and LTβR-mediated signals are absolutely required for DC development, no compensatory mechanisms may be available. As a result, the observed loss of lymph nodes and DCs are direct consequences of the knockouts. In contrast, since LTβ knockouts can show

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occasional development of cervical or mesenteric lymph nodes, it suggests that there are some compensatory mechanisms available. For example, the ligand LIGHT expressed on activated T cells may have some weak LTβ-like activity (Mauri et al., 1998). As a result, occasional T-cell activation in the head or intestine may be able to drive development of lymphoid tissue in sites draining those regions. Similarly, DCs may be induced to develop in the spleen of LTβ knockout mice through interactions with occasional LIGHT-expressing T cells. The increased proportion of CD8α DCs in the spleen seems to parallel the situation in RelB and Ikaros knockouts, where these compensatory pathways appear to be most efficient in driving development of the CD8α phenotype.

DC DEVELOPMENT AND SECONDARY LYMPHOID TISSUE DCs do not spontaneously arise in any tissue; the phenotypically mature DCs are found primarily in the spleen, thymus and secondary lymphoid tissue. It is not yet clear whether these cells appear in the lymphoid tissues as fully mature DCs that are drawn in, or whether they are recruited to the lymphoid tissue as precursor forms, then induced to differentiate in situ through interactions with T lymphocytes. Published reports appear to support both possibilities. In one case, spleen DC subsets appeared to be normal in mice lacking various lymphocyte subsets (e.g. RAG, TCRα, BCR knockouts), and the myeloid and lymphoid subsets were even found to home in their proper orientation around central arterioles in the presumptive white pulp even in the absence of lymphocytes (Crowley et al., 1999). By contrast, in a separate study, DC development appeared to be deficient in the absence of T cells (Shreedhar et al., 1999). Thus, in order to study the relationship between DC development and their homes in lymphoid tissue, we have begun an analysis of DC recruitment into newly developing secondary lymphoid tissue.

The study of secondary lymphoid tissue development is made difficult by the fact that the development of lymph nodes is not easily manipulated during fetal life, although some studies have successfully used LT-blocking reagents to interfere with some fetal lymph node development (Ettinger et al., 1996). Interestingly, lymphoid tissue development can also be triggered in mice postnatally by the expression of specific transgenes. Thus, expression of LTα by islet beta cells in the pancreas (RIP-LT) can give rise to lymphocytic infiltrates that can organize in a manner resembling secondary lymphoid tissue (Kratz et al., 1996; Sacca et al., 1998). These infiltrates contain a high frequency of activated CD44hi T lymphocytes, and in that respect they may parallel the situation seen in autoimmune diabetes, where islet reactive T-cell infiltrates also appear to organize spontaneously into lymphoid tissue with induction of HEV (Hanninen et al., 1993; Yang et al., 1997) and segregation of distinct T cell/DC and B compartments (Lo et al., 1993; Scott et al., 1994; Degermann et al., 1994). The activated phenotype of T lymphocytes in RIP-LT mice and in spontaneous autoimmune diabetes may drive mechanisms that are distinct from those driving lymphoid tissue development in the absence of T-cell activation. To examine this in detail, we developed a new model in which secondary lymphoid tissue develops postnatally under conditions where development can be followed over time, and experimental manipulations can be imposed. The model was generated by driving expression of the chemokine TCA4/SLC (Tanabe et al., 1997; Gunn et al., 1998a) using the rat insulin promoter (Ins-TCA4 transgenic mice). Our interest in this chemokine began with the observation that this chemokine was expressed strongly in lymph node HEV and the T-cell compartment of lymph nodes, and it was also expressed very strongly in the medullary compartment of the thymus, including in a subset of medullary epithelium expressing a fucose bound by the lectin UEA-1 (Tanabe et al., 1997). The relationship of this chemokine to lymphoid tissue development was further strengthened by

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the observation that RelB knockout mice, which lack the thymus medullary compartment and all secondary lymphoid tissue, also lack expression of TCA4/SLC (Tanabe et al., 1997). Studies on the function of TCA4/SLC have found that it binds to the chemokine receptor CCR7, which is expressed by naïve T cells and mature DCs, with lower expression on B cells (Sallusto et al., 1998). Thus, the accumulation of mature thymocytes to the thymic medulla may in part be driven by chemotactic recruitment of maturing cells from the cortex. Indeed, as thymocytes mature, they acquire expression of the CCR7 receptor (Suzuki et al., 1999). In Ins-TCA4 transgenic mice (Fan et al., 2000), the expression of the chemokine is initially limited to islet beta cells, and so pancreatic islet tissue begins to accumulate lymphocytes and DCs. The early infiltrates show an interesting pattern, as we first find CD4 and CD8 T cells in compact clusters within the center of the islets. These clusters are associated with mature phenotype CD11c DEC-205 DCs, as might be expected on the basis of CCR7 expression in mature DCs. Remarkably, while B cells are thought to express CCR7, they are not associated with these T cell/DC clusters, and instead are found initially as scattered cells at the perimeter of the islet tissue. Thus, even at this early stage, T-cell and B-cell compartment segregation appears to be mediated through differential behavior in response to the chemokine TCA4/SLC. As the transgenic mice get older, the islet infiltrates expand, and we now find the induction of high endothelial venule structures, expressing both MAdCAM-1 and PNAd. In addition, a network of stromal reticular cells (expressing the marker ER-TR7) develops to provide a scaffold for the islet infiltrate. B-cell follicles can now be distinguished as a distinct compartment at the perimeter of the infiltrates, with detectable expression of the follicular DC marker FDC-M1. In essence, the recruitment of lymphocytes and DCs by the chemokine TCA4/SLC was sufficient to drive the development of secondary lymphoid tissue. The DCs did not accumulate at random within the islet infiltrates, and instead

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began from the start as part of a well-organized T-cell dependent compartment. As an indicator of the complete development of the secondary lymphoid tissue, macrophages expressing F4/80 were excluded from these lymphoid collections, and were only found at the perimeter of the infiltrates.

IS RECRUITMENT OF DC INTO SECONDARY LYMPHOID TISSUE DEPENDENT ON T LYMPHOCYTES? In our studies on DC subsets in normal and knockout mice (Crowley et al., 1999), we found that the CD8α and CD8α subsets were present in relatively normal proportions in the spleen of mice deficient in T cells (TCRα knockout), B cells (IgM knockout) and all lymphocytes (RAG1 knockout). In all of these cases, DCs were found in their appropriate anatomic locations within the white pulp, with the CD8α subset in the marginal zone, and the CD8α subset located around the central arteriole of the white pulp. Interestingly, this relationship was retained even in RAG1 knockout mice in which there were no lymphocytes at all to separate the DC subsets; the DC subsets were found in concentric rings around central arterioles with the CD8α cells holding their place on the inside ring. It would appear that this arrangement must be maintained by a gradient of morphogens, focused on the central arteriole. Whether this gradient is formed by specific adhesion molecules or soluble factors is not yet clear. Precedence in other developmental systems suggests a gradient of soluble factors, though a standing gradient may be maintained by binding of the soluble factors to the extracellular matrix. One prime candidate for the soluble factor is the chemokine TCA4/SLC, since it appears that high level expression of this chemokine also insures the localization of DCs and T cells to the appropriate compartment of lymph nodes (Gunn et al., 1998a, 1999). In the spleen TCA4/SLC expression is

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not so prominent in the central arteriole, but expression by stromal cells anchored to the central arteriole may perform the same function. The concentric arrangement of CD8α and CD8α DCs in the white pulp might be accomplished in the same way that B cells were recruited to the perimeter of islet infiltrates in Ins-TCA4 transgenic mice. Thus, one prediction would be that the CD8α subset should show either lower level expression of CCR7 or a different migratory behavior in response to the chemokine as compared with the CD8α subset. As reagents specific for CCR7 become available we will be able to test this notion, but in the meantime it is suggestive that in plt/plt mice deficient in TCA4/SLC expression, DC localization appears to be random within the spleen (Gunn et al., 1999). Yet while the development and localization of DCs in the spleen (a ‘primary lymphoid tissue’) appears to be normal in the absence of lymphocytes, can the same be true in secondary lymphoid tissue? That is, can the expression of the chemokine TCA4/SLC by itself (e.g. on HEV) be sufficient to recruit DCs? Studies on migration of Langerhans cells from the skin suggest that activated DCs express the CCR7 receptor and are drawn directly into the draining lymphatic vessels by SLC expressed by lymphatic endothelial cells (Kellermann et al., 1999; Saeki et al., 1999). Thus, recruitment into islets expressing TCA4/SLC may also be dependent only on the expression of CCR7 by DCs. Based on these studies, one would predict that the Ins-TCA4 transgene backcrossed to the RAG1 knockout should still induce recruitment of DCs to the islets. Surprisingly, the islets of these mice showed no recruitment of infiltrating cells of any kind (Fan et al., 2000). Since both phenotypic subsets of DCs were present in the spleen, the inability to recruit these cells to islets could reflect a failure of these DCs to express CCR7. Alternatively, DCs may indeed be recruited to islets, but in the absence of lymphoid tissue or other factors they might only pass through the tissue and fail to accumulate in significant numbers. In both cases, it is possible that the presence of other cells such as T

lymphocytes may be required for the accumulation of DCs. This was tested in part by backcrossing the Ins-TCA4 transgene to Ikaros-null mice. The targeted disruption of the gene encoding the transcription factor Ikaros has a number of effects on the immune system (Georgopoulos et al., 1997). This transcription factor is considered to be critical in lymphoid cell development, and so Ikaros knockout mice entirely lack B lymphocytes. T lymphocytes are also absent during fetal development, though later expression of the related factor Aiolos appears to permit development of T cells beginning in the perinatal period. DCs are reduced in number, but the CD8α subset appears to be present in the spleen (Wu et al., 1997). Perhaps due to the defects in fetal lymphocyte development or other defects, the Ikaros knockout mice fail to generate any peripheral lymph nodes at all (Wang et al., 1996). Despite the absence of peripheral secondary lymphoid tissue in Ins-TCA4/Ikaros knockout mice, islets formed new lymphoid tissue, complete with high endothelial venules expressing MAdCAM-1 and PNAd, and ER-TR7 reticular stromal cells (Fan et al., 2000). Contained within these lymphoid structures were CD4 and CD8 T cells, and CD11cCD8αDEC-205 DCs. Thus, the presence of T lymphocytes was sufficient for the induction of both secondary lymphoid stroma and CD8α DCs even in mice lacking normal peripheral lymph nodes.

T-CELL INDUCTION OF LYMPHOID TISSUE AND DC: LYMPHOTOXIN-MEDIATED EVENTS? The induction of lymphoid stromal development and DC recruitment appears possible through the action of naïve T cells (in Ins-TCA4 mice) or activated T cells (autoimmune T cells in diabetes, nonspecifically activated cells in RIPLT mice). While it is not clear that naïve and activated T cells act through the same mechanisms, studies suggest that one common pathway may be through lymphotoxin-mediated signals. For

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example, as discussed above, mice deficient in lymphotoxin β (LTβ) have been found to lack peripheral lymph nodes, although mesenteric or cervical nodes can sometimes be found. Mice deficient in lymphotoxin α and mice deficient in the LTβ receptor both have even more profound deficits in lymph node development. To test whether lymphotoxin ligands were required for the development of lymphoid tissue in the islets of Ins-TCA4 mice, we backcrossed the transgene to LTβ knockout mice. In preliminary studies (unpublished results), we found that the transgenic TCA4/SLC was still effective in recruiting large numbers of T and B lymphocytes to the islets. However, lymphoid stromal cell development (HEV, and ER-TR7 reticular stroma) was not induced, so islets were surrounded only by loose collections of lymphocytes without separate T- and B-dependent compartments. Even more striking, the infiltrates failed to recruit significant numbers of DCs; thus, CD11c and DEC-205 were not detectable in the infiltrates, even though cells with this phenotype were detectable in significant numbers in the spleen. As discussed above, in the organized islet infiltrates of regular Ins-TCA4 transgenic mice, the DCs were limited to the T cell-dependent regions, while F4/80 macrophages were excluded to the perimeter of the infiltrate. In contrast, infiltrates in Ins-TCA4/LTβ knockout mice had scattered F4/80 cells throughout. Whether these cells represent CCR7 DC precursors or a

Undifferentiated Stroma

random infiltration by neighboring tissue macrophages is not clear. Thus, in the backcrosses of the Ins-TCA4 transgene to both RAG1 knockout and LTβ knockouts, deficiencies in T cells or their signaling molecules were associated with a failure of the TCA4/SLC to recruit DCs to islets, even though mature phenotype DCs were present in the spleen. These conclusions may help explain the potential disparity in the observations that RAG1 and T cell knockouts contain normal DC subsets, yet functional studies on DCs indicate a dependence on T cells for full functional maturation. The T cell–DC interaction may be critical in situations where T-cell responses are most likely to be triggered, such as in lymph nodes draining the skin.

SUMMARY: ARE SECONDARY LYMPHOID TISSUE DCs PART OF AN ACTIVE DIALOGUE WITH T CELLS? The results from our studies can be fitted into a model of secondary lymphoid tissue development (Figure 9.1). The left section of the figure shows that within normal peripheral tissues, stromal cells such as vascular endothelium and fibroblasts may perform some routine functions, but also retain potential as precursors of more specialized cells. In the middle section of the figure, events

In fetal development: CD4CD3LTβα4β7cell In diabetes: Activated T cell In Ins-TCA4 mice: Dendritic cell Naive CCR7 LTβ Tcell or precursor

vascular endothelium

LTβR

MAdCAM-1 LTβR 

FIGURE 9.1

Differentiated Lymphoid tissue Genes induced: HEV



fibroblasts/ mesenchyme

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LTβR

Lymphoid Dendritic cells Stromal Reticulum

RelB, CD8α CCR7 BP-3, ER-TR7 TCA4/SLC

Follicular FDC-M1, FDC-M2 Dendritic cells BLC

Model of secondary lymphoid tissue development (see text for details). DENDRITIC CELL BIOLOGY

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can lead to recruitment of T lymphocytes (e.g. autoimmune responses, or the Ins-TCA4 transgene). The proximity of these T cells expressing the LTβ ligand (or a related ligand such as LIGHT) induces differentiation in three distinct stromal cell types. First, vascular endothelium can differentiate into high endothelial venules, and become specialized for recruiting additional lymphocytes and DCs through expression of the chemokine TCA4/SLC and addressin molecules such as MAdCAM-1 and PNAd. Second, mesenchymal cells can be induced to differentiate into specialized stromal reticulum. These cells not only provide structure to the lymphoid tissue, but also can express additional TCA4/SLC. Third, T cell interactions with DCs or their precursors appears to be important in causing their recruitment and persistence within the developing lymphoid tissue. In the right section of the figure, we now see the fully developed lymphoid tissue. The proper orientation of the various stromal cells and lymphocytes has been established, in part due to the opposing gradients of chemokines (TCA4/SLC from HEV drawing T cells and DCs in one direction, BLC from follicular DCs drawing B cells in the opposite direction (Gunn et al., 1998b)). The HEV is established as the anchor of the T cell/DC compartment, and continuous interaction between T cells and DCs is possible. The implication from these studies and the model is that secondary lymphoid tissue DCs are in some way seduced into staying in the lymph nodes by an active engagement with T cells. This is of course a two-way dialogue; studies on lymphocyte homeostasis have suggested that the persistence of naïve T cells in the periphery is dependent on low-level stimulation of T cells by self MHC antigens, most likely on DCs.

CONCLUDING REMARKS An outstanding wine can only be properly appreciated in the context of the finest food; in the same way, the complexities of DC function can be best appreciated in the context of a comprehensive understanding of lymphocytes and

lymphoid tissues. As we can see from the studies discussed here, DC development and function is becoming ever more integrated with other components of the immune system: studies on the role of hematopoietic transcription factors indicates how the plasticity of the developmental program drives different DC phenotypes in different situations. Studies on the development of organized secondary lymphoid tissue show how DC recruitment and development is critically shaped by a series of environmental factors, including their lively and engaged discussions with T lymphocytes. DC-ologists will be merged with T-cell-ologists and B-cell-ologists, and we will all once again be immunologists!

ACKNOWLEDGEMENTS We thank Dr Monica Carson for helpful discussions and critical review of the manuscript. These studies were supported by grants from the National Institutes of Health, the Juvenile Diabetes Foundation International, and the International Human Frontier Science Program. This is manuscript No. 13174-IMM from The Scripps Research Institute.

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10 Mobilization, migration and localization of dendritic cells Jonathan M. Austyn University of Oxford, John Radcliffe Hospital, Oxford, UK

A journey of a thousand miles begins with a single step. Chinese proverb

INTRODUCTION

DC subsets Studies of DCs grown from progenitors cultured in the presence of various cytokine combinations have revealed distinct pathways for development of DC subsets, including Langerhans cells (LCs), myeloid DCs, monocyte-derived DCs, and lymphoid DCs (Austyn, 1998). After production in the bone marrow (or fetal liver) it is likely that the in vivo counterparts of the first two subsets, and probably the third, enter nonlymphoid tissues where they reside transiently before migrating to secondary lymphoid tissues. Their principal function appears to be the induction of immunity to foreign antigens (Young and Steinman, 1996; Cella, et al., 1997; Steinman and Inaba, 1999). In contrast, lymphoid DC progenitors probably enter the thymus and develop into DCs that play a crucial role in the induction of thymic (central) tolerance. Recent evidence also indicates that a subset of DCs possibly related to lymphoid DCs may enter secondary lymphoid tissues directly from

Dendritic cells (DCs) are responsible for the initiation of many immune responses against foreign antigens and most likely play a central role in the induction of tolerance to self antigens. These functions can be ascribed to different subsets of DCs and to different stages in their life history. This chapter summarizes current understanding of four essential stages in the in vivo immunobiology of DC subsets responsible for the initiation of immunity: recruitment into nonlymphoid tissues; mobilization from these sites; migration through lymph and blood; and localization within secondary lymphoid tissues. It will also address the recruitment and direct entry to lymphoid tissues of DC subsets that may induce self tolerance and/or regulate immune responses to foreign antigens. Some general principles will first be outlined.

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the blood. These cells may have regulatory functions that contribute to the induction or maintenance of extrathymic (peripheral) tolerance to self antigens. An important difference between DC subsets in lymphoid tissues is therefore whether or not they are derived from DCs that entered nonlymphoid tissues. Further information on the origin of these DC subsets is in Chapters 1–4 and 6.

DC maturation In general, freshly isolated DCs from nonlymphoid tissues can internalize foreign antigens by pinocytosis (soluble antigens) and phagocytosis (particles); macropinocytosis has also been demonstrated, for example in monocytederived DCs (Steinman and Swanson, 1995; Austyn, 1996; Lanzavecchia, 1996). Freshly isolated DCs such as LCs from skin have a developed endolysosomal system for generation of peptides from internalized proteins, and the isolated cells exhibit a high biosynthetic rate for MHC class II and invariant chain molecules, presumably for loading with antigenic peptides and expression as foreign peptide–MHC complexes at the plasma membrane. In general, DCs isolated from nonlymphoid tissues have little or no expression of co-stimulatory molecules such as CD40, CD80 and CD86, and they have little or no capacity to initiate responses of naïve T cells in vitro. Cells with these properties can be termed ‘immature’ or ‘processing’ DCs. Profound phenotypic and functional changes can occur during culture of DCs isolated from nonlymphoid tissues. For example, in the presence of GM-CSF, cultured LCs lose the capacity to internalize and process antigens, and to synthesize MHC class II and invariant chain molecules. In contrast, membrane expression of foreign peptide–MHC class II complexes is increased, the cells express higher levels of costimulatory molecules such as CD40, CD80 and CD86, and they can now initiate responses of naïve T cells in vitro. Furthermore, they secrete cytokines (e.g. IL-12) that polarize T-cell responses (e.g. induction of TH1 cells). In many respects these cells resemble DCs that can be

isolated from secondary lymphoid tissues such as spleen and lymph nodes (LN). Cells with these properties can be termed ‘mature’ or ‘costimulatory’ DCs. The development of cells from an immature, processing stage towards a more mature, co-stimulatory stage is generally referred to as ‘maturation’ or ‘activation’.

Leukocyte migration and chemotaxis Collectively, all leukocytes have the capacity to migrate through body fluids and to localize within different anatomical compartments. Examples include naïve T cells that recirculate between lymph and blood, naïve T and B cells that localize in secondary lymphoid tissues, activated T cells that access peripheral sites of inflammation, and plasma cells that localize in part to bone marrow. In the classical paradigm for leukocyte migration from vascular to extravascular sites, the cells roll on vascular endothelium (weak adhesion mediated in part by selectins), can be signalled to arrest, flatten and adhere strongly to endothelial cells (strong adhesion mediated in part by integrins), and subsequently undergo diapedesis into the underlying tissue. An unusual feature of DCs is their capacity for reverse transendothelial migration (TEM) from nonlymphoid tissues into the blood. The directed movement, or chemotaxis, of leukocytes to particular compartments occurs in response to both classical chemoattractants, such as bacterial N-formyl peptides (fMLP) and complement components (C5a), and to a family of chemoattractant cytokines or chemokines (CKs). Most of the latter fall into one of two families, the CC (β) and CXC (α) chemokines, although a single C chemokine (lymphotactin) and an unusual membrane-bound molecule with a terminal CX3C domain (fractalkine) have been identified (Schall and Bacon, 1994; Hedrick and Zlotnik, 1996; Schluger and Rom, 1997; Homey and Zlotnik, 1999). Many of these molecules are produced at inflammatory sites (e.g. inflamed endothelium), while others are produced constitutively in certain tissues including secondary lymphoid tissues. It is therefore

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possible to distinguish broadly between ‘inducible’ or ‘inflammatory’ chemokines, versus ‘constitutive’, chemokines respectively. Much has been learnt of the response of DCs to different classes of chemokines (see Table 10.1).

RECRUITMENT TO NONLYMPHOID SITES DC progenitors can enter nonlymphoid tissues and develop into cells with specialized capacities for uptake and processing of foreign antigens. These cells are localized in topologically external epithelial sites, such as skin epidermis and the mucosae of the respiratory, gastrointestinal and urogenital tracts. Cells with similar functions are also present in the interstitial spaces of vascularized organs such as heart and kidney. However, in normal circumstances, DCs are absent from ‘immunologically privileged’ sites such as central cornea, the bulk of the central nervous system (CNS), and the testis. With a few exceptions, the functions of DCs in nonlymphoid tissues have been elucidated mainly after isolation of the cells and studies of their properties in vitro. For example, much information has come from studies of Langerhans cells (LCs) from skin epidermis, and of DCs isolated from interstitial spaces of vascularized organs such as heart and kidney. The isolated cells have a high rate of biosynthesis of MHC class II molecules and invariant chain in vitro, and they express high levels of foreign peptide–MHC complexes at the cell surface. However, these may not be features of DCs residing in normal, unperturbed tissues. For example, interstitial DCs in rat heart express MHC class II molecules but lack the invariant chain (Saleem et al., 1997), suggesting that the cells in situ may not normally produce peptide–MHC complexes unless they are stimulated, for example by a local inflammatory response. As noted above, distinct DC progenitors can give rise to cells resembling either LCs or interstitial DCs (reviewed in Cella et al., 1997; Austyn, 1998; Steinman and Inaba, 1999). Under defined

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culture conditions (e.g. in the presence of GMCSF plus TNFα), LCs can be generated from human CD34 CLA progenitors that give rise to CD1a cells via a committed CD14 CD1a precursor. In contrast, myeloid DCs are derived from CD34 CLA progenitors that give rise to CD1a cells via a bipotential CD14 CD1a precursor that can alternatively differentiate into macrophages in the presence of M-CSF (Caux et al., 1996). The presence of TGFβ appears to promote growth of human LCs in culture (Strobl et al., 1996), and LCs are absent from the skin of mice lacking a functional TGFβ gene, whereas DCs are present in their lymphoid tissues (Borkowski et al., 1996a). A converse phenotype to that of TGFβ knockout mice is reportedly seen in mice lacking relB or Ikaros transcription factors (Burkly et al., 1995; Wang et al., 1996; Wu et al., 1997). In vivo, these progenitors may enter epithelial tissues or the interstitial spaces of vascularized organs, respectively, in order to maintain basal levels of DCs during the steady state. The requirement for a distinct progenitor to LCs, as opposed to interstitial DCs, may reflect the fact that these cells require specialized properties in order for them to gain access to nonvascularized sites such as skin epidermis. Moreover, they may elicit different types of immune response. For example, myeloid DCs, but not LCs, can modulate B-cell responses (Caux et al., 1997) and there is some evidence that LC can preferentially induce cytotoxic T cells. Under defined culture conditions, it is also possible to generate monocyte-derived DCs. Culture of CD14CD1a human peripheral blood monocytes in GM-CSF plus IL-4 or IL-13 induces the development of CD14lowCD1a cells (Young and Steinman, 1996; Cella et al., 1997; Austyn, 1998), whereas culture in M-CSF results in generation of macrophages. Therefore monocytes may be recruited to inflammatory sites and develop into DCs or macrophages depending on the local cytokine milieu or other factors such as the presence of particulate antigens. In this respect, monocytes could thus provide a rapidly mobilized source of DC progenitors during inflammation and infection. Therefore,

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Systematic name

Chemokines and receptors expressed by dendritic cells and lymphocytes Chemokines

Chemokine receptors

I-309 MCP-1 MIP-1α MIP-1Β RANTES

Yes (line) Yes Yes Yes

CCL7

MCP-3

CCL9, CCL10 CCL13 CCL15 CCL17

MIP-1γ

Yes Yes

CCL23 CCL25 CXCL8

MCP-4 MIP-5 TARC, dendrokine PARC, MIP-4 ELC, MIP-3β LARC, MIP-3α SLC,6Ckine MDC, DCtactin-β (ABCD-1) MPIF-1 TECK IL-8

CXCL9 CXCL10 CXCL11 CXCL12 CXCL13 CX3CL1

MIG IP-10 I-TAC SDF-1α , SDF-1Β BLC, BCA-1 Fractalkine

CCL18 CDL19 CCL20 CCL21 CCL22

Yes

Chemokine receptor

TH1, TH2

CCR8 CCR2 CCR1, CCR5 CCR5 CCR1, CCR5 or CCR3 CCR1, CCR5 or CCR3 Unknown

B TH1, TH2 TH1, TH2

Yes Lymphoid DC? Yes

Immature

TH2

Unknown CCR7 CCR6 CCR7 CCR4

Mature Immature Mature Mature

Naïve T, B TH2

CCR2, CCR3 CCR9 CXCR1, CXCR2

Immature Immature Immature

TH2, thymocytes Thymocytes B

CXCR3 CXCR3 CXCR3 CXCR4 CXCR5 (BLR1) CX3CR1

Plasmacytoid monocytes Plasmacytoid monocytes Plasmacytoid monocytes Mature

TH1 TH1 TH1 Naïve T, TH1, B B Activated T

TH1, TH2, GC-B

No

Immature Immature

TH2 TH1 TH1, B TH1 TH1 or TH2 TH1 or TH2 Naïve T, activated T TH2

TH1

(line) Yes No Yes (membrane)

Immature Immature Immature Immature

Expression by, or response of, lymphocytes

CCR2, CCR3 CCR1, CCR3 CCR4

Yes Yes No Yes

Expression by, or response of, dendritic cells

TH2 Naïve T Naïve T, B

A summary of chemokines and receptors known or believed to be expressed by DCs and/or lymphocytes, based either on detection of mRNAs or proteins within the cells, or on observation of chemotactic responses of the cells. Not necessarily all chemokines are expressed by all types of DC, and there may be species differences (for example, CCR6 may not be expressed on mouse DCs). CCR6 is expressed on mouse Peyer’s Patch CD11CCO8α - CD116DC . Unless stated, chemokine and receptor expression on immature versus mature DC refers principally to Langerhans cells and/or myeloid DCs and/or monocyte-derived DCs. Only predominant patterns are indicated (for example, TH1 cells have been reported to produce MDC, and to express low levels of CCR4, but in both cases they appear to be expressed predominantly by TH2 cells). For a full human nomenclature, see Homey and Zlotnik (1999) on which this table is based.

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CCL1 CCL2 CCL3 CCL4 CCL5

Secretion by lymphocytes

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human DCs can develop from CD14CD11c progenitors (monocytes). In contrast, LCs and myeloid DCs may develop from circulating CD14CD11c progenitors that have been identified in human blood, whereas lymphoid DCs may originate from CD14CD11c progenitors.

Constitutive recruitment Presumably, tissue-derived signals are responsible for ‘constitutive’ recruitment of DC progenitors from the blood in the normal steady state. However, in contrast to inflammatory situations (see below), little is known of this process. It is possible that, in the absence of infection per se, local flora and fauna (e.g. skin and gut commensal bacteria) provide a basal level of (subclinical) inflammatory stimuli to which progenitors can respond. Alternatively, DC-attracting chemokines could be produced constitutively and at low levels in normal tissues. An example could be MIP-3α which is produced by keratinocytes and venular endothelial cells in apparently normal skin; it has been demonstrated that CD1a LC precursors express the corresponding chemokine receptor (CCR6) and respond selectively to this chemokine in vitro, whereas CD14 precursors of DCs and monocytes do not (Charbonnier et al., 1999).

Recruitment during inflammation and infection Recruitment of DC progenitors DC recruitment to nonlymphoid tissues in response to acute inflammatory stimuli has been particularly well documented in the case of rat lung and airways. For example, a wave of DC enters these sites in response to local challenge with a variety of agents, including bacterial, viral and soluble protein antigens (McWilliam et al., 1996, 1997). One chemokine that could be involved in recruitment is MIP-3α, which is expressed in inflamed and mucosal tissues, and for which lung DCs express receptors (Power et al., 1997). Expression of MDC has also been demonstrated in a mouse model of lung inflam-

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mation and it has been suggested that this chemokine may be involved in transit or retention of leukocytes, perhaps including DCs, within the tissue (Gonzalo et al., 1999). Similar responses are likely to occur in other nonlymphoid tissues. For example, DC progenitors enter heart and kidney after systemic administration of LPS to mice (Roake et al., 1995b), presumably in response to cytokines or chemokines induced by this agent. DCs may also be recruited into the brain, from which they are normally absent. For example, in rat, infiltrates of cells resembling DCs have been identified within CNS allografts (Lawrence et al., 1990) and an influx of cells expressing the OX62 antigen (a marker of DCs) has been described in certain inflammatory situations (Matyszak and Perry, 1996). In mouse tumour models, it has been suggested that MIP-1α may be responsible for DC infiltration into sites of GM-CSF production (Kielian et al., 1999), and MCP-3 has also been implicated in recruitment to perivascular sites (Fioretti et al., 1998). In the above examples it has not been clearly established whether or not the cells are recruited into the tissues as DC progenitors, or as monocytes that subsequently differentiate into DCs. However, certain DC subsets do express counterligands for molecules expressed by inflamed endothelium. These include: PSGL-1 which binds E-selectin (CD62E) and P-selectin (CD62P) (Laszik et al., 1996); VLA-4 and VLA-5 (CD49d/CD29, CD49e/CD29) which bind VCAM-1, CD62E and/or fibronectin (Strunk et al., 1997); and CLA which binds CD62E (Strunk et al., 1997). Expression by DC subsets of the neural cell adhesion molecule L1, which binds the αvβ3 integrin, and gp40, the murine homologue of human epithelial cell adhesion molecule, has also been reported (Borkowski et al., 1996b; Pancook et al., 1997). Recruitment of monocytes and differentiation to DCs Recently it has been demonstrated that monocytes can differentiate into LN DCs in vivo (Randolph et al., 1999). It was previously shown

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that human monocytes can differentiate into DCs in an in vitro model of transendothelial migration (TEM), particularly following a phagocytic stimulus in the absence of exogenous cytokines. Subsequently, it was shown that an intracutaneous injection of latex microsphere into mouse skin resulted in monocyte infiltration at the site (Randolph et al., 1999). The monocytes phagocytosed the particles and subsequently migrated to regional LNs, where they acquired the characteristic phenotype of DCs. Control experiments clearly established that the latex-laden cells acquired the particles within the skin, before migration, and that they could be distinguished from a minor subset of Langerhans that acquired (very few) particles before also migrating to the nodes. Furthermore, the former cells were not observed in lymph nodes of monocyte-deficient op/op mice. These observations are reminiscent of those from other studies demonstrating the recruitment of DCs to rat liver sinusoids after systemic injection of latex microspheres, and their subsequent blood–lymph translocation and migration to coeliac LNs (Matsuno et al., 1996). It has been suggested that the acquisition and presentation of particulate (as opposed to soluble) antigens may be a major role of DCs derived from monocytes. Role of chemokines As described in Chapters 6, 14, 15 and 20, an increasing number of studies are addressing stimuli that induce chemotaxis of human DCs in vitro. Some examples have already been noted. For convenience, a summary of the various chemokines that can act on DCs at different stages of maturation is in Table 10.1. There are differences depending on the precise source of DC, for example whether the cells were generated from CD34 progenitors or from CD14 blood monocytes, and not all chemokines may be active on any given DC population. However, in general, immature DCs can respond to ‘inducible’ or ‘inflammatory’ chemokines such as MCP-1, MIP-1α, MIP-1β, RANTES, MCP-3, MIP-5, MIP-3α and IL-8, and it seems probable

that they attract DCs into and/or within nonlymphoid sites of inflammation (Dieu et al., 1998; Lin et al., 1998; Sozzani et al., 1997b; Vecchi et al., 1999). Interestingly, and as indicated in Table 10.1, many of these chemokines also preferentially attract activated T cells of TH1 or TH2 type, suggesting that they contribute to the coordinated regulation of DC–T cell interactions in peripheral sites. After maturation, DCs lose their responsiveness to inducible chemokines due to receptor desensitization or receptor downregulation, and they become responsive to a set of ‘constitutive’ chemokines as discussed later. Maturing DCs can also secrete higher levels of many chemokines to which immature DCs as well as naïve and/or activated T cells can respond. In addition to chemokines, immature DCs undergo chemotaxis towards other ‘endogenous’ mediators, such as C5a (Sozzani et al., 1995; Morelli et al., 1996), platelet-activating factor (Morelli et al., 1996; Sozzani et al., 1997a) and IL16 produced by activated T cells (Kaser et al; 1999). They also undergo chemotaxis towards ‘exogenous’ mediators such as bacterial formyl peptides (Sozzani et al., 1995), HIV-derived Mtropic gp120 (Lin et al., 2000) and, reportedly, HIV-Tat (Benelli et al., 1998). This indicates that DCs can be attracted to sites of tissue injury, inflammation and infection, presumably facilitating their acquisition of foreign antigens and initiation of immune responses. At the same time, however, recruitment of DCs by viruses such as HIV may contribute towards more generalized infection of these cells, and certain viruses such as HSV may subvert the immune response by inhibiting the response of DCs towards chemokines (Chan et al. 1999; Salio et al., 1999).

MOBILIZATION FROM NONLYMPHOID SITES As a general rule, DCs are mobilized from peripheral sites in response to inflammation and infection, and they subsequently migrate via lymph or blood into secondary lymphoid tissues. These events are accompanied by maturation of the cells from an immature, processing

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stage towards a more mature, co-stimulatory stage (Austyn, 1996, 1998; Cella et al., 1997; Reis e Sousa, et al., 1999). In addition, inflammatory stimuli and microbial agents can recruit DCs to nonlymphoid tissues, presumably to replace the mobilized populations.

Mobilization from epithelia Numerous studies have documented a role for the inflammatory cytokines IL-1β and TNFα in migration of epidermal Langerhans cells from both human and mouse skin (e.g. Enk et al., 1993; Cumberbatch et al., 1997a; Cumberbatch et al., 1997b; Cumberbatch and Kimber, 1999; Stoitzner et al., 1999); the latter cytokine also promotes survival of LCs in culture. IL-1β, which is produced in skin exclusively by epidermal LCs, appears to act in an autocrine manner on LCs which express IL-1R type I (and type II receptors at lower levels) (Cumberbatch et al., 1998). IL-1β also acts in a paracrine fashion via the production of TNFα which appears to function predominantly via TNFR II (p75) rather than TNFR I (p55), at least in mice (Wang et al., 1997; but note that epidermal LCs were not examined directly in this study); both receptors have, however, been implicated in migration of corneal LCs in response to TNFα (Dekaris et al., 1999). Mobilization of LCs from the epidermis in response to these cytokines can be inhibited by IL-4, which downregulates TNFR II (p75) in human (Takayama et al., 1999), and by IL-10, which may inhibit production of both TNF and IL-1β in mice (Wang et al., 1999). In addition TGFβ, which is required for LC development in vitro and in vivo, prevents maturation of LCs in response to IL-1, TNFα and LPS, but not to CD40 ligation (Geissmann et al., 1999). IL-1α, produced by epidermal cells in response to staphylococcal enterotoxin A (SEA) has also been implicated in depletion of LCs from epidermis in response to this agent (Shankar et al., 1999). As noted later, systemic administration of LPS mobilizes DCs from rat gut. Molecular events during DC mobilization are becoming increasingly understood. For example during maturation and/or migration, epidermal

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LCs have been shown to downregulate E-cadherin, which may facilitate detachment from keratinocytes (Cumberbatch et al., 1996); to utilize α6 integrin molecules, perhaps for adhesion to laminin in the basement membrane (Price et al., 1997); and to secrete type IV collagenase (matrix metalloproteinase 9) presumably to enable crossing of the basement membrane (Kobayashi et al., 1999) during egress from the epidermis. In addition, LCs upregulate expression of particular splice variants of CD44 (v4, v5, v6, v9) during maturation (Kobayashi et al., 1999). Perhaps surprisingly, LC depletion can also be inhibited by injection into skin explants of mAbs or agonists specific for p-glycoprotein (MDR); this molecule was previously only known as a transporter that contributes to multidrug resistance to chemotherapeutic agents (Randolph et al., 1998b).

Mobilization from vascularized tissues Microbial products, such as bacterial lipopolysaccharide (LPS), or cytokines elicited by this agent such as TNFα or IL-1, also play a central role in DC maturation and mobilization of DC populations from vascularized tissues (Austyn, 1996). For example, systemic administration of LPS induces depletion, presumably by mobilization of DCs from mouse heart and kidney, and as noted earlier, recruits DC progenitors to these sites (Roake et al., 1995a, 1995b). Hence DCs may be mobilized from peripheral sites in response to both endogenous mediators (inflammatory cytokines) and exogenous agents (microbial products), although it is not yet clear to what extent each plays a role. Certainly, DC subsets are known to express toll-like receptors (TLR-4) for LPS of Gram-negative bacteria. Maturation, and perhaps migration, of DCs can also be induced by mycolic acid of Gram-positive bacteria, and by exposure to mycobacteria and stappylococcal enterotoxin A, both of which induce TNF or IL-1 secretion (Shankar et al., 1996; Thurnher et al., 1997). When DCs are mobilized from vascularized tissues, they may enter the blood and travel to the spleen and/or migrate via afferent lymph

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into regional lymph nodes. An apparently unique property of DCs is their capacity to undergo reverse TEM, migrating across the vascular or perhaps lymphatic endothelium, respectively, in an abluminal to luminal direction. When human monocytes were placed in a collagen matrix containing latex microspheres and a monolayer of endothelial cells (EC) was cultured on top of the matrix, the monocytes were observed to develop into DCs and to undergo reverse TEM across the endothelial monolayer and out of the matrix (Randolph et al., 1998a). It has been suggested that this in vitro model mimics the events that occur when monocytes are recruited into tissues and subsequently develop into DCs in response to a phagocytic stimulus in the absence of exogenous cytokines, before mobilization from the tissue and migration into the blood or lymph. Reverse TEM of monocyte-derived DCs has also been studied using monolayers of human vascular EC (D’Amico et al., 1998): adhesion to resting ECs required CD11a and CD11b, but not CD11c; adhesion to activated ECs required VLA-4 and CD18; and, in the presence of fibronectin, both VLA-4 and VLA-5 were utilized; while CD31 was involved in TEM. In addition, both the urokinase plasminogen activator receptor (uPAR) and MIP-1α are involved in TEM and extracellular matrix invasion by CD14CD34 DC progenitors in vitro (Ferrero et al., 2000).

Role of DC-derived chemokines in efferent responses As noted above, maturing DCs can secrete a variety of different chemokines. It is most likely that DCs commence maturation in peripheral tissues before their migration out of these sites and therefore they may produce chemokines at this stage. For example, LCs stimulated by LPS in vitro can produce chemokines such as MIP-1α, MIP-1β, RANTES and IL-8 (Lore et al., 1998), which in vivo could result in recruitment of immature DCs and replacement of the mobilized populations. In addition, chemokines produced by maturing DCs in peripheral sites

may facilitate the recruitment of activated T-cell subsets and hence contribute to the effector phase of immune responses. For example, in addition to the chemokines just noted, MDC is produced by LCs at sites of contact sensitization and by dermal DCs in atopic dermatitis, (Tang and Cyster, 1999; Vestergaard et al., 1999), and TARC is secreted constitutively by lung LCs (Table 10.1). In addition, maturing mouse LCs synthesize ABCD-1 which induces chemotaxis of activated T cells (Ross et al., 1999).

MIGRATION VIA LYMPH AND BLOOD This section describes three migratory pathways for DCs mobilized from peripheral sites: from epithelial tissues to regional nodes via afferent lymph; from vascularized organs to spleen via blood; and from liver sinusoids to celiac nodes via a blood–lymph translocation.

Migration from epithelial sites to lymph nodes Many studies have demonstrated the presence of DCs, termed veiled cells, in afferent (but not efferent) lymph draining skin sites, and it is also possible to obtain such cells from central lymph after mesenteric lymphadenectomy (Austyn and Larsen, 1990). Migration of these cells into regional nodes is likely to be essential for the initiation of immune responses against antigens derived from epithelial tissues. Isolated DCs migrate from skin to regional nodes The capacity of DCs to migrate from skin sites to regional lymph nodes has been clearly demonstrated in a number of tracking studies. DCs isolated from lymphoid tissues, such as mouse spleen (Kupiec-Weglinski et al., 1988), cultured LCs (Liddington and Austyn, unpublished studies) or rat pseudoafferent lymph (Fossum, 1988), have been labelled with fluorochromes or radioisotopes, injected into footpads, and subsequently detected in popliteal

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nodes and localized to T-cell areas. Similar findings have been made using DCs grown in culture from mouse blood and bone marrow progenitors (Inaba et al., 1992; Lappin et al., 1999), and subcutaneous injection of DCs, but not granulocytes, derived with a contact sensitizing agent was shown to induce contact hypersensitivity responses (Lappin et al., 1999). In a primate model, DCs were generated from chimpanzee blood monocytes by culture in GM-CSF and IL4, and fluorochrome-labelled before subcutaneous injection. Subsequently fluorochromelabelled cells were detectable in the T areas of draining lymph nodes for at least 5 days (Barratt-Boyes et al., 1997). These findings, however, contrast with those from a human study in which monocyte-derived DCs labelled with indium-111 were injected into skin of cancer patients (Morse et al., 1999). While a small percentage of radiolabel was detected in regional nodes of some patients after intradermal administration, no lymph node activity was detected after subcutaneous injection. One variable may have been the relative maturation state of the cells in these two studies, though this is not clear. Epidermal LCs migrate to lymph nodes; dermal DCs migrate to spleen Using mouse skin transplants, and mouse and human skin explants, it has been shown that LCs migrate from the epidermis into dermal lymphatics and out of the tissue, and that migration is accompanied by phenotypic and functional maturation of the cells (Larsen et al., 1990b; Lukas et al., 1996). Presumably, LCs are mobilized from the epidermis in response to endogenously derived mediators (e.g. IL-1, TNFα). Furthermore, after skin grafting between allogeneic or congenic strains of rats and mice, small numbers of donor-derived cells can be detected in regional lymph nodes of the recipients (Richters et al, 1999; Austyn and Liddington, unpublished observations). Marked enlargement of the lymph nodes in allogeneic, but not congenic, recipients suggests a response of alloreactive T cells to the administered DC

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(Richters et al., 1999). Interestingly, after skin grafting in the mouse, rare donor-derived cells were also detected in recipient spleen (Liddington and Austyn, unpublished studies). Since these cells were not detected after injection of purified LCs into a subcutaneous pocket to mimic cells entering the bed of a graft, a dermal origin seems likely. The relative contribution of these cells, versus LCs, to initiation of rejection is unclear. When contact sensitizers such as FITC were topically applied to mouse skin, many DCs labelled with the contact-sensitizing agent could be isolated from regional lymph nodes (e.g. Macatonia et al., 1987; Kripke et al., 1990; van Wilsem et al., 1994). The isolated cells were found to induce antihapten responses after adoptive transfer to naïve recipients. In addition, adoptive transfer of cell suspensions from mouse lymph nodes draining skin allografts to which contact sensitizers were applied (see below) was found to induce donor-specific responses in the naïve recipients (Kripke et al., 1990). The interpretation of these and other related observations has been that (1) epidermal LCs acquire contact sensitizers locally before migrating to regional lymph nodes where they induce contact sensitivity reactions, and (2) the DCs accumulating in lymph nodes are derived from epidermal LCs. In apparent contradiction to the foregoing conclusions, after topical application of FITC to skin grafts before or after restoration of lymphatic connections, many FITC-labelled DCs were detected in regional nodes, but exceedingly few were donor-derived (Liddington and Austyn, unpublished studies). Furthermore, estimates of the numbers of LCs that migrated from the skin grafts indicated that these were not sufficient to account for the numbers of FITC-labelled cells detected within regional nodes. Therefore, either FITC can travel directly from skin into regional nodes for acquisition by (recipient) DCs, and/or the contact sensitizer could be acquired by infiltrating (recipient) dermal DCs that then migrate to the nodes. In other studies, injection of TNFα or IL-1 into mouse skin resulted in increased numbers of DCs within regional

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lymph nodes (e.g. Cumberbatch et al., 1997b). This correlated with depletion of epidermal LCs from the site of injection, and both depletion of LCs and accumulation of lymph node DCs were inhibited by administration of blocking cytokine-specific mAbs. However, it is possible that the majority of lymph node DCs were derived from DC progenitors that were recruited into the dermis and/or the cells were recruited directly from the blood into the lymph nodes. There are other indications that DCs can migrate from skin dermis into nodes. For example, migration of dermal cells from skin has been shown to contribute to contact sensitization in mouse (Sato et al., 1998). In addition, when Leishmania parasites were injected into mouse skin they were internalized by presumptive dermal DCs, and parasite-bearing DEC-205 cells were subsequently detected in T-cell areas of regional lymph nodes and persisted for long periods (Moll et al., 1995). The latter findings should also be viewed in the light of observations noted earlier that monocytes can be recruited after injection of particulates into skin, prior to migration and development into lymph node DCs. DCs migrate from gastrointestinal epithelia to regional nodes Following mesenteric lymphadenectomy of rats, DCs that would normally migrate from the gut wall to the mesenteric lymph nodes can be isolated from central lymph after cannulation of the thoracic duct. There is a continuous, low level flux of such cells draining from the gut, probably from the lamina propria (Maric et al., 1996). However, the cell output is markedly increased following systemic administration of LPS (MacPherson et al., 1995). This indicates either that the resident population of DCs is induced to leave the tissue and/or that DC progenitors are recruited to the gut wall prior to exit in the mesenteric lymph. DCs migrating from the gut wall can acquire and present foreign antigens delivered directly into the gut (Liu and MacPherson, 1995). Recently it has been demonstrated that these cells normally contain

apoptotic bodies derived from intestinal epithelium, and it has been suggested that these DCs could potentially contribute to induction of self tolerance (Huang et al., 2000; Steinman et al., 2000).

Migration from vascularized tissues to spleen Migration of DCs mobilized from vascularized tissues via blood to spleen may be essential for the initiation of immune responses against antigens in these sites. However, the relative importance of this route, compared with migration of DCs in lymph draining from vascularized organs into regional nodes, is not yet clear. Isolated DCs migrate from blood to spleen The capacity of DCs to migrate from blood to spleen has been demonstrated in a number of tracking studies. DCs isolated from lymphoid tissues, such as mouse and rat spleen (Austyn et al., 1988; Kupiec-Weglinski et al., 1988; Oluwole et al., 1991) have been labelled with fluorochromes or radioisotopes, injected intravenously, and subsequently detected in spleen and localized to T-cell areas. Similar findings have been made using DCs grown in culture from mouse bone marrow progenitors (Lappin et al., 1999; Suri and Austyn, unpublished studies). Intravenously injected DCs are excluded from the thymus, and from peripheral lymph nodes, although one study (Fossum, 1988) has documented entry to coeliac nodes of DCs derived from rat pseudoafferent lymph perhaps because of their capacity to undergo a blood–lymph translocation via liver sinusoids. An important question is at what stage DCs acquire their capacity to migrate from blood to spleen. Recent studies indicate that stably immature, maturation-resistant DCs cultured from mouse bone marrow in the presence of low concentrations of GM-CSF, as well as mature DCs cultured in various combinations of GMCSF, IL-4, TNFα and/or LPS can enter spleen to a similar extent after intravenous administration, and home to T-cell areas (Suri and Austyn,

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unpublished observations). This suggests that DCs acquire their specialized capacities for homing to lymphoid tissues, and localization within distinct compartments, early in development. One report has also documented that immature (GM-CSF-cultured) bone marrowderived DCs home to splenic T-cell areas after intravenous transfer to allogeneic recipients and that these cells upregulate co-stimulatory molecules (Fu et al., 1996). Whether the injected cells were already committed to maturation, or this was induced by local factors (e.g. cytokines), is not yet clear. DCs migrate from vascularized tissues via blood to spleen The capacity of DCs to migrate from vascularized organs and tissues via blood to spleen has primarily been demonstrated in transplantation settings. After transplantation of fully vascularized cardiac allografts in the mouse (Larsen et al., 1990a), DCs disappeared from the tissue over a period of 1–3 days. Concomitantly, donorderived cells were detected in recipient spleens. Donor cells were first detected in spleen 1 day after transplantation and maximum numbers were present between 2 and 4 days, but they were no longer detectable at day 6. Since the spleen lacks an afferent lymphatic supply, these cells were derived from the blood and presumably represented DCs that had migrated from the transplants. Related observations were made in a rat limb transplantation model (Codner et al., 1990). DCs from vascularized epithelial sites such as the dermis of skin may also migrate via blood to spleen. It seems likely that the blood migration pathway is physiologically important for the induction of immune responses against foreign antigens in vascularized sites. However, a flux of cells resembling DCs was noted in afferent lymph draining sheep renal transplants, presumably en route to regional lymph nodes (reviewed in Austyn and Larsen, 1990). The relative importance of these respective pathways is at present unclear.

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Migration via liver sinusoids to regional lymph nodes Coeliac lymph nodes may be a specialized site for the induction of immune responses against particulate antigens in blood (Austyn, 1996). The evidence for this has come from studies of rats that were injected intravenously with latex particles after coeliac lymphadenectomy had been performed (Matsuno et al., 1996; Kudo et al., 1997). A key finding was that particle-laden DCs that would normally traffic in coeliac lymph could be isolated from the central lymph of these rats, but they were not detected in mesenteric lymphadenectomized animals. Following injection of particulates, DCs are recruited to the liver sinusoids (Matsuno et al., 1996). The signals for recruitment are not yet clear, but one possibility is that Kupffer cells (liver macrophages) phagocytose particles from the bloodstream and then elaborate cytokines or chemokines in response to the phagocytic load that attract DC progenitors; a precedent for this comes from the finding that DCs from coeliac lymph can adhere to Kupffer cells in frozen sections of liver (Kudo et al., 1997). Within the liver sinusoids, the DCs then appear to internalize the particles, before maturing and migrating into coeliac lymph. At this stage, the DCs are unable to phagocytose further particulates in vitro, but they express costimulatory activity that would enable them to initiate immune responses within the coeliac nodes. Alternatively, monocytes may be recruited to the sinusoids and develop into DCs in response to the phagocytic stimulus before migration. Movement of DCs from the liver sinusoids into coeliac lymph involves a blood–lymph translocation, presumably via the space of Disse. This translocation has been demonstrated directly following intravenous injection of DCs isolated from coeliac lymph (Kudo et al., 1997). When cells that had phagocytosed particles in vivo were injected into the blood, they were subsequently localized to the paracortical regions of coeliac lymph nodes. Furthermore, when allogeneic DCs were injected they were found to be associated with proliferating T cells which they

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had presumably activated in these sites. Whether or not DCs originating from other tissues, such as the interstitial spaces of vascularized organs, can also undergo such a blood–lymph translocation is not yet known, although DCs derived from mesenteric lymph can migrate from the blood into the coeliac nodes (Fossum, 1988).

LOCALIZATION IN SECONDARY LYMPHOID TISSUES This section reviews information on the localization of DC subsets derived from peripheral nonlymphoid tissues, such as interstitial DCs that have migrated to spleen, or Langerhans cells that have migrated from skin.

DCs in spleen DC subsets in spleen Two major DC subsets have been identified in mouse spleen with distinct phenotypes (Vremec and Shortman, 1997; Steinman et al., 1997; Anjuere et al., 1999). One subset can be identified in ‘bridges’ that interrupt the marginal zone (MZ) between the red and white pulp. ‘Nests’ of DCs have been demonstrated in these regions, apparently extending into the peripheral T-cell areas but being clearly excluded from the B-cell follicles (Steinman et al., 1997). The phenotype of these cells, termed for convenience ‘MZDCs’, is probably CD11c CD8α DEC205(NLDC145)HSA (CD24)CD11b33D1, similar to that of the bulk population of DCs that can be isolated following mechanical disruption of spleen. In contrast, a phenotypically distinct subset can be identified in central T-cell areas. The phenotype of these cells, which appear to represent interdigitating cells (IDCs), is probably CD11cCD8αDEC205(NLDC145)HSA(CD24) CD11b 33D1, similar to that of a population of DCs that can be isolated from collagenase digests of spleen. MZDCs appear to be relatively immature in phenotype and function, whereas

IDCs appear to be mature (e.g. Steinman et al., 1997). Localization in spleen It is currently believed that MZDCs represent DCs that have entered spleen from blood after mobilization from vascularized tissues, whereas at least some IDCs may be representatives of the lymphoid DC subset that does not enter peripheral sites. It is also believed that MZDCs eventually migrate into central T-cell areas, presumably intermingling with IDCs. Evidence for these ideas comes from a number of studies. First, studies tracing the fate of DCs that were isolated from mouse spleen, labelled with a fluorochrome (H33342), and injected intravenously into syngeneic recipients, revealed that the cells localized to distinct subcompartments at different times after transfer (Austyn et al., 1988). At 3 hours the majority of the labelled cells were found to be present in the red pulp, but by 24 hours they were present in T-cell areas (PALS) of white pulp and were clearly excluded from B-cell follicles. In more detailed kinetic studies (Suri and Austyn, unpublished observations), bone marrow-derived DCs were labelled with fluorochromes (such as CFSE or PKH26) and injected intravenously to syngeneic mice. At 2 hours labelled cells were scattered within marginal zone and red pulp, at 8 hours they were preferentially localized to marginal zone ‘bridges’, and by 24 hours the vast majority were situated in T areas. Thus DCs appear initially to enter the spleen from the blood from the marginal zone (and/or red pulp). Consistent with this idea, fluorochrome-labelled splenic DCs can adhere specifically to the marginal zone of spleen in cryosections (Austyn et al., 1988). Second, after transplantation of vascularized cardiac allografts, donor-derived DCs were detected in recipient spleens (Larsen et al., 1990a). An initially unexpected finding was that these cells were not localized within T-cell areas 1 or 2 days after transplantation (Larsen et al., 1990a). Instead, they were present in B-cell areas of (peripheral) white pulp, or close to the border B-cell and T-cell areas. However, the DCs were

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found to be associated primarily with CD4 but not CD8 T cells in these areas. These observations suggested that movement of the allogeneic DCs through the tissue might be retarded when they encountered alloreactive T-cells, and their association with CD4 T-cells in B-cell areas might contribute to the generation of alloantibody responses. Later studies confirmed these observations and also revealed that donorderived DCs do in fact enter T-cell areas, but only at later time points (3–5 days) after transplantation (Roake et al., 1995b). Third, after systemic administration of LPS to mice, DCs appear to move from marginal zone nests to the PALS, and to undergo functional maturation as assessed in vitro by their reduced ability to capture antigens and enhanced costimulatory activity (De Smedt et al., 1996). In addition, studies have been performed using mAbs specific for peptide–MHC complexes that are expressed by cells in spleen after intravenous immunization (Reis e Sousa and Germain, 1999). After injection of purified protein, only splenic B cells expressed detectable levels of peptide–MHC complexes. However, coadministration of LPS resulted in expression of complexes on DCs that appeared to move from marginal zone into T-cell areas.

DCs in lymph nodes DCs subsets in lymph nodes Three populations of DCs have been identified in mouse lymph node cell suspensions (Vremec and Shortman, 1997; Anjuere et al., 1999). A major subset of CD8αDEC-205CD24CD11b cells resembles MZDCs of spleen and could represent cells that have recently entered the subcapsular space of lymph nodes from nonlymphoid tissues via afferent lymph. A second subset of CD8αDEC-205CD24CD11b cells resembles IDCs of spleen and may correspond to IDCs in paracortical regions (T-cell areas) of lymph node; these cells appear to be relatively mature in phenotype and function. A third subset, trace levels of which are also present in splenic DC suspensions, has the CD8aDEC-

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205CD24CD11b phenotype; the origin of this subset is unknown. A major problem in interpreting the lineage relationship between these subsets (and those in spleen) is that phenotypic changes corresponding to a transition from, for example, MZDCs to IDCs have not generally been observed in culture, apart from the observations that epidermal LCs acquire expression of CD8α and may downregulate 33D1 on maturation (Anjuere et al., 1999, 2000; Herouet et al., 1999). As reviewed elsewhere, follicular dendritic cells (FDCs) within germinal centres (GCs) retain immune complexes on the cell surface and are involved in affinity maturation of B cells. In humans, a distinct subset of CD4CD11c CD3 DCs has also been identified in these sites (Grouard et al., 1996). These germinal centre DC (GCDCs) are present in both the dark and light zones of germinal centres in human tonsils, spleen and lymph nodes, and are associated with GC T cells. It seems likely that GCDCs can induce or sustain the response of memory T cells in germinal centres, and maintain the germinal centre reaction during secondary T cell-dependent responses of B cells. In addition, GCDCs (as well as IDCs in T-cell areas) produce the DC-CK1 chemokine that is chemotactic for naïve CD45RA T cells (Adema et al., 1997). Evidence has also been obtained suggesting that ligation of OX40 on T cells by DCs that express OX40L can result in accumulation of activated CD4 T cells in B-cell follicles (Brocker et al., 1999). Hence DCs may regulate both primary and secondary T-dependent B-cell responses in vivo. Localization in lymph nodes As described above, there is good evidence that DCs can migrate from peripheral sites (e.g. skin or gut wall) via afferent lymph into regional nodes, presumably via subcapsular sinuses. DCs appear to be retained within a given node and it is unlikely that they subsequently migrate via efferent lymph into the next node in a chain. Instead the cells become localized in the paracortical and interfollicular regions (T areas).

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Studies of the migration of antigen-pulsed DCs into lymph nodes of transgenic mice have shed light on the subsequent fate of these cells. When peptide-pulsed DCs were injected into footpads of mice that had been reconstituted with antigen-specific transgenic T cells, they were observed to home to paracortical regions and to cluster with the transgenic T cells, suggesting that antigen presentation could occur. Peptidepulsed DCs persisted within nodes for 2 days but then disappeared, whereas DCs that had not been pulsed with specific peptide homed to the same regions (but did not associate with transgenic T cells) and persisted for at least 4 days. Conceivably the peptide-pulsed DCs were eliminated by the T cells they activated.

DC subsets in Peyer’s patches DC subsets similar to those in spleen and lymph nodes have been described in mouse Peyer’s patches (Kelsall and Strober, 1996; Anjuere et al., 1999). One, resembling DCs of splenic MZ, is situated in the subendothelial dome, while another, resembling IDCs, is situated deeper in central T areas.

Signals for localization of DCs in secondary lymphoid tissues As discussed earlier, immature DCs undergo chemotaxis to a distinct set of inducible or inflammatory chemokines (see Table 10.1). During maturation in vitro, the cells lose their responsiveness to these chemokines due to receptor desensitization and/or downregulation. For example, DCs downregulate mRNA and/or protein expression for CCR1, CCR2, CCR5 and CCR6 (Sozzani et al., 1997b; Dieu et al., 1998; Lin et al., 1998; Vecchi et al., 1999) when stimulated by cytokines such as TNFα, exposed to LPS or subjected to CD40 ligation; responses can be rapid (within 1 hour of stimulation). The maturing DCs then upregulate expression of receptors for constitutive chemokines (see Table 10.1), particularly CCR7 (the receptor for ELC/MIP-3β and SLC/6Ckine) and CXCR4 (the receptor for SDF-1); receptor

upregulation is usually slower (24–48 hours after stimulation). Whereas receptors for inducible chemokines can be downregulated on immature DCs by exposure to the respective chemokine, expression of receptors for constitutive chemokines on mature DCs is refractory to such stimulation (Sallusto et al., 1999). Since constitutive chemokines are expressed in secondary lymphoid tissues, it is most likely that these molecules contribute to the homing to and/or localization of DCs within secondary lymphoid tissues. For example, SLC is produced within various subcompartments of secondary lymphoid tissues including T areas, and by lymphatic endothelium of multiple organs (Gunn et al., 1998). In the case of skin, SLC has been demonstrated within dermal lymphatics containing LCs that are probably migrating from the tissue (Saeki et al., 1999), suggesting that this chemokine may guide the LC along the lymphatics and thence to the T areas of regional nodes. Interestingly, constitutive chemokines also preferentially attract naïve T cells, suggesting that they may facilitate DC–T cell interactions within secondary lymphoid tissues during the induction of immune responses. In the plt mutant mouse there is no production of SLC in lymphoid tissues and, apparently as a result, there are no naïve T cells or DCs in these sites (Gunn et al., 1999). Entry of DCs from blood to spleen is a T-dependent process, since radiolabelled DCs did not enter spleens of nude mice unless they were reconstituted with T cells (Kupiec-Weglinski et al., 1988), but the basis for this is not yet known. Migration to the spleen also requires expression of membrane lymphotoxin by mouse DCs (Wu et al., 1999). As noted earlier, maturing DCs secrete a variety of different chemokines.These include the ‘constitutive’ chemokines ELC/MIP-3β and PARC/DC-CK1 (Hashimoto et al., 1999; Imai et al., 1999;Sallustoetal.,1999)thatarechemo-tacticfor naïve T cells. Therefore, DCs may produce these chemokines within secondary lymphoid tissues to facilitate their interactions with T cells and the induction of immune responses. Such interactions may also be promoted by expression on DCs of the membrane-bound CX3C chemokine

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fractalkine, which is reportedly upregulated in response to CD40 ligation (Kanazawa et al., 1999; Papadopoulos et al., 1999).

LYMPHOID DCs Lymphoid DC progenitors DCs are localized in the medulla and perhaps the corticomedullary junction of thymus. There is evidence that these cells can induce negative but not positive selection of developing thymocytes, and that they are required for induction of thymic (central) tolerance of T cells to self antigens (e.g. Brocker et al., 1997). Thymic DCs, but not bone marrow- derived DCs, can secrete the chemokine TECK, which is chemotactic for thymocytes (and for DCs), presumably via expression of CCR9 (Vicari et al., 1997), perhaps facilitating their interactions with developing T cells within this organ. A major population of phenotypically homogeneous DCs can be isolated from mouse thymus. The phenotype of these cells (CD11cCD8αDEC-205CD24CD11b33D1) is similar to that of DC subsets that can be isolated from spleen and thymus, although trace levels of cells with different phenotypes can also be detected (Vremec and Shortman, 1997; Anjuere et al., 1999). An important characteristic of the major thymic DC subset is the expression of CD8 as αα homodimers (see below). There is no evidence that DCs can migrate from nonlymphoid tissues into the thymus, and it is probable that at least the majority of thymic DCs originate from a distinct progenitor that seeds the thymus and generates cells of the lymphoid DC subset. A thymic lymphoid progenitor has been isolated from mouse thymus that resembles the haematopoietic stem cell except for expression of the Sca-2 antigen and low levels of CD4 (Wu et al., 1996). These CD4lo cells can generate CD8α DCs, T cells, B cells and natural killer (NK) cells when they are adoptively transferred to irradiated recipients. A downstream cell has also been isolated that resembles a pro-T cell

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and which gives rise to T cells and DCs. Further downstream, at the pre-T cell stage, the capacity to generate DCs appears to be lost and the cells become committed to the T cell lineage. Both CD8α DCs and T cells are absent from mice lacking a functional Ikaros gene, which encodes a transcription factor required for development of lymphoid cells (Wang et al., 1996; Wu et al., 1997), and from mice lacking a functional Notch1 gene. In humans, a CD34CD38lo progenitor has been isolated from thymus that has the capacity to differentiate into DCs, T cells and NK cells, and which is distinct from the pluripotent stem cell (Res et al., 1996). A CD34CD10CD45RA progenitor has also been isolated from human fetal and adult bone marrow that can generate DCs, T cells, B cells and NK cells (Galy et al., 1995). Limiting dilution analysis appears to prove that this progenitor is truly multipotent. These and other observations indicate that a developmentally distinct subset of lymphoid DCs originates from bone marrow-derived progenitors that may seed the thymus.

Lymphoid DCs in secondary lymphoid tissues As noted earlier, major subsets of CD8α DCs, of similar phenotype to the predominant DC population in thymus, can be isolated from mouse spleen and lymph nodes. A lymphoid origin for these cells was initially suggested from the fact that CD8α DCs are absent from spleen and lymph nodes of mice with a dominant negative Ikaros gene (Wu et al., 1997), although surprisingly CD8α DCs (but not LCs) are also absent. These cells appear to have regulatory properties in vitro. For example, CD8α DCs isolated from mouse spleen have been reported to activate CD4 T cells but then to induce apoptosis of the activated T cells in a Fas-dependent manner (Suss and Shortman, 1996), and to activate CD8 T cells but to limit T-cell proliferation apparently because they stimulate little IL-2 production (Kronin et al., 1996). They may also skew T-cell responses towards a TH2 profile of cytokine secretion after administration in vivo,

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although this may not be a physiologically relevant route if, for example, these cells do not normally traffic through peripheral, nonlymphoid tissues (see below). In addition, CD8α DCs isolated from mouse lymph nodes express high levels of a self peptide–MHC complex, and can induce T-cell apoptosis after antigen presentation to a T-cell hybridoma (Inaba et al., 1997). It has therefore been suggested that these presumptive lymphoid DCs could be involved in regulation of responses to foreign antigens and/or the induction of tolerance to self antigens. There is evidence for a distinct subset of DCs in human lymphoid tissues that may represent lymphoid DCs. A circulating CD11cCD3CD4CD45RA DC progenitor has been identified in human blood (O’Doherty et al., 1994; Thomas and Lipsky, 1994). Phenotypically similar cells are also present in paracortical regions of lymph nodes. They were originally termed ‘plasmacytoid T cells’, but are now sometimes referred to as ‘plasmacytoid monocytes’. After isolation from tonsils, these cells develop into DCs when they are cultured with IL-3 and stimulated with CD40 ligand (Grouard et al., 1997). The fact that these cells can be detected within the lumen and walls of high endothelial venules (HEVs) within lymph nodes originally suggested that they enter the tissue directly from the bloodstream; in contrast, DCs originating from peripheral sites (e.g. LCs and myeloid DCs) enter nodes via subcapsular sinuses. A lymphoid origin for these cells has not yet been convincingly demonstrated, but it is intriguing that a subset of blood cells with similar phenotype has been shown to give rise to CD3CD4TcRαβ mature T cells (Bruno et al., 1997). One puzzle is that plasmacytoid monocytes are rarely detected in uninflamed lymph nodes. Therefore, it is not easy to draw parallels between these cells and the major subset of presumptive lymphoid DCs in mouse. More recently plasmacytoid monocytes have been shown to produce large amounts of IFNα after exposure to inactivated HSV or influenzavirus, or stimulation by CD40 ligation (Cella et al., 1999; Siegel et al., 1999). These cells, and

their presumptive circulating progenitors, have been shown to express the immunoglobulin-like transcript (ILT) receptor ILT3, but not ILT1. Interestingly those in blood express CD62L that may mediate their adhesion to HEV endothelium, whereas those in tonsil lack CD62L expression, suggesting this molecule may be cleaved after entry to lymph nodes. In addition, the blood progenitors express CXCR3 (the receptor for MIG, IP-10 and I-TAC; Table 10.1) suggesting that these cells can be recruited to lymph nodes in inflammatory situations. In contrast to these (CD11c) ILT3/ILT1 cells, a subset of (CD11c) ILT3/ILT1 cells has also been identified in human blood that seems to be related to myeloid DCs; these cells have heterogeneous expression of CD62L, which is lost on culture, and they lack CXCR3 (Cella et al., 1999). In summary, plasmacytoid monocytes and naïve T cells are recruited from the blood into lymph nodes via HEVs. Both cell types express CD62L, and plasmacytoid monocytes also express CXCR3, while naïve T cells express CCR7, the receptor for SLC that is known to be expressed by HEVs (Gunn et al., 1998). In contrast, DCs migrating in afferent lymph from the periphery enter nodes via the subcapsular sinuses. These cells probably do not express CD62L, and their expression of CCR7 may, at least in part, permit localization within T areas where SLC is also produced. Hence, the molecular basis for the recruitment of different DC subsets to secondary lymphoid tissues, and their localization within particular compartments, is becoming increasingly clear.

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11 Regulation of antigen capture, MHC biosynthesis, and degradation by dendritic cells Russell D. Salter and Xin Dong University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania, USA

We must eat to live and live to eat. Henry Fielding

INTRODUCTION

type that allows for efficient T-cell stimulation. DC maturation is regulated by uptake and processing of antigens, and many forms of antigen can directly induce DC maturation or contain immunostimulatory compounds such as lipopolysaccharides (LPS). Following uptake by DCs, processed antigens must bind to MHC proteins to provide a ligand which, together with costimulatory molecules, can trigger T-cell activation. How DCs take up, process and present antigens via class I and class II MHC proteins will be discussed below (Figure 11.1).

Dendritic cells (DCs) are highly potent antigenpresenting cells (APCs) that are required to initiate most primary immune responses. In the periphery, processing DCs have the ability to capture antigen through a variety of means, and then mature into presenting DCs with a pheno-

MACROPINOCYTOSIS Macropinocytosis is an actin-driven endocytic process in which plasma membrane ruffles fuse to form vesicles that nonselectively include solutes and antigens present in the fluid phase (Sallusto et al., 1995; Watts, 1997). Vesicles formed range in diameter from 0.5 to 2 µm, which is in contrast to micropinocytosis, in

FIGURE 11.1 Multiple mechanisms for antigen uptake by dendritic cells. Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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which clathrin-coated vesicles of approximately 0.15 µm in size are formed (Watts and Marsh, 1992; Robinson et al., 1996). This process allows cells to continually take up large amounts of extracellular fluid containing soluble antigens. Immature DCs are particularly active at macropinocytosis compared with other cells, accumulating nearly an entire cell volume over a 1-hour period (Sallusto et al., 1995; Norbury et al., 1995). Exogenous antigens captured by DCs via macropinocytosis are directed in endocytic vesicles to specialized antigen-processing compartments called MIIC, which contain MHC class II proteins and other molecules required for peptide loading. In contrast, macrophages ingest about 25% of their own volumes in fluid each hour. However, internalized solutes are delivered primarily to lysosomes for degradation, rather than loading into class II molecules (Sallusto et al., 1995; Watts, 1997). The reason for this difference between DCs and macrophages in vesicular trafficking is unknown. Aquaporins, a family of water channel proteins, have recently been shown to play an essential role in control of fluid phase uptake in DCs. Specifically, aquaporins 3 and 7 found on the cell surface of DCs participate in transporting and concentrating soluble antigens during macropinocytosis (de Baey and Lanzavecchia, 2000). To measure macropinocytosis, small molecules which can be taken up without interaction with cell surface receptors are typically used as markers. Uptake of lucifer yellow or horseradish peroxidase is widely used for quantitation of fluid phase pinocytosis (Racoosin and Swanson, 1992; Sallusto et al., 1995). Fluorescently-labeled dextran is also commonly used for this purpose, but it is also a ligand for the mannose receptor on human DCs (Sallusto et al., 1995). Internalization of these labeled markers occurs at 37C, and can be inhibited by cytochalasin D, which causes actin depolymerization, or amiloride, which blocks membrane ruffling (Norbury et al., 1995). Brefeldin A has no effect on macropinocytosis, which contrasts with its ability to inhibit clathrin-dependent receptor-mediated endocytosis (Racoosin and Swanson, 1992; Watts and Marsh, 1992).

Rho family GTPases Cdc42 and Rac1 play an important role in regulating macropinocytosis in DCs, and also in nonclathrin-dependent phagocytosis (West et al., 2000; Garrett et al., 2000). Clostridium difficile toxin B, which inactivates Cdc42, inhibits uptake of solutes by immature bone marrow-derived DCs following microinjection. Conversely, endocytically inactive mature DCs can be induced to take up solutes by stimulation with the GTPase activator SopE from Salmonella typhimurium (Garrett et al., 2000). However, inactivation of Cdc42 failed to affect uptake in DCs isolated from the spleen, and in addition, active Rac was unable to reactivate macropinocytosis in mature splenic DCs (West et al., 2000). Antigen uptake may thus be regulated by distinct mechanisms in different DC subsets.

PHAGOCYTOSIS Definition and types Phagocytosis is a process by which particles, microbes or fragments of dead cells are engulfed and internalized, usually by specific membrane receptors. In the immune system, phagocytosis provides an important means for uptake and processing of antigens for presentation to T cells, in addition to its nonimmune role as a clearance process (Robinson et al., 1996; Watts, 1997). Phagocytosis is actin-mediated and ATPdependent, and requires the action of a number of small GTP-binding proteins (Brown, 1995; Franc et al., 1998). Two types of phagocytosis have been identified in macrophages, with distinct morphologies and biological consequences (Caron and Hall, 1998). Type I phagocytosis is mediated by Fc receptors, requires Cdc42 and Rac, and results in a respiratory burst and an inflammatory response. Ruffling of the plasma membrane during engulfment of particles is characteristic of the type I process. Type II phagocytosis is mediated by complement receptors such as CR3, requires Rho, and does not induce a respiratory burst or inflammatory response. Particles are seen to ‘sink’ directly into

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the plasma membrane, and membrane ruffling is not observed in the type II process (Franc et al., 1999). Other receptors which have been suggested to mediate type II phagocytosis are CD14 (Devitt et al., 1998), integrin receptors such as αvβ3 and αvβ5 integrins (Albert et al., 1998; Finnemann and Rodriguez-Boulan, 1999), CD36 (Ryeom et al., 1996; Albert et al., 1998), scavenger receptors (Fukasawa et al., 1996; Platt et al., 1996), and a phosphatidylserine receptor (Fadok et al., 2000). Each of these receptors have been implicated in the uptake of apoptotic bodies by macrophages or DCs (Platt et al., 1998; Gregory, 2000), which is consistent with recent reports demonstrating that phagocytic clearance of apoptotic cells does not induce inflammation (Gallucci et al., 1999; Sauter et al., 2000; Salio et al., 2000). Class I-restricted T-cell responses to viral antigens can be induced by DCs following uptake of apoptotic debris from infected cells (Albert et al., 1998). This suggests that additional signals may be delivered by some sources of apoptotic cells that can trigger maturation of DCs and production of proinflammatory cytokines.

Phagocytosis of antigen-coated particles by DCs For the study by Caron and Hall (1998) described above, fixed red blood cells were coupled to IgG or coated with C3b to generate particles which were internalized by macrophages. Red blood cells were then directly counted in vesicles within phagocytes. Phagocytosis of fluorescent latex beads following uptake by macrophages and DCs can be measured either by microscopy or by flow cytometry (Ramachandra et al., 1998; Randolph et al., 1998). The latter technique is useful for analysing uptake of beads by large numbers of cells, but is more accurate if cells are separated over protein gradient layers to wash off beads that are loosely attached to the cell surface. By measuring bead uptake in the presence of cytochalasin D, internalized beads can be distinguished from those that are bound to the cell surface. In addition, iron oxide particles have been used to quantitate phagocytosis, and can

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be quantitated microscopically after uptake (Falo et al., 1995; R.D. Salter and X. Dong, unpublished data). Soluble antigens conjugated to iron oxide particles are presented very efficiently by DCs to both class I- and class II-restricted T cells (Falo et al., 1995; Shen et al., 1997). Particulate antigens are presented to T cells approximately 100-fold more efficiently than the same antigen given in soluble form to DCs (Shen et al., 1997). Immature and mature DCs can each internalize these particles and process and present antigens to T cells. Both types of DC are also capable of internalizing latex beads, suggesting that phagocytic mechanisms continue to operate following DC maturation (R.D. Salter and X. Dong, unpublished data). This suggests that uptake of these particles can occur independently of Cdc42, which is inactivated in mature DCs.

Phagocytic receptors on DCs Both mouse and human DCs express Fcγ receptors (Schmitt et al., 1990a; de la Salle et al., 1997). Mouse bone marrow-derived DCs express low to moderate affinity FcγRII or FcgγRIII receptors (Regnault et al., 1999; Machy et al., 2000). Human DCs isolated directly from blood express FcγRI (CD64) (Fanger et al., 1997), a high-affinity IgG receptor, while following in vitro culture, monocyte-derived DCs downregulate CD64 and begin to express FcγRII (CD32) (Sallusto et al., 1994; Dong et al., 1999), a low- to moderateaffinity IgG receptor. CD32 is also expressed on Langerhans cells (Schmitt et al., 1990b; Esposito-Farese et al., 1995). Both receptors are involved in uptake of antibody-coated bacteria (Aderem and Underhill, 1999). Although this has been directly demonstrated for other cell types, antibody-mediated phagocytosis has not been directly shown for DCs. Complement receptors can stimulate phagocytosis of C3bi-opsonized bacteria, either alone or together with Fc receptors (Carroll, 1998). CR1 and CR3 are expressed on subsets of DCs, notably on follicular dendritic cells (Reynes et al., 1985; Yoshida et al., 1993), where these receptors can act together with Fc receptors to facilitate antigen uptake. It is less

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clear whether complement receptors facilitate antigen uptake by other types of DCs. Uptake of particulate antigens may require involvement of several different receptors and be subject to complex regulation.

RECEPTOR-MEDIATED ENDOCYTOSIS Soluble antigens can be captured by a wide variety of specific receptors on the surface of DCs, and internalized for processing and presentation by class I and class II molecules. Antigens bind to receptors and cluster in coated pits, and are then internalized in clathrin-coated vesicles, which are about 0.1 µm in diameter, in an ATP-dependent process. This mechanism of uptake is distinct from phagocytosis or macropinocytosis mentioned earlier, in that it does not require actin filaments, but does depend on certain Ras proteins (AlvarezDominguez and Stahl, 1998; Cox et al., 2000). Endocytic receptors on DCs can be classified broadly into two distinct types based on where they localize intracellularly. One type, exemplified by the macrophage mannose receptor (or mannose receptor) is similar to transferrin receptor in delivering bound ligands to early endosomal compartments and then recycling back to the plasma membrane (Taylor, 1997; Stahl and Ezekowitz, 1998). The second type, of which Fc receptors are examples, do not recycle, but instead are degraded together with bound ligands in lysosomes in most cell types. In DCs, however, antigens internalized by Fc receptors may be partially degraded and exported from the endocytic vesicle to avoid complete destruction of potential epitopes (Rodriguez et al., 1999). Receptor-mediated endocytosis is a very efficient way to selectively target antigens to the class II processing pathway. Both Fc receptors and mannose receptor can enhance the efficiency of presentation of soluble antigens by 100–1000-fold (Engering et al., 1997; Tan et al., 1997, Cella et al., 1999). Presentation is most efficient when immature DCs are allowed to internalize antigens, and are then induced to mature

with proinflammatory stimuli such as LPS or TNFα (Sallusto et al., 1995). Upon maturation, many endocytic receptors on DCs are downmodulated. What regulates this process is unclear. It is likely to be controlled by small GTPases of the Rab family, which direct endocytic receptors to different compartments (Alvarez-Dominguez and Stahl, 1998; Cox et al., 2000). Since receptor-mediated endocytosis is not actin-dependent, its regulation in DCs may differ from that of macropinocytosis, although the two processes are both downregulated following DC maturation (Sallusto et al., 1995). It should also be mentioned that some endocytic receptors, such as Fc receptors, can mediate uptake of opsonized particles as well as nonparticulate immune complexes. How the same receptor functions in these mechanistically distinct pathways of uptake is unresolved. Different types of endocytic receptors will be discussed individually below. The involvement of each receptor has been determined by blocking uptake of ligands with competitive inhibitors, such as receptor-specific antibodies. In addition, brefeldin A can be used to block receptor-mediated endocytosis and to distinguish it from other types of uptake.

Fc receptors Expression of different Fcγ receptors on DCs was described in the previous section on phagocytosis. Antigens in immune complexes taken up by these Fc receptors can be presented by class II MHC much more efficiently than in soluble form (Lanzavecchia, 1996). In addition, CD32 has been shown to mediate uptake of antigens for presentation to class I molecules in DCs (Regnault et al., 1999). Interestingly, the FcRassociated γ chain is required both for class I presentation and for triggering maturation of mouse DCs by immune complexes. In addition, uptake of antigens by distinct CD32 isoforms may lead to differences in epitope processing, as was shown in B cells (Amigorena et al., 1998). DCs also express high-affinity receptors for IgE (FcεRI) which can induce immune response to allergens. Langerhans cells, dermal DCs and

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monocyte-derived DCs have each been shown to express this receptor (Rajakulasingam et al., 1997; Maurer et al., 1998; Shibaki, 1998; Kahlert et al., 2000). Whether the Fc receptor-associated γ chain is required for function of the receptor in DCs is unclear.

C-type lectin receptors A broad family of structurally related lectins require calcium for their binding, and thus are called C-type lectins. A number of these molecules are important for immune function. On DCs, these include endocytic receptors which bind carbohydrate antigens, and internalize them for presentation via class II or CD1 pathways. The best-studied such receptor is the macrophage mannose receptor, which is expressed on both macrophages and DCs (Stahl and Ezekowitz, 1998). In addition, a related protein called DEC-205 is expressed on DCs and several other cell types (Jiang et al., 1995). Other C-type lectin receptors which may play a role in endocytosis of antigens have been identified, but their expression and function on DCs have not been confirmed.

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fluorescein labeled for measuring uptake by cells (Sallusto et al., 1995). Chemically synthesized compounds such as polyacrylamide coupled to fucose or mannose can also be used to analyze receptor binding specificity, as shown in Figure 11.2. An example of a ligand for mannose receptor which is relevant to immune responses to pathogens is lipoarabinomannan (LAM), which is present in mycobacteria (Prigozy et al., 1997). Binding of LAM to mannose receptor on DCs triggers uptake of the

Mannose receptor Mannose receptor is expressed on immature human monocyte-derived DCs, but not on Langerhans cells (Mommaas et al., 1999). In mice, no DC subsets have been identified which express mannose receptor, and its expression appears limited to macrophages and some endothelial cells (Linehan et al., 1999). Studies using macrophages or purified protein have shown that mannose receptor binds to ligands containing mannose, fucose, N-acetylglucosamine, and less well, glucose (Lennartz et al., 1987). In vivo, it is likely that relevant ligands for mannose receptor are glycoproteins or glycolipids with multiple sugar moieties that are considerably more complex than the ligands usually used to study mannose receptor specificity, typically proteins conjugated to single types of sugars. One of the most widely used of these nonphysiologic ligands is mannosylated BSA, which is typically

FIGURE 11.2 Uptake of fluorescent ligands by human monocyte-derived DCs. CD14-positive adherent cells from peripheral blood were cultured for 5 days in AIM-V medium with GM-CSF and IL-4 to generate DCs with an immature phenotype, as assessed by surface marker expression (data not shown). DCs were washed and resuspended in medium containing 20 µg/mL of FITC-conjugated ligands indicated on the x-axis for 1 hour at 37C. After washing, cells were fixed and analyzed by flow cytometry. Uptake of BSA was enhanced by mannosylation as previously demonstrated (Sallusto et al., 1995). Polyacrylamide (PAA) coupled to the indicated monosaccharides were also analyzed, and show a pattern of uptake which is consistent with mannose receptor-mediated binding (fucose, mannoseN-acetylglucosaminegalactose). No cell-associated fluorescence was seen when cells were incubated at 4C, indicating that ligands were internalized by cells. This was confirmed by microscopy. PAA ligands were obtained from Dr N. Bovin (Russian Academy of Sciences, Moscow) and are available commercially from Glycotech, Rockville, Maryland, USA or Syntesome, Munich, Germany.

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bacterium, and also may direct the antigen to endocytic compartments where LAM can bind to CD1b molecules for presentation to specific T cells. Cell surface molecules on other pathogens, including gram-positive and gram-negative bacteria, yeast and amastigotes of Trypanosoma cruzi, can also interact with mannose receptor, which presumably facilitates pathogen uptake (Kahn et al., 1995; Ezekowitz et al., 1991). It has also been shown recently that agalactosyl IgG, which is produced during some disease states, can bind to mannose receptor and be internalized efficiently by DCs (Dong et al., 1999). This may allow for uptake and presentation of antigens which are bound to agalactosyl IgG antibodies. The intracellular trafficking of mannose receptor allows for highly efficient uptake of antigens. After ligand binding, the complex is rapidly internalized into early endosomes, where ligands dissociate and the receptor is quickly recycled to the cell surface. Over a 1-hour period, fewer than 106 mannose receptors on DCs are capable of internalizing up to 2  107 molecules of ligand, suggesting that recycling is very rapid (Sallusto et al., 1995). It is not clear whether antigens taken up in this way are directed to antigenprocessing compartments such as the MIIC after dissociation from mannose receptor. Following maturation of human monocytederived DCs induced by inflammatory stimuli, endocytic uptake of ligands by mannose receptor stops, although the protein can still be detected at the cell surface for several days with antibodies (Dong et al., 1999). Receptor activity, thus, appears to be regulated independently of surface expression, at least transiently. DEC-205 Another member of the C-type lectin family, DEC-205 is expressed on both mouse and human DCs (Jiang et al., 1995; Kato et al., 1998; Guo et al., 2000). The binding specificity of this receptor is unknown, although its ability to function in antigen presentation was established by coupling polyclonal DEC-205-specific antibodies to a peptide epitope, which then

facilitated uptake and presentation of the epitope by mouse DCs to T cells (Jiang et al., 1995). Expression of this receptor at the surface of DCs is downregulated during maturation, but not as strikingly as seen with mannose receptor. It will clearly be important to define the binding specificity of the receptor to determine how it may function in antigen uptake by DCs.

Scavenger receptors Scavenger receptors are a family of proteins which can bind chemically modified lipoproteins, including oxidized or acetylated lowdensity lipoproteins, denatured proteins, chemically modified proteins such as maleylated BSA, and unopsonized gram-positive bacteria (Rigotti et al., 1997; Bansal et al., 1999). There are three different groups of scavenger receptors: A, B, C. Of these, members of the A and B groups appear to be potentially important for antigen uptake by DCs. In human monocytederived DCs, Cla-1, a B group receptor, is expressed upon differentiation, and is downregulated after LPS treatment (Buechler et al., 1999). No direct role in antigen presentation for Cla-1 has been demonstrated. Another member of the B group, CD36, has been shown together with αvβ5 integrin to facilitate uptake of apoptotic bodies by immature human DCs (Albert et al., 1998). Scavenger receptors can take up maleylated antigens for presentation by class I molecules (Bansal et al., 1999). Antigen presentation by macrophages and B cells, but not DCs, were addressed in this latter study.

Induction of immunity by stress response-induced proteins It was shown nearly a decade ago that the stressinduced protein gp96 isolated from tumor cell lines was able to induce CD8 T-cell responses against the tumor during in vivo or in vitro priming (Srivastava et al., 1994). This initially puzzling observation has now been shown to be due to transfer of peptides bound to gp96 molecules from the tumor into an APC, where the peptides can enter the class I pathway. Precisely how

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this form of antigen transfer occurs is unknown, but recently progress has been made in defining cell surface receptors for gp96, and also for hsp70 and hsp90, which allow for their internalization by APCs such as DCs and macrophages (Binder et al., 2000; Singh-Jasuja et al., 2000a). Receptors for stress proteins have not yet been directly identified, but binding of gp96 and hsp70 to APCs is saturable and self-competitive, but not cross-competitive (Binder et al., 2000; Singh-Jasuja et al., 2000a). The latter finding suggests that gp96 and hsp70 interact with distinct receptors. In DCs, following binding to the plasma membrane, both proteins localize to coated pits, and are rapidly internalized into an endocytic compartment containing class I and II MHC (Singh-Jasuja et al., 2000a). It is not known if class I loading with peptides occurs here, or if epitopes must be transferred to the ER either directly or via the cytosol. Uptake of gp96 induces DC maturation and downregulation of its receptor, suggesting that it is regulated similarly to mannose receptor, scavenger receptor, and other antigen uptake processes (SinghJasuja et al., 2000b). In addition to uptake of soluble molecules such as gp96 and hsp70, DCs can also internalize exosomes, in which hsc73 is concentrated (Thery et al., 1999). These observations demonstrate that DCs have specialized means for internalizing and processing peptidecontaining stress proteins, which presumably broaden the range of exogenous antigens which they can present.

SYNTHESIS AND TRAFFICKING OF CLASS II MHC PROTEINS Class II MHC biosynthesis Biosynthesis of class II proteins has been studied extensively in human B lymphoblastoid cell lines and in mouse B-cell lymphomas, using metabolic radiolabeling of class II proteins in these cells followed by immunoprecipitation with specific antibodies. During their intracellular maturation, class II dimers associate

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with the invariant chain, DM, and in some cells DO, both of which facililate trafficking or loading of peptide antigens into class II dimers (Denzin et al., 1997). The invariant chain associates with class II subunits in the ER, and then chaperones the class II complex to the MIIC compartment, where peptide loading occurs (Newcomb and Cresswell, 1993). In the MIIC, the invariant chain is proteolytically cleaved by cathepsins B, D and S, resulting in class II complexes containing a fragment of the invariant chain called CLIP (Riberdy et al., 1992). DM, which localizes to this site, then interacts with the class II–CLIP complex and facilitates removal of CLIP, generating class II dimers which are receptive to other peptides, including potential antigens (Amigorena et al., 1995). Following stabilization by binding an appropriate peptide ligand, class II trimers are transported to the plasma membrane. In this way, newly synthesized class II molecules bind peptides derived from endocytosed proteins in all APCs expressing class II molecules.

Recycling pathways In addition, however, class II molecules can recycle from the plasma membrane and pass through the MIIC compartment, providing a further opportunity for antigen loading to occur. First identified in B cell lines, this pathway is very robust in immature DCs, and is weak or absent in mature DCs (Cella et al., 1997) By passing through the MIIC multiple times, class II molecules are frequently exposed to the contents of endocytic compartments of an APC, and efficiently interact with peptides from newly internalized pathogens. Following maturation induced by an inflammatory stimulus such as LPS, recycling stops rather abruptly, and class II molecules are transported to the cell surface (Cella et al., 1997). This final exocytic step is mediated by specialized CIIV vesicles containing class II proteins and co-stimulatory molecules, which remain in clusters at the plasma membrane for an extended period (Turley et al., 2000). Nonrecycling class II molecules at the surface of mature DCs have a half-life of about

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100 hours, which appears sufficient to maintain epitope expression while the DC moves from the periphery to secondary lymphoid organs. In contrast, recycling class II molecules have a relatively short half-life of about 16 hours, possibly a consequence of their frequent exposure to a proteolytic environment (Cella et al., 1997). The mechanism by which class II recycling stops following maturation is not clear, but is temporally linked to the downregulation of endocytic capacity by DCs, which was discussed in a previous section.

Intracellular proteases for class II antigen processing Upon arriving in the MIIC, newly exported class II–invariant chain complexes are exposed to proteases which degrade the invariant chain to CLIP, which remains bound to class II dimers until removed by DM. This cleavage occurs in step-wise fashion, and is mediated by several different cathepsins, with cathepsin S being required for generation of CLIP (Driessen et al., 1999). The importance of this enzyme for class II trafficking in DCs was demonstrated using cathepsin S knockout mice. Class II molecules accumulate in a late endocytic compartment and their movement to the cell surface is impaired in these mice. Cathepsin S activity is regulated during DC maturation (Pierre and Mellman, 1998). In immature DCs, much of the total class II is directed towards lysosomes, due to inefficient removal of the the lysosomal sorting signal of the invariant chain. This is a result of low levels of cathepsin S activity in these cells, which is caused by synthesis of an endogenous cathepsin S inhibitor called cystatin C. Following DC maturation, cystatin C expression decreases, which elevates cathepsin S activity, allowing efficient cleavage of the invariant chain bound to class II and promoting transport to the cell surface. These studies demonstrate that cathepsin S is crucial for proper trafficking of class II in DCs. It remains unclear whether cathepsin S or other proteases are required for generating antigenic peptides which bind to class II.

Cell surface loading of antigens into class II In both of the previously mentioned pathways, class II molecules bind antigens inside DCs in specialized endocytic compartments. More recently, it has been shown that class II molecules on the surface of DCs can also bind antigens by a process which requires DM to facilitate peptide exchange or editing (Santambrogio et al., 1999a; Arndt et al., 2000). Interestingly, both DM and ‘empty’ class II dimers were shown to be present on the surface of immature DCs, but not on mature DCs, suggesting that cell surface peptide binding would be limited to peripheral sites in the body during inflammation, and would not take place during DC migration to secondary lymphoid organs (Santambrogio et al., 1999b). It was further shown that proteases secreted by immature DCs can degrade intact proteins into fragments that can bind class II molecules and stimulate T cells. Since secreted proteases might not generate the same repertoire of peptides as in the MIIC, there is a possibility that neoepitopes could potentially be generated by this pathway, triggering autoimmunity. However, DCs express high levels of CD13/aminopeptidase N on their plasma membrane (Dong et al., 2000), which has been shown in a murine B cell model to be capable of processing longer peptides bound to class II molecules into epitopes (Larsen et al., 1996). CD13 and unidentified cell surface carboxypeptidases on DCs could potentially trim longer peptides generated by extracellular proteases into optimal epitopes. If binding of these longer fragments depends upon the presence of the appropriate class II binding motif, then perhaps the final peptide products found after trimming would not be different from those generated within the cell.

SYNTHESIS AND TRAFFICKING OF CLASS I MHC PROTEINS As with class II molecules, B-cell lines and genetic mutants bearing defects in antigen-processing

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machinery have contributed greatly to the current understanding of the function of classical class I (class Ia) proteins. Class Ia (hereafter referred to as class I) proteins consist of a polymorphic heavy chain, encoded by HLA-A, B, or C loci in humans and by H-2K, D, or L loci in mice, and an invariant subunit, β2-microglobulin (β2m). Heavy chains and β2m assemble in the ER following association of the heavy chain with calnexin, an ER resident protein which aids the folding of many newly synthesized glycoproteins (Degen and Williams, 1991). A second chaperone, calreticulin, then binds to the class I dimer, and guides association with tapasin, a protein whose function is to facilitate binding of the class I complex to TAP, the transporter of antigenic peptides (Ortmann et al., 1997). Once bound to TAP, newly synthesized class I proteins are receptive to peptides translocated from the cytosol across the ER membrane. After stably binding a peptide in its cleft, the class I molecule dissociates from the peptide-loading complex and is transported through the Golgi and to the plasma membrane. In contrast to class II MHC, there appears to be no specialized adaptation in DCs of the class I biosynthetic pathway. In DCs, calnexin, calreticulin and tapasin all participate in class I biosynthesis, and most peptides which bind to class I molecules are provided by the proteasome (R.D. Salter and X. Dong, unpublished data). Class I molecules do not recycle from the plasma membrane, and have similar lifespans in mature and immature DCs, with a half-life of roughly 16 hours (Cella et al., 1997). This is comparable to other cells which have been examined, and suggests that in contrast to class II molecules, class I molecules in both immature and mature DCs need to be synthesized continually to expose epitopes to T cells. This would occur readily for epitopes derived from endogenous proteins, such as those made during viral infections, which would be synthesized, then degraded into peptides which could bind to newly synthesized class I proteins. However, it is unclear whether exogenous antigens taken up by DCs, for example in apoptotic bodies, could persist in the cytoplasm and provide a contin-

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ued source of antigen for loading into new class I molecules following DC maturation. An endocytic compartment exists in a mouse DC cell line called FSDC in which intact antigen is stored for prolonged periods and released into the cytosol for presentation by class I molecules (Lutz et al., 1997). Such a compartment has not been identified in primary DC cultures however.

Cross-priming Presentation of exogenous antigens by class I molecules on DCs, which forms the basis for cross-priming, results from transfer of the contents of endocytic compartments to the cytosol (Rodriguez et al., 1999). How this antigen transfer occurs is not well understood, but it appears to be selective for the size of the antigen. It is also not clear why certain receptors are capable of facilitating cytosolic transfer more readily than others, or why DCs have this capacity while other cells do not. Differences in localization of distinct receptors within the endocytic pathway, or in other aspects of receptor function may be important. Additional work is needed to resolve the precise details by which epitopes from exogenous antigens are provided to class I molecules in DCs.

Proteolysis of antigens for class I presentation As first demonstrated for other cell types, proteasomes generate most of the peptides which bind to class I MHC in DCs. During DC maturation, proteasome subunits LMP-2, LMP-7 and MECL-1 are upregulated, resulting in the production of almost exclusively immunoproteasomes (Macagno et al., 1999). This may be beneficial for presentation of certain epitopes, as it has been demonstrated that the cleavage specificity of immunoproteasomes can favor production of peptides with appropriate binding motifs for a number of class I molecules. However, it has also been shown that immunoproteasomes are incapable of producing an epitope from a tumor antigen which can be generated by the standard proteasome in DCs (Morel et al., 2000).

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It may be critical to regulate the levels of each type of proteasome to present certain antigens effectively.

TRAFFICKING AND ANTIGEN PRESENTATION BY CD1 MOLECULES CD1 molecules expressed on DCs are important for presentation of nonpeptide antigens to T cells (Kawano et al., 1997). CD1 heavy chains are similar to class Ia heavy chains in overall structure, and assemble with β2m in the ER, but have structurally distinct binding clefts in the α1 and α2 domains which allow them to interact with a broad range of non-peptide antigens containing a lipid moiety. Humans have five distinct CD1 proteins (a–e) while only two forms (CD1d1 and d2) have been identified in mice. CD1a, CD1b and CD1c (or group 1 CD1) molecules have been shown to provide a protective function against infection by intracellular pathogens, such as mycobacteria (Stenger et al., 1997). Following phagocytosis mediated in part by mannose receptor interaction with glycolipid components of mycobacteria, infected cells can be lysed by group 1 CD1-restricted T cells (Prigozy et al., 1997). CD1d (or group 2 CD1) molecules have been shown to stimulate NKT cells to regulate production of IgG directed against parasites (Schofield et al., 1999). Distinct patterns of expression of CD1 are found on different subsets of DCs. CD1a is present on Langerhans cells, dendritic cell subsets, and binds lipid and glycolipid antigens in early recycling endosomes (Sugita et al., 1999). CD1b, which is found on a variety of cell types including DC, binds a different set of lipid antigens in late endocytic compartments molecules (Sugita et al., 1999). CD1c has an expression pattern similar to that of CD1b, but is widely distributed throughout the endocytic pathway, allowing for a broad sampling of lipids from different regions within the cell (Sugita et al., 2000). CD1d molecules are expressed on a number of cell types, and bind exogenously provided α-galactosylceramide, a sphingolipid derivative isolated from a marine

sponge, as well as a variety of phospholipids found within cells (Brossay et al., 1998; Gumperz et al., 2000). Whether lipid antigens must be processed before binding to CD1 molecules is unknown. Inhibitors of endosomal acidification can block presentation of lipid antigens by CD1b, but not CD1a molecules, which suggests that antigens might require processing before binding to some CD1 proteins (Sugita et al., 1999). Alternatively, the lower pH in the endocytic pathway may induce a conformational change in the CD1b-binding cleft which increases its receptivity to antigens. If some lipid antigens are processed before binding to CD1 molecules, it will be interesting to determine if DC maturation affects this process, as it does for the proteases involved in class II MHC antigen presentation.

DIFFERENT SUBSETS OF DCs HAVE DISTINCT CAPACITIES FOR ANTIGEN PROCESSING DCs found throughout the body exhibit a wide range of phenotypes and functional capabilities. The most intensively studied DCs in terms of antigen processing capacity are bone marrowderived CD34 DCs from the mouse, and human monocyte-derived DCs expanded in vitro with GM-CSF and IL-4. Although the validity of the latter cell type as a model for DCs was initially questioned, more recently monocytes were shown to acquire the properties of DCs both in vitro without cytokine stimulation and in vivo (Randolph et al., 1998, 1999). Both of these types of DCs express myeloid lineage markers, and show marked differences in function with, for example, lymphoid DCs from the spleen. Differences between DCs in vivo and those cultured for use in antigen-presentation assays are worth noting. Langerhans cells in the skin have poor endocytic capacity, and lack the ability to internalize antigen via mannose receptor, which is in contrast to human monocytederived DCs (Mommaas et al., 1999). A novel Ctype lectin, langerin, is expressed on Langerhans

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cells however, and induces formation of Birbeck granules (Valladeau et al., 1999, 2000). It is possible that langerin functions in antigen capture similarly to mannose receptor, but this is not proven. A number of differences exist between lymphoid DC subpopulations isolated from the spleen and myeloid DCs, as summarized elsewhere in this volume. Of relevance to studies of antigen processing, splenic CD8 DC populations from mice are considered to be poorly phagocytic, but with high capacity to take up soluble antigens via pinocytosis (Pulendran et al., 1997). Precisely how these different DC subsets vary in terms of the internalization and antigen-processing machinery, and what effect this might have upon their ability to function as APCs, will be the subject of future investigations.

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12 Development and testing of dendritic cell lines Akira Takashima and Hiroyuki Matsue University of Texas Southwestern Medical Center, Dallas, Texas, USA

Jan Klein said, when admiring a painting by Gaugin and then comparing a work of art to a work of a scientist, that we all contribute by placing chips in the vast mosaic of science. Some of us may be fortunate to place a piece that fits well and is prominent, while others may place smaller pieces and, unfortunately, some may not fit well and may even fall off. Paul M. Allen, Kenneth M. Murphy, Robert D. Schreiber and Emil R. Unanue (1999). Immunity 11, 649–651

INTRODUCTION

thymus, lymph nodes, tonsils, epidermis and other epithelial tissues, dermis and other connective tissues, peripheral blood, and afferent lymphatics (Hart, 1997). Substantial progress has been made over the last several years in the development of experimental protocols for the isolation of relatively large numbers of DCs directly from tissues and from short-term cultures (Caux et al., 1992; Inaba et al., 1992; Strunk et al., 1994; Sallusto et al., 1995; Maraskovsky et al., 1996). Moreover, several stable DC lines and clones have been established from various tissues of mice, thus providing useful experimental tools for studying the biology of DCs at molecular and biochemical levels and for the establishment of new DC-based immunotherapies. In this chapter, we will describe immunological features of those long-term DC lines and their utilities in biomedical research.

Immunologically naïve T cells can be activated most efficiently or even exclusively by special subsets of antigen-presenting cells, termed dendritic cells (DCs) (Hart, 1997; Banchereau and Steinman, 1998; Maurer and Stingl, 1999). DCs are equipped with all the phenotypic and functional features that are required for achieving this task; these features include: (1) antigen uptake and processing in MHC class I and class II cascades, (2) surface expression of the MHC–peptide complexes, (3) expression of adhesion molecules and co-stimulatory molecules, (4) migration of the T-cell area in draining lymph nodes, (5) secretion of selected chemokines and cytokines, and (6) delivery of full activation signals to T cells. Members of the DC family have been identified in virtually all the tissues and organs tested such as spleen, Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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ESTABLISHMENT OF STABLE DENDRITIC CELL LINES Immortalization of DCs by oncogenes Pagia and coworkers established the first stable DC line, termed CB1, by introducing an immortalizing oncogene env AKR-myc MH2 into splenic DCs isolated from newborn DBA2 mice (Paglia et al., 1993). The same strategy was then used to establish other immortalized DC lines from Balb/c spleen (D2SC/1 line) (Granucci et al., 1994) and from day 17–19 fetal skin of (DBA/2  C57BL/6)F1 mice (FSDC line) (Girolomin et al., 1995). Volkmann et al. (1996) established a ‘conditionally’ immortalized DC line from bone marrow of mice that expressed a thermolabile mutant transgene encoding the SV40 large T antigen. The resulting line, termed tsDC, expressed the above oncogene and thus continued to proliferate only when maintained at the permissive temperature range of 33–37C. When exposed to a higher temperature (39C), however, this line stopped dividing and began to differentiate into ‘mature’ DCs (Volkmann et al., 1996). Shen et al. (1997) developed a series of DC clones (DC1.2, DC2.4, DC4.1) by introducing the GM-CSF gene into the C57BL/6 bone marrow cells, followed by infection with a retrovirus encoding myc and raf oncogenes. Other immortalized DC lines include: the JAWS II line from C57BL/6 bone marrow (Brossart, 1997; Butz and Bevan, 1998), the AG series from skin and brain of the AG129Sv/Ev mice that lack both type I and type II interferon (IFN) receptors (Nunez et al., 1997, 1999), and the DDC/B210K line from CD.1 bone marrow and the DDC/B301 line from Balb/c bone marrow (Rowden et al., 1995). Of these oncogene-immortalized DC lines, the ones established in Ricciardi-Castognoli’s laboratory (CB1, D2SC/1 and FSDC) have been most well characterized (Table 12.1).

Generation of leukemic DC cultures Rasko et al. (1997) established a tumorigenic proDC line from mice with myeloid leukemia. Briefly, these investigators first infected the GM-CSF

transgenic mice with Molony murine leukemia virus and then developed a multipotential leukemic cell line, termed DMG36, from spleens of these leukemia animals. This line, which differentiated under certain conditions into macrophages, neutrophils and eosinophils, began to exhibit many characteristic features of DCs upon stimulation with TNFα and TNFγ (Rasko et al., 1997). Similarly, Friend virusinduced, murine erythroleukemic cell line FBL3 differentiated into DCs shortly after stimulation with GM-CSF (Cao et al., 1998). These observations in mice suggest that one may be able to generate human DC lines from leukemic cells. In fact, DC-like cells were generated from a patient with acute myelogenous leukemia (AML) by culturing peripheral blood mononuclear cells (PBMCs) in the presence of GMCSF, TNFα, stem cell factor (SCF) and IL-6 (Santiago-Schwarz et al., 1994). Likewise, PBMCs from patients with chronic myelogenous leukemia (CML) developed morphological, phenotypic and functional characteristics of DCs when cultured in the presence of GM-CSF, IL-4 and TNFα (Choudhury et al., 1997). More recently, Choudhury et al. (1999) reported that PBMCs from 18 out of the 19 patients with AML differentiated into DCs during short-term culture in the presence of GM-CSF plus IL-4 in combination with either TNFα or CD40L. In an independent study, differentiation of acute leukemia blasts into DCs was observed in 12/15 cases (Robinson et al., 1998). Finally, DC lines with Langerhans cell-like phenotype were established from a patient with histiocytic lymphoma (Nunez et al., 1998). Although those leukemia-derived DCs remain viable and functional only for relatively short periods (up to a few weeks), these cells expressing not only features of DCs, but also leukemia-specific antigens may have significant clinical potential, especially as a vaccine to initiate protective immunity against leukemia cells.

Establishment of growth factordependent DC lines In 1994, Elbe et al. established three growth factor-dependent, long-term DC lines from fetal

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Establishment of DC lines D2SC/1

FSDC

80/1

XS52

XS106

D1

Origin Mouse strain Donor age Donor tissue

DBA/2 newborn spleen

Balb/c not described spleen

(DBA/2C57BL/6) F1 fetus skin

C3H fetus skin

Balb/c newborn epidermis

A/J newborn epidermis

C57BL/6 adult spleen

Immortalization Oncogene

() env AKR-myc MH2

() env AKR-myc MH2

() env AKR-myc MH2

()

()

()

()

Culture condition Medium FCS Growth factor

RPMI-1640 10% ()

Iscove’s MDM 5% ()

RPMI-1640 10% ()

RPMI-1640 10% IL-2 Con A

RPMI-1640 10% GM-CSF NS Sup

RPMI-1640 10% GM-CSF NS Sup

Iscove’s MDM 5% GM-CSF 3T3 Sup

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CB1

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mouse skin. The 80/1 and 86/2 lines were generated from C3H mice by repeated stimulation with IL-2 and Con A, whereas the 18 line was established from Balb/c mice in the presence of GMCSF. These lines required IL-2 plus Con A or GMCSF alone for their continuous growth, and they resembled Langerhans cells in fetal skin by the lack of surface MHC class II expression (Elbe et al., 1994). We generated stable DC lines from newborn Balb/c mouse skin by culturing epidermal cells in the presence of GM-CSF and culture supernatants collected from Pam 212 keratinocytes. Interestingly, all the cultures that contained colonies of growing DCs were contaminated with fibroblastic cells, and the DC colonies stopped dividing when the fibroblast contaminants were removed from the cultures, suggesting that those fibroblasts were secreting DC growth factor(s) (Xu et al., 1995a). In fact, we were able to restore the growth of DC colonies by the addition of supernatants collected from the independently expanded fibroblast cultures (termed NS lines) (Schuhmachers et al., 1995), and more than 20 stable DC lines and clones (XS series) have been established by using the combination of GM-CSF and the NS fibroblastconditioned medium (Xu et al., 1995a, 1995b). More recently, we established a fully mature line XS106 from the epidermis of newborn A/J mice by using the same growth factors. Unlike the original XS series, which were generated from TABLE 12.2

‘adherent’ epidermal cell populations, the XS106 line was established from a ‘nonadherent’ population (Timares et al., 1998). Winzler et al. (1997) established a splenic DC line (D1) from adult C57BL/6 mice by using the combination of GMCSF and the NIH 3T3 fibroblast-conditioned medium. Of these growth factor-dependent DC lines, the 80/1, XS52, XS106 and D1 lines are now being used by many investigators (Table 12.1).

PHENOTYPIC AND FUNCTIONAL PROPERTIES Surface phenotype As shown in Table 12.2, substantial differences are observed in the surface phenotypes of different DC lines (Paglia et al., 1993; Elbe et al., 1994; Granucci et al., 1994; Xu et al., 1995a; Lutz et al., 1995; Altenschmidt et al., 1996). For example, the expression level of surface MHC class II molecules was relatively high to modest in the CB1, D2SC/1, XS106 and D1 lines; minimal in the FSDC and XS52 lines; and totally undetectable in the 80/1 line. CD80 was expressed by the CB1, D2SC/1, XS106 and D1 lines, but undetectable on the FSDC, 80/1 and XS52 lines. With regard to DC markers, DEC-205 was detected only in the CB1 lines, whereas constitutive expression of CD11c was observed in all DC

Surface phenotypes of DC lines

MHC I MHC II CD80 CD86 CD40 CD11a CD11b CD11c CD18 CD54 CD24 CD32 CD44 DEC-205

CB1

D2SC/1

FSDC

80/1

XS52

XS106

D1

   NT NT    NT     

   NT NT NT NT  NT NT  NT  NT

    NT NT   NT     

    NT   NT      

             

      NT  NT  NT NT  NT

        NT    NT 

NT = not tested

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lines except for the XS52 and XS106 lines. Importantly, each DC line changes the surface phenotype dramatically depending upon the culture conditions. For instance, FSDC cells elevated the expression of MHC class II molecules and CD54 following IFNγ treatment (Lutz et al., 1996), and XS52 cells increased the expression of MHC class II and CD86 upon stimulation with lipopolysaccharide (LPS) or following extended culture with GM-CSF alone (Xu et al., 1995c; Takashima and Kitajima, 1998). Stimulation of D1 cells with LPS, TNFα or IL-1β upregulated the expression of MHC class II, CD86, CD40 and CD54 (Winzler et al., 1997). Thus, the observed heterogeneity in surface phenotype between the above DC lines most likely reflected the differences in the tissues of their derivation (e.g. skin versus spleen, and fetal versus adult animals), in cytokine compositions in the growth media and/or in their states of maturation.

Growth factor requirement The oncogene-immortalized cell lines CB1, D2SC/1 and FSDC continued to proliferate in conventional culture media (RPMI-1640 or Iscove’s MDM) with 5 or 10% FCS in the absence of added growth factors (Paglia et al., 1993; Granucci et al., 1994; Girolomoni et al., 1995). By contrast, the XS52, XS106 and D1 lines required GM-CSF and fibroblast-conditioned media for their maximal proliferation and long-term survival (Xu et al., 1995a; Winzler et al., 1997; Timares et al., 1998). Proliferative responsiveness of these DC lines to GM-CSF is consistent with the current notion that GM-CSF serves as a primary growth factor for DCs (Witmer-Pack et al., 1987; Hart, 1997; Banchereau and Steinman, 1998; Maurer and Stingl, 1999). With respect to the identity of DC growth factor(s) secreted by fibroblasts, we observed that: (a) NS fibroblasts as well as 3T3 fibroblasts expressed mRNA for CSF-1; (b) neutralizing antibodies against CSF-1 receptor (CD115) blocked the ability of the NS supernatant to promote XS52 cell growth; and (c) XS52 cells proliferated maximally in response to recombinant CSF-1 (Takashima et al., 1995). These observations suggest a paracrine mecha-

169

nism in which CSF-1 produced by fibroblasts supports the growth of DCs. In fact, surface expression of CD115 has been confirmed for both XS52 and D1 lines (Takashima et al., 1995; Kitajima et al., 1995a; Winzler et al., 1997), and relatively large numbers of DCs have been expanded from fetal mouse skin by using the combination of GM-CSF and CSF-1 (Jakob et al., 1997). Thus, CSF-1 appears to serve as a second growth factor for at least some DC populations. By testing 28 different cytokines individually and in combinations, we further characterized growth factor responsiveness of XS52 cells. IL-4 and IL-13 were both found to promote the growth of XS52 cells significantly, albeit to a lesser extent than that observed for GM-CSF and CSF-1. When tested in combination with GM-CSF and/or CSF1, IFNγ showed a potent ability to inhibit the proliferation of XS52 cells, with IL-6, IL-10 and TGFβ1 showing lesser degrees of inhibition (Yokota et al., 1996). Girolomoni et al. (1995) observed that the immortalized DC lines also responded to selected growth factors when tested in serum-free medium. GM-CSF, but not CSF-1, promoted the maximal proliferation of both FSDC and D2SC/1 lines, whereas IL-4 supported the growth of D2SC/1 cells, but not of FSDC cells. IFNγ inhibited spontaneous proliferation of both FSDC and D2SC/1 cells (Girolomoni et al., 1995). In summary, some of the DC lines maintain the growth factor dependence, thus providing a unique opportunity to study cytokinemediated growth regulation of DCs (Table 12.3).

Endocytotic potential and antigen processing Although the CB1 line showed very little zymosan phagocytosis compared with macrophages (Paglia et al., 1993), other DC lines exhibited potent endocytic capacities. For example, the FSDC line was found to be even more effective than macrophages in the pinocytosis of FITC-conjugated ovalbumin (OVA) and dextran. The pinocytotic potential of FSDC cells was enhanced further by GM-CSF and IL-4, whereas it was downregulated by IFNγ (Lutz et al., 1996). D2SC/1 cells ingested heat-killed E. coli into

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Growth-regulatory factors for DC lines D2SC/1

FSDC

80/1

XS52

XS106

D1

Growth factor

GM-CSF IL-4

GM-CSF

IL-2

GM-CSF CSF-1 IL-4 IL-13

GM-CSF CSF-1

GM-CSF

Growth-inhibitory factor

IFNγ

IFNγ

NT

IFNγ IL-6 IL-10 TGFβ1

NT

NT

NT = not tested

phagosomes (Riva et al., 1996). XS52 cells phagocytosed latex particles efficiently, and this activity was downregulated by IFNγ (Kitajima et al., 1996a). Likewise, D1 cells pinocytosed FITC-OVA and FITC-dextran with high efficiency (Winzler et al., 1997), and they also phagocytosed latex particles and even bacteria (Rescigno et al., 1998). Thus, many DC lines resemble immature DCs in their endocytotic potential. Working with FSDC cells, Lutz et al. (1997) have identified that the internalized antigens are stored in unique retention compartments distinct from endosomes or lysosomes. These compartments are mildly acidic and associated with lysosome-associated membrane protein 1 (LAMP-1), cathepsin D, and MHC class II molecules, suggesting their role in storing antigens without massive degradation (Lutz et al., 1997). Other studies showed that MHC class II molecules in D1 cells were mainly concentrated in the LAMP-1 lysosomal compartments, from which newly synthesized MHC class II αβ dimers were rapidly transported to the cell surface (Regnault et al., 1998). Therefore, these DC lines are capable of incorporating and processing exogenous antigens for MHC class II presentation. The same DC lines can also process the internalized antigens for the presentation on the MHC class I molecules. The D2SC/1 line as well as the MHC class II-negative DC line 18 incorporated viral particles and presented the viral antigens to MHC class I-restricted CTL clones efficiently (Schirmbeck et al., 1995; Bachmann et al., 1996). Working with D1 cells, Rodriguez et al. (1999) identified a pathway through which DCs

transport the internalized antigens to the cytosol for MHC class I-dependent presentation. Fluorescence-labeled protein antigens initially accumulated in the LAMP-1 endocytic vesicles, whereas they were subsequently translocated into the cytosol and processed by the proteosome in D1 cells. By contrast, the same antigens remained in the endosomal compartments in macrophages, indicating DC-specificity (Rodriguez et al., 1999). In summary, these DC lines maintain the characteristic properties of immature DCs to internalize, process and present exogenous protein antigens in either a MHC class I- or class II-restricted manner.

Cytokine and cytokine receptor profiles Long-term DC lines have been used by many investigators to study cytokine profiles expressed by DCs (Table 12.4). The CB1 line secreted IL-1β andTNFαfollowingLPSstimulation, albeit in relatively small amounts (Paglia et al., 1993). The D2SC/1 line produced higher amounts of IL-1β andTNFα after LPS treatment, and it also secreted TGFβ1 in response to stimulation with GM-CSF (Granucci et al., 1994). Both D2SC/1 and FSDC lines secreted IFNα and IFNβ when stimulated with virus and/or bacteria (Eloranta et al., 1997). The XS52 line constitutively expressed mRNAs for IL-1α, IL-1β, IL-7, TNFα, CSF-1, IFNα and macrophage inflammatory protein (MIP)-1α (Xu et al., 1995c). At the protein level, XS52 cells secreted relatively large amounts of IL-1β, IL-6, IL-12 (p40), TNFα and small, but detectable, amounts

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TABLE 12.4

Cytokine profiles of CD lines

Cytokine production

CB1

D2SC/1

FSDC

XS52

D1

IL-1β TNFα

IL-1β TNFα TGFβ1 IFNα IFNβ

IFNα IFNβ

IL-1β IL-6 IL-10 IL-12 (p40) IL-18 TNFα MIP-1γ

IL-1β IL-6 IL-12 (p70) TNFα

of IL-10, following LPS stimulation or upon antigen-specific interaction with the CD4 T cell line (Ariizumi et al., 1995; Kitajima et al., 1995b, 1996b; Torii et al., 1997a). In addition, XS52 cells constitutively secreted MIP-1γ in biologically relevant amounts, and the culture supernatant from XS52 cells promoted chemotactic migration of T cells in vitro (Mohamadzadeh et al., 1996a). More recently, Stoll et al. (1998) reported that XS52 cells secreted IL-18 (originally called IFNγ-inducing factor) and that XS52 culture supernatant significantly augmented the secretion of IFNγ by T cells. D1 cells secreted IL-1β, IL-10 and TNFα upon internalization of apoptotic bodies (Rovere et al., 1998, 1999), and TNFαpretreated D1 cells elab-orated the biologically active form (p70) of IL-12 upon antigen-specific interaction with T cells (Winzler et al., 1997). Taken all together, these results document the ability of DC lines to produce proinflammatory cytokines (IL-1β, IL-6 andTNFα),T cell-attracting chemokines (MIP-1α and MIP-1γ), and TH1/TH2regulatory cytokines (IL-10, IL-12 and IL-18). With respect to the cytokine receptor (R) profiles, the 80/1 line expressed IL-2R α chain (CD25) and IL-2R β chain (CD122), consistent with the fact that IL-2 promoted the growth of this DC line (Elbe et al., 1994). Although XS52 cells expressed neither CD25 nor CD122 at detectable levels, they expressed the IL-2R γ chain or the common cytokine receptor γ (CD132) (Xu et al., 1995a; Mohamadzadeh et al., 1996b). The two lines (XS52 and D1) that were established in the presence of fibroblastconditioned media (containing CSF-1) expressed the CSF-1R (CD115) (Winzler et al., 1997; Kitajima et al., 1995a). The findings that the

FSDC, XS52 and D1 lines responded functionally to IL-1β, IL-4, IL-10, TNFα, GM-CSF and/or IFNγ suggest that these DC lines must express relevant receptors for respective cytokines. Thus, some DC lines maintain the ability to secrete a wide variety of cytokines and to respond to exogenous cytokines in the microenvironment.

T cell-stimulatory capacity The CB1, FSDC, XS52, XS106 and D1 lines have been shown to be capable of presenting complex protein antigens to CD4 T cells in vitro (Paglia et al., 1993; Xu et al., 1995a; Lutz et al., 1996; Winzler et al., 1997; Mutini et al., 1999). Moreover, the FSDC and XS52 lines also presented chemical antigens (i.e. reactive haptens) to hapten-reactive T-cell clones efficiently (Girolomoni et al., 1995; Caceres-Dittmar et al., 1995; Lutz et al., 1996). In primary mixed leukocyte reactions (MLR), the CB1, 80/1, XS106 and D1 lines constitutively exhibited a potent ability to activate naïve T cells isolated from allogeneic animals, whereas the FSDC and XS52 lines exhibited this activity only after stimulation with selected cytokines (Paglia et al., 1993; Russell et al., 1993; Elbe et al., 1994; Xu et al., 1995a; Girolomoni et al., 1995; Winzler et al., 1997; Yamada and Katz, 1999). These in vitro observations indicate that some, if not all, DC lines maintain the ability to present various forms of antigens to immunologically naïve T cells, i.e. the hallmark of DCs as antigen-presenting cells. Many studies also documented the in vivo potential of DC lines to initiate antigen-specific immune responses. Following administration of the CB1 cells that had been pulsed in vitro with

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DNFB or FITC, the recipient animals developed significant ear swelling responses to the same hapten upon challenge (Paglia et al., 1993). This capacity to initiate contact hypersensitivity reactions has also been observed for the FSDC and 80/1 lines (Girolomoni et al., 1995; Kolesaric et al., 1997) and for the XS52 line but only after stimulation (Yamada and Katz, 1999). Administration of FSDC or 80/1 cells that had been pulsed with heat-inactivated Equine herpes virus-1 (EHV-1) induced protective immunity against subsequent challenge with live virus, as evidenced by the prevention of severe clinical symptoms (including death of infected animals) and by reduced viral load (Steinbach et al., 1998). Injection of the D2SC/1 cells that had been loaded with β-galactosidase (β-Gal) resulted in the priming of β-Gal-specific CD4 T cells (Paglia et al., 1996). Likewise, D2SC/1 cells pulsed with Borrelia burgdorferi antigens primed spirochete-reactive T cells in vivo (Altenschmidt et al., 1996). Injection of the MHC class II-negative DC line 80/1 line (derived from C3H mice) into the H-2-disparate mice induced allo-specific transplantation immunity, as evidenced by significantly accelerated rejection of C3H-derived skin grafts (Lenz et al., 1996). The XS106 cells that had been pulsed with DNFB or OVA primed the syngeneic mice to exhibit ear swelling responses to DNFB or footpad swelling responses to OVA, respectively (Matsue et al., 1999a). These results are consistent with the central dogma that DCs play crucial roles in the induction of hypersensitivity responses to environmental antigens, transplantation immunity against allografts, and protective immunity against infectious pathogens and tumors.

USES OF DENDRITIC CELL LINES IN BIOMEDICAL RESEARCH Molecular/biochemical analyses of DC biology Long-term DC lines have served as useful tools for studying the biology of DCs at molecular and

biochemical levels. For example, we have employed subtractive cDNA cloning in an attempt to identify the genes that are expressed selectively by DCs. cDNAs prepared from the XS52 line were hybridized with excess amounts of mRNAs isolated from the J774 macrophage line, and a DC-specific cDNA library was then constructed from the cDNA clones that remained unhybridized. Following three rounds of screening of this library, we have identified two unique type II membrane-integrated molecules, termed dectin-1 and dectin-2, both of which contained conserved carbohydrate recognition domain motifs of C-type lectins (Ariizumi et al., 2000a, 2000b). This project, which required large numbers ( 109 cells) of pure DC populations, would not have been feasible in the absence of a stable DC line. DC lines have been also used to study nitric oxide (NO) synthesis in DCs. Qureshi et al. (1996) reported that XS52 cells expressed inducible NO synthase (iNOS) and produced NO in response to LPS stimulation. LPS-induced NO synthesis by XS52 cells was augmented significantly by IFNγ and downregulated by IL10 (Qureshi et al., 1996). Working with FSDC cells Cruz et al. (1999) subsequently characterized the signaling pathway for LPS-induced NO synthesis. LPS stimulation rapidly triggered nuclear translocation of NFκB and subsequently upregulated iNOS expression. A specific inhibitor of Janus kinase-2 (JNK2) blocked both LPS-triggered NFκB translocation and LPSinduced NO synthesis. These observations suggest that LPS activates the NFκB pathway in a JNK2-dependent mechanism, leading to the iNOS upregulation, followed by NO production (Cruz et al., 1999). Munder et al. (1999) reported recently that arginine was metabolized not only into NO by iNOS, but also into ornithine and urea by arginase I in D2SC/1 cells. Interestingly, the balance between the iNOS pathway and the arginase pathway was regulated differentially by TH1 and TH2 cytokines, with IL-4 and IL-10 augmenting the arginase I activity selectively, whereas IFNγ and TNFα elevated solely iNOS activity (Munder et al., 1999). Once again, these studies performed at the biochemical level

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document the usefulness of long-term DC lines for studying the biology of DCs.

Mechanisms of DC maturation The DC lines described in this chapter differ substantially in their steady states of maturation (Table 12.5). As judged by the surface expression of MHC class II and co-stimulatory molecules, by the endocytotic potentials, and by their capacity to activate naïve T cells in allogeneic MLR, they may be classified into: (a) mature DCs (XS106), (b) immature DCs (CB1, D2SC/1 and 80/1), and (c) early DC precursors (FSDC and XS52). These DC lines, in turn, have provided a unique opportunity to study mechanisms that regulate DC maturation. Following stimulation with GM-CSF, CB1 cells began to secrete IL-1β and TGFβ1 and they also elevated the in vitro ability of MHC class IIrestricted antigen presentation. Moreover, their in vivo ability to induce contact hypersensitivity responses in animals was also augmented significantly by pretreatment with GM-CSF (Paglia et al., 1993; Granucci et al., 1994). FSDC cells exhibited a potent T cell-stimulatory capacity only after treatment with GM-CSF or IFNγ (Girolomoni et al., 1995). XS52 cells elevated the surface expression of MHC class II molecules as well as their T cell-stimulatory capacity in MLR when cultured with GM-CSF alone, in the absence of added NF fibroblast supernatant (Xu et al., 1995c). More recently, Yamada and Katz TABLE 12.5

(1999) have induced further maturation of XS52 cells by using the combination of IL-4, IL-1β, TNFα and agonistic anti-CD40 monoclonal antibody. The XS52 cells cultured for several days under these conditions expressed high levels of MHC class II, CD80, CD86, CD40 and CD11c, and they exhibited potent T cell-stimulatory capacities in allogeneic MLR as well as in the contact hypersensitivity assay in vivo (Yamada and Katz, 1999). D1 cells upregulated the surface expression of MHC class II, CD86 and CD40 following stimulation with IL-1β or TNFα (Winzler et al., 1997). TNFα-triggered maturation of this DC line was also accompanied by depolymerization of F-actin filaments, diminished pinocytotic capacity, loss of vinculin-containing adhesive structure, and elevated motility. These observations illustrate mechanisms by which secreted cytokines regulate DC maturation. DC maturation is also regulated by other stimuli. Upon stimulation with LPS, XS52 cells elevated rapidly the expression of CD86 and, at the same time, they also diminished the expression of CD115 and their phagocytotic and adhesive capacities (Takashima et al., 1995; Kitajima et al., 1995a, 1996a). Interestingly, LPS-triggered maturation of XS52 cells was inhibited by calcitonin gene-related peptide (CGRP) (Asahina et al., 1995; Torii et al., 1997b). These results, together with the recent report on the expression of neuropeptide receptors by XS52 cells (Torii et al., 1997c), suggest a novel downregulatory mechanism of DC maturation. D2SC/1 cells exhibited

Maturational states of DC lines CB1

D2SC/1

FSDC

80/1

XS52

XS106

D1

Maturational state

immature

immature

early precursor

immature

early precursor

mature

immature

Maturation stimuli

GM-CSF

B.burgdorferi

GM-CSF IFNγ

NT

GM-CSF NT IFNγ IL-4 IL-1β TNFα anti-CD40 CD4 T cells LPS

NT = not tested

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IL-1β TNFα CD4 T cells apoptotic body S.gordonii immune complex LPS

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mature features after treatment with live Borrelia burgdorferi (Altenschmidt et al., 1996), and D1 cells underwent maturation following LPS stimulation or upon phagocytosis of Staphylococcus gordonii or apoptotic tumor cells (Rovere et al., 1998; Rescigno et al., 1998). Moreover, D1 cells, which express FcγRIIb1, FcγRIIb2 and FcγRIII, underwent phenotypic maturation after treatment with an immune complex composed of OVA and anti-OVA antibodies (Regnault et al., 1999). Taken together, these observations illustrate the ability of DCs to undergo maturation in response to exposure to relevant antigens, such as pathogenic microorganisms, apoptotic tumor cells and the immune complex. XS52 cells exhibited many of the features of mature DCs upon antigen-specific interaction with T cells. Within 24 hours of co-culturing with CD4 T-cell clones in the presence of relevant antigens, XS52 cells began to secrete proinflammatory cytokines, elevated the expression of CD86, diminished the expression of CD115, and lost their phagocytotic capacity. None of these changes were observed when XS52 cells were co-cultured with T cells or antigens alone (Kitajima et al., 1995a, 1995b, 1996a). A similar observation was also made for D1 cells, which secreted IL-12 (p70) upon antigen-specific interaction with T cells (Winzler et al., 1997). These changes may represent a critical transition of DCs, termed T cell-mediated terminal maturation, that takes place during antigen presentation. Interestingly, dexamethasone at relatively low concentrations inhibited T cell-mediated maturation of XS52 cells, suggesting a mechanism by which glycocorticoids suppress cellular immune responses (Kitajima et al., 1996b). More recently, we observed that XS52 cells eventually underwent apoptosis in CD95dependent and independent fashions after antigen-specific interaction with CD4 T cells. Not only does this phenomenon of T cell-mediated DC apoptosis document the terminal phase of DC maturation, it also suggests a unique downregulatory mechanism that prevents the interminable activation of T cells by antigen-bearing DC (Matsue et al., 1999b).

Development of DC-based immunotherapies DC lines have been used to develop several new technologies that are potentially applicable for the treatment of human disease. In conventional DC-based vaccines, DCs are pulsed with soluble protein antigens or antigenic peptides in vitro and then administered to animals. Bachmann et al. (1996) reported that viral antigens in particulate forms or in association with cell debris were presented by D2SC/1 cells predominantly on the MHC class I molecules for efficient CTL activation. Similar observations have been made for the MHC class II-negative, GM-CSF-dependent DC line 18 (Schirmbeck et al., 1995). Rovere et al. (1998) showed that D1 cells phagocytose apoptotic tumor cells expressing OVA and present OVA-derived peptides efficiently to both MHC class I- and class IIrestricted T cells. Rescigno et al. (1998) used genetically modified bacteria for MHC class Iloading of antigens. D1 cells phagocytosed recombinant Streptococcus gordonii producing OVA, processed OVA by TAP-dependent mechanisms, and presented the antigenic peptides to the class I-restricted T-cell clone with extremely high efficiency, 106-fold higher than that for soluble OVA. Regnault et al. (1999) showed that the efficiency of MHC class I-restricted presentation of OVA by D1 cells can be augmented 104-fold by conjugating OVA with anti-OVA antibodies, revealing a pathway in which antibody-captured antigens are processed by DC for MHC class I presentation. It is known that antigen-specific CTL responses are inducible with extremely high efficiency by conjugating antigenic peptides to heat shock proteins (HSP). Working with D2SC/1 cells, Arnold Schild et al. (1999) uncovered the molecular basis for the HSP-mediated MHC class I presentation. Gold-labeled HSP first bound to the clathrin-coated pits on D2SC/2 cells. The surface bound HSP were subsequently endocytosed and co-localized with MHC class I molecules in early and late endosomal structures, illustrating a pathway in which HSP– peptide conjugates are endocytosed through HSP receptors and processed for MHC class I

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SUMMARY

presentation. These strategies (i.e. use of particulate antigens, apoptotic bodies, recombinant bacteria or HSP–peptide conjugates) may improve the clinical efficacy of DC-based vaccines to initiate antigen-specific CTL responses. Song et al. (1997) demonstrated that DCs can be loaded genetically with tumor-associated antigens. Administration of the XS52 cells that had been transfected with the β-Gal gene using an adenoviral vector induced the generation of β-Gal-specific CTL responses. Working with a βGal-transduced colon carcinoma cell line as a model, they observed further that a single s.c. injection of β-Gal-transfected XS52 cells was sufficient for the induction of protective immunity against lethal challenge with live tumor cells (Song et al., 1997). By introducing the MUC-1 gene into D2SC/1 cells using a retroviral vector, Henderson et al. (1998) also observed the generation of MUC-1-specific CTL responses in vivo. We employed a gene gun to introduce the α1antitrypsin (AAT) gene into XS106 cells. AATtransduced XS106 cells, but not AAT-transduced fibroblasts, induced the development of a wide variety of immune responses to AAT, including CTL activity, humoral responses, and delayedtype hypersensitivity responses (Timares et al., 1998). These observations illustrate a new strategy of ‘genetic’ antigen loading of DCs. Paglia et al. (1996) introduced the GM-CSF gene into D2SC/1 cells by using a retroviral vector and tested their in vivo ability to initiate antigenspecific CTL activity. GM-CSF-transduced DCs loaded in vitro with β-Gal proteins induced β-Gal-specific CTL responses, whereas nontransduced DCs induced no detectable responses (Paglia et al., 1996). We introduced the IL-4 gene into XS106 cells by using the polyamidoamine dendrimer-based system (SuperFectTM). When pulsed in vitro with KLH and then injected into syngeneic animals, the IL4-transduced XS106 cells induced TH2-biased immune responses to KLH (Hayashi et al., 2000). These observations indicate that the magnitude and direction of immune responses induced by DC-based vaccines can be manipulated experimentally by introducing selected cytokine genes into DCs.

Fernandez et al. (1999) reported the unique ability of D1 cells (as well as bone marrowderived DCs) to activate natural killer (NK) cells. Not only did D1 cells elevate the in vitro cytotoxicity of NK cells against NK-sensitive YAC-1 targets, they also delayed significantly the growth of AK7 tumor cells in nude mice upon local injection into the tumor site (Fernandez et al., 1999). Finally, we have created ‘killer’ DCs by introducing the CD95L gene into XS106 cells by gene gun. The resulting CD95L-transduced XS106 cells, when pulsed with OVA, showed the potent ability to kill OVAreactive CD4 T cells in vitro. When injected into syngeneic mice, the OVA-pulsed CD95LXS106 cells suppressed footpad swelling responses to OVA, without affecting the responsiveness to an irrelevant antigen, hen egg lysosome (HEL). Conversely, HEL-pulsed CD95LXS106 cells suppressed HEL responses, but not OVA responses, documenting antigen-specificity. Moreover, DNFB-pulsed CD95L-XS106 cells suppressed contact hypersensitivity responses to DNFB. We believe that killer DCs represent a novel immunosuppressive strategy that is designed to selectively eliminate pathogenic T cells that recognize a given antigen (Matsue et al., 1999a). Taken together, it is evident that long-term DC lines have provided extremely useful tools for the development of new DCbased immunotherapies.

SUMMARY Over the last several years, several stable DC lines have been established from mice. These lines include oncogene-immortalized CB1, D2SC/1 and FSDC lines and growth factordependent 80/1, XS52, XS106 and D1 lines. These lines maintain the original features of DCs by virtue of surface phenotype, endocytic potential, antigen-processing capacity, cytokine and cytokine receptor profiles, and the ability to initiate antigen-specific immune responses. These DC lines have provided extremely useful tools for studying the biology of DCs at molecular and biochemical levels. Moreover, some

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of the DC lines have allowed the investigators to develop new DC-based immunoregulatory protocols that are potentially applicable for the treatment of cancer, infectious disease and autoimmune disease.

ACKNOWLEDGEMENTS We thank many colleagues who established and characterized the XS series of DC lines. These individuals include Drs Kiyoshi Ariizumi, Paul R. Bergstresser, Gisela Caceres-Dittmar, Satoru Hayashi, Stephen A. Johnston, Toshiyuki Kitajima, Keiko Matsue, Mansour Mohamadzadeh, Laura Timares, Shan Xu and Koichi Yokota. We also thank Pat Adcock for her excellent secretarial assistance in the preparation of this manuscript. This study was supported by grants (RO1-AR35068, RO1-AR43777, and RO1-AI43262) from the National Insitutes of Health.

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Munder, M., Eichmann, K., Modolell, M. et al. (1999). J. Immunol. 163, 3771–3777. Mutini, C., Falzoni, S., Ferrari, D. et al. (1999). J. Immunol. 163, 1958–1965. Nunez, R., Ackermann, M. and Suter, M. (1997). Adv. Exp. Med. Biol. 417, 425–432. Nunez, R., Sanchez, M., Wild, P. et al. (1998). Immunol. Lett. 61, 33–43. Nunez, R., Grob, P., Baumann, S. et al. (1999). Immunol. Cell. Biol. 77, 153–163. Paglia, P., Girolomoni, G., Robbiati, F. et al. (1993). J. Exp. Med. 178, 1893–1901. Palgia, P., Chiodoni, C., Rodolfo, M. et al. (1996). J. Exp. Med. 183, 317–322. Qureshi, A.A., Hosoi, J., Xu, S., Takashima, A. et al. (1996). J. Invest. Dermatol. 107, 815–821. Rasko, J.E.J., Metcalf, D., Alexander, B. et al. (1997). Leukemia 11, 732–742. Regnault, A., Rescigno, M., Ricciardi-Castagnoli, P. et al. (1998). J. Immunol. 161, 2106–2113. Regnault, A., Lankar, D., Lacabanne, V. et al. (1999). J. Exp. Med. 189, 371–380. Rescigno, M., Citterio, S., Thèry, C. et al. (1998). Proc. Natl Acad. Sci. USA 95, 5229–5234. Riva, S., Nolli, M.L. and Lutz, M.B. (1996). J. Inflamm. 46, 98–105. Robinson, S.P., English, N., Jaju, R. et al. (1998). Br. J. Haematol. 103, 763–771. Rodriguez, A., Regnault, A., Kelijmeer, M. et al. (1999). Nature Cell Biol. 1, 362–368. Rovere, P., Vallinoto, C., Bondanza, A. et al. (1998). J. Immunol. 161, 4467–4471. Rovere, P., Sabbadind, M.G., Vallinoto, C. et al. (1999). Arthritis Rheum. 42, 1412–1420. Rowden, G., Dean, S. and Colp, P. (1995). Adv. Exp. Med. Biol. 378, 39–41. Russell, S.M., Keegan, A.D., Harada, N. et al. (1993). Science 262, 1880–1883. Sallusto, F., Cella, M., Danieli, C. et al. (1995). J. Exp. Med. 182, 389–400. Santiago-Schwarz, F., Coppock, D.L., Hindenburg, A.A. et al. (1994). Blood 84, 3054–3062.

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13 Dendritic cell-derived exosomes Clotilde Théry 1, Joseph Wolfers 2, Armelle Regnault 1, Fabrice André 2, Nadine C. Fernandez 2, Graça Raposo 3, Sebastian Amigorena 1 and Laurence Zitvogel 2 1

INSERM, Institut Curie, Paris, France Institut Gustave Roussy, Villjuif, France 3 CNRS, Institut Curie, Paris, France

2

Someday, after we have mastered the winds, the waves, the tide and gravity, we shall harness for God the energies of love. Then, for the second time in the history of the world, man will have discovered fire. Teilhard de Chardin

INTRODUCTION

Subsequently, Raposo et al. (1996, 1997a, 1997b), reported that EBV-transformed B (EBV-B) lymphocytes and mast cells also secrete similar membrane vesicles. In professional APCs, multivesicular late endosomes are sites of MHC class II peptide loading (West et al., 1994; Morkowski et al., 1997; Pierre and Mellman, 1998). Accordingly, the presence of functional MHC class II–peptide complexes on exosomal membranes was confirmed using CD4 T-cell hybridomas in vitro (Raposo et al., 1996). More recently, we found that dendritic cell (DC)-derived exosomes accumulate not only MHC class II–peptide complexes but also MHC class I molecules that trigger T cell-mediated antitumor immune responses in vivo (Zitvogel et al., 1998). In order to unravel the molecular bases of the immunostimulatory effects of DCderived exosomes, the conditions of their production as well as their protein composition were analyzed.

Exosomes were originally described as small membrane vesicles of 50–90 nm diameter released from reticulocytes during maturation into red blood cells to eliminate transferrin receptors ( Johnstone et al., 1987). Electron microscopy studies (Pan et al., 1985) have suggested that exosomes do not result from the budding of plasma membrane but rather originate from late multivesicular endosomes (MVBs). The small vesicles accumulated in the lumen of MVBs are thought to arise from the inward budding of the endosome limiting membrane. During this process a small portion of cytosol is trapped inside the vesicle (Trowbridge et al., 1993; van Deurs et al., 1993). Upon direct fusion of the MVB with the plasma membrane, the internal membrane vesicles, called ‘exosomes’, are released into the extracellular milieu (Pan et al., 1985) (Figure 13.1).

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FIGURE 13.1 Schematic representation of the endocytic and exocytic pathway in antigen-presenting cells. Ultrastructural studies performed in EBV-B cells revealed that the vast majority of MHC class II molecules reside in late endocytic compartments, called MHC class II compartments (MIICs). MIICs encompass both multilamellar structures and/or multivesicular bodies (MVBs). MVBs represent a meeting point between the endocytic and the exocytic pathways. In electromicroscopy studies, the late endosomes appear as 200–300 nm membrane compartments containing in their lumen variable amounts of small 60–90 nm vesicles. The internal vesicles of the MVBs seem to arise from the budding, into the endosomal lumen, of a portion of the external limiting membrane of the MVB. During this invagination process, a sorting of certain cytosolic and membrane proteins may occur. The limiting membrane fuses with the plasma membrane, resulting in the exocytosis of the internal vesicular content, the exosomes, into the extracellular milieu. TGN, trans-Golgi network.

TUMOR PEPTIDE-PULSED DC-DERIVED EXOSOMES INDUCE TUMOR GROWTH SUPPRESSION IN TUMOR-BEARING MICE The initial investigations were performed in mouse DCs. As shown by electron microscopy (EM), exosomes are secreted from immature DCs (bone marrow (BM)-derived DC or D1 splenic dendritic cell line, Winzler et al., 1997) displaying numerous and characteristic MVBs containing internal vesicles labeled with antiMHC class II antibodies. Whole-mount EM studies of the exosome pellets purified by differential ultracentrifugation of DC supernatants showed a population of vesicles (Figure 13.2)

FIGURE 13.2 Whole-mount immuno-electron microscopy of D1-derived exosomes. The 100 000 g pellets obtained after differential ultracentrifugation are composed of small vesicles with a diameter varying from 50 to 90 nm, intensely labeled with an anticlass II antibodies (protein A-gold 15 nm, Pag 15). Bars  250 nm.

morphologically similar (cup-shaped) to exosomes released by human monocyte-derived DC (MD-DC) (Zitvogel et al., 1998) but appearing more heterogeneous in size than those secreted by EBV-B cells. Western blot analyses showed that mouse D1-derived exosomes are enriched in MHC class II molecules compared with D1 lysates, and devoid of Ii and calnexin (ER-resident proteins). Like the EBV-B cellsderived exosomes (Raposo et al., 1996; Escola et al., 1998), D1- and BM-DC-derived exosomes migrate on a continuous sucrose gradient at a density of 1.14–1.20 g/mL, confirming their vesicular nature (Théry et al., 1999). We tested the capacity of these vesicles to induce T cell-mediated immune responses in vivo. BM-DCs cultured in IL-4  GM-CSF (Zitvogel et al., 1996) loaded with acid-eluted tumor peptides were previously shown to mediate specific antitumor immune responses. P815 is an immunogenic but aggressive mastocytoma, syngeneic of DBA/2 (H-2d), for which

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very few effective immunotherapies on day 10-established tumors have been reported. Acid-eluted tumor (P815) peptides were pulsed onto syngeneic mouse BM-DCs as previously described (Zitvogel et al., 1996). Exosomes were prepared from the DC supernatants by differential ultracentrifugation and used for in vivo immunization. As reported in Zitvogel et al. (1998), therapy of day 10-established P815 tumors (50–90 mm2 in size) was carried out using a single intradermal (i.d.) administration of 3–5 µg of exosomes per mouse, corresponding to the 24-hour production of 1–5 million BM-DCs ex vivo. Within a week, tumor growth stopped in the groups receiving exosomes derived from autologous tumor peptide-pulsed DCs and 40–60% of mice were tumor-free at day 60. These animals had a long-lasting immune response and rejected a lethal tumor challenge with P815 but not with the syngeneic leukemia L1210 (L. Zitvogel, unpublished results). Groups of mice immunized with exosomes derived from self-splenic peptide-pulsed DCs showed no effect on tumor growth as compared with control mice groups. Therefore, P815 peptide-pulsed DC-derived exosomes promoted tumor regression. Similar antitumor effects were achieved in the day 3–4 established TS/A tumor model. These antitumor effects were not found in athymic Nu/Nu counterparts, indicating that T cells are required for the exosome-induced antitumor immune responses. Splenocytes from mice that rejected P815 tumors following immunization with exosomes were harvested at day 90 and cultured for 5 days in the presence of irradiated B7.1-expressing P815 cells to enhance specific precursor frequency. These effector cells were tested in a 4-hour51Cr release assay against the autologous tumor cells P815 (H-2d), against the irrelevant H-2d leukemia L1210, and YAC cells. Significant specific lytic activity on P815 was achieved in splenocytes from exosomeimmunized mice (Zitvogel et al., 1998). Interestingly, none of the spleens from littermates spontaneously rejecting P815, or bearing growing P815 tumours display cytolytic activity against P815 under the same conditions

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(Zitvogel et al., 1998). Therefore, a single injection of exosomes derived from DCs pulsed with the relevant peptides efficiently primed specific antitumor CTL responses in vivo. In order to assess the usefulness of DCderived exosomes as vectors for the in vivo priming of peptide-specific CTL responses, we tested the CTL activity of splenocytes collected from mice immunized with H2b-BM-DC pulsed with OVA peptides. Specific responses were achieved that were comparable with that mediated with OVA peptide  incomplete Freund adjuvant i.d. (J. Wolfers, unpublished results).

EXOSOMES BEAR MHC CLASS I–PEPTIDE COMPLEXES THAT ARE FUNCTIONAL IN VITRO AND IN VIVO As reported in Zitvogel et al. (1998), human MDDCs secrete antigen-presenting vesicles similar to murine DC-derived exosomes. In a human in vitro model system, we determined whether exosomes may directly stimulate HLA-A2restricted MART1-specific CD8 CTLs clones (Dufour et al., 1997). HLA-A2 human (day 6) MD-DCs were pulsed with MART-1/MelanA(26-35) peptides for 24 hours and exosomes were isolated from the DC culture supernatants by differential ultracentrifugation. MART1/A2 exosomes were incubated with the CTLs with or without HLA-A2 DCs. Exosomes could not directly trigger IFNγ production from the CTL clone. In contrast, when pulsed onto DCs, they efficiently stimulate these clones (significant IFNγ, TNFα production). Carryover peptides could be ruled out since exosomes secreted from HLA-A2 DCs pulsed with similar peptides had no effect. In addition, when MART1/A2 exosomes were pulsed onto HLA-A2 DCs, similar IFNγ production could be achieved from the specific clones (Wolfers et al., 2000). It is noteworthy that Grommé et al. (1999) have recently described the presence of MHC class I molecules in late endocytic compartments of EBV-β cells and melanoma cells which may account for

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peptide loading onto class I molecules in a TAPindependent manner.

OVERALL PROTEIN COMPOSITION OF EXOSOMES To help our understanding of the molecular basis of the effects exosomes have on the immune system, a characterization of the protein content of exosomes was carried out in murine DCs (Théry et al., 1999). BM-DCs and D1 were metabolically labeled with [35S]methionine/ cysteine and, after a cold wash, metabolically labeled exosomes contained in the 24-hour culture supernatant were harvested by ultracentrifugation. The amount of radioactivity recovered in exosomes was 0.17  0.1% of the radioactivity incorporated into cells. The protein composition of exosomes from metabolically labeled cells was analyzed by SDS-PAGE and autoradiography, and compared with that of whole cells. As shown in Figure 13.3, exosomes from BM-DCs and the D1 cell line display a unique protein pattern very different from the protein pattern of whole cells. All the proteins present in exosomes comigrated on a continuous sucrose gradient at the expected vesicular density (1.15 g/mL) (Théry et al., 1999). The same protein pattern was obtained reproducibly with different preparations of BMDC- or D1-derived exosomes, showing that these vesicles represent a new subcellular compartment resulting from the active selection of a defined set of proteins. The specificity of the mechanism of exosome production was also confirmed by the observation that this production is regulated: only immature DCs efficiently secrete exosomes, a strong reduction of exosomes production was observed upon activation by LPS (Théry et al., 1999). In order to identify the exosomal proteins, the major bands of D1-derived exosomes preparations run onto a 8–15% gradient SDS gel were excised, trypsin digested and the resulting peptides analyzed using matrix-assisted laser desorption ionization time-of-flight mass spec-

FIGURE 13.3 Protein pattern of metabolically labeled BM-DCs and D1 cells and exosomes. Exosomes (Ex) purified from the supernatant of metabolically labeled D1 cells or BM-DCs were run on a 11% SDS gel, together with lysate (including both cytosolic and membrane components) obtained from the whole cells (Cell). Exosomes contain a subset of proteins distinct from the whole cell proteins (compare Ex and Cell from each cell type). D1 and BM-DCs whole cells on the one hand, and exosomes on the other, show very similar protein patterns. This validates the choice of D1 as the model system used to analyze the protein composition of exosomes.

trometry (MALDI-TOF-MS) and subsequently compared with the theoretical tryptic peptide profiles of known proteins from the databases ( Jensen et al., 1996). Among the identified proteins, three are transmembrane (Mac-1 α chain,

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MHC class II β chain, CD9), one is secreted and peripherally associated to membranes (MFG-E8), and four are cytosolic (Gi2α, annexin II, gag from MRV provirus, hsc73). Consistent with the proposed origin of exosomes, most proteins identified in exosomes have been described elsewhere in association with endocytic compartments. Western blots and immunoprecipitation studies concluded that MHC class II and CD9 are enriched by 10 fold in exosomes compared with total cells, Mac-1 was enriched by 5–10-fold, annexin II and hsc73 by 2–3-fold. In immuno-EM studies it appeared that, in contrast to MHC class II, CD9 and Mac-1 α chain, hsc73 and annexin II are not exposed at the surface of exosomes but contained within the lumen. Figure 13.4 shows a schematic representation of our current view of the structure of exosomes, and of the potential roles of the proteins they contain. The cytosolic proteins found in exosomes (annexin II, Gi2α) are most likely involved in exosome formation or biogenesis. The presence of hsc73 in exosomes is also very interesting in light of the reported effect of this protein in antitumor immunity (Srivastava et al., 1998). The membrane proteins found in exosomes are

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potentially involved in T-cell activation (MHC class I, II, CD86), or in the association of exosomes to target cells (MFG-E8, Mac-1, CD9). The most abundant exosomal protein, MFG-E8, originally described at the surface of milk fat globules (Stubbs et al., 1990) has two EGF-like domains with an integrin-binding motif (ArgGly-Asp), and a phosphatidylserine-binding, factor VIII-like domain. This suggests that MFGE8 may bind exosomes through phospholipids. The human and bovine homologues of MFG-E8 also interact with αvβ3 and αvβ5 integrins (Andersen et al., 1997; Taylor et al., 1997), suggesting that MFG-E8 addresses exosomes to cells expressing these integrins. CD9 is the major tetraspanin in mouse D1-derived exosomes. CD9 also associates with other transmembrane proteins and is a cofactor potentiating the interaction between EGF-like growth factors and their receptors (Higashiyama et al., 1995). The identification of Mac-1 α chain coprecipitating with its β chain counterpart together with data from immuno-EM studies suggest that this β2 integrin is functional on the surface of exosomal membranes and might bind efficiently to ICAM-1- or ICAM-2-expressing cells.

FIGURE 13.4 A speculative model for the protein structure of DC-derived exosomes. The molecular structure has been examined using western blot analysis, immunoelectron microscopy, metabolic labeling experiments and electrophoresis (SDS-PAGE) followed by trypsin digestion and MALDI-TOF-MS analyses of the peptides (Théry et al., 1999). The topology of membrane proteins is based on previous studies on exosome membrane orientation (Pan et al., 1985; Raposo et al., 1997a) and on EM observations of whole-mounted exosomes (Théry et al., 1999). The most abundant proteins are involved in MHC-restricted T-cell induction, in the targeting of exosomes to other cells of the immune system and in vesicule biogenesis. DENDRITIC CELL BIOLOGY

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DISCUSSION Ultrastructural studies of the possible transport routes of lysosomal constituents to the cell surface in not only B cells but also DCs revealed that late endosomal compartments, i.e multivesicular bodies displaying intraluminal membrane vesicles, can fuse with the plasma membrane in an exocytic fashion and release, in the extracellular environment, 60–90-nm diameter vesicles called ‘exosomes’ (Raposo et al., 1996, 1997a, 1997b; Escola et al., 1998; Zitvogel et al., 1998; Kleijmeer et al., 1998). We reported that antigen-loaded DC-derived exosomes bear functional class I–peptide complexes that allow CTL priming in vivo and established tumor growth suppression in various murine experimental model systems (Zitvogel et al., 1998). Exosomes can be obtained in relatively large quantities (1 µg/million DCs per 24 hours using Bradford assay, up to 0.25% of protein turnover in D1) from the culture media of immature DCs (d5 mouse BM-DCs in GM-CSF  IL-4, growth factor-dependent D1 line in the absence of activating stimuli, or CD83 MD-DC from human peripheral blood monocytes) following ultracentrifugation at 100 000 g of the culture supernatants. We characterized exosomes by morphological and biochemical criteria. The membranes pelleted at 100 000 g, analyzed by immunoelectron microscopy, represented a population of vesicles that resembled those isolated from the cell culture supernants of EBV-B cells, labeling for MHC class I, class II, CD86 and lysosomal-associated tetraspan molecules (Escola et al., 1998; Kleijmeer et al., 1998). Exosome abundantly overexpressed MHC class II, tetraspanins such as CD63/CD81, CD82 and CD86 molecules as compared with plasma membrane. Endoplasmic reticulum components were not detected in western blotting using anti-calnexin and anti-gp96 antibodies (Théry et al., 1999). Flotation on a continuous sucrose gradient revealed that all the exosomal proteins are found in membrane vesicles floating to a density of 1.13–1.20 g/mL. Exosome preparations did not significantly carry over retroviruses, plasma membrane fragments,

microsome constituents or apoptotic bodies (Zitvogel et al., 1998; Théry, unpublished results). The extensive characterization of the molecular composition of D1-derived exosomes has been examined by Théry et al., (1999). Even though exosome release has been associated with clearance of transferrin receptors, reticulocyte maturation and differentiation into an erythrocyte pathway (Johnstone et al., 1987), the physiological relevance of exosome secretion and function in vivo are still a matter of debate. Our data imply an immunostimulatory role of this exosome release and show that (1) their protein composition differs from that of plasma membrane with selective enrichments with proteins of endosomal origin, (2) exosome secretion by DCs seems to be regulated and active in that upon maturation, it is strongly reduced, (3) their immunogenicity in tumorbearing mice could be accounted for by the functionality and enrichment of the MHC class I and II–peptide complexes as well as HSPs (hcs73). They also bear large amounts of CD86 and CD9 tetraspanins as well as integrins such as Mac-1 α chain that may facilitate MHCrestricted immune responses. Importantly, the most abundant exosomal protein, MFG–E8, binds to integrins αvβ3 and αvβ5 expressed on immature DCs and macrophages and could target DC-derived exosomes to other APCs. In line with this assumption, our unpublished data show that MHC class I–peptide complexes presented on exosomes are functional onto CTL clones only when APCs are co-cultured with exosomes plus T cells. It is conceivable that following a danger signal/stress (Bell et al., 1999), the DCs might not be able to migrate to the T cell-enriched area of the lymphoid organs after antigen uptake and processing. In such circumstances, DCs might be able to secrete surrogate antigen-presenting vesicles that could prime and/or boost immune reponses at the periphery. In line with such a hypothesis, our preliminary data show that incubation of DCs with IL-10 allows secretion of larger numbers of immunogenic vesicles in vivo. Should exosomes be delivered at ports of entry where immature DCs are sentinels, they might be able to modulate innate

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effector functions since we showed that exosomes trigger IFNγ production from resting NK and NKT cells in vitro. However, it is also conceivable that clusters of MHC class I and II– peptide complexes with CD86 presented on exosomes represent an amplification system for the DCs to facilitate sustained serial triggering of TCR in the T cell-enriched areas. In line with this hypothesis, IFNγ is a helper cytokine which enhances the immunogenicity of exosomes in tumor-bearing mice (J. Wolfers, unpublished results). Further studies are needed to further unravel the mechanisms of exosome-mediated antitumor effects. These data support the use of DC-derived exosomes for cancer immunotherapy as a novel dendritic cell-free therapeutic cancer vaccine and suggest that exosomes may represent a physiological means of communication between DCs and T lymphocytes.

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14 Dendritic cells as recipients of cytokine signals Jonathan Cebon, Ian Davis, Thomas Luft and Eugene Maraskovsky Ludwig Institute for Cancer Research, Melbourne Tumor Biology Branch, Austin & Repatriation Medical Centre, Heidelberg, Victoria, Australia

Who are YOU? said the Caterpillar . . . Alice replied, rather shyly, I—I hardly know, sir, just at present— at least I know who I WAS when I got up this morning, but I think I must have been changed several times since then. Lewis Carroll, Alice’s Adventures in Wonderland

INTRODUCTION

systems that have defined the roles of these cytokines include in vitro studies, animal models and to a more limited extent clinical research. Because of the critical interdependence between DCs and the cells with which they interact, it has not always been possible to distinguish the direct from indirect effects of cytokines. Furthermore, many experimental systems are highly artificial and, given the sensitivity of DCs to environmental manipulation, it can be difficult to know how physiologically relevant an observation may be. Nonetheless, using knockout and transgenic mice as well as observing clinical effects, the impact of cytokine signals on DCs is now better understood. Complex mixtures of cytokines have been used for optimal production of DCs from various precursors (Saunders et al., 1996). It is not clear, however, that these conditions are those which are involved in DC generation in vivo. For example, although GM-CSF and IL-4 allow immature

Dendritic cells (DCs) display extraordinary plasticity, transforming from one form to another in order to perform a variety of discrete and highly specialized functions. To control these transformations, complex regulation is required throughout the DC life cycle. Soluble and membrane-associated signals maintain homeostasis, enable fine control of phenotype and function through maturation and migration and finally, orchestrate immune responses during inflammation or immune challenge. Among these regulatory molecules, cytokines, chemokines, prostaglandins and products of diseased tissues (derived from pathogens or distressed cells) have all been shown to have potent effects on DC form and function. Cytokines implicated in the process of DC ontogeny, maturation and activation are shown in Table 14.1 and Figure 14.1. Experimental Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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TABLE 14.1 Cytokines used in generating DCs in vitro References

DCs generated in GM-CSF plus IL-4 require further stimulus for mature phenotype

Inaba et al., 1993 Sallusto and Lanzavecchia, 1994

IL-4

DCs generated in GM-CSF plus IL-4 require further stimulus for mature phenotype Enhances DC1 maturation; kills DC2

Sallusto and Lanzavecchia, 1994 Rissoan et al., 1999

IL-13

Can be used in place of IL-4 Causes maturation of monocyte-derived DCs grown in GM-CSF and IL-4

Piemonti et al., 1995 Alters et al., 1999 Sato et al., 1999

TNFα

Important in maturation Important in differentiation from hematopoietic progenitors

Sallusto and Lanzavecchia, 1994 Zhang et al., 1997

Flt-3L

Increase in DCs generated from hematopoietic progenitors with GM-CSF and TNFα

Rosenzwajg et al., 1998

c-kit ligand (stem cell factor, Steel factor)

Increase in DCs generated from hematopoietic progenitors with GM-CSF, TNFα and other combinations

Rosenzwajg et al., 1998 Caux et al., 1997 Saunders et al., 1996 Zhang et al., 1997

TGFβ1

Promotes generation of Langerhans cells from monocytes in vitro

Geissmann et al., 1998

IL-3

DCs from thymic precursors May be important for lymphoid DCs expressing IL-3Rα

Saunders et al., 1996

IL-6

Promotes generation of DCs from thymic precursors Blockade of IL-6 decreases generation of DCs from CD34 progenitors; no effect on mature DC function

Saunders et al., 1996 Santiago-Schwarz et al., 1996

IL-1β

Absence of IL-1β  TNFα leads to poor DC cluster formation in GM-CSF/DCs

Saunders et al., 1996

IFNα

Acceleration of DC maturation

Luft et al., 1998b

IL-10

Inhibition of DC generation from monocyte precursors Inhibition of antigen presentation, enhancement of endocytosis, implying immature phenotype Potentiates effect of IL-4 on DC1 and DC2

Buelens et al., 1997 Allavena et al., 1998

Important when GM-CSF absent

Saunders et al., 1996

IL-7

Rissoan et al., 1999

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GM-CSF

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Cytokine

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FIGURE 14.1

189

Cytokine regulators of dendritic cell ontogeny and function.

DCs to be generated from peripheral blood mononuclear cells (Sallusto and Lanzavecchia, 1994) it is difficult to imagine circumstances where this would occur in the appropriate context in vivo. There are two broad ways in which cytokines may regulate DCs. First, specific cytokines allow expansion of DC precursors. Absence of these factors will lead to deficiencies in particular DC subtypes. These deficiencies may not be absolute and might be functional rather than numerical. The effects of overexpression of these cytokines may be difficult to predict. Second, cytokines are able to influence DC differentiation from precursors, maturation and activation. In this case, over- or underexpression of particu-

lar cytokines may have important effects on the numbers of DCs within tissues or, more importantly, on the potential functional capabilities of DCs. This may in turn lead to impaired or enhanced antigen presentation and a consequent functional phenotype.

CYTOKINE REGULATION OF DCs In vitro studies The studies listed in Table 14.1 show that diverse cytokines can be used to generate DCs in vitro from monocytic precursors, CD34

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hematopoietic progenitors or thymic progenitors. Some apparently similar cytokines, for example IL-4 and IL-13, can have slightly different effects: IL-13 can replace IL-4 in combination with GM-CSF to allow production of DCs from monocytes, however IL-13 also causes maturation of the DC phenotype, whereas IL-4 does not (Sato et al., 1999). The importance of some other cytokines may be intuitively less obvious, although they may still play critical roles in vitro and possibly in vivo. IL-6 and IL-7 may be important cofactors, produced by the complex mixture of cells that is present during the in vitro culture of DCs from their precursors (Saunders et al., 1996). Interferon α allows acceleration of maturation of DCs from CD34 hematopoietic progenitors (Luft et al., 1998b), an observation we are incorporating into our own clinical trials of DCs. Conversely, IL-10 may inhibit the maturation of DCs in vitro (Allavena et al., 1998; Rissoan et al., 1999).

In vivo studies Analysis of the hematopoietic development of other leukocyte subsets indicates that the control mechanisms for these processes are likely to be complex and redundant. In vitro studies provide clues as to which cytokines may be critical in vivo. However, gene knockout studies have shown that apparently important factors are in some cases redundant for normal cell function and may have little effect on the overall phenotype of the animal. In contrast, other factors, perhaps previously unsuspected, play important roles. This is certainly the case in the control of DC differentiation. Cytokine knockout studies provide perhaps the purest method of determining the importance of particular molecules in the first levels of DC ontogeny. Studies in which cytokines are present at abnormally high levels, such as in the transgenic mouse or when exogenous cytokines are administered, can also provide important information although it is more difficult to control for redundant pathways. Table 14.2 lists studies in which the genes for

specific cytokines or their receptors have been disrupted and in which the effects on different DC populations have been evaluated. Some models in which high levels of certain cytokines are provided by gene transfection or by exogenous administration are also listed. TGF β The TGFβ1 gene has been deleted in mice using targeted gene disruption (Borkowski et al., 1996, 1997). In mice in which this signaling pathway has been disrupted, skin Langerhans cells (LCs) are completely absent (Borkowski et al., 1996). Follicular CD11c DCs are not affected. The defect is due to a direct effect on LCs rather than on marrow progenitors, since TGFβ1/ marrow gives rise to LCs when transferred into lethally irradiated wild-type recipients in which paracrine TGFβ1 secretion is intact (Borkowski et al., 1997). Inactivation of the TGFβ type II receptor by a dominant negative mutation affects TGF-β1 signaling in keratinocytes (Borkowski et al., 1997). These mice have normal numbers of LCs, indicating that the effect of TGFβ1 disruption on LCs is a result of effects directly on LCs or their precursors and is not caused by an epidermal abnormality. However, local TGFβ1 expression in the epidermis is probably also important. TGFβ1/ mice express low levels of mRNA for IL-1 and TNFα, two cytokines known to be important in mediating migration of LCs out of the epidermis (Borkowski et al., 1997). LT The major cytokine pathway involved in follicular DC (FDC) production is lymphotoxin (LT) (Alimzhanov et al., 1997; Fu et al., 1997; Koni et al., 1997; Matsumoto et al., 1997; Pasparakis et al., 1997; Alexopoulou et al., 1998; Fütterer et al., 1998; Berger et al., 1999; Ito et al., 1999a; Stoitzner et al., 1999). Targeted disruption of the LTα (Fu et al., 1997) or LTβ genes (Alimzhanov et al., 1997) or the LTβ receptor (Endres et al., 1999) leads to a complete lack of FDCs although skin LCs retain intact numbers

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TABLE 14.2 Cytokines relevant to DC ontogeny and function in vivo References

Targeted disruption

Normal numbers of DCs Intact T-cell allostimulatory capacity Impaired anti-KLH IgG2a production Impaired CD4 anti-KLH responses Possible skew towards lymphoid DC production

Saunders et al., 1996 Wada et al., 1997

Targeted disruption of common β chain

Decrease in number of DCs DCs can be produced in absence of GM-CSF, IL-3 and IL-5 signals

Vremec et al., 1997

Normal DC numbers in nodes and Peyer’s patches Langerhans cells present, migration intact Abnormal follicular DC organization

Pasparakis et al., 1997 Stoitzner et al., 1999

GM-CSF/IL-3/IL-5

TNFα

TNF receptor

Targeted disruption of p55 or p75

Absent follicular DC and germinal centers (p55) Langerhans cells present, migration intact in p55 knockout but impaired in p75 knockout

Pasparakis et al., 1997 Stoitzner et al., 1999 Wang et al., 1997 Fu et al., 1997 Matsumoto et al., 1997

Lymphotoxin α

Targeted disruption

Lack of follicular DCs Can be partly overcome by transgenic TNFα Langerhans cells present, migration intact Increased B16F10 melanoma growth Normal NK numbers but impaired recruitment

Alexopoulou et al., 1998 Stoitzner et al., 1999 Fu et al., 1997 Ito et al., 1999b

Lymphotoxin β

Targeted disruption

Complete lack of follicular DCs Disorganized splenic architecture Absent Peyer’s patches and peripheral lymph nodes (some mesenteric and cervical nodes) Low IgA, low-affinity maturation of IgG Impaired antiviral responses Effects mediated via TNF-R1 and LTβR

Koni et al., 1997 Pasparakis et al., 1997 Fütterer et al., 1998 Alimzhanov et al., 1997 Berger et al., 1999

Lymphotoxin β receptor

Targeted disruption

Absent follicular DCs, Peyer’s patches, all lymph nodes

Fütterer et al., 1998 Endres et al., 1999

IL-4

Targeted disruption

DCs present in lymph nodes but impaired response to herpes simplex virus, lower la expression

El-Ghorr and Norval, 1999

TGFβ1

Targeted disruption of ligand or type II receptor

Absent Langerhans cells CD11c DCs present in lymph nodes Defect is due to effect of TGFβ1 on Langerhans cells, no effect on marrow progenitors

Borkowski et al., 1996 Borkowski et al., 1997

Flt-3L

Targeted disruption

Marked reduction in DCs

McKenna et al., 2000

M-CSF/CSF-1

Natural M-CSF mutation (op/op)

Normal numbers in skin, spleen, lymph nodes, Peyer’s patches. No information on function

Witmer-Pack et al., 1993 continued

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Cytokine

Type of study

Comments

References

No direct data regarding DCs. Mice capable of producing IgG2a, suggesting intact DC/B cell crosstalk

McKenzie et al., 1998

Absent lymph nodes and Peyer’s patches

Cao et al., 1995

IL-12

p40 disruption

DCs present, unable to stimulate IFNγ production, not correctable by IL-18

Stoll et al., 1998

IL-10

Targeted disruption

Increased Langerhans cell migration in response to FITC contact sensitization

Wang et al., 1999

50% increase in number of blood DCs

Vremec et al., 1997

Retroviral or plasmid transfection of DC or tumor (cell lines or fresh tumor samples)

Tumor expression of GM-CSF leads to expansion of immature DCs that may suppress CD8 T-cell function in some models but enhance it in others Human GM-CSF-transduced renal cell cancer or melanoma vaccines induce DC infiltration Transgenic GM-CSF expression in DCs enhances antigen presentation

Bronte et al., 1999

cIL-10

Retroviral gene transfer into tumor cells

Inhibition of effect of GM-CSF on accumulation of DCs within tumor

Qin et al., 1997

vIL-10

Retroviral transfection of DCs

Inhibition of maturation Impaired T-cell responses Impaired alloresponses

Takayama et al., 1998

MCP-1

Transgenic expression in epidermis

No spontaneous skin inflammation Exaggerated contact hypersensitivity response Increase in Langerhans cells

Nakamura et al., 1995

Subcutaneous

Marked increases in circulating and tissue DCs Enhanced antitumor immunity

Maraskovsky et al., 1996 Borges et al., 1999

GM-CSF

Subcutaneous Intradermal

Expansion of DC numbers in spleen Induction of migration of Langerhans cells from epidermis to dermis with functional maturation

Daro E et al., 2000 Smith et al., 1998

IL-1β

Intradermal

Enhancement of MHC class II expression and T-cell stimulation by Langerhans cells Induction of migration to draining nodes

Enk et al., 1993a Cumberbatch et al., 1997

TNFα

Intradermal

Langerhans cells induced to migrate to draining nodes

Cumberbatch and Kimber, 1992 Cumberbatch et al., 1997

G-CSF

Subcutaneous

Six-fold increase in circulating DCs

Romani et al., 1996

Transgenic or gene transfer studies GM-CSF Transgenic GM-CSF

Exogenous cytokine Flt-3L

Hanada et al., 1996 Lee et al., 1997 Simons et al., 1997 Soiffer et al., 1998 Curiel-Lewandrowski et al., 1999

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Targeted disruption of γc chain

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TABLE 14.2 (Continued )

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and function. These mice also lack Peyer’s patches and peripheral lymph nodes, although some mesenteric and cervical lymph nodes persist in LTβ and LTβR knockout mice but not LTα knockout mice (Koni et al., 1997). This discrepancy in the phenotype of lymph node deficiency might be due to the presence of another receptor capable of transducing trimeric LTα3 signals. Mice deficient in LT have severely perturbed immune systems. They have high levels of IgM, low levels of serum IgA and low-affinity maturation of IgG (Fu et al., 1997). They have impaired antiviral responses (Berger et al., 1999). Antitumor responses are also impaired (Ito et al., 1999). NK numbers are intact but NK recruitment into tumor sites is reduced (Ito et al., 1999).

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TNFα and LT form trimeric complexes that bind to their respective receptors. LTα homotrimers (LTα3) or TNF homotrimers signal via the TNF p55 and p75 subunits (Ware et al., 1995). LTα2β1 trimers can bind the TNF p55/p75 complex, while the LTβ receptor recognizes only LTα1β2 trimers and not LTα2β1, TNFα homotrimers or LTα3 (Crowe et al., 1994; Ware et al., 1995). Thus, the TNF receptor is not required for LTβ signaling. LTβR-deficient mice have a different phenotype to mice lacking LTα or LTβ, implying that other factors may also signal via LTβR (Fütterer et al., 1998). Therefore, the individual LT and TNF ligands, as well as the subunits of their respective receptors, probably have distinct and important roles in the control of FDC production.

TNFα Signaling via the TNFα receptor also appears to be important in the genesis of FDCs in a complex fashion. The development of splenic FDCs depends on expression of LTβ and TNFα by B cells and LTβR by marrow stromal cells (Matsumoto et al., 1997; Endres et al., 1999). Mice in which the TNFα gene has been disrupted show normal numbers of DCs in lymph nodes, Peyer’s patches and LCs in skin, although the FDC network is disorganized (Pasparakis et al., 1997; Stoitzner et al., 1999). LTα/ mice that are also transgenic for TNFα show partial correction of the LTα/ phenotype (Alexopoulou et al., 1998). Disruption of the p55 or p75 subunits of the TNFα receptor cause specific abnormalities. Inactivation of p55 leads to absent FDCs and Peyer’s patches but normal LC numbers. This is due to effects on cells outside the bone marrow, since reconstitution of these animals with wild-type marrow does not reverse the defect (Matsumoto et al., 1997). Conversely, when LTα/ mice are reconstituted with bone marrow cells lacking p55, FDC organization is restored, again showing that expression of p55 by FDC precursors in the marrow is not required. LCs are present when p75 is inactivated but their migration is impaired. These different phenotypes may be explained by the complex pathways of LT signaling. Both

IL-4 IL-4 is important in vitro in the control of DC differentiation from monocytes (Sallusto and Lanzavecchia, 1994) or CD34 progenitor cells (Luft et al., 1998a). Mice in which the IL-4 gene has been disrupted show normal numbers of DCs in lymph nodes but have impaired responses to herpes simplex virus infection and lower expression of Ia on DC (El-Ghorr and Norval, 1999). Disruption of the common gamma chain required for signaling via the receptors for IL-2, IL-4, IL-7, IL-9 and IL-15 gives a much more severe phenotype, with absent lymph nodes and Peyer’s patches (Cao et al., 1995). GM-CSF It is somewhat surprising that certain other cytokines do not appear to be essential for DC development in vivo. GM-CSF, thought to be an important factor in the development of myeloid DCs, is redundant: deletion of the genes for the ligand or for the signaling component of the receptor do not significantly decrease DC production. DCs from GM-CSF knockout mice are present in normal numbers and are capable of inducing normal allostimulatory responses. However, in vivo responses to xenoantigens are abnormal. When these mice are immunized with

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keyhole limpet hemocyanin (KLH), anti-KLHIgG generation is impaired. There is a specific defect in IgG2a production suggesting a failure of production of TH1 cytokines, implying impaired function of DCs in vivo at the level of B cell costimulation. In addition, anti-KLH CD4 T-cell proliferative responses are impaired in these animals. These knockout studies may also provide clues to other DC developmental pathways, such as those for lymphoid DCs (Vremec et al., 1997). Therefore, although gross measures of DC numbers and function are intact, more subtle defects in DC function can be detected. IL-3 and IL-5 share a common receptor subunit with GM-CSF. Disruption of this common β chain receptor subunit blocks the activity of all three cytokines. In this setting, absolute DC numbers are reduced but DCs are still present (Vremec et al., 1997), implying that other signaling pathways can replace signaling via the common β chain. Conversely, transgenic expression of GM-CSF in tumor cells leads to DC expansion (Hanada et al., 1996; Bronte et al., 1999), although antigen presentation by these DCs may be impaired (Bronte et al., 1999). Similarly, systemic administration of a chemically stable form of recombinant GM-CSF increases the numbers of myeloid-related DCs in vivo (Daro et al., 2000), whilst overexpression of GM-CSF within the DCs themselves leads to enhanced antigen presentation (Curiel-Lewandrowski et al., 1999). Others Other factors also seem to play a role in determining numbers of DCs and their precursors, although the phenotype of the resulting DCs may not be appreciably different. Deletion of the gene for Flt-3 ligand causes a marked reduction in splenic DC numbers (McKenna et al., 2000). Conversely, exogenous administration of certain cytokines (e.g. Flt-3L, G-CSF or GM-CSF) can lead to striking increases in DC numbers in the spleen or peripheral blood (Maraskovsky et al., 1996; Borges et al., 1999; Daro E et al., 2000). It is also possible to inhibit DC function and perhaps to render them more tolerogenic. The

immunosuppressive cytokine viral IL-10 (vIL10), when transfected into DCs, can cause inhibition of DC maturation, inhibition of T-cell responses to DC stimulation, and impaired alloresponsiveness (Takayama et al., 1998). Disruption of the p40 chain of IL-12 does not interfere with the production of DCs, however they are unable to stimulate IFNγ production by T cells, a defect that cannot be reversed with IL-18 (Stoll et al., 1998). Other factors such as MCP-1 may also enhance the reactivity of DC subtypes such as LCs (Nakamura et al., 1995). In conclusion, the effects of cytokines on DC progenitors and on differentiated DCs are complex. Some factors have been shown to be critical in the production or functional activity of certain DC subsets. Other factors, thought to be important from in vitro studies, appear to be redundant in vivo.

CYTOKINE REGULATION OF DC MATURATION AND ACTIVATION Mobilization of an immune response involves recruitment of innate and subsequently specific effectors. In response to ‘danger’ signals, tissues release proinflammatory cytokines or other mediators that can mediate DC maturation and migration. These mature APCs can induce quiescent T cells to become activated (Matzinger, 1994). The cytokine response to pathogens is best characterized by TNFα and IL1β which are released during cellular damage. Both cytokines are required for the maturation of murine LCs (Cumberbatch et al., 1997). Contact allergens are known also to induce IL-1β release by LCs (Enk et al., 1993a) and TNFα release by keratinocytes (Holliday et al., 1997). LPS is a potent direct activator of McDCs, an effect mediated by surface or soluble CD14 (Verhasselt et al., 1997). Furthermore, TNFα is released in response to LPS from T lymphocytes (Del Prete et al., 1989), monocytes, macrophages (Kornbluth and Edgington, 1986) and DCs (Avice et al., 1999). A number of proinflammatory molecules can

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synergize with TNFα and IL-1β and potentiate DC maturation in vitro. Such factors include IL-6 (Jonuleit et al., 1997), IFNα (Luft et al., 1998b) (see below), PGE2 (Rieser et al., 1997), monocyte-conditioned medium (which contains many proinflammatory cytokines) (Reddy et al., 1997) and free oxygen radicals (Rutault et al., 1999). Particulate antigens crosslinking mannose receptors (Shibata et al., 1997) and Igcomplexed antigen crosslinking Fc receptors (Regnault et al., 1999) are also capable of stimulating DC maturation. The notion that some of these processes may have an autocrine or paracrine component is supported by the observation that IFNα (Milone and FitzgeraldBocarsly, 1998) and TNFα (Polat et al., 1993) secretion is increased in response to mannose receptor crosslinking. Other triggers for DC maturation may be independent of cytokine signals such as engagement of cell adhesion molecules/complement receptors such as CR3 and CR4 (CD11b and CD 11c), CpG DNA and necrotic cells. Clearly the process of DC maturation is complex and the net interaction of an ensemble of factors contributes to different aspects of maturation and activation. For example, IFNγ was shown to inhibit the expression of CD80 on Langerhans cells (Ozawa et al., 1996) and counteract IL-4 effects on DC differentiation (Lardon et al., 1997), whilst enhancing CD40L-induced IL-12 secretion (Kalinski et al., 1999). PGE2 and TNFα were reported to induce IL-12 secretion (Rieser et al., 1997), however, other groups reported a shift of PGE2-activated DCs towards the induction of TH2-type immune responses due to the reduced capacity of PGE2-activated cells to secrete IL-12 (Kalinski et al., 1998). This highlights the importance of microenvironmental signals that provide a ‘context’ at the time and place of antigen exposure, which in turn shapes the outcome of the immune response.

Effects of type I interferons on DC maturation

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order to investigate cytokines involved in DC maturation. In these GM-CSF, TNFα and IL-4 driven cultures, immature DCs develop by day 14 and spontaneously mature if cultured for a further 14 days (Luft et al., 1998a). Of 18 cytokines tested in this system, only type I IFNs (α or β) were capable of accelerating maturation, so that within 3 days a majority of the large-sized cells in culture expressed CD80, CD83, CD86 and started to downregulate CD1a and CD11b (Luft et al., 1998b). This effect of IFNs was dependent on TNFα being present and was concentration-dependent in a range between 10 and 1000 U/mL for three different type I IFNs (α2a, α8 and β). Consistent with the phenotypic changes, DCs exposed to IFNα showed increased T-cell stimulatory capacity (Luft et al., 1998b). By further investigating the effects of type I interferons, we demonstrated enhanced activity on the maturation of monocyte-derived DCs in the presence of TNFα and PGE2 (T. Luft, submitted) as well as enhancing CD40L-induced IL-12p70 secretion by monocyte-derived DCs (McDC) (T. Luft, submitted). In addition, type I interferons have recently been shown by others to induce the maturation of murine DCs as well as having adjuvant effects on the priming of immune responses in vivo (Gallucci, 1999). These results suggest that type I IFNs might enhance the effect of TNFα on DCs in vivo. Therefore, in addition to their multiple immunomodulatory effects during viral infections, type I interferons may also regulate immune responses at the level of the antigenpresenting cell. This may help to explain the autoimmune phenomena associated with the use of IFNα in hepatitis and cancer patients (Gisslinger et al., 1992). It may also provide an additional mechanism for the anticancer effects of IFNα in a variety of tumors (Kirkwood, 1998). While IFNs are being used as nonspecific immunotherapy for cancer and as antiviral therapy, this effect on DC function suggests a further possible clinical application for IFNα as a vaccine adjuvant.

We have used a novel serum-free culture system to produce DCs from CD34 progenitor cells in DENDRITIC CELL BIOLOGY

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Cytokines inhibiting DC differentiation and maturation The study of culture conditions to produce DCs from precursors has provided insight into cytokine environments that interfere with DC differentiation. A number of inhibitory cytokines acting during early phases of DC differentiation have been described, including vascular endothelial growth factor (VEGF) (Gabrilovich et al., 1996), IL-10 (Allavena et al., 1998), IL-6 and M-CSF (Menetrier-Caux et al., 1998), and PGE2 (Kalinski et al., 1997). IL-10 and PGE2 also impaired or biased the maturation of immature DCs (Enk et al., 1993b; Kalinski et al., 1997), and a similar effect was reported for TGFβ1 (Geissmann et al., 1999; King et al., 1998). In addition, exposure of DCs to TGFβ1, PGE2 or IL-10 deviated the immune system toward a TH2-type response (Kalinski et al., 1997; King et al., 1998). These factors can also be secreted by tumors and may therefore contribute to our understanding of how established tumors evade immune responses.

Cytokines which activate DCs Although cytokines such as TNFα, IL-1β and IFNα are potent inducers of DC maturation they are not sufficient to induce secretion of cytokines such as IL-12, which in turn is an important inducer of IFNγ and thus of the TH1type immune responses. Rather, IL-12 is induced by CD40L, a TNF family member expressed on activated T helper cells (Koch et al., 1996) or by pathogens and their derivatives (e.g. LPS, CpG-DNA motives, dsRNA) (Koch et al., 1996; Henderson et al., 1997; Jakob et al., 1998; Saurwein-Teissl et al., 1998; Sparwasser et al., 1998; von Stebut et al., 1998; Marriott et al., 1999). DCs encounter CD40L after maturation and migration into lymph nodes, or by interacting with pre-primed T helper cells or non-T cells (activated platelets, inflamed endothelium) at the site of infection, so the final activation of DCs, characterized by secretion of IL-12 can be regarded as a separate stage of development. CD40L crosslinking alone is suboptimal for

induction of IL-12 secretion. Rather, additional signals produced by T cells during cognate interactions with DCs, such as IFNγ (Snijders et al., 1998) or IL-4 (Hochrein et al., 2000) are essential to enhance CD40L induced IL-12 secretion. A role for IFNγ in potentiating IL-12 secretion is substantiated by the finding that IFNγ can prime the promoter of IL-12 (Ma et al., 1996). However, IFNγ−deficient mice are capable of producing unimpaired levels of functional IL-12, suggesting that there is also an alternative, IFNγindependent mechanism for IL-12 induction (Scharton-Kersten et al., 1996). In addition to and independent of the CD40L pathway, pathogens or their derivatives such as Leishmania (von Stebut et al., 1998), whole influenza virus (Saurwein-Teissl et al., 1998) and living bacteria (Henderson et al., 1997; Marriott et al., 1999) or their derivatives (e.g. LPS (Verhasselt et al., 1997), dsRNA (Cella et al., 1999), bacterial DNA motifs (CpG) (Jakob et al., 1998; Sparwasser et al., 1998)) are capable of directly inducing the secretion of IL-12. Therefore, two mechanisms exist to induce full DC activation: the T cell-dependent, CD40L pathway and a T cell-independent pathway induced by pathogens.

Interaction of T cell and DC: mutual activation The interaction between DCs and T lymphocytes induces signals leading to activation in both cell types. The contact area between DC and T cell forms a supramolecular aggregation cluster (SMAC) composed of a peripheral region rich in cell adhesion molecules and a central region involving TCR/MHC molecules (Monks et al., 1998). The contact between DC and T cell is initiated by cell adhesion molecules, and is further stabilized by co-stimulatory molecules (CD80/CD86) interacting with CD28 on T cells, TCR–MHC interactions, CD4/CD8–MHC(α3) interactions (Shaw and Dustin, 1997). These intimate contacts allow prolonged engagement, which is an important requirement for full Tcell activation. It has been suggested that up to 4 hours ofTCR engagement is necessary to induce

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IL-2 secretion in naïve T cells (Shaw and Dustin, 1997). Further, the tight contacts between DCs and T cells would allow efficient and specific transfer of small quantities of cytokines required for T-cell activation and the priming of the T cell’s cytokine repertoire. This is a critical mechanism for maintaining specificity given that the lymph node can simultaneously host multiple immune responses involving T helper cells of different cytokine repertoires (TH1 and TH2). Indeed, it has been demonstrated that antibody responses against two different strains of viruses (each separately inducing either IgA or IgG) can coincide in the same lymph node without affecting each other. Since the two viruses infected different levels of the airway epithelium, different DCs would have presented their antigens. This indicates that opposing immune responses against different pathogens from different sites can simultaneously occur within the same lymphoid organ with each being regulated independently of the other (Sangster et al., 1997). DCs in turn receive activation signals from T cells by crosslinking CD40 with CD40L provided on the surface of activated T helper cells. It has recently been shown that these interactions involve DCs, T helper cells and CTLs forming immune regulatory loops (Stuhler et al., 1999). For CTL activation, T helper cells can be replaced if DCs were activated previously by CD40L (Ridge et al., 1998). This suggests that the main function of T helper cells at this stage is to activate DCs via CD40L. In response to activation with CD40L and IFNγ, McDCs secrete much higher levels of cytokines than would apparently be required during tightly controlled focal immune responses within the T-cell areas (Snijders et al., 1998). This paradox may reflect a distinct function of McDCs in the recruitment of the innate immune response at primary sites of inflammation rather than a specific role involving the priming of immune responses within lymphoid tissue. DCs are more potent stimulators of naïve Tcell proliferation than monocytes, macrophages or B lymphocytes (Van Voorhis et al., 1983). They are also the most potent presenters of soluble antigens to MHC class II-restricted T helper cells

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(Guéry et al., 1996). There are numerous publications suggesting that DCs might be directly involved in priming the cytokine repertoire of responding T-cell populations (Macatonia et al., 1995; Kalinski et al., 1997; Stumbles et al., 1998; Maldonado-Lopez et al., 1999; Pulendran et al., 1999). In particular, DC subpopulations (Stumbles et al., 1998) or DCs matured in the presence of factors such as IL-10 (Liu et al., 1998) or PGE2 (Kalinski et al., 1997) have been associated with TH2-biased T cell responses in vitro. However, little is known about the complex factors that condition DC maturation, the interaction of several DC subpopulations or T-cell subpopulations during an immune response in vivo. In addition to factors acting on DCs, cytokines that act directly on the priming of naïve T helper cells, such as type I interferons (Rogge et al., 1998) will also shape the developing immune response. As mentioned above, cytokines such as IFNα do not appear to perform nonidentical functions during murine and human immune responses (Cho et al., 1996; Rogge et al., 1998). Therefore, the understanding of how DCs are involved in the regulation and fine tuning of human immune responses should preferentially rely on the study of human immune functions.

CYTOKINE ADMINISTRATION: PRECLINICAL AND CLINICAL OBSERVATIONS As shown in Figure 14.1, many cytokines affect DC development from myeloid and lymphoid precursor cells. The definition and characterization of DC subsets is still evolving, so the precise role of these cytokines in the evolution of DC subsets remains unclear. Nonetheless, among these cytokines, key roles appear to be played by Flt-3 ligand, GM-CSF and IL-4. It is worth reemphasizing that it is difficult to ascribe an in vivo role for these agents on the basis of in vitro observations, or the effects of pharmacological doses of cytokines administered to experimental animals or to human patients in the clinic. These cytokines are discussed here in greater detail:

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Flt-3 ligand Flt-3 ligand as a DC growth factor in mice Flt-3 ligand (Fms-like tyrosine kinase receptor 3 ligand) stimulates the proliferation of stem and progenitor cells through binding to the Flt-3 receptor, which is a type III receptor tyrosine kinase member of the PDGF family (Brasel et al., 1996; Lyman and Jacobsen, 1998). Although Flt3L is expressed by multiple cell types in many tissues, the receptor is restricted to hematopoietic progenitors, early B cells and myeloid cells (Rasko et al., 1995). Daily subcutaneous injections of mammalian cell-expressed recombinant human Flt-3L for 10 days resulted in a dramatic increase in the numbers of hematopoietic progenitors in the bone marrow, peripheral blood and spleen (Brasel et al., 1996). Examination of the effects of systemic Flt-3L on more mature, lineage-committed cells indicated that the Flt-3L-mediated expansion and mobilization of bone marrow progenitors was accompanied by an increase in myeloid lineage cells, early B lymphocytes and NK cells (data not shown), but no apparent effect on T lymphocytes, erythroid, or megakaryoctye lineage cells (Brasel et al., 1996). Closer examination of the tissues in Flt-3L-treated mice demonstrated that Flt-3L dramatically increases the number of DCs in several sites, including the spleen, lymph nodes, thymus, Peyer’s patch, as well as in the circulation, lungs and liver (Maraskovsky et al., 1996).

DC subsets mobilized by Flt-3L in mice The DC subsets generated in Flt-3L-treated mice resemble the corresponding DC subsets identified in normal mice (Maraskovsky et al., 1996; Pulendran et al., 1997). In particular, both the myeloid-related (CD11cCD11bbrightCD8α) and lymphoid-related (CD11cCD11bloCD8α) DC subsets were expanded following Flt-3L administration. These DCs expressed high levels of CD11c and MHC class II, as well as intermediate levels of CD40 and CD86 (Maraskovsky et al., 1996; Pulendran et al., 1997). The majority of

CD11cCD11bloCD8α subset cells expressed high levels of DEC-205, a marker constitutively expressed on the lymphoid-related DC subset residing in the T-cell areas of lymphoid tissues. In contrast, CD11cCD11bloCD8α lacked DEC205 expression but did express myeloid markers (F4-80 and 33D1) and were located in the marginal zones of the spleen (Pulendran et al., 1997). Both DC subsets were capable of antigen uptake and presentation and could efficiently prime antigen-specific CD4 and CD8 T cells efficiently in vitro (Daro et al., 2000). However, the lymphoid subset can be induced to secrete much higher levels of IL-12 than the myeloidrelated population (Pulendran et al., 1997). Flt-3L as a DC growth factor in humans DCs represent a minor fraction (1%) of peripheral blood mononuclear cells (1%) (PBMCs) in humans and can be distinguished from other mature cell lineages by their characteristic dendritic morphology, the lack of surface expression of CD3, CD14, CD19 and CD56, and high expression of CD1b/c, CD4, CD11c, CD33 and HLADR. In addition, a DC precursor subset lacking CD11c expression and expressing high levels of surface IL-3Rα (CD11c DCs) and previously referred to as the ‘plasmacytoid T cell’ has been identified in lymphoid tissue and peripheral blood (Grouard et al., 1997; Olweus et al., 1997). The effect of Flt-3L treatment on expansion of DC has been examined in a randomized, placebo controlled, double-blind study. Twenty healthy human volunteers received daily subcutaneous doses of either placebo or Flt-3L at five dose levels (10, 25, 50, 75 or 100 µg/kg/day) for 14 consecutive days. Each dose group consisted of three Flt-3L-treated individuals and one placebo control. Samples of peripheral blood (PB) were collected every 2 days for 21 days and analyzed by flow cytometry for changes in the distribution of various leukocyte populations. Flt-3L treatments were welltolerated by all subjects at all doses tested compared with the placebo controls, and resulted in increased numbers of white blood cells (WBCs), PBMCs and CD14 monocytes, but not lympho-

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cytes (Maraskovsky et al., 2000). Flt-3L expanded the numbers of CD11cCD14 DCs at all doses tested. These cells were rare in the blood of placebo-treated individuals but were increased, on average, to 14% of PBMCs following Flt-3L administration, a 40-fold increase in the numbers of circulating DCs (Maraskovsky et al., 2000). As was reported in mice, Flt-3L-generated human DCs were phenotypically immature, expressing low to negligible levels of CD80, CD83 and CD86 and intermediate levels of HLA-DR (Maraskovsky et al., 2000) which was rapidly upregulated following in vitro culture (E. Maraskovsky, unpublished observations). However, they were efficient stimulators of naïve T-cell proliferation (Maraskovsky et al., 2000) indicating that they were functionally competent antigen-presenting cells. Finally, Flt-3L was also able to increase the numbers of CD11cIL-3Rα DCs representing the ‘plasmacytoid’ DC subpopulation. This indicates that, as in mice, Flt-3L has the unique capacity of expanding the numbers of multiple DC subsets in vivo but not significantly altering their maturation state. It now remains to be established whether this dramatic increase in DC numbers in vivo can enhance immune responses to a variety of candidate vaccine antigens, as has been observed in mice.

Granulocyte-macrophage colony stimulating factor (GM-CSF) GM-CSF promotes the expansion and differentiation of hematopoietic progenitor cells, particularly cells of the granulocytic and monocyte/ macrophage lineage (Metcalf, 1989). It has more recently been shown to have potent effects on antigen-presenting cells, including dendritic cells (DCs) and Langerhans cells (Caux et al., 1992; Inaba et al., 1993). In vitro DCs can be generated in GM-CSF together with TNFα and/ or IL-4 (Caux et al., 1992; Inaba et al., 1993) although these cells are relatively immature and require further stimulation in order to fully mature (Sallusto and Lanzavecchia, 1994). In animal gene transfer models, GM-CSF gene transfer into poorly immunogenic tumors led to rejection

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of those tumors (Tepper and Mule, 1994), an observation which stimulated further evaluation of the role of GM-CSF as an anticancer agent, particularly in its role as regulator of DC action. Subsequent clinical studies have shown that GMCSF gene transfer can significantly enhance immunologic responses against human cancers, particularly melanoma in humans (Ellem et al., 1997; Soiffer et al., 1998). Nasi et al. (1999) documented an increase in dermal and tumorinfiltrating DCs following GM-CSF injection, although no clinical benefit was documented. In contrast Si et al. (1996) reported that direct injection of GM-CSF into subcutaneous melanoma metastases caused regression of injected and noninjected tumor deposits (Si et al., 1996). Animal studies have indicated that GM-CSF enhances immune responses to protein vaccination with an efficacy comparable to that of complete Freund’s adjuvant, and can also induce responses against otherwise nonimmunogenic peptides (Disis et al., 1996). As a consequence of such studies, GM-CSF has been used as a cytokine ‘adjuvant’ in conjunction with vaccination. Recent studies using peptide vaccination in cancer patients has confirmed that GM-CSF can enhance immune responses to peptide antigens in patients with breast cancer (Disis et al., 1999) and melanoma (Jager et al., 1996). In this latter study, clinical responses were seen in HLA-A2 patients with metastatic melanoma treated with systemic GM-CSF and intradermal peptides derived from the melanoma differentiation antigens melanA/MART-1, tyrosinase and gp100 (Jager et al., 1996). The optimal method for administering GM-CSF is unknown since the administration of GM-CSF injections at a distant site (thereby acting as a systemic cytokine adjuvant) appeared to enhance immune effects qualitatively and quantitatively (Jager et al., 1996). GM-CSF at lower doses co-administered with peptides also demonstrated induction of antigen-specific immune responses (Disis et al., 1999). It therefore seems likely that the effects were to both enhance DC function systemically (perhaps by acting on precursor populations) and locally, though effects on LCs and/or dermal DCs.

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Interleukin 4 Since the combination of GM-CSF and IL-4 was shown to be effective for DC generation in vitro, this combination was tested in patients with advanced cancer. The patient cohort that received GM-CSF alone (at a dose of 2.5 µg/kg) experienced an increase in CD14 cells. In contrast the addition of IL-4 at doses ranging from 0.5 to 6.0 µg/kg resulted in downregulation of CD14 and the upregulation of HLA-DR and CD11c, consistent with an increase in the antigen-presenting cell phenotype. These cells were functionally more effective in endocytic assays and T-cell stimulation in an MLR. In addition, there was an increase in CD83-positive DCs in peripheral blood. Several clinical responses were seen, suggesting that this combination of cytokines may not only increase APCs in situ, but may mediate anticancer responses under some circumstances (Roth et al., 2000). Several clinical trials with IL-4 alone have also been reported. Occasional transient clinical responses have also been reported, although these studies were generally carried out in the era before the effect of IL-4 on DCs was appreciated. As a result there are no comprehensive reports of effects on DCs in these patients (Davis et al., 2000).

CONCLUSION In summary, the clinical application of cytokines aimed at manipulating DC activity in vivo is only just beginning. It seems likely that DC numbers and function can be increased, however this is a potentially complex task. Understanding how DCs respond to their environment and to pathogens and how their functional state regulates the developing immune response is critical to their appropriate clinical use. There are many DC subsets and states of maturation currently being defined – and DCs in each of these states potentially have very different functions. Further, the regulators of these subsets are produced in tightly controlled microenvironmental niches. The clinical administration of these

cytokines is therefore crude and nonphysiological. Nonetheless, carefully conducted clinical trials supported with detailed laboratory analysis should lead to improved methods for enhancing (or inhibiting) immune responses and resultant therapeutic benefit.

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15 Dendritic cells and chemokines Silvano Sozzani, Paola Allavena and Alberto Mantovani Istituto di Ricerche Farmacologiche Mario Negri, Milan, Italy

Quali colombe dal desio chiamate con l’ali alzate e ferme al dolce nido seguon per l’aere, dal voler portate . . . Dante, Inferno As mating doves that love calls to their nest glide through the air with motionless raised wings borne by the sweet desire that fills each breast – just so . . . Translation by John Ciardi

Inflammatory signals exert a dual influence on the migration of DCs. On the one hand they increase recruitment of immature DCs at sites of infection or inflammation; on the other hand, they induce the maturation of DCs and promote their egress from tissues towards secondary lymphoid tissues (Macpherson et al., 1995; Roake et al., 1995; De Smedt et al., 1996). Chemotactic factors and, in particular, the chemotactic cytokines called chemokines, play a crucial role in regulating the trafficking of various leukocytes, including DCs, in health and disease. Here we will discuss how DCs are both a target and a major source of chemokines, and how in vitro and in vivo results have led to the formulation of a paradigm for how these molecules direct the traffic of DCs. A review of chemokines is beyond the scope of this chapter; the reader is referred to Rollins (1997), Luster (1998), Mantovani (1999a, 1999b) for an introduction to the chemokine world.

INTRODUCTION Migration is an essential component of the function of dendritic cells (DCs). DCs need to localize in tissues to capture antigen and to travel to lymphoid organs, where they fulfill their antigen-presenting function in an appropriate cellular and structural context. Different pathways of DC migration in vivo have been defined, as discussed elsewhere in this book. For instance, in vivo recruitment of DCs was first observed at sites of allergy and transplanted tissues (Austyn and Larsen, 1990; Banchereau and Steinman, 1998). Inhaled pathogens induce a very rapid mobilization of DCs in the airway epithelium (McWilliam et al., 1994, 1996). After i.v. injection of inert particles, cells with DC characteristics are first recruited to the hepatic sinusoid and then translocate to the hepatic lymph (Matsuno et al., 1996).

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CHEMOKINE RECEPTORS AND RESPONSIVENESS IN RESTING, IMMATURE DCs: RECRUITMENT AND POSITIONING IN NONLYMPHOID TISSUES DCs are capable of in vitro migration under a variety of in vitro conditions. They also can cross endothelial cell monolayers after interacting with the luminal side of endothelial cells as well as migrate basal to apical (reverse transmigration) (D’Amico et al., 1998; Randolph et al., 1998). The latter in vitro assay mimicks the process whereby DCs leave peripheral tissues by entering blood or lymphatic vessels to travel to secondary lymphoid organs. DCs respond to chemoattractants representative of different classes of chemotactic agonists, including lipid attractants (platelet-activating factor, PAF), formyl peptides (representative of bacterial proteins) and C5a generated by the complement cascade (Sozzani et al., 1995, 1997a, 1997b). Moreover, they respond to a distinct set of chemokines (Sozzani et al., 1995). DCs obtained from circulating monocytes by culture with GM-CSF and IL-4 or IL-13 were used in the original description (Sozzani et al., 1995) of chemokine activity on this cell population, but subsequent studies have extended the analysis to cells obtained from CD34 hematopoietic precursors and to Langerhans cells (LCs). Chemokines have been classified in constitutively expressed and inflammatory or inducible molecules, though these two realms overlap to a considerable extent (Rollins, 1997; Luster, 1998; Mantovani, 1999a, 1999b). DCs express receptors for and respond to both constitutive and inducible chemokines (Table 15.1). The prototypic constitutively expressed chemokine SDF-1 is active on DCs, which express the cognate receptor CXCR4 (Sozzani et al., 1997b). Similarly, MDC, originally described as a constitutively expressed chemokine (Godiska et al., 1997), is a potent attractant for DCs, being 100-fold more active than it is on monocytes (Godiska et al., 1997). Accordingly, DCs express CCR4 (Sozzani et al., 1999). It should be noted

TABLE 15.1 Chemokine receptors and responsiveness in in vitro differentiated DCs Receptors

Immature DCs

Mature DCs

Ligands

CXCR4





SDF-1α, SDF-1β

CCR1





MIP-1α, RANTES MIP-5, MCP-2, -3

CCR2





MCP-1, -2, -3, -4

CCR4





TARC, MDC

CCR5





MIP-1α, MIP-1β, RANTES, MCP-2

CCR6

a



MIP-3α

CCR7





MIP-3β, SLC

fMLPR





fMLP

PAFR





PAF

C5aR





C5a

Expressed only in CD34 precursor-derived DCs and LCs. a

that MDC is also produced in a regulated way by monocytes, T cells and DCs themselves (see below). The responsiveness of immature DCs to constitutively expressed chemokines is probably important for their localization and positioning in tissues under normal conditions. Immature DCs, generated in vitro from monocytes, express a unique repertoire of inflammatory chemokine receptors (CCR1, CCR5, CCR6) (Sozzani et al., 1999). These receptors bind a pattern of ‘inflammatory’ chemokines, including RANTES, MCP-3, MIP-1α, MIP-1β, MIP-5 (Table 15.1). Purified circulating DCs were also found to express CCR1, CCR2, CCR3, CCR5 and CXCR4, by PCR analysis (Ayehnie et al., 1997). Interestingly, plasmacytoid DCs (CD11c) express CXCR3, while monocyte DCs and blood DCs (CD11c) do not (Cella et al., 1999). LCs purified from skin or generated in vitro from CD34 precursors are characterized by the

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expression of CCR6, the receptor for MIP-3α, also known as LARC or exodus, in addition to the receptors expressed by monocyte DCs (Power et al., 1997; Greaves et al., 1997). Thus, the presence of CCR6 differentiates between monocyte DCs and CD34 DCs, and the latter cells may be representative, in this respect, of LCs. Monocyte DCs were also reported to express the orphan receptor chemR23, although the relevance of this protein in DC migration is still unknown (Samson et al., 1998). The ability of precursor cells and immature DCs to migrate in response to a pattern of chemotactic signals is probably relevant for the accumulation of immature DCs into nonlymphoid tissues in normal conditions and during the early phase of inflammation, when local production of chemokines is strongly induced. The specificity of MIP-3α for LCs, and its constitutive production by keratinocytes may represent one of the homing signals for the positioning of these cells in the epidermis (Charbonnier et al., 1999). Defensins have recently been shown to bind CCR6 and have agonist activity (Yang et al., 1999). These molecules, produced by phagocytes and other cells, may therefore attract immature Langerhanstype DCs at sites of infection. The role of MCP-1 and IL-8, two chemokines that bind CCR2 and CXCR1/2, respectively, is at the moment uncertain. These receptors are expressed b immature DCs but contradictory results were reported in terms of their biological activity (Xu et al., 1996; Sozzani et al., 1997b; Rubbert et al., 1998). A murine DC cell line and DCs differentiated from mouse bone marrow CD34 precursors showed a pattern of responsiveness to chemokines very similar to that observed with human cells (Foti et al., 1999; Vecchi et al., 1999). The only major difference between mouse and human is the responsiveness to MCP-1. Human DCs express CCR2, the MCP-1 receptor, and bind labeled MCP-1, but show little or no response in chemotaxis assays (Sozzani et al., 1995, 1997b; Xu et al., 1996). In contrast, mouse DCs generated from bone marrow precursors readily migrate in response to mouse MCP-1 (Vecchi et al., 1999). It is of note that transgenic mice overexpressing

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MCP-1, under the keratin promoter, have local accumulation of cells with DC morphology in the basal layer of the epidermis (Nakamura et al., 1995). This accumulation may result from direct recruitment of DCs or migration of monocytes and local differentiation to DCs. Chemokine receptor expression in DCs has been investigated recently in relation to HIV infection, as these cells may represent an important port of entry of the virus and a vehicle for HIV-1 transmission (Cameron et al., 1992; Graziosi and Pantaleo, 1998). In vitro cultured DCs and circulating blood DCs express both CCR5 and CXCR4, the two main coreceptors for M-tropic (R5) and T-tropic (X4) strains of the virus, respectively (Granelli-Piperno et al., 1996; Sozzani et al., 1997b; Ayehnie et al., 1997). Immature DCs selectively replicate R5 HIV (Granelli-Piperno et al., 1998), and immature LCs express only CCR5 and select R5 HIV strains in vivo (Zaitseva et al., 1997; Reece et al., 1998). LCs and immature DCs are present at mucosal sites of virus transmission and selective entry and replication of R5 HIV could explain the restriction of virus phenotype during virus infection before seroconversion.

MIGRATION TO LYMPHOID ORGANS: THE SWITCH PARADIGM Maturation of DCs can be induced in vitro by a variety of factors, in particular by LPS and the inflammatory cytokines TNF and IL-1. Engagement of CD40 and TRANCE/RANK with their respective ligands expressed on activated T cells also leads to maturation and activation of DCs (Cella et al., 1997; Banchereau and Steinman, 1998). In vivo, antigen uptake and the exposure to immune and inflammatory agonists cause the rapid mobilization of DCs from peripheral nonlymphoid tissues and activate the maturation process in these cells (Austyn, 1996; Banchereau and Steinman, 1998). Previous studies performed with mononuclear phagocytes and lymphocytes have shown that cell activation is often associated with a drastic

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change of the repertoire of expressed chemokine receptors. In particular, proinflammatory agonists (e.g. LPS, IL-1 and TNF) induce a downmodulation of certain CC chemokine receptors in monocytes, while anti-inflammatory signals, such as IL-10, have an opposite effect (Sica et al., 1997; Tangirala et al., 1997; Xu et al., 1997; Sozzani et al., 1998b; Penton-Rol et al., 1998). Recently, it was shown that signals that induce maturation of DCs also affect their migratory ability. Exposure of DCs to LPS, IL-1 and TNF, or culture in the presence of CD40 ligand, induced a rapid (1 hour) inhibition of chemotactic response to MIP-1α, MIP-1β, MIP-3α, RANTES, MCP-3 and fMLP (Dieu et al., 1998; Lin et al., 1998; Sallusto et al., 1998; Sozzani et al., 1998a; Yanagihara et al., 1998; Foti et al., 1999). Receptor desensitization by endogenously produced chemokines could be responsible for this effect, however, the reported desensitization to C5a and fMLP, two chemotactic factors that cannot be produced by activated DCs, implicates additional agonist-independent mechanisms (Sozzani et al., 1998a; Sallusto et al., 1998). As previously observed in phagocytes (Lloyd et al., 1995; Sica et al., 1997), inhibition of chemotaxis was followed, with a slower kinetics, by the reduction of membrane receptors and by the downregulation of mRNA receptor expression (Dieu et al., 1998; Granelli-Piperno et al., 1998; Sallusto et al., 1998; Sozzani et al., 1998a). Concomitantly, the expression of CCR7 and the migration to their constitutive ligands, MIP-3β and SLC, were strongly upregulated, with a maximal effect at 24 hours. Receptor expression and chemotactic response to other constitutive chemokines, such as SDF-1 (CXCR4) and MDC (CCR4), were not downregulated or increased during maturation of (Sallusto et al., 1998; Lin et al., 1998; Vecchi et al., 1999). Upregulation of CCR7 in DCs migrating to secondary lymphoid organs is a crucial event for the correct localization of these cells in T-cell areas. MIP-3β and SLC are specifically expressed in T cell-rich areas of tonsils, spleen and lymph nodes, where mature DCs home, to become interdigitating DCs (Dieu et al., 1998; Ngo et al., 1998; Willimann et al., 1998). Within the T-cell

area, SLC is expressed by stromal cells, as indicated by the severe reduction of expression in stromal cell-deficient mice, such as the lymphotoxin α/ (Ngo et al., 1999). SLC is also expressed in high endothelial venules, the site of entry of naïve T lymphocytes and DCs (Gunn et al., 1998). In contrast, MIP-3β is apparently mainly produced by interdigitating DCs (Ngo et al., 1998). The crucial role of SLC/MIP-3β and CCR7 is clearly reflected in mice deficient for these proteins. In mice homozygous for an autosomal recessive mutation, paucity of lymph node T cells (plt), naïve T cells fail to home to secondary lymphoid organs. The plt mutation is associated with a defective expression of SLC in lymphoid organs. DCs from these mice fail to accumulate in the spleen and T-cell areas of lymph nodes (Gunn et al., 1999). Similarly, CCR7/ mice showed a defective architecture of secondary lymphoid organs and a defective homing of DCs and lymphocytes (Förster et al., 1999). Overall, these findings provide a model for DC trafficking in which inflammatory chemokines acting through CCR1 and CCR5 may function as signals to localize DC precursors to nonlymphoid organs. After antigen uptake, immune/ inflammatory stimuli induce DC maturation and loss of responsiveness to the inducible cytokines locally produced. This unresponsiveness may play a permissive role, allowing DCs to leave peripheral tissues. Meanwhile, the slower upregulation of CCR7 prepares the cells to respond to MIP-3β and SLC (Gunn et al., 1998) expressed in lymphoid organs.

CHEMOKINE PRODUCTION BY DCs DCs also represent a source of chemokines in vitro and in vivo (Table 15.2). In vitro, immature DCs constitutively produce MDC, MIP-1γ and DC-CK1/PARC/MIP-4 (Mohamadzadeh et al., 1996; Adema et al., 1997; Godiska et al., 1997). Furthermore, thymic DCs selectively express TECK, a CC chemokine active on thymocytes, macrophages and DCs (Vicari et al., 1997).

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TABLE 15.2

The chemokine repertoire of DCs

Class

Chemokine

Constitutively expressed

MDC, MIP-1γ, DC-CK1, TECKa

Induced/ augmented

MDC, TARC, MCP-1, MIP-1α RANTES, IL-8, fractalkine, MIP-3β

a

Detected in thymic DCs.

Production of chemokines by DCs is strongly increased when these cells are induced to differentiate by proinflammatory stimuli or engagement of CD40. In vitro, mature DCs produce conspicuous amounts of MCP-1, MIP-1α, RANTES and IL-8 (Caux et al., 1994; Zhou and Tedder, 1995; Sallusto et al., 1998). Mature DCs also produce very high concentrations of MDC and TARC. In spite of the finding that interdigitating DCs produce MIP-3β in the lymph nodes (Dieu et al., 1998; Ngo et al., 1998; Tang and Cyster, 1999), this chemokine is not released in vitro, suggesting that additional signals present in situ might be important for its induction (unpublished data). In vivo, MDC was detected by in situ hybridization in mature DCs (Tang and Cyster, 1999) and the protein is strongly produced by CD83 cells in the skin of atopic dermatitis patients (Vulcano et al., 2001). Recently, fractalkine, a chemokine present in both a membrane-anchored and a soluble form, was found on the membrane of DCs generated in vitro (unpublished observations) or purified from spleen and epidermis, and produced in vivo by mature DCs (Kanazawa et al., 1999; Papadopoulos et al., 1999). In general, DCs are very efficient producers of chemokines, being able to release one or two logs higher concentrations than mononuclear leukocytes, endothelial cells and fibroblasts.

CHEMOKINES AND DC DIFFERENTIATION AND ACTIVATION The ability of chemokines to interfere with the in vitro cytokine-driven differentiation of monocytes into DCs was investigated in a limited

207

number of studies. Chemokines were added in the culture medium, together with GM-CSF and IL-13, at the beginning of the 7-day culture. Phenotypic analysis of the resulting population revealed no significant effect of RANTES and SDF-1 on DC differentiation as far as the expression of CD86 and CD83 was concerned. Moreover, MDC and MIP-3β did not affect terminal maturation of DCs induced by LPS when added at day 6 of differentiation together with LPS (Allavena and Sozzani). Chemokines were also tested for their ability to modulate cell functions typically related to DCs. Neither short (1 hour) nor long (24 hours) pretreatments of monocyte DCs with a number of chemokines, including MCP-3, MIP-1α, MIP-3β, RANTES, MDC and SDF-1, modified the ability of these cells to take up fluorescent labeled dextran (Sozzani et al., 1997b). Chemokine-treated DCs were also tested as APCs in MLR assays. No significant alteration of the immunostimulatory properties of DCs was observed with DCs pretreated with MCP-3, RANTES, MIP-1α, MIP-3β and SDF-1. Moreover, the addition of AOPRANTES and Met-RANTES in the MLR assay to antagonize CCR1, CCR3 and CCR5 did not modify the APC activity of DCs. These results are in contrast to those reported by Sato et al. (1999), which suggested that CC chemokines may play a role in the full maturation of DCs as functional APCs. These results indicate that chemokines, at least at the doses optimal for chemotaxis assay, have no relevant effect on DC differentiation from precursors and on their biological functions, other than cell migration.

CHEMOKINE RECRUITMENT OF DCs IN TUMORS A stromal reaction which includes leukocytes and mesenchymal cells characterizes neoplastic tissues (Mantovani, 1994). Cells with DC characteristics are present in tumors. They have been classically studied as S100 cells; detailed analysis of maturation stage, expression of costimulatory molecules and antigen-presenting capacity has only been done in a limited number

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of recent studies (Furukawa et al., 1985; Nomori et al., 1986; Tsujitani et al., 1987; Schroder et al., 1988; Ambe et al., 1989; Giannini et al., 1991; Nakano et al., 1992; Inoue et al., 1993; Zeid and Muller, 1993; Bethwaite et al., 1996; Thurnher et al., 1996; Coventry et al., 1997; Maehara et al., 1997; Nestle et al., 1997; Wright-Browne et al., 1997; Coppola et al., 1998; Goldman et al., 1998; Ishigami et al., 1998; Hillenbrand et al., 1999; Lespagnard et al., 1999; Kleeff et al., 1999; Scarpino et al., 2000). Tumor-associated DCs (TADCs) generally show an immature phenotype with high CD1a and low co-stimulatory molecules CD80, CD86 and CD40 (Chaux et al., 1996; Enk et al., 1997; Nestle et al., 1997; Viac et al., 1997; Bell et al., 1999; Scarpino et al., 2000). Accordingly, in the few studies conducted so far it was found that TADCs have defective allostimulatory activity (Chaux et al., 1996; Nestle et al., 1997; Viac et al., 1997). In a recent careful analysis in breast cancer it was found that immature langerin DCs are interspersed in the tumor mass, whereas more mature CD83, DC-LAMP DCs are confined to the peritumoral area (Bell et al., 1999). As for breast carcinoma, TADCs in papillary thyroid carcinoma had an immature phenotype, but they tended to localize at the invasion front of the tumor (Scarpino et al., 2000). Interestingly, this distribution was clearly different from that of tumor-associated macrophages (TAMs) which were evenly scattered in the tissue (Scarpino et al., 2000). In papillary thyroid carcinoma, prominent accumulation of DCs in draining lymph nodes has been described, occasionally raising the diagnostic issue of Langerhans cell histiocytosis (Schofield et al., 1992; Thompson et al., 1996; Safali et al., 1997; Lindley et al., 1998; Scarpino et al., 2000). One can therefore assume that in this tumor DC traffic through the tumor to regional lymph nodes actually occurs. The stimuli driving the recruitment and selective distribution of DCs in human tumor tissues have not been definitively identified. The CC chemokine MIP-3α/LARC/exodus is a potent attractant of Langerhans-type DCs, but not of monocyte-derived DCs which do not express the

cognate receptor CCR6. MIP-3α has been shown to be expressed in pancreatic carcinoma, renal cancer, breast carcinoma and papillary thyroid carcinoma (Bell et al., 1999; Kleeff et al., 1999; Scarpino et al., 2000; M.C. Dieu-Nosjean, personal communication). MIP-3α was expressed by carcinoma cells as well as TAMs in pancreatic cancer (Bell et al., 1999; Kleeff et al., 1999; Scarpino et al., 2000). Immature DCs express receptors (e.g. CCR1 and CCR5) for inflammatory chemokines and RANTES has been suggested to be a major attractant for immature DCs produced by papillary thyroid carcinoma (Scarpino et al., 2000). In this tumor, the hepatocyte growth factor amplifies chemokine production by interacting with the Met receptor (Scarpino et al., 2000). The low responsiveness of human DCs to MCP-1, as well as the differential distribution of TADCs and TAMs (Scarpino et al., 2000) suggests that this chemokine, which plays an important role in macrophage recruitment, is not a critical determinant of the recruitment and localization of TADCs. While the actual significance of TADCs for human tumor progression in the absence of immunotherapeutic intervention remains uncertain, circumstantial evidence in experimental models suggests that DCs may represent a tool and target for antitumor strategies. Inoculation of murine melanoma or colon carcinoma modified to release GM-CSF, IL-4 or IL-12 (Armstrong et al., 1996; Levitsky et al., 1996; Stoppacciaro et al., 1997; Nanni et al., 1998) results in local accumulation of DCs with a mature phenotype and their localization in lymph nodes. TADCs in engineered tumors have potent antigen-presenting capacity (Chiodoni et al., 1999). It is likely, but not proven, that cytokine-induced chemokine production is responsible for DC recruitment and trafficking in engineered tumors. More direct evidence for the potential of chemokines to guide DC recruitment was obtained in tumors transfected with MCP-3, a potent attractant for immature DCs (Fioretti et al., 1998). MCP-3 is active in vitro in inducing DC chemotaxis (Sozzani et al., 1995, 1997b; Xu et al., 1996). After MCP-3 gene transfer, P815 mastocytoma cells grew, but subse-

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quently underwent rejection (Fioretti et al., 1998). MCP-3-elicited rejection was associated with increased leukocyte infiltration. Analysis of leukocyte subsets infiltrating tumor tissues revealed that macrophages, T cells, eosinophils and neutrophils were increased in MCP-3transfected tumors. DCs (i.e. DEC-205, MHC class II and CD11c cells) did not substantially increase in the tumor mass, but accumulated in peritumoral tissues and were found in proximity of blood vessels. This peculiar localization has also been found in human carcinoma, and improved patient survival was indeed correlated with peritumoral – rather than intratumoral – DC infiltration (Goldman et al., 1998). MCP-3transfected tumor cells grew normally in nude mice. In immunocompetent mice, MCP-3elicited rejection induced resistance to subsequent challenge with parental cells, and antibodies against CD4, CD8 and IFNγ, but not against IL-4, inhibited tumor rejection. Thus, MCP-3 gene transfer elicits tumor rejection by activating type 1 T cell-dependent immunity. It is tempting to speculate that altered trafficking of antigen-presenting cells, which express receptors and respond to MCP-3, together with recruitment of activated T cells, underlies activation of specific immunity by MCP-3-transfected cells. Therefore gene transfer of a DC chemotactic cytokine in a mouse tumor indicates that these molecules indeed have the potential to regulate DC trafficking in vivo, though the possibility that MCP-3 may act via secondary mediators cannot be formally discarded. As discussed above, TADCs that infiltrate human neoplastic parenchymas generally have an immature phenotype. When present, mature DCs accumulate in peritumoral tissues, even after chemokine gene transfer (Fioretti et al., 1998). This immature phenotype of TADCs may reflect lackofeffectivematurationsignalsinsitu,prompt migration to lymph nodes of mature cells or the presence of maturation inhibitors in the tumor context. These elements may be operative to different degrees in different tumors. For instance, in papillary thyroid cancer there is evidence for substantial accumulation of Langerhans-type DCs in regional lymph nodes, presumably reflecting

migration from the tumor (Schofield et al., 1992; Thompson et al., 1996; Safali et al., 1997; Lindley et al., 1998; Scarpino et al., 2000). Many tumors produce IL-10, IL-6 and M-CSF. IL-10 has been shown to block the differentiation and maturation of DCs, including TADCs (De Smedt et al., 1997; Buelens et al., 1997; Qin et al., 1997; Allavena et al., 1998). Inhibition of DC differentiation from bone marrow precursors has also been observed with IL-6 and M-CSF (Menetrier-Caux et al., 1998). One couldspeculatethatimmatureDCsmaymaintain tolerance to tumor antigens and that TADCs, in analogy with TAMs (Mantovani, 1994), may in some tumors promote tumor progression and dissemination.

CONCLUDING REMARKS The responsiveness to chemokine and nonchemokine attractants is a crucial determinant of the migratory properties of DCs essential for effective antigen presentation. A now large phenomenology is comfortably accommodated in the framework of the ‘chemokine receptor switch’ paradigm. Studies with gene transfer or targeting have yielded results compatible with this general paradigm. Initial studies in tumor models suggest that modulation of DC traffic may be a valid strategy for the upregulation and orientation of specific immunity.

ACKNOWLEDGEMENTS This study was supported by Associazione Italiana per la Ricerca sul Cancro (AIRC); special project AIDS (40A.0.66), and ‘Italy-U.S. Program on Therapy of Tumor’ from Istituto Superiore Sanità; National Research Council (CNR) Target Project Biotechnology.

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16 Studies of endocytosis Wendy S. Garrett and Ira Mellman Yale University School of Medicine, New Haven, Connecticut, USA

Stimulate the phagocytes! Drugs are an illusion! The Doctor’s Dilemma George Bernard Shaw

INTRODUCTION

TYPE OF ENDOCYTOSIS

Dendritic cells (DCs) are the pre-eminent antigen (Ag)-presenting cells of the immune system. In the peripheral tissues, they act as sentinel cells capable of a variety of antigen capture mechanisms including receptor-mediated endocytosis, macropinocytosis and phagocytosis. DCs possess several specializations relevant to antigen uptake. Both the features and distribution of their endocytic compartments, MHC class II (MHC II) compartments and Langerhans cell granules (LCGs), and their modes and levels of endocytic uptake are modulated upon maturation. A consideration of the endocytic machinery of DCs should provide information for a variety of questions in DC biology as well as supplying needed insight for the design of DC-based vaccine strategies. This chapter provides a brief summary of the literature on DC endocytosis, a review of the current methods used to study endocytosis, and a discussion of the developmental regulation of the endocytic pathway in DCs.

The study of endocytosis and its link to immunity began on a beach in Messina around 1879. It was there that Ilya Metchnikoff performed his famous phagocytosis experiment with a rose thorn and a starfish larva. Endocytosis (cell internalizing) may be subdivided into phagocytosis (cell eating) and pinocytosis (cell drinking). The different forms of endocytosis will be considered as will their implications for DCs (Figure 16.1).

Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

Phagocytosis Phagocytosis is a receptor-mediated, actin- and ATP-dependent phenomenon that is triggered by the binding of particles or organisms to specific plasma membrane receptors (Silverstein et al., 1977). Actin assembles around the particle when ligands expressed by the particle bind to a specific class of cell surface receptors. F-actindriven pseudopods engulf the particle, which is

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FIGURE 16.1 There are several types of endocytosis in mammalian cells. DCs utilize these mechanisms to internalize antigens and maintain cellular homeostasis. Immature DCs are particularly adept at internalizing antigens through phagocytosis and macropinocytosis. The actin cytoskeleton is important for phagocytosis and macropinocytosis. Phagocytosis and receptor-mediated endocytosis utilize receptors to selectively internalize receptor-specific antigens.

then internalized in a cytoplasmic phagosome. Mannose, β−glucan, Fc, scavenger and complement receptors can transduce the signals essential for phagocytic internalization (Brown, 1995). Members of the rho family of GTPases have been implicated in the actin-mediated cytoskeletal rearrangements that occur concomitant with particle uptake. Different GTPases may be involved with different types of phagocytosis. For example, FcR phagocytosis is mediated by Cdc42 and Rac1 (Cox et al., 1997), while complement receptor-mediated phagocytosis is dependent upon Rho (Caron and Hall, 1998). However, the precise roles of these GTPases remain uncertain. One recent study has demonstrated a role for Cdc42 and Rac1 in the recruitment of F-actin to the phagocytic cup (Cox et al., 1997), another suggests that Cdc42 and Rac1 do not function in F-actin recruitment but rather serve distinct functions in the formation of the phagocytic cup. Inhibition of Cdc42 resulted in the formation of arm-like structures supporting the particulate ligands while inhibition of Rac1 resulted in the enclosure of particulate ligands within thin membrane protusions that did not fuse (Massol et al., 1998). There are additional regulatory pathways involved in phagocytosis. FcR-associated phago-

cytosis involves a tyrosine kinase-dependent pathway. Several cytoskeletal proteins, including paxillin, which associate with early phagosomes are tyrosine kinase substrates that are phosphorylated in response to Fc receptor ligation (Greenberg et al., 1994) Phagocytosis provides a means for the internalization of a broad range of receptor-bound antigenic ligands. DCs seem specially engineered for the internalization and presentation of phagocytic ligands as evidenced by the observation that the antigen-presenting capacity of DCs is much higher for bead-adsorbed antigen than for the soluble form of the antigen (Scheicher et al., 1995). Indeed, DCs exhibit a remarkable efficiency in the generation of MHC class II epitopes derived from the phagocytosis of cellular fragments. Phagocytosis of I-E MHC class II molecules from cellular fragments was 1000–10 000-fold more efficient in generating MHC class II–peptide complexes than uptake of preprocessed I-E peptide (Inaba et al., 1998). Furthermore, resident lymphoid DCs phagocytose short-lived migratory DCs and present antigens internalized by these migratory DCs (Inaba et al., 1998). The importance and consequences of this phenomenon probably reflect the fact that particles, e.g. cells and bacteria, are the most physiological forms of antigen DCs encounter during their lifetimes. Although there are many differences between the machinery driving phagocytosis and receptor-mediated endocytosis, a series of papers have shown several reciprocal similarities. Dynamin-2, a dynamin GTPase involved in receptor-mediated endocytosis, is also necessary for phagocytosis (Gold et al., 1999). In endocytosis, dynamin functions in the scission step of clathrin-coated vesicle formation. During phagocytosis, dynamin-2 was found on forming phagosomes. Although it is not entirely clear how dynamin-2 acts in the final budding of nascent phagocytic vacuoles, it does appear to be functional in the process. Inhibition of dynamin-2, via the expression of its dominant negative mutation, blocked membrane extension around particles (Gold et al., 1999). The role of dynamin-2 and possibly the Rho GTPases

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may indicate the involvement of continued membrane remodeling and vesicle traffic in phagocytosis.

Pinocytosis Pinocytosis encompasses both fluid-phase uptake and receptor-mediated endocytosis. Fluid-phase uptake includes micropinocytosis and macropinocytosis: two distinct forms of solute uptake. Micropinocytosis involves internalization of macromolecules less than 0.2 µm in diameter via both clathrin-coated and uncoated vesicles (Steinman et al., 1983) Clathrin coat-mediated endocytosis Most receptor-bound ligands are internalized via a clathrin coat-mediated process. Certain membrane receptors, e.g. FcγR, are efficiently clustered in clathrin-coated vesicles. These receptors usually have an internalization signal, a tyrosine or dileucine motif, in their cytoplasmic domain. Such signals work by binding to adaptor proteins to which the clathrin coat attaches. Coats consist of a clathrin cage, a trimer of heterodimers (clathrin heavy and light chain) and a heterotetrameric adaptor complex (Robinson, 1994; Schmid, 1997). An important component of the clathrin-coated vesicle machinery is dynamin, a GTPase, which is required for coated pits to pinch off as coated vesicles. After internalization in a clathrincoated pit, receptor–ligand complexes have two possible fates (Mellman, 1996). The complexes can traffic through the endosomal–lysosomal pathway encountering lower and lower pH and increasing concentrations of proteases and hydrolytic enzymes, e.g. FcR complexes (Mellman, 1996). Alternatively, the receptor– ligand complexes can dissociate within the endosomal network where they can be sorted. Upon sorting to the recycling arm of the endocytic pathway, receptors can return to the cell surface where they can initiate multiple rounds of ligand internalization, e.g. mannose receptor (Mellman, 1996). Clathrin coat receptormediated endocytosis provides a selective and

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efficient means of concentrating the internalization of specific ligands. It is necessary for the uptake of nutrients and other components essential for the maintenance of cellular homeostasis (Mellman, 1996). Macropinocytosis Macropinocytosis is mediated by the actin cytoskeleton and is independent of clathrin and membrane receptors. Macropinosomes are heterogeneous in size, ranging from 0.2 µm to 5 µm in diameter. Three differentially regulated processes are essential for macropinocytosis: actin cytoskeleton-driven ruffle formation, closure of the ruffle into a vesicle, and dissociation of actin filaments from the vesicle so that it can participate in intracellular trafficking and fusion with other vesicles (Swanson andWatts, 1995). In contrast to micropinocytosis, the capacity for constitutive macropinocytosis may be restricted to a select number of cells. Several agonists are capable of inducing macropinocytosis. EGF induces macropinocytosis in fibroblasts while PMA stimulates the process in macrophages (Swanson and Watts, 1995). Studies examining Salmonella typhimuriuminduced macropinocytosis have demonstrated that cdc42 is essential: dominant negative cdc42 expressed in the host cell can block macropinocytosis of Salmonella typhimurium (Chen et al., 1996). Bacterial products of the Salmonella typhimurium type III secretion system appear to activate cdc42 in the host cell, resulting in membrane ruffling (Galan and Bliska, 1997). Cells usually require inductive signals to macropinocytose and downregulate macropinocytosis in the absence of these stimulating factors. However, immature DCs constitutively macropinocytose (Sallusto et al., 1995). Macropinocytosis is a nonselective form of endocytosis. It has been unclear how internalized ligands are concentrated in macropinosomes. However, preliminary experiments suggest that aquaporins (water channels) may play a role by mediating water egress from macropinosomes (de Baey and Lanzavecchia, 2000).

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The fate of macropinosomes depends on the cell type. In EGF-stimulated A431 cells, a human epitheloid cell line, macropinosomes may fuse with each other but fail to deliver their contents to early or late endosomes (Hewlett et al., 1994). In macrophages, macropinosomes gradually acquire lysosomal markers progressing through three stages: a transferrin receptor-positive phase (TfnR), a TfnR, rab 7, mannose 6-phosphate receptor (M6PR) phase, and TfnR, rab7, M6PR, cathepsin L, lgpA, and lgpB phase before fusing with the tubular lysosomal compartment (Racoosin and Swanson, 1993). In a fetal skin-derived dendritic cell line (FSDC) (Girolomoni et al., 1995), macropinosomes acquire proteases and decrease pH but do not possess all lysosomal markers (Lutz et al., 1997), suggesting that a macropinosome-derived compartment may function as an antigen retention compartment. However, it is not clear how faithfully this process is preserved in primary DCs. Macropinocytosis is a nonselective form of uptake by which a relatively large volume of antigen-rich solute can be internalized. It is an appealing concept that sentinel DCs in the periphery may be able to internalize vast quantities of a variety of antigens and eventually prime naïve T cells to mount an immune response. Two studies have demonstrated that macropinocytosis in mouse GM-CSF bone marrow DCs can result in cross-presentation. Ovalbumin internalized by immature DCs by macropinocytosis was presented by class I molecules to CD8 ovalbumin-specific T-cell clones and horseradish peroxidase was shown to move from macropinosomes to the cytosol (Norbury et al., 1997; Brossart and Bevan, 1997). DCs may have evolved a specialized endosome-to-cytosol translocation apparatus for the selective export of internalized antigen to the cytosol (Rodriguez et al., 1999). Cross-presentation or cross priming involves the presentation of exogenous antigens by the class I molecules in addition to the presentation of the antigen by the class II molecules (Bevan, 1976; Carbone and Bevan, 1990). Investigations into the role of actin regulatory proteins in DC macropinocytosis have demonstrated that while loss of gelsolin expres-

sion disrupts dermal fibroblast membrane ruffling, DCs generated from gelsolin knockout mice are capable of macropinocytosis and cross-priming (West et al., 1999) Potentially, macropinocytosis may provide a unique pathway in DCs for efficient channeling of antigenic peptide to both the MHC class I and MHC class II pathways. The presence of cell surface receptors, e.g. mannose receptor, may increase the efficiency of macropinocytosis if a number of receptors with bound ligands are rapidly internalized in a macropinosome. However, there is scant evidence that macropinocytosis actually serves a purpose in situ. The epidermis is a tight mesh-like network of keratinocytes in which LCs interdigitate. It seems unlikely that LCs are capable of generating large membrane ruffles while in the skin. Perhaps, macropinocytosis is an in vitro activity which simply reflects an inherent capability to generate the pseudopod extensions that engulf phagocytic ligands. There are several similarities between phagocytosis and macropinocytosis. Both are dependent on remodeling of the actin cytoskeleton and cdc42 is an important effector in regulating these changes (Chen et al., 1996; Greenberg et al., 1994). There are also morphological similarities between the generation of macropinocytic ruffles and phagocytic cups. However, the receptor dependence of phagocytosis and ligand–receptor-mediated zippering of the phagocytic pseudopods around the particle demonstrate that the two are distinctive processes that may, however, share several mechanistic similarities.

Potocytosis Potocytosis is a form of endocytosis specialized for the internalization of GPI-anchored membrane proteins. Caveolae, first identified and called plasmalemmal vesicles by Palade, are cave-like, variably shaped invaginations of the plasma membrane involved in podocytic internalization of small molecules. The internalization of 5-methyltetrahydrofolate via the folate receptor has been the model system for ligand– receptor studies of potocytosis. Caveolin-1, -2, -3

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are the specific proteins which biochemically define caveolae. While early studies of caveolae focused on its role in transcytosis, present research trends have focused on investigations of how the caveolar membrane system is involved in the compartmentalization of signal transduction (Anderson et al., 2000). The importance of potocytosis as a method of antigen uptake in DCs is unknown. Respiratory syncytial virus (RSV ) antigens have co-localized with caveolins in RSV-infected cattle DCs (Werling et al., 1999). However, whether RSV potocytosis is involved in generating DC-driven immunity is uncertain. Caveolins are often found on the plasma membrane in lipid microdomains called rafts. Lipid rafts are an area of intensive interest for a variety of signal transduction questions including antigen presentation (Monks et al., 1998; Viola and Lanzavecchia, 1999; Viola et al., 1999). Whether caveolae function in DCs and other hematopoietic cells in the organization of cell signaling and transduction events is worthy of further investigation.

ENDOCYTIC RECEPTORS Below is a brief summary of the numerous receptors that are expressed by dendritic cells during their development and that are capable of mediating receptor-mediated endocytosis and in several cases phagocytosis as well.

Mannose/β glucan receptor The mannose receptor (MR) is expressed on immature DCs (Reis e Sousa et al., 1993; Sallusto et al., 1995; Engering et al., 1997). MR contains multiple carbohydrate-binding domains and internalizes a variety of glycoproteins. The receptor binds a range of oligosaccharides with different affinities: D-mannose = L-fucose >> DN-acetylglucosamine >= D-glucose >> D-xylose >>> galactose (Pontow et al., 1992). Unlike FcR, MR recycles and can mediate multiple rounds of ligand internalization via pH-sensitive ligand dissociation (Pontow et al., 1992). MR plays an

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important role in host defense-mediated phagocytosis of unopsonized targets and internalization of a variety of microorganisms that have exposed mannosylated glycoproteins: Candida albicans, Leishmania promastigotes and Pneumocystis carinii (Ezekowitz and Hoffman, 1996). In antigen-presentation experiments, MR enhanced the ability of DCs to present low concentrations (109–1012 ng/mL) of soluble mannosylated ligands (Engering et al., 1997). Similarly, mannosylation of peptides resulted in 200–10 000-fold higher antigen-presentation efficiency when compared with unmannosylated peptides (Tan et al., 1997). Agalactosyl IgG, found at elevated levels in the serum of rheumatoid arthritis and other autoimmune disease patients, is internalized by mannose receptors (Dong et al., 1999). Mannose receptors also internalize antibody and antigen–antibody complexes enzymatically modified to expose Nacetylglucosamine residues (Dong et al., 1999). Mannose receptor is now being exploited as a site of entry for gene delivery using mannosyl polyethylenimine DNA conjugates (Diebold et al., 1999a, 1999b). Expression of mannose receptor in human monocyte-derived DCs appears to be developmentally regulated. Exposure to the maturation factors, LPS or TNFα, reduced the capacity of DCs to present both mannosylated and nonmannosylated ligands by 10–100-fold (Engering et al., 1997). Endocytosis in general is reduced with maturation and MR expression itself is also downregulated. In contrast, the ‘tolerogenic’ cytokine IL-10 (Longoni et al., 1998) and the glucocorticoid immunosuppressants dexamethasone (Piemonti et al., 1999) and methylprednisolone (Vanderheyde et al., 1999), appear to upregulate expression of mannose receptor and increase the internalization of the mannose receptor ligand FITC-dextran.

DEC-205 receptor DEC-205 receptor, a member of the C-type lectin receptor family, is an orphan receptor (Jiang et al., 1995; Kato et al., 1998). Initial experiments

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using rabbit polyclonal antibodies directed against the ectodomain of the receptor demonstrated that it is internalized by receptormediated endocytosis and is delivered to class II multivesicular structures (Jiang et al., 1995). Rabbit antibodies specific for the DEC-205 ectodomain were presented to reactive T-cell hybridomas with 100-fold more efficiency than nonspecific rabbit antibodies (Jiang et al., 1995). Although total DEC-205 expression increases with maturation, cell surface expression decreases during development (M. Nussenzweig, personal communication). Downmodulation during development and preliminary data on its antigen-presenting function suggest that it may serve as an antigen receptor. The increasing levels of expression with development and intracellular localization suggest an additional role.

FcR Both murine and human epidermal Langerhans cells express high-affinity receptors for IgE molecules (Wang et al., 1992; Bieber et al., 1992; Maurer et al., 1996; Stingl and Maurer, 1997) and low-affinity receptors for IgG molecules (FcRII) (Schmitt et al., 1990; Esposito-Farese et al., 1995). The high-affinity receptor for IgE (FcεRI) is expressed on Langerhans cells, dermal DCs (Wang et al., 1992) and peripheral blood (PB) DCs (Maurer et al., 1996). On DCs, FcεRI functions to augment allergen presentation in an IgE-dependent manner. Peripheral blood human DCs, Langerhans cells and dermal DCs lack FcεR1β and express FcεRI as a multimer of FcεR1α and FcεRIγ chains (Maurer et al., 1996). Immature murine Langerhans cells express two membrane anchored isoforms of FcγR, FcγRIIb2 (CD32b2) and FcγRIII (CD16). LCs are capable of internalizing IgG–Ag complexes via receptormediated endocytosis by their cell surface FcγRs. Human blood dendritic cells and cultured monocyte-derived DCs express both FcγRI (CD64) and FcγRII (CD32) and are capable of phagocytosis via these receptors (Fanger et al., 1996). The FcRs expressed by LCs and DCs may not be internalized for the destruction of

opsonized particles in phagolysosomes but rather for antigen processing and presentation.

Complement receptor The complement system plays a role in enhancing a number of immune phenomena: phagocytosis of pathogens, inflammation, antigen focusing and cell lysis (Burke and Gigli 1980). There are several receptors for components of the complement system: C3b, C3bi, C3d and C4b receptors. Many cells express C3b receptor (C3bR) and some subsets of these cells express a form of C3bR that crossreacts with C4b. Receptors specific for C4b are more limited in expression (eosinophils, peripheral B lymphocytes, neutrophils). LCs express C3bR but do not express C4bR (Stingl and Maurer, 1997). LCs also express C3biR and downregulate expression after 1–3 days in culture (Schuler and Steinman, 1985). The significance of complement receptors on DCs is an issue of speculation, for example, some have proposed that C3R may allow for antigen bridges between DCs and naïve T cells expressing C3 (Burke and Gigli, 1980).

Scavenger receptor There are three known classes of scavenger receptors: A, B and C. Initial studies on the scavenger receptors focused on their binding of oxidized and acetylated low-density lipoprotein. However, they are capable of binding a variety of negatively charged molecules (Rigotti et al., 1997). The first scavenger receptors identified were the class A type I and II receptors which are expressed on macrophages. Class A receptors are also expressed on DCs (S. Gordon, personal communication). Recently, scavenger type A receptors have been implicated in the clearance of apoptotic cells in the thymus (Platt et al., 1996). Class B scavenger receptors (SR-Bs) are members of the CD36 protein family and bind negatively charged liposomes, HDL (Rigotti, 1997), acetylated and oxidized LDL, and apoptotic cells (Fukasawa et al., 1996). Human monocyte-derived DCs developmentally regulate the expression of CD36 (Albert et al., 1998a).

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CD36 and the αvβ5 integrin function as coreceptors for the phagocytosis of apoptotic cells. Phagocytosis of apoptotic bodies and cells through these receptors results in cross-priming and presentation to CTLs and may be involved in the generation of cross-tolerance as well (Albert et al., 1998b). Class C receptors have been cloned from and are expressed by Drosophila embryonic hemocytes and macrophages and the Schneider L2 cell lines. Preliminary characterization of this receptor demonstrates affinity for acetylated LDL and polyanionic ligands (Pearson et al., 1995). The expression of scavenger receptors has not been characterized for all the different dendritic cell populations.

MEASUREMENT OF ENDOCYTOSIS Experimental approaches Assays for endocytosis are conceptually simple: cells are incubated in the presence of a ligand and then the uptake of the ligand is measured. The complexity of these assays using DCs is introduced in four basic ways: the preparation of the cell population, the choice of the ligand, the conditions under which the cells are exposed to the ligand, and how ligand uptake is assessed. The techniques used to assess ligand uptake can be qualitative and/or quantitative. Fluorescently conjugated ligands in conjunction with flow cytometry and immunofluorescence microscopy can be used to examine uptake in viable and fixed populations of cells. Antibodies or enzymatic reactions can be employed to follow the uptake of a ligand using immunohistochemical techniques at the light and electron microscope level. In addition, fluorometric substrates can be used to quantitate enzymatic ligands. The quantity of internalized versus exocytosed ligands and the rates of internalization and exocytosis are easily deduced from data analysis of these techniques. Subcellular fractionation and free flow electrophoresis are standard cell biological techniques utilized in studying the intracellular

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trafficking of ligands through the endocytic pathway when the ligand can be assessed by an enzymatic activity or antibody-mediated detection system. Alternatively, antigen presentation assays have demonstrated the uptake and processing of a ligand when appropriate T-cell clones or hybridomas are available.

Probes/reporters FM 3-25 FM 3-25 (N-(3-triethylammoniumpropyl)-4(4-(dioctadecylamino)styryl)pyridinium,di-4chlorobenzenesulfonate) and FM 4-64 N-(3triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl)pyridinium dibromide are vital, lipophilic styryl dyes that have been used to assay for endocytosis. The dye molecules insert into the outer leaflet of the plasma membrane where they fluoresce when excited (Vida and Emr, 1995). FM3-25 and FM4-64 are tracers for membrane turnover and plasma membrane dynamics. These dyes may be quite helpful in understanding the maturation of macropinosomes in DCs or studies in situ. Lucifer yellow The potasssium salt of Lucifer yellow CH (LY) is a hydrophilic tracer for fluid-phase pinocytosis with a peak absorbance at 425 nm and peak fluorescence emission at 528 nm. At 488 nm, the excitation wavelength of an argon laser, the emission is rather low, ~700 cm1M1. Lucifer yellow CH bears a carbohydrazide group allowing for aldehyde fixation, which allows for immunocytochemical analysis. In macrophages, LY is not degraded and is nontoxic at concentrations up to 6 mg/mL (Swanson et al., 1985). Although LY has been used successfully in labeling the endocytic pathway of macrophages and PBMC GM-CSF  IL-4 DCs, LY membrane impermeance should be evaluated by examining for cytosolic leakage in different cell types (Swanson, 1989). Intense light excitation of lysosomes loaded with LY occasionally results in

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lysosomal explosion and leaking of LY into the cytoplasmic space (Swanson 1989). Dextrans Dextrans are electro-neutral, hydrophilic branched polymers of poly-D-glucose that do not cross biological membranes. Fluoresceinated dextrans (F-DXs) are commercially available. Molecular Probes (Eugene, Oregon, USA) provides a ranges of fluorescein dextrans (mol. wt 3000–2 000 0000) with a high degree of dye substitution (2–4 dye molecules 40 000 mol. wt: 3–6 dye molecules 70 000 mol. wt). Commercially available F-DXs may contain free fluorescein which is highly adsorptive and potentially cytotoxic. Gel-exclusion chromatography or dialysis should be used to reduce the levels of unconjugated fluorescein. Seminal studies using FITC-DXs demonstrated their usefulness in studies of fluid-phase pinocytosis (Berlin and Oliver, 1980). In DCs, dextrans will bind the mannose receptor (Sallusto et al., 1996). DCs have been pulsed with 0.1–10 mg/mL F-DX and typically assayed for uptake using three basic methods: cell detergent lysates have been analyzed in fluorimeters, live or fixed DCs have been assayed by flow cytometry, and cells have been examined using fluorescence or confocal microscopy. Quantitative measurement of intracellular F-DX are often inaccurate because fluorecein fluorescence varies as an inverse function of pH. Thus visualization of F-DX in the acidic milieu of endosomes and lysosomes often reflects a fluorescence overshoot. Fluorescence overshoot with acidic pHs is less of a problem with Texas red and rhodamine-conjugated dextrans and proteins. However, these fluors are more adsorptive (Swanson et al., 1989). Horseradish peroxidase Horseradish peroxidase (HRP) is a glycoprotein that is absent from most mammalian cells. It can be employed as both a fluid-phase and receptormediated probe. HRP was demonstrated to be a ligand for MR (Stahl et al., 1978). It is a conven-

ient tracer for immunocytochemistry; in situ, HRP can be detected by reaction with diaminobenzidine. Its concentration can be measured by colorimetric assay using O-phenylenediamine (OPD) and hydrogen peroxide (Straus, 1964). Polyclonal sera directed against HRP are commercially available and are useful for fluorescence microscopy imaging. HRP has been a standard tracer for the endocytic pathway for almost four decades (Steinman et al., 1974). It is relatively nontoxic and has only been demonstrated to perturb the endocytic pathway in thioglycollate-stimulated macrophages (Swanson et al., 1985). Nonfluorescent, nonenzymatic probes/reporters There are a number of probes that have been used in endocytic assays which have not been mentioned explicitly. These ligands have been used in conventional light microscopy and ultrastructural studies. Colloidal carbon, latex microscospheres/beads, paramagnetic latex and iron-containing beads have been valuable tools in a variety of uptake assays. Paramagnetic latex and iron-containing beads can be used in conjunction with magnets to isolate different cell populations or to separate subcellular organelles. Pathogens, including bacteria, mycobacteria and yeast, have been used to ask questions about host/pathogen interactions as well as endocytosis. Such organisms can be easily labeled directly with FITC, TRITC or biotin or indirectly with appropriate antibodies.

ENDOCYTIC CAPACITIES OF DENDRITIC CELLS Initially, endocytosis was of interest to those studying DCs because it provided a means to functionally characterize and differentiate DCs from other splenic or epidermal cells. Three basic techniques dominated studies of dendritic cells endocytosis: electron microscopy, flow cytometry and antigen-presentation assays. Until 1992, bulk culturing method for the generation of developmentally homogeneous DCs did

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not exist and previous to this time, endocytosis assays were performed on LCs or DCs in situ, or from short-term culture of tissue explants/isolates. (For reviews see: Steinman and Swanson, 1995; Austyn, 1996; Cella et al., 1997b).

Electron microscopy studies Early analysis of the endocytic capacities of LCs was performed at the ultrastructural level using ferritin or horseradish peroxidase (HRP) as an uptake tracer. These studies demonstrated that LCs could use endocytosis to internalize exogenous protein but that LCs were weakly endocytic compared with keratinocytes (Nordquist et al., 1966; Wolff and Schreiner, 1970; Sagbiel, 1972). Initial studies of the endocytosis of soluble HRP, colloidal carbon and thorium dioxide by splenic DCs in vitro and in vivo demonstrated poor DC uptake of soluble endocytic markers compared with macrophages (Steinman and Cohn, 1974). The LCs and DCs that these groups examined would be characterized as mature by the morphological phenotyping that is employed today. Thus, these experiments demonstrated that mature DCs have low endocytic capacity. Phagocytosis by sentinel DCs, such as LCs and organ DCs, have been studied in detail. DCs isolated from mouse spleen, lymph nodes, thymus, liver (Steinman and Cohn, 1974; Steinman et al., 1980), and rat solid organs and tissue (Hart and Fabre, 1981) demonstrated low levels of phagocytosis. As observed in a transmission electron microscopy (EM) study, LCs isolated from epidermal suspensions phagocytose zymosan and latex microspheres (Reis e Sousa et al., 1993). In vivo antigen-uptake studies in rats and sheep have contributed to the understanding of DC antigen uptake and migration in an intact immune system. Following subcutaneous injection of HRP in complete Freund’s adjuvant (CFA) and boosting, HRP immune complexes were visualized within veiled DCs (Hall and Robertson, 1984). EM studies demonstrated that dendritic cells can internalize Borrelia burgdorferi by coiling phagocytosis (Filgueira et al., 1996).

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In freshly isolated and cultured human peripheral blood DCs, the trafficking of pulse chased BSA-gold was compared. The antigen trafficked through the endocytic pathway and accumulated in class II endosomal/lysosomal compartments similar to those observed in B cells (Nijman et al., 1995). Ultrastructural studies have been important in demonstrating that in DCs, phagocytosis is not only required for the uptake of immune complexes, bacteria, fungi and yeast but also for the clearance of dead cells. Several EM studies have demonstrated that IDCs phagocytose necrotic and apoptotic cells. Allogeneic lymphocytes have been visualized within IDCs in lymph node and spleen (Fossum and Rolstad, 1986; Fossum et al., 1984) and irradiated necrotic thymocytes have also been identified within IDCs (Duijvestijn et al., 1982). In murine vaginal epithelia during late metestrus and early diestrus, LCs phagocytosis apoptotic vaginal epithelial cells (Parr et al., 1991). Perhaps, the phagocytosis of apoptotic cells in this milieu functions to tolerize the immune system to cells at a mucosal interface that undergo cyclical changes.

Video microscopy Cell biologists have used video microscopy to observe biological phenomenon in real time for decades. Technological innovations and cost reductions are making this technique more accessible to many investigators in a variety of disciplines. Live cell imaging has been used in recent years to follow membrane transport events, e.g. the transport of MHC class II molecules (Wubbolts et al., 1996), and has been increasingly employed to follow the dynamic changes occurring during antigen presentation in T cells and APCs (Monks et al., 1998; Underhill et al., 1999; Grakoui et al., 1999). Fluorochromecoupled Fab fragments of antibodies directed against topologically accessible epitopes can be used to follow the trafficking of proteins. The advent of green fluorescent protein (GFP) and the related spectrum of fluorescent proteins (blue fluorescent proteins (BFP), yellow

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fluorescent proteins (YFP), cyan fluorescent proteins (CFP)) has also helped to popularize video microscopy. Chimeric protein fusions with N- or C-terminal GFPs are often fully functional and can be employed to elucidate trafficking events or dynamics of protein–protein interactions at the visual or biophysical level. The use of such reporters in DCs is not trivial. DCs are not easily transfectable, however, adenoviral or retroviral expression systems have been used with success in DCs (Dietz and Vuk-Pavlovic, 1998; Gasperi et al., 1999; Zhong et al., 1999). Even cDNA microinjection is not straightforward. Often, levels of expression that may suffice for immunological read-outs are not high enough for visually based assays.

Flow cytometry Flow cytometry which can assay bulk populations at the level of a single cell, is an amenable system for the analysis of heterogeneous DC preparations. Bulk populations of homogeneous cells are required for the subcellular fractionation and spectrophotometry/fluorimetry traditionally used to accurately determine the localization of ligands in different compartments of the endosomal–lysosomal system. DCs isolated from mouse spleen and incubated for 24 hours had heterogeneous endocytic activities in flow cytometry assays (Levine and Chain, 1992). The uptake of Staphylococcus aureus, latex microspheres and zymosan was investigated in freshly isolated Langerhans cells and splenic DCs using flow cytometry (Reis e Sousa et al., 1993). Flow cytometry was also used to follow the internalization of FITC by sentinel DCs isolated from heart and kidney (Austyn et al., 1994). Macropinocytosis and mannose receptormediated endocytosis are extremely efficient means for internalizing macromolecules in immature DCs from the GM-CSF, IL-4 PBMC culture system (Sallusto et al., 1995). Mannose receptor mediated the saturable uptake of F-DX as judged by competition studies using antimannose receptor antibody, mannan, mannosylated BSA and EDTA. LY uptake was non-

saturable in the flow cytometry assays and was inhibited by cytochalasin D (actin cytoskeleton inhibitor) and amiloride (Na/H pump inhibitor), suggesting that its uptake is mediated by macropinocytosis. Cord blood CD34 hematopoietic progenitor cells treated with GM-CSF and TNFα differentiate into two distinct DC lineages in culture: an LC-type DC and a CD14-derived DC (Caux et al., 1996). The two lineages have similar functional capacities in T-cell activation assays but differences in their endocytic capacities (Caux et al., 1997). The CD14 DCs bound immune complexes while the LC-type lineage DCs demonstrated greatly reduced immune complex binding in a FACS-based assay (Caux et al., 1997). The endocytic activity of CD14-derived DCs was approximately 10-fold higher than the LC-type DCs. Mannose receptor mediated the uptake of F-DX and HRP in both lineages (Caux et al., 1997).

Antigen-presentation assays In the past decade, several groups have explored the antigen capture function of DCs by assaying for antigen presentation. The majority of these studies do not directly address the mechanisms by which DCs take up antigen but are instructive for a number of reasons. Freshly purified splenic DCs could internalize, process and present keyhole limpet hemocyanin (KLH). The processing of KLH was chloroquine-sensitive, suggesting that an acidified intracellular compartment, an endosome or lysosome, was necessary for processing (Chain et al., 1986). Freshly isolated epidermal LCs present soluble, intact myoglobin (Romani et al., 1989). In LCs, antigen processing of myoglobin and conalbumin was inhibited by both chloroquine and cycloheximide. This observation suggests that both an acidified compartment and newly synthesized proteins were required for processing. Antigen could be retained in cultured, antigen-pulsed LCs for up to 2 days in an immunogenic form (Romani et al., 1989; Puré et al., 1990). Splenic DCs efficiently generated and presented immunogenic fragments of intact hen egg lysozyme

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(HEL) to an I-Ak restricted T cell hybridoma-specific HEL. When splenic DCs and LPS blasts were used as APCs in presentation assays, DCs required 100-fold less HEL to induce the same level of T-cell proliferation as with LPS blasts (De Bruijn et al., 1992). LCs process and present phagocytic ligands to antigen-specific T cells, e.g. Leishmania major (Moll et al., 1995). Mouse bone marrow-derived GM-CSF cultured dendritic cells phagocytose particulate antigen and can process and present bacille CalmetteGuérin (BCG) to antigen-specific T cells (Inaba et al., 1993). In addition to phagocytosing a variety of particles and pathogens, DCs phagocytose apoptotic cells. Human PBMCs internalize fluinfected apoptotic macrophages and present derived antigens via both the MHC class I and class II pathways (Albert et al., 1998).

DEVELOPMENTAL REGULATION Recent studies have confirmed a concept inferred from earlier studies, namely that endocytosis is a developmentally regulated process in dendritic cells. Immature dendritic cells constitutively macropinocytose, endocytose and phagocytose. In response to stress and ‘danger’, in the form of explantation, pathogen and inflammatory stimuli (Matzinger, 1994), DCs mature and undergo striking morphological and functional changes (Banchereau and Steinman, 1998). One functional alteration is the downregulation of endocytosis. A shutdown of endocytosis is consistent with DCs changing from antigen sentinels to T-cell primers. A sentinel DC with its high endocytic capacity and with an additional T-cell stimulatory capacity trafficking through or located in a lymph node might result in autoimmunity. Studies examining the internalization and presentation of myoglobin in fresh versus cultured LCs and splenic DCs demonstrated that processing and presentation of exogenous protein was lost during culture, suggesting that endocytosis was developmentally regulated (Romani et al., 1989).

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Particulate uptake and presentation studies in mouse bone marrow DCs clearly indicated that progenitor/early DCs pulsed and chased with BCG were much more efficient in presentation assays than pulsed and chased mature DCs (Inaba et al., 1993). Studies of DCs in the peripheral hepatic lymph established that DCs go through a transitory phagocytic stage and that they down regulate their phagocytic activity when they enter their migratory stage (Matsuno et al., 1996). Investigations of CD34-derived LC lineage DCs and CD14 lineage DCs suggested that LC endocytic activity is temporally restricted to an early stage of their development, ‘day 6’, while CD14 lineage DCs vigorously endocytose for a longer duration, ‘day 8–13’. Maturational stimuli downregulate macropinocytosis and receptor-mediated endocytosis. Treatment of PBMC-derived dendritic cells with TNFα, CD40L, Il-1β and LPS, inflammatory stimuli, resulted in a marked reduction of endocytosis (Sallusto et al., 1995). Ceramide has emerged as a downstream effector and intracellular signal common to pathways mediated by the above inflammatory stimuli (Sallusto et al., 1996). Ceramide inhibited the uptake of FDX and LY in PBMC-derived DCs (Sallusto et al., 1996). Recently, the ceramides have been shown to prevent membrane recruitment of the Rho GTPases (Abousalham et al., 1997). These findings may, in part, explain the endocytic modulatory effects of ceramide since the Rho GTPases have been implicated in regulating phagocytosis and macropinocytosis. Treatment with TNFα, IL-1β and CD40L decreased endocytosis and increased intracellular levels of ceramide. However, ceramide did not upregulate MHC class II and co-stimulatory molecules. This finding is inconsistent with the observation that upregulation of cell surface MHC class II molecules in mature DCs results from a shutdown of endocytic recycling (Cella et al., 1997a). Thus ceramide cannot be the only intracellular signaling molecule responsible for downregulating endocytosis. Indeed, our preliminary results have suggested a role for Cdc42 in this process (Garrett et al., 2000).

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SPECIALIZATIONS OF THE DC ENDOCYTIC PATHWAY Regulation of the MHC class II pathway Antigen-processing compartments (Plate 16.2) have been a subject of intensive research and much debate in recent years (Neefjes et al., 1990; Harding et al., 1991; Peters et al., 1991; Amigorena et al., 1994; Tulp et al., 1994; West et al., 1994; Pierre and Mellman, 1998b). The discovery of novel endocytic compartments enriched in MHC class II molecules proved a major conceptual advance in the cell biological basis of antigen processing (Mellman, 1996). Examinations of MHC class II compartments in DCs have been carried out at the level of fluorescence (Sallusto and Lanzavecchia, 1994) and immunoelectron microscopy (Nijman et al., 1995). Two recent studies have examined the organization of MHC class II pathways during the in vitro development of DCs (Cella et al., 1997a; Pierre et al., 1997). These studies provided insight into how the antigen-processing machinery of DCs is a dynamic device, reconfiguring itself in response to environmental stimuli (Watts, 1997). In the human PBMC GM-CSF IL-4 DC system, MHC class II molecules in immature DCs are rapidly internalized and recycled (t1/2 10 hours), however, MHC class II molecules in DCs exposed to maturational factors have a MHC class II half-life 100 hours and increase their synthesis of MHC class II (Cella et al., 1997a). As the DCs mature, the MHC class II half-life appears to increase as a result of a general shutdown of the recycling of MHC class II. Previous studies have observed a downregulation in DC endocytosis in response to inflammatory stimuli (Sallusto et al., 1995). The increased MHC class II synthesis and half-life results in a large pool of long-lived peptide-loaded MHC class II molecules on the cell surface awaiting recognition by the appropriate T cell. In mouse bone marrow GM-CSF DC system, three distinct developmental stages differentiated by their MHC class II compartmentalization have been identified (Pierre et al., 1997).

MHC class II in immature DCs accumulates in lysosomes instead of trafficking to the cell surface. As maturation progresses, MHC class II redistributes to peripherally located vesicles that resemble CIIV (reviewed in Pierre and Mellman, 1998b; described in Amigorena et al., 1994). Finally, in fully mature DCs, peptide-loaded MHC class II traffics to the cell surface ready to stimulate relevant T cells. The changes in MHC class II membrane transport are tightly linked to the developmentally distinct stages of the DC and the maturational program is initiated by inflammatory stimuli, LPS, TNFα and IL-1β. Remarkably, in the absence of inflammatory stimuli, immature DCs will sequester antigens in their lysosomes and MHC class II will traffic to these lysosomes, but these antigens will not be loaded onto these newly synthesized MHC class II molecules (Inaba et al., 2000). However, the delivery of an inflammatory stimulus results in the loading of these MHC class II molecules with these antigens and the trafficking of these antigenic peptide-MHC class II complexes to the cell surface via CIIV (Inaba et al., 2000; Turley et al., 2000). Insight into the mechanisms controlling some of these trafficking events has come from two groups who have examined the role of the protease cathepsin S. Using gene disruption of cathepsin S and cathepsin S active site-directed probes, cathepsin S was shown to play an important regulatory role in the trafficking of MHC class II molecules to the cell surface (Driessen et al., 1999). Regulation of Ii chain proteolysis by cathepsin S was shown to control the transport of newly synthesized MHC class II molecules in mouse bone marrow-derived GM-CSF cultured mouse DCs. Furthermore, the proteolytic activity of cathepsin S was shown to be developmentally regulated by the protease inhibitor cystatin C (Pierre and Mellman, 1998a). In immature DCs, when cathepsin S activity is low because of cystatin C, MHC class II–Ii complexes are transported to lysosomes. However, with maturation, cathepsin S activity increases because of a modification in cystatin C activity. Then cathepsin S can efficiently proteolyze invariant chain, and MHC class II can be loaded

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with peptide and transported to the plasma membrane. In addition to this protease inhibitor–protease regulatory mechanisms, DCs have evolved other specializations to make them especially suited for antigen processing and presentation.

DC endosomes and cross-priming In 1976, Bevan first identified the process of cross-priming, wherein exogenously derived antigen, antigens not expressed by the APC, can be presented on MHC class I molecules. This cross-presentation phenomenon has been documented time and again since that time, primarily in macrophages (Shastri and Gonzalez, 1993; Huang et al., 1994; Norbury et al., 1995; Carbone et al., 1998). Recently, it has been asked whether cross-presentation occurs in DCs. It has been demonstrated by several groups-to-occur using phagocytosed apoptotic cells (Albert et al., 1998), soluble macropinocytosed ovalbumin (Norbury et al., 1997), and receptor-mediated internalized immune complexes (Regnault et al., 1999). Numerous studies have proposed various models to explain the cross-presentation phenomenon (reviewed in Yewdell et al., 1999). A recent study has demonstrated a constitutive endosome-to-cytosol transport mechanism that is restricted to DCs, is antigen-specific and size-selective (Rodriguez et al., 1999). Perhaps, DC endosomes bear specialized translocators to facilitate the endosome-tocytosol transport of internalized antigen for the purpose of cross-priming. Whether all DC endosomes express this presumptive translocator or merely a subset, whether this process is constitutive in DCs and inducible in other bone marrow-derived cells (e.g. macrophages), and the molecular identity of these translocators all remain to be determined (Watts, 1999).

CIIVs: specialized transport vesicles In addition to specialized conventional endosomes, DCs have evolved endocytic transport vesicles to ferry peptide-loaded MHC class II molecules to the plasma membrane. These

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vesicles, CIIV, originally identified in A20 B cells (Amigorena et al., 1994) have been found in a slightly different incarnation in dendritic cells. DC CIIVs, unlike B cell CIIVs, lack any antigenprocessing machinery and are not accessible to endocytic ligands or receptors. DC CIIIVs are specialized compartments, containing costimulatory molecules (CD86), peptide-loaded MHC class II molecules, and MHC class I molecules, that exist during the intermediate stage of DC development (Turley et al., 2000). The disappearance of CIIV correlates with transition to the mature stage of DC development and the appearance of high cell surface levels of peptideloaded MHC class II and co-stimulatory molecules. Preliminary experiments suggest that this process is actin-dependent (Turley et al., 2000). CIIV biogenesis is probably a selective sorting and budding event from lysosomes, and the trafficking of membrane proteins to and from CIIVs is under active investigation.

Langerhans cell granules There has been a longstanding interest in the function of LCGs or Birbeck granules. Their origin and purpose remain elusive. The two most commonly debated origins are secretory and endocytic. In the secretory model, the granules bud from the Golgi stacks and fuse with the plasma membrane, exocytosing their putative contents, while in the endocytic model, the granules invaginate from the plasma membrane, bringing antigens and/or nutrients into the cell (Hashimoto and Tarnowski, 1968; Bartosik, 1992; Bucana et al., 1992). The evidence supporting an endocytic function for LCGs has at times seemed contradictory. While some groups found that HRP did not accumulate in LCs (Wolff and Schreiner, 1970), others demonstrated the accumulation of HRP in LCGs (Bartosik, 1992). LCGs appear to concentrate lectins and lectin conjugates, such as conalbumin and ferritinconjugated conalbumin, but did not accumulate ferritin (Takigawa et al., 1985). Ferritin immune complexes were also internalized and accessed LCGs (Takigawa et al., 1985). Using immunogold histochemistry and electron microscopy, the

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trafficking of CD1 and HLA-DR antibodies was followed in the endocytic pathway of LCs. At successive time points, the antibodies localized to coated vesicles, Birbeck granules, endosomes and lysosomes, providing evidence that LCGs are part of the endocytic pathway (Hanau et al., 1987). Although LCGs do not appear to be essential for the normal function of human LCs (Mommass et al., 1994), many have speculated that LCGs play a role in antigen sequestration, accounting for the ability of LCs to retain antigen in an immunogenic form for several days. A type II Ca2 lectin with mannose-binding specificity has been identified that is exclusively expressed by LCs; it is called langerin (Valladeau et al., 2000). Langerin is associated with LCGs and, surprisingly, transfection of langerin into fibroblasts cells results in the appearance of structures resembling LCGs by electron microscopy (Valladeau et al., 2000). Langerin was identified by its reactivity to the monoclonal antibody DCGM4, which was generated by immunization with human in vitro-generated DCs derived from CD34 stem cells. It is a 40-kDa protein and was cloned using DCGM4 expression cloning (Valladeau et al., 1999). The Lag antibody, a reagent used for decades to immunolocalize LCGs and label LCs, is reactive to an intracellular epitope of the langerin protein (Valladeau et al., 1999). The identification of langerin and its documented cellular functions as a potential antigen receptor and inducer of LCGs should prove useful for the elucidation of LCG biogenesis and function.

CONCLUSIONS Over the past 25 years, numerous studies have contributed to an emerging understanding of endocytosis in DCs. While initial studies suggested that DCs were weakly endocytic, the development of bulk culturing methods has helped to establish that early/immature DCs are endocytically active and that endocytosis is developmentally downregulated in DCs. Bulk culturing methods have facilitated careful studies of DC developmental and cell biology in

recent years. Endocytic internalization, recycling assays and subcellular fractionation have been instrumental in elucidating the nature of the endocytic specializations that make DCs the pre-eminent APCs of the immune system. Video microscopy is emerging as an exciting technique for visualizing antigen-presenting functions and membrane transport phenomena in DCs. Many questions remain regarding the characteristics of the novel endocytic organelles present in DCs. There have been new studies suggesting that DC possess endosomes specialized for endosome-to-cytosol antigen transport and other recent experiments have characterized endocytic pathway vesicles specialized for the transport of the antigen-presentation machinery. Recently, the protease cathepsin S and its protease inhibitor cystatin C were found to contribute to the regulation of MHC class II trafficking. While there have been many advances in our understanding of endocytosis and the endocytic pathway in DCs, many questions still remain. The nature of the signals involved in the downregulation of endocytosis and in the developmental remodeling of the MHC class II trafficking pathway is still an active area of inquiry.

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PLATE 16.2 MHC class II molecules (purple) are synthesized in the endoplasmic reticulum (ER). MHC class II molecules are associated with the invariant chain (blue) during transport to the endocytic pathway. Peptide is loaded onto MHC class II molecules within the endocytic pathway. The specific routes by which peptide-loaded MHC class II molecules reach the cell surface involve lysosomes and the nonlysosomal CIIV (transport vesicles that carry peptide loaded MHC class II, co-stimulatory molecules and MHC class I). Antigens (pink) internalized by DCs by endocytosis may also be loaded onto MHC class I molecules. DCs are particularly adept at this so-called cross-presentation of antigens. Endosomes and Birbeck granules may possess channels, allowing for the transfer of endocytosed antigens to the cytosol, that facilitate cross-presentation.

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17 Imaging dendritic cells: a primer Glenn D. Papworth, Donna Beer Stolz and Simon C. Watkins University of Pittsburgh, Pittsburgh, Pennsylvania, USA

A series of static pictures from sections, which is the problem that plagues the morphologist in studying most [regenerating] systems. Elizabeth Hay

INTRODUCTION

effecting the biology of the cells themselves. In the past this goal would have been extraordinarily difficult to achieve. However, developments in microscopic technology have empowered modern microscopic imaging such that it is now possible to undertake these seemingly forbidding tasks. The dendritic cell (DC) represents an ideal target for these types of analyses and implementation of the various microscopic technologies to the study of this cell type is of considerable importance to maximizing our understanding of the biology of these cells. The goal of this chapter is to provide a primer of the methods that we have developed or use to facilitate the use of microscopic imaging tools for the study of DC form and function. This will allow laboratories that do not specifically focus on the application of microscopy tools to perform useful quantitative analyses of DC biology using these approaches. Essentially each approach is discussed, standard methods and examples are given and the limitations of each approach

In the post genome era of biomedical research, understanding the functionality of molecules at the tissue, cellular and subcellular level will become predominant. In this era we must move beyond static ‘snapshots’ of cellular state to an understanding of the biology of cells over time and in three-dimensional space. Furthermore, within this cellular environment it is expected that we will be able to study the regulation of expression, the functional role(s) and interactions of multiple different molecules concurrently and to determine the effects of these molecules on cell development, organization and fate over extended periods of time. To perform this type of study it is necessary to involve an entire repertoire of microscopic analyses, from high-resolution analyses of cell shape and molecular distribution using electron microscopy to multiparametric analyses of cells at the light microscope level which allow definition of molecular distribution and behavior without Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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defined. The chapter is divided into sections which deal with light (fixed cell, tissue and confocal) and electron (scanning and transmission) microscopies. Due to space limitations the discussion is limited to fixed cell methods routinely in use in our laboratory, and we have not attempted to include any description of live cell techniques. It is hoped that individuals will be able to implement DC imaging solutions within their own laboratories if the methods within this chapter are followed carefully.

LIGHT MICROSCOPY Preparation methods Perfusion fixation In situ perfusion fixation is essential when fixing large tissue samples, and is particularly useful for fixing soft tissues. When used in conjunction with further post-dissection fixation as described, perfusion fixation gives better morphological preservation than post-dissection fixation alone. The fixative used and method of perfusion, however, depends on the species, tissue and antigen/s to be studied. The example given here is a protocol for perfusion fixation of a mouse in a fluorescence immunohistochemical study of in vivo migration of injected dendritic cells, focusing on pancreas and spleen (Feili-Hariri et al., 1999). Immediately following euthanasia of the mouse, expose the heart carefully to minimize bleeding. Insert a syringe (23G needle) filled with ~20 mL of PBS (phosphate-buffered saline: 8 g/L NaCl, 0.2 g/L KCl, 1.15 g/L Na2HPO47H2O, 0.2 g/L KH2PO4, pH 7.4) into the left ventricle. Cut open the right ventricle for drainage, and slowly inject the PBS continuously into the heart until little blood is seen leaving the right ventricle. Good perfusion can be seen in any visible blood-rich organs slowly turning a white-gray color. Insert a syringe into the hole in the left ventricle made by the first syringe, and slowly perfuse with ~20 mL of freshly prepared 2% paraformaldehyde (4C). During perfusion,

muscle spasms will occur indicating passage of fixative to the extremities, following perfusion the animal should be rigid. Tissues are then removed and fixed on ice according to the appropriate protocol (as described below). Paraformaldehyde fixation and sucrose infusion Prior to cryosectioning, fixation and subsequent sucrose infusion improves morphological preservation of tissues. Allow fixation to proceed at 4C for an empirically determined time ranging from 15 min to 4 hours (generally 1 hour will suffice), depending on the tissue. As fixatives can destroy antigenicity, for immunohistochemical studies the fixation time required should be minimized so as to prevent loss of antibody recognition of epitopes. Place dissected tissue in labeled 20-mL snap-cap glass vials (for very small samples use silanized glass vials). Fill the vials with freshly prepared (2% for immunohistochemistry) paraformaldehyde (PFA) fixative at 4C. Following fixation, wash tissue samples twice in PBS. Infuse samples in 30% sucrose in PBS overnight prior to rapid freezing (see below). Quick freezing and storage of tissue in preparation for sectioning Optimally, freezing must be sufficiently rapid to stop artifacts caused by ice crystals forming in tissue during the freezing process. If tissues are slow frozen by placing directly in a freezer or by direct immersion in liquid nitrogen, the periphery of the tissue will appear to be full of holes (like swiss cheese) due to the formation of ice crystals. The reason ice forms when tissue is directly immersed in liquid nitrogen is that the cold nitrogen boils when it comes into contact with the warm tissue, thereby generating a gaseous, insulating shell around the tissue which slows freezing considerably. Thus a transition fluid which is cold enough to ensure rapid freezing, but which will not boil in the presence of the relatively warm tissue sample must be used. Two commonly used options are liquid propane or isopentane. Though liquid propane

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does freeze tissue very rapidly and is the optimal transition fluid, considerable care must be taken when propane is used as it is very volatile and flammable. Generally we use isopentane for safety reasons. Our general method for freezing is to chill a 500-mL Pyrex beaker by immersion in a Dewar flask or expanded polystyrene box filled with liquid N2. Fill beaker with isopentane. Label one end of a filter paper strip with pencil and place specimen on the other end with forceps. Immerse specimen in cold (but liquid) isopentane for about a minute. If the isopentane becomes solid it can be warmed slightly with a conductive metal object, for example a large screwdriver or copper rod. Specimens may be stored in either liquid nitrogen or in a 80C freezer, generally we store at 80C as large tissue blocks often crack under liquid nitrogen. Note that the orientation of the tissue must be noted appropriately when freezing, as it is often difficult to later determine orientation in frozen tissues. Cryosectioning Paraffin and methacrylate sections are substrates which give better morphological detail of cells and tissues than cryosections. Unlike cryosections, however, in sections prepared using these techniques native protein form is not preserved as the tissue must be dehydrated, infused with paraffin wax or plastic and then heated for an extended period of time. This will often change the conformation of proteins within the specimen and immunohistochemical markers do not generally work on this type of specimen. For detailed procedures for use of cryostats, the reader should consult literature regarding the specific instrument being used, however some general rules are appropriate to ensure good morphological preservation. Cutting temperature range is usually 30C to 20C depending on the tissue. Softer tissues (eg. pancreas, spleen) are cut at colder temperatures than harder tissue (e.g. muscle). Cryostat temperature must be stabilized for 30 minutes before use after the chosen temperature is reached or temperature is altered. Wrinkled sec-

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tions suggest the cryostat is too warm; brittle sections suggest too cold. Preparation of cultured dendritic cells for immunocytochemistry Three types of preparations of cultured dendritic cells are commonly used for fluorescent immunolabeling: cytospins, DCs plated onto coverslips or suspensions of DCs prepared for cryosectioning. Ideally, cells plated onto coverslips are the best substrate for labeling methods, as an appropriate native cellular morphology is maintained. However, as DCs are only weakly adherent, care must be taken when preparing and labeling cells in this fashion. For routine labeling we commonly use cytospins of DCs, though cells prepared in this fashion show little native morphology. More recently we have developed a method which allows preparation of DCs for cryosectioning from suspensions. Cytospins Cytospins essentially rely on pelleting a thin monolayer of cells onto a glass slide within a centrifuge. As a process, this is very damaging to cell morphology, though it is generally the only way to prepare nonadherent cells for immunocytochemical analysis, unless labeling is done in suspension. As DCs are weakly adherent to appropriately prepared substrates, and can be labeled if treated correctly we try to avoid the use of cytospins unless absolutely necessary. The most critical variables to consider when preparing cytospins are the spin speed and the number of cells used. A major advantage of this technique is that relatively few cells need to be used. For DCs we generally use about 1  104 cells; if more cells than this are used the resultant preparation will appear clumped as cells are spun down on top of each other. However, it is possible to use as few as 1  103/ slide for this technique. To optimize morphology spin speed and time must be controlled. For DCs we prepare samples at 600 RPM for 6 minutes. To ensure an optimal preparation the slides are polylysine coated prior to use or, more conveniently, Superfrost slides (Fisher,

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Pittsburgh PA, USA) can be used. Once the cytospin is complete the cells must be washed with PBS and used immediately; if they are to be stored they should be dehydrated in cold methanol (20C) air dried and stored at 80C. An important thing to recognize is that the process of preparing cytospins inevitably permeabilizes cells, thus if it is important to distinguish between surface and cytoplasmic antigen this technique cannot be used. Cell adhesion method DCs will adhere to glass, though only weakly. However for many immunocytochemical experiments in which resolution must be optimized, or if there is a need to distinguish between cell surface and cytoplasmic labeling, this is the method of choice. Generally we rely on crosslinking the DCs to coverslips which have been coated with either gelatin or polylysine. To this end coverslips (we use 12-mm-round coverslips) are coated in poly-L-lysine (Sigma, St. Louis, MO, USA) or 2% gelatin in water and allowed to air dry. Cold (4C) DCs are plated onto the coverslips at high concentration in a small volume (generally 5  104 in 25 µL of proteinfree medium or PBS). When cold, DCs do not attach; however, as they are warmed to 37C they will attach to substrates and show a typical dendritic morphology. The cells are warmed and left in a moist chamber for 10–15 minutes. Subsequently they are lightly fixed in 2% paraformaldehyde for 10 minutes and then washed in PBS. The process not only fixes the cells but also ensures that they remain adhered to the coverslip. If this fixative does not provide adequate adhesion to the coverslip, it is possible to add a crosslinking fixative at low concentration, in this case we use 2% paraformaldehyde containing 0.01% glutaraldehyde for 10 minutes. As mentioned above, an important reason for preparing cells by adhesion is that, depending on the objective of the experiment, experimental protocols can be designed to detect antigens associated with the DC membrane, or antigens contained within the cell cytoplasm. Caution should also be exercised since epitopes can be altered by fixation and antibodies may not bind

under all fixation protocols. The basic protocol described above can be used to selectively label cell surface antigens as no permeabilization has been performed. Alternatively, to label cytoplasmic antigens, following the light paraformaldehyde fixation described above, the cells are postfixed for a further 10 minutes in 2% PFA/0.1% Triton X-100 in PBS on ice which will permeabilize cell membranes. Following fixation, pipette off fixative and wash (5 min/wash) twice in 4C PBS and proceed as described in the basic labeling protocol below. Preparation of cell suspensions for cryosectioning When an abundant source of cells is available (at least 106 cells) it is reasonable to prepare a suspended pellet of cells which can then be cryosectioned as described earlier. Cool 1065  106 DCs in media (for example in 15-mL Falcon tubes) on ice. Centrifuge cells for 5 minutes at 800  g, 4C, in a tabletop centrifuge. Pipette off culture media and resuspend cells in cold PBS. Centrifuge to a pellet, pipette off PBS, and fix by resuspending cells (15 minutes on ice) in 1–2 mL 2% paraformaldehyde. Centrifuge cells to a pellet for 5 minutes at 800  g, 4C, pipette off fixative, and resuspend cells in 250 µL of 4C PBS containing 10-µm colored latex beads (Molecular Probes, Eugene, OR, USA) at a similar concentration to the cells. The cells are then spun through Histogel (Fisher, Pittsburgh, PA, USA) in a microfuge tube, fixed and sucrose infused as described for cryosections above. Once frozen, the location of the cells within the Histogel may be identified by the presence of colored beads which are of a similar size and density to the cells. Fixation methods So far throughout this chapter we have used 2% paraformaldehyde as the primary fixative for all experiments. The reason for this is simple: it provides adequate fixation with mimimal disruption of cellular morphology and protein structure. In our laboratory we tend to avoid the use of dehydration fixation unless absolutely necessary. The reasons are that during

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dehydration, cytoplasmic proteins often precipitate out onto cytoskeletal elements and give an apparently erroneous localization, and commonly antibodies may not recognize their epitopes following this fixation method. On the other hand, while the combination of paraformaldehyde fixationfollowedbyparaformaldehydeandTriton X-100 does permeabilize cells, it may not be sufficient if nuclear antigens or DNA laddering (as found in apoptosis) is being studied. Furthermore,ifcellspreparedascytospinsaretobestored for an extended period of time, they should also be fixed by dehydration prior to placing in a 80C freezer. Thus on occasion, dehydration fixation is warranted.

Colorimetric and fluorescence immunohistochemical labeling methods Fluorescence immunohistochemistry depends on the binding of a primary antibody which may be conjugated to a fluorochrome or detected by a fluorescent secondary antibody (as is described below). This method is quantitative; it allows comparative subcellular optical detection of multiple protein localizations within cultured DCs or tissue sections, and the localization of the fluorescent label is constrained by the binding site of the primary antibody. In contrast, colorimetric methods such as immunoperoxidase are not quantitative, in that the labeling intensity depends not only on the amount of antibody binding, but also on the length of time that the sections are incubated in the reactive substrate. We find that in inexperienced hands this often leads to false positives. Furthermore, the reaction product is diffusible, such that the method is certainly not high resolution (at best cellular resolution may be achieved). Finally, it is generally impractical to perform multicolor labeling experiments using these techniques. While skilled immunocytochemists are able to perform double-labeling experiments using colorimetric methods, this is quite difficult and generally prone to error. To this end we avoid colorimetric methods unless absolutely necessary.

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Basic labeling technique for all types of cells/tissues The following labeling methods are common to cells and tissue sections. Single-labeling experiments involve cells or sections being reacted with a primary antibody and then a secondary antibody–fluorochrome conjugate. By replacing primary and secondary antibodies with mixtures of two primary and two secondary antibodies, multicolor immunofluorescence of multiple antigens on the surface or within the cytoplasm of DCs can be performed. Such multilabel experiments require that the secondary antibodies be directed against primary antibodies of different species and be conjugated to fluorochromes of differing excitation and emission wavelengths. A simple labeling regimen for cells and tissues that is routinely used in our laboratory can be employed with satisfactory results on most biological samples. It must be realized that this technique is not amenable to all antibodies, and modifications are often required. Remove frozen sections or cytospins from freezer and place in a humidified chamber (a plastic slide box with wet tissues in the bottom is ideal for this purpose). If cells on coverslips are to be used, the coverslips are inverted onto puddles of buffer on clean parafilm. Delineate the sections or cells on the slide using a PAP pen (Research Products Intl Corp., Mount Prospect, IL.). This will provide a hydrophobic barrier about the cells or tissue sections and minimize the antibody volume used, but is not necessary if cells are plated onto coverslips. Rinse cells or sections three times in PBS containing 0.5% bovine serum albumin (BSA), 0.15% glycine (PBG buffer). Incubate in 5% nonimmune goat or donkey serum in PBG buffer for 30 minutes at room temperature. This is done using a vacuum pipette to remove buffer from one side of the slide while fresh buffer is introduced from the other side of the section. In the case of cells grown on coverslips, the coverslips are simply transferred from one drop of buffer or reagent to another fresh drop on the same piece of parafilm with blunt forceps. Generally the blocking

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serum used is from the same species used to generate the secondary antibodies. Primary antibodies diluted in PBG buffer are cleared by centrifugation at 13 500g, 4C for 2 minutes, then added to samples at dilutions empirically determined for 1 for 2 hours at room temperature, or overnight at 4C. Between 25 and 50 µL is used for each slide. Dilutions for antibodies are sometimes provided by suppliers, but are more often not, so dilution series must be done initially to determine the proper dilution for each antibody. Sections are then washed five times in PBG buffer. Fluorescently tagged secondary antibodies, diluted in PBG buffer and cleared by centrifugation at 13 500 g, 4C for 2 minutes, are then added to the sections for 1 hour at room temperature. The samples are then washed three times in PBG buffer, three times in PBS, and coverslipped using gelvatol (see below for recipe and method). Negative controls are very important internal labeling controls. Some controls we routinely use are omission of primary antibody, followed by addition of secondary antibody. Isotype controls involve use of nonimmune purified antibodies, either pre-immune rabbit IgG (for rabbit polyclonal antibodies), or monoclonal antibodies such as mouse IgG1, IgG2a, IgG2b, IgM, etc., identical to the primary antibodies used in the study. Additionally, control antibodies should be employed at the identical concentrations used in the study. Labeling for increased sensitivity The sensitivity of immunofluorescent staining can be increased by enhancement using streptavidin and biotin. A centrifuged, diluted biotin–secondary antibody conjugate is applied to the primary antibody-labeled sections as per the basic protocol and washed 3 times in PBS (15 min per wash). A streptavidin–fluorochrome conjugate is then applied, also as per the basic protocol.

Slide mounting technique Lay clean coverslips on paper towels and place a drop of aqueous mounting material (molviol or

gelvatol are our preferred materials; see below for gelvatol recipe and preparation) in the middle of the coverslip. Invert slides on coverslip (do not apply pressure as this will damage the section). Leave slides for 30 minutes at room temperature or in the fridge overnight to allow gelvatol to harden (under aluminum foil to keep light out). Mounted slides can be stored at 4C in a closed slide box. In the case of cells mounted on coverslips, the reverse procedure is used, in that the gelvatol is placed on a slide and the coverslip gently placed onto the gelvatol. Gelvatol preparation To prepare gelvatol we use a modification described in Watkins (1989): Slowly add 20 g of polyvinyl alcohol in 5-g increments every hour to a beaker of stirring PBS. Keep beaker covered. Transfer beaker to cold room and stir overnight. The next morning, remove beaker from cold room and continue to stir at room temperature. Slowly add 3 g polyvinyl alcohol and a few crystals of sodium azide and continue to stir until dissolved (this could take an additional day). Finally add 50 mL glycerol and stir until solution is uniform. Centrifuge gelvatol at 10 000–30 000 g for 1 hour to remove particulates. Store final product in 3–10-mL syringes in the refrigerator. Choosing fluorochromes Using modern fluorochromes such as the carbocyanine (Amersham) dyes or Alexa (Molecular Probes) dyes, we do not recommend the use of antifade reagents. They are not necessary given the high quantum efficiency of the dyes in use. We strongly suggest that investigators follow Table 17.1 when choosing dyes.

FLUORESCENCE AND CONFOCAL MICROSCOPY DCs can be identified according to many different phenotypic characteristics, related both to their tissue of origin and stage of differentiation (Clark and Hart, 1999). As such, different

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TABLE 17.1 Dye families and suitability for fluorescence imaging Dye color (Emission)

You should use:

You should not use:

Blue, no really good dyes

AMCA

Cascade blue

Alexa 488

FITC

Green

Red Far red

Cy2 Bodipy FL Oregon green Cy3 Alexa 562 Cy 5

Phycoerythrin TRITC

populations can be fluorescently immunolabeled as described earlier, using antibodies to one or more molecules expressed by the cells (e.g. FITC-conjugated monoclonal antibody (mAb) specific for CD80, PE-conjugated mAb specific for CD40; Feili-Harari et al., 1999). Fluorescence microscopy can then be used to examine the specific locations of different populations of endogenous DCs within sections of various tissues (e.g. spleen, pancreas (BarrattBoyes et al., 1997), tonsil (Zhong et al., 1999)) and their interactions with other types of cells within the tissue. Fluorescence microscopy of immunohistochemically labeled tissue sections can also be used to track the migration of DCs differentiated in vitro, following in vivo administration (e.g. Barratt-Boyes et al., 1997). Labeling of administered DCs with an appropriate lipophilic fluorescent marker (e.g. DiIC18(5), see Plate 17.1) can also allow long-term cell tracking and thus delineation of DCs expressing specific cell marker phenotypes from the total population of injected DCs. Fluorescence confocal microscopy and fluorescence light microscopy of DC preparations do not differ significantly in terms of the theory of appropriate fluorescent multilabeling. Both imaging techniques essentially face the same limitations, in that good multiple fluorescence signal isolation necessitates good spectral separation of the emission wavelengths of the multiple fluorophores. Ideally, multiple fluorophores would also strongly absorb light at a coincident

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excitation wavelength. The distinct advantage that confocal microscopy has over both brightfield and fluorescence light microscopy is that confocal can provide depth discrimination in a sample. The optical sectioning capabilities of confocal microscopy enable threedimensional morphological data to be obtained on appropriately fluorescent-immunolabeled antigens on dendritic cells. Thus, for example, it is possible using confocal microscopy to obtain more detailed information of the interdigitation of DCs with other cell types in various tissues. If fixed laser illumination, pinhole diameter and photomultiplier settings are used, sections that have been stained equivalently can also be compared in terms of relative staining intensity for different antigenic phenotypes (e.g. Feili-Hariri et al., 1997). By rejecting fluorescent emissions from planes other than the plane of focus within a specimen, confocal microscopy also allows for clearer imaging in thicker tissue sections, and indeed raises the potential for in vivo imaging of dendritic cell populations. Many fluorophores commonly used for immunolabeling of dendritic cells excite and emit at visible or ultraviolet wavelengths (e.g. DiIC18(5), exc.644nm/em.663nm). Such wavelengths are, however, attenuated by absorption and scattering in living tissue, meaning that the degree of excitation of the focal plane and the fluorescent emission returning to the confocal detector is reduced. Thus the ability of conventional confocal microscopy to penetrate deep into living tissue is limited.

ELECTRON MICROSCOPY (EM) Transmission electron microscopy of cells isolated in suspension This technique is useful for freshly sorted or isolated dendritic cells or cultured dendritic cells. DCs have unique ultrastructural characteristics that set them apart from other mesenchymal cells, including leukocytes such as monocytes, T and B cells (Figure 17.2). Typically, a visible pellet is recommended, and consists of approximately

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FIGURE 17.2 Transmission electron micrograph of murine DC, showing typical DC morphologic features. Cytoplasmic veils are clearly seen at the cell surface (arrows). Furthermore, the cytoplasm shows relatively few granules when compared with macrophages, for example.

5  105–106 cells. However, if acquisition of this amount of cells is problematic, it is possible to proceed by processing ‘blindly’, assuming that the small number of cells will remain adherent to the edge of the microfuge tube. In this case, we recommend that the cells be pelleted with reference to the tab on the flip cap of the microfuge tube. If the tab is oriented towards the outside of the fixed angle microfuge rotor when the cells are pelleted, they will remain there throughout processing and will serve as a reference for sectioning. Alternatively, if a swinging bucket microfuge rotor is available, the cells will remain attached to the bottom.

Protocol Note of caution regarding TEM reagents: Most reagents used for EM protocols are fixatives and thus are irreversible chemical crosslinkers. As a result, all fixatives, such as glutaraldehyde, paraformaldehyde and osmium tetroxide, dehydration solvents, such as propylene oxide, as well as post-stains, such as those containing lead and uranium salts, are toxic and should be handled with gloved hands. Fixatives and solvents should be used only under a working, certified fume hood. All reagents should be disposed of in accordance with your institution’s hazardous waste disposal protocol.

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Isolated suspensions of DCs are pelleted in a 0.5 or 1.5-mL microfuge tube at 100 g, supernatant removed, then resuspended 2.5% glutaraldehyde in PBS (8 g/L NaCl, 0.2 g/L KCl, 1.15 g/L Na2HPO4 7H2O, 0.2 gm/l KH2PO4, pH 7.4). Cells are repelleted at 100 g and maintained in the pellet for 1 hour to overnight at 4C. Pellets are washed 3 times in PBS then post-fixed in 1% OsO4, 1% K3Fe(CN)6 for 1 hour. Following three PBS washes, the pellet is dehydrated through a graded series of 30–100% ethanol, 100% propylene oxide then infiltrated in 1 : 1 mixture of propylene oxide : Polybed 812 epoxy resin (Polysciences, Warrington, PA, USA) for 1 hour. After several changes of 100% resin over 24 hours, the pellets are embedded in molds, cured at 37C overnight, followed by additional hardening at 65C for 2 more days. Thick sections (300 nm) for light microscopy are heated onto glass slides and stained for 1 minute with filtered 0.5% Toluidine blue in 1% aqueous sodium borate, then rinsed in water. Stained thick sections can be coverslipped with Permount (Fisher, Pittsburgh, PA, USA) for archiving. Ultrathin (60 nm) sections are collected on 200mesh copper grids, stained with 0.2 µm filtered 2% uranyl acetate in 50% methanol for 10 minutes, washed three times in 0.2 µm filtered ultrapure water then stained in 0.2 µm filtered 1% aqueous lead citrate for 7 minutes. Sections are then ready for viewing using transmission electron microscopy.

Transmission electron microscopy of dendritic cells grown on tissue culture dishes, filter inserts or coverslips This technique is useful when cells are required to be attached to a substratum, which is often the case when examining DC interacting with other cells such as T cells, or monolayers of cells grown on tissue culture filter inserts. If removal of cells from the surface is disadvantageous, then they can be processed and removed from the dish or coverslip once they are fully cured and embedded. The embedded cells can then be sectioned en face or cut out and re-embedded for cross-sectioning. Filters can be removed

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from their supports once they are infiltrated with resin, then embedded and cured in molds and cross-sectioned. Protocol Cells grown on tissue culture plastic dishes or coverslips are fixed in 2.5% gluaraldehyde in PBS overnight at 4C. Monolayers are then washed in PBS three times then post-fixed in aqueous 1% osmium tetroxide, 1% Fe6CN3 for 1 hour. Cells are washed three times in PBS then dehydrated through a 30–100% ethanol series then several changes of Polybed 812 embedding resin (Polysciences, Warrington, PA, USA). Note that since cells are grown on solvent-soluble polystyrene tissue culture dishes, the propylene oxide steps described in the pellet processing above must be avoided. Resin-filled BEEM capsules are inverted over the cells in the dishes and are cured overnight at 37C, then cured for 2 additional days at 65C. BEEM capsules are snapped from the dish and inspected to assure that cells detach with the resin. At this point, cells can be processed for en face sectioning or cut and re-embedded for cross-sectioning.

SCANNING ELECTRON MICROSCOPY (SEM) DCs are often best identified from other leukocytes using surface topology afforded by scanning electron microscopy (SEM). This technique allows the visualization of the membrane ‘veils’ or ‘dendrites’, that readily distinguish DCs from other cell types (Figure 17.3, see also Lu et al., 1994). SEM requires a solid support so DCs must be lightly adherent to coverslips in order to visualize the membrane veils. Depending on the type of SEM, sample stubs are amenable to 18 or 12-mm-diameter round coverslips (Fisher Scientific, Pittsburgh, PA, USA). Live DCs are placed onto these coverslips in medium at 37C for 5–10 minutes. Cells are checked using phase light microscopy to assure that they have adhered to the coverslip. When attached, the medium is removed and cells are fixed in 2.5%

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FIGURE 17.3 Scanning electron microscopy shows a typical DC morphology. In both panels of this image typical DCs are shown. The most classical morphologic feature of the cells is the presence of veils as defined by arrowheads.

glutaraldehyde in PBS overnight at 4C. Samples are washed three times in PBS, post-fixed for 1 hour in aqueous 1% osmium tetroxide, then washed three times in PBS. Samples are dehydrated through a graded ethanol series (30–100%), further dehydrated by three additional 15-minute washes with absolute ethanol then critical point dried (Emscope CPD 750, Ashford, Kent, UK). Samples are mounted onto aluminum stubs then sputter coated with gold/paladium (Hummer VI, Technics West, San Jose, CA, USA). Samples are ready to be viewed on a scanning electron microscope. If cells are already fixed in suspension, it is often difficult to afix the cells to a solid support. However, we have had varied success with the following method. Wash fixed cells with PBS, then drop the concentrated DC suspension onto a 3% gelatin-coated coverslip. Let cells drop by gravity through the gelatin for about 10–15 minutes. Look to see that the cells are on the coverslip by phase light microscopy. Very gently remove the supernatant, then gently add 2% paraformaldehyde in PBS to the cells. Inspect the coverslips using phase light microscopy to determine if cells have adhered to the coverslip. Let fix for 10 minutes. Proceed as above for liveattached DCs.

IMMUNOELECTRON MICROSCOPY (IEM) Immunoelectron microscopy is an essential tool for evaluating subcellular localization of molecules within cells at nanometer resolution. This technique is especially useful for colocalization studies involving two or more proteins. A wide variety of immunology reagents, including primary antibodies, and a large selection of variously sized (1–30 nm) colloidal gold conjugated secondary antibodies directed toward many immunoglobulin species are now available to examine DCs using these sophisticated imaging techniques. Additionally, new protocols developed using nano-gold particle technology exhibit increased penetration into tissue and cells. The nano-gold particles can then be silver enhanced to increase their size and enhance visualization of the molecule in question.

IEM protocol DCs are pelleted in a 0.5 or 1.5-mL microfuge tube, fixed by resuspension in 2% paraformaldehyde, 0.01% glutaraldehyde in 0.1 M PBS and stored at 4C for 1 hour. Cells are pelleted and resuspended in a small volume (10 µL) of 3%

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gelatin in PBS (maintained at 37C to keep gelatin liquid). The cell–gelatin pellets are solidified at 4C for 10–15 minutes, then fixed in 2% paraformaldehyde, 0.01% glutaraldehyde in 0.1 M PBS for 10 minutes. The pellet is loosened, and allowed to stand for 10 additional minutes in the fixative. Fixative is removed and cells are infiltrated in cryoprotectant solution. Cryoprotectants include 2.3 M sucrose in PBS or PVP:sucrose solution described by Tokuyasu (1989) and prepared using 1.1 M sodium carbonate in PBS (11.66 g NaCO3 qs to 100 mL with PBS), 4.0 mL; 2.3 M sucrose in PBS (78.66 g sucrose to 100 mL with PBS), 80.0 mL; polyvinylpyrrolidone, 20.0 g. The PVP is added to the sodium carbonate and mixing begins. Then the sucrose solution is added and mixed until completely dissolved. The cryoprotectant is stored in plastic transfer pipettes at 20°C. Cells are incubated in cryoprotectant overnight at 4C, then small chunks of gelatin– cell mixture are flash frozen on ultracryotome stubs under liquid nitrogen and stored in liquid nitrogen until use. Ultrathin sections (70– 100 nm) are cut using an ultramicrotome with cryo-attachment, lifted on a small drop of 2.3 M sucrose and mounted on Formvar-coated copper grids. Sections are washed three times with PBS, then three times with PBS containing 0.5% bovine serum albumin and 0.15% glycine (PBG buffer) followed by a 30-minute incubation with 5% normal goat serum in PBG. (As with fluorescence microscopy, the blocking serum is the secondary host serum, so either goat or donkey serum are the blocking sera). Sections are labeled with primary antibodies for 1 hour at room temperature. Dual labeling requires primary antibodies produced in two separate species (i.e. rabbit and mouse). Sections are washed four times in PBG and labeled with appropriate gold-conjugated secondary antibodies (Amersham, Arlington Heights, IL or Jackson ImmunoResearch Laboratories, West Grove, PA, USA), for 1 hour at room temperature. Sections are washed three times in PBG, three times in PBS, then fixed in 2.5% glutaraldehyde in PBS for 5 minutes, washed twice in PBS then

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washed six times in ddH2O. Sections are poststained in 2% neutral uranyl acetate, for 7 minutes, washed three times in ddH2O, stained for 2 minutes in 4% uranyl acetate, then embedded in 1.25% methyl cellulose. After drying grids overnight, samples are ready to be viewed using TEM.

GENERAL LITERATURE RESOURCES Light microscopy Harlow, Ed and Lane, David (1999). Using Antibodies: A Laboratory Manual. New York: Cold Harbor Laboratory Press. Spector, D.L., Goldman, R.D. and Leinwand, L.A. (eds) (1998). Cells, A Laboratory Manual, Vol. 3: Subcellular Localization of Genes and their Products. New York: Cold Harbor Laboratory Press. Watkins, S.C. (1989). ‘Immunohistochemistry, Unit 14.6’. In: Ausubel, M. et al. (eds) Current Protocols in Molecular Biology, Vol. 2. Chichester: John Wiley and Sons.

Electron microscopy The protocols listed describe procedures that work reproducibly in our laboratory, but are by no means the only protocols known to give satisfactory results when examining DC ultrastructure. For other procedures, as well as methods describing proper use of EM hardware, such as sectioning, critical point drying and sputter coating, as well as alternative staining techniques, we refer the reader to other very informative general sources. Buzzola, John J. and Russell, Lonnie D. (1999). Electron Microscopy: Principles and Techniques for Biologists. Boston: Jones and Barlett. Griffiths, Gareth (1993). Fine Structure Immunocytochemistry. Berlin: Springer-Verlag. Hayat, M.A. (ed.) (1995). Immuno-Gold Silver Staining: Principles, Methods and Applications. Boca Raton: CRC Press. Maunsbach, Arvid B. and Afzelius, Bjorn A. (1999). Biomedical Electron Microscopy: Illustrated Methods and Interpretations. San Diego: Academic Press.

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REFERENCES Barratt-Boyes, S.M., Watkins, S.C. and Finn, O.J (1997). J. Immunol. 158, 4543–4547. Clark, G.J. and Hart, D.N.J. (1999). In: Lotze, M.T. and Thomson, A.W. (eds). Dendritic Cells: Biology and Clinical Applications, San Diego: Academic Press, pp. 555–577. Feili-Harari M., Dong, X., Alber, S.M., Watkins, S.C., Salter, R.D. and Morel, P.A. (1999). Diabetes 48, 2300–2308. Lu, L., Woo, J., Rao, A.S. et al. (1994). J. Exp. Med. 179, 1823–1834. Tokuyasu, K.T. (1989). Histochem. J. 21, 163–171. Watkins, S.C. (1989). In: Skedman, J.G. et al. (eds) Current Protocols in Molecular Biology, New York: J. Wiley and Sons. Zhong, R.-K., Donnenberg, A.D., Zhang, H-F, Watkins, S., Zhou, J-H. and Ball, E.D.(1999). J. Immunol. 163, 1354–1362.

MATERIALS Light microscopy Although sources are not listed here, primary antibodies are available through many vendors, check primary literature for sources. The following list our preferred vendors for secondary reagents, dyes and general consumables

Jackson ImmunoResearch Laboratories, 872 West Baltimore Pike, PO Box 9, West Grove, PA 19390, USA. www.jacksonimmuno.com (secondary antibodies). Molecular Probes. PO Box 22010, Eugene, OR, USA. www.probes.com (secondary antibodies, fluorophores). Fisher Scientific, Pittsburgh, PA, USA. www.fishersci.com (coverslips, tissue culture material, cryochemicals). Sigma, PO Box 14508, St. Louis, MO 63178, USA. www.sigma-aldrich.com (general chemicals).

Electron microscopy Reagents and materials are purchased from suppliers specializing in electron microscopy. Electron Microscopy Sciences, 321 Morris Road, Box 251, Fort Washington, PA 19034 USA. www.emsdiasum.com Energy Beam, 11 Bowles Road, PO Box 468, Agawam, MA 01001, USA. www.ebsciences.com Ernest F. Fullam, Inc. 900 Albany Shaker Rd. Latham, NY 12110, USA. www.fullam.com Polysciences, 400 Valley Road, Warrington, PA 18976, USA. www.polysciences.com Ted Pella, Inc. PO Box 492477, Reading, CA 960492477, USA. www.tedpella.com Tousimis Research Corporation, PO Box 2189, Rockville, MD 20847, USA. www.tousimis.com

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PLATE 17.1 (a, b) These figures are derived from previous studies of cell tracking in primates in vivo (Barratt Boyes et al., 1997). Fluorescent microscopy of cryosectioned inguinal lymph node of chimpanzee 48 hours following subcutaneous injection of cultured DCs labeled with the lipophilic carbocyanine dye DiIC18(5). Sections were labeled with Hoechst 33322 (blue) to show cell nuclei, and monoclonal antibody (Cy3, shown as green in this digital print) to (a) CD3 or (b) HLA-DR. Dendritic cells double labeled with DiIC18(5) (red) and green indicates expression of (a) CD3 or (b) HLA-DR in subcutaneously administered cultured DCs. Bar = 100 microns. PLATE 17.1 (c) Confocal microscopy of cryosectioned spleen of mouse 48 hours following intravenous injection of GM DCs (bone marrow-derived DCs cultured with GM-CSF) labeled with Cy5-DiIC18(5). Sections were labeled with monoclonal antibody (FITC, green) specific for CD80. The two-color confocal images were derived from integration of images of the Cy5-DiIC18(5) signal (red) and the green signal of CD80-positive cells. Cells labeling positive for both the Cy5 dye and CD80 can be seen as orange cell profiles (arrows).

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18 NK cells Nadine C. Fernandez 1, Carole Masurier 1, Magali Terme1, Joseph Wolfers1, Eugene Maraskovsky 2 and Laurence Zitvogel1 1

Institut Gustave Roussy, Villejuif, France; 2 Austin and Repatriation Medical Centre, Heidelberg, Australia

If you think something so small cannot make a difference, try going to sleep with a mosquito in the room. Unknown

INTRODUCTION

T cells but also by directly modulating CD40dependent B-cell activation (Dubois et al., 1997). In contrast, there are as yet few reports describing direct interactions between DCs and NK cells. In addition, most of them have analyzed the lysis of DCs by activated NK cells (Chambers et al., 1998). In this chapter, after a brief introduction to NK cells, we will show that DCs are also capable of interacting with resting NK cells.

During the last few years, significant progress has been made in the understanding of both dendritic cell (DC) and natural killer (NK)-cell biology. The regulation of NK-cell functions has been studied in innate immune responses. Although there is accumulating evidence on signals that negatively regulate NK-cell functions, our understanding on the mechanisms leading to NK-cell activation is still sketchy. In addition to providing a first line of defense against pathogens, innate immunity dictates adaptive responses to antigen (Medzhitov and Janeway Jr, 1997). The role of DCs in modulating adaptive immune responses has been extensively investigated. Following antigen capture, these cells have been shown to be a cornerstone of antigen-specific T-cell primary and secondary immune responses (Banchereau and Steinman, 1998). In addition, DCs are essential for the development of antibody responses not only as a consequence of priming of naïve helper Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

NK CELLS Natural killer (NK) cells are CD3-negative lymphocytes which do not express antigen-specific receptors at their surface. In humans, CD56 serves as a tag to distinguish NK cells from other non-T lymphocytes, whereas in the mouse and rat, NKR-P1 molecules subserve the same function (Rolstad and Seaman, 1998). These nonclassical lymphocytes contribute to the host defense mechanisms by direct cytotoxicity of infected or transformed target cells (Trinchieri,

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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1989). Through their ability to exert their functions without prior immunization, they have been recognized as cellular actors of the innate immune system. The importance of NK cells goes beyond their capacity to lyse specific target cells. NK cells are also capable of releasing cytokines and chemokines that promote an inflammatory immune response (Bancroft, 1993). The production of IFNγ by NK cells plays an important role in regulating T cells, promoting a Th1 immune response. However, the mechanisms underlying the influence of NK cells on the subsequent adaptive immune response are not exactly known. It is possible that an interaction with a professional antigen-presenting cell (APC) such as DCs is involved. While the mechanisms used by NK cells to discriminate between target cells and normal cells remained poorly understood, great progress has been made recently. There is an emerging consensus that NK cells are regulated by a fine balance between positive and negative signals derived from membrane receptors that determine their ability to exert their functions in response to immune challenge. The cytotoxic activity of NK cells can be triggered by the interaction of several surface molecules. Activating receptors include those specific for MHC class I molecules and those specific for non-MHC ligands (Raulet, 1996; Lanier et al., 1997; Long and Wagtmann, 1997; Moretta et al., 1998). This latter family include integrins (such as lymphocyte function-associated antigen-1), members of the immunoglobulin superfamily (such as CD2, CD16, 2B4, NKp30 (Pende et al., 1999), NKp46 (Sivori et al., 1997) and NKp44 (Vitale et al., 1998)) and members of the C-type lectin superfamily (such as CD69, NKRP1). At present, there is no evidence for a unique NK cell-specific receptor that is responsible for initiating the cytolytic response; rather, it is likely that different receptors may be used depending upon the activation state of the NK cell and the availability of the relevant ligands on the target cell. In contrast to the limited understanding of the molecules responsible for positive signaling in NK cells, the involvement of receptors in negative signaling is well-defined.

First, an inverse correlation has been established between the expression of surface MHC class I molecules and the susceptibility to NK cell-mediated lysis (Ljunggren and Kärre, 1990). This led to the ‘missing self hypothesis’ which has been confirmed by the characterization of multiple NK-cell inhibitory receptors that recognize allelic forms of MHC class I molecules (Moretta et al., 1996; Lanier, 1998). These receptors prevent NK cells from lysing normal self cells expressing a cognate class I ligand. Consequently, NK cells lyse target cells in which class I expression is reduced or extinguished. This may allow the elimination of autologous cells in which expression of one or more class I alleles is prevented due to viral infection, mutation or transformation (Ljunggren and Kärre, 1985). Some tumor cells expressing a high level of autologous class I molecules are, nevertheless, sensitive to cytolysis by NK cells, indicating that class I deficiency is not the only mechanism of target cell discrimination by NK cells. NK cells are sentinels in the immune system, ever vigilant in scanning the cellular environment for an imbalance in the appropriate activating and inhibitory signals.

DC-MEDIATED TRIGGERING OF RESTING NK CELLS It has been demonstrated previously that the initiation of host-protective mechanisms against pathogens (viruses, parasites) involves macrophages, DCs and NK cells that may interact either locally at the ports of entry or distally, but most likely through soluble mediators such as IFN, IL-12 and TNFα. However, our understanding of how these components are functionally orchestrated to provide resistance during the course of infections or tumor spreading remains sketchy. Orange and Biron (1996) showed that murine cytomegalovirus induces IFNα/β-dependent NK-cell cytotoxicity and IL12-dependent NK-cell IFNγ production that inhibit viral replication. They suggested that bone marrow-derived NK cells are found in the splenic marginal zone during the course

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of the viral infection where they may receive co-stimulatory signals from accessory cells to produce high levels of IFNγ. A potential source of these signals are myeloid DCs which reside in the splenic marginal zone (Pulendran et al., 1997). Moreover, migration, inflammatory and protective antiviral responses mediated by NK cells into the virally infected livers were shown to be macrophage inflammatory protein (MIP)-1α dependent (Salazar-Mather et al., 1996, 1998). The location of the so-called accessory cells, the requirement for MIP-1α and IL-12 imply that DCs may be instrumental in triggering NK activity in these viral infections. In the course of leishmaniasis, Scharton-Kersten et al. demonstrated that IL-12 drives both NK-cell cytotoxicity activity and IFNγ production (SchartonKersten et al., 1995). Reis e Sousa et al. showed that DCs, NK cells and IL-12 are necessary for mice to survive acute toxoplasmosis (Reis e Sousa et al., 1997). They observed that DCs increase in numbers and redistribute to T-cell areas of the spleen. DCs, and not macrophages, in spleen are the key initial producers of IL-12 in response to Toxoplasma gondii antigens or LPS. Therefore, it has been suggested by Johnson and Sayles that DC, via IL-12, are critical in initiating innate NK cell-dependent and T-cell responses to Toxoplasma (Johnson and Sayles, 1997). NK-cell activation has the potential of contributing to beneficial immunoregulatory mechanisms critical in promoting early defenses against pathogenic agents and tumor metastasis. However, NK-cell overexpression of cytokines may be detrimental to the host. Therefore, their activation must be tightly regulated. Little is known about (1) the in vivo pathways modulating NK-cell activation in a tumorbearing host and (2) the NK cell activation threshold necessary to shape the antitumor immune responses. We now report a molecular crosstalk between murine DC and resting NK cells that is relevant in regulating innate antitumor immune responses in vivo (Fernandez et al., 1999a).

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Flt3 ligand (FL) induces tumor regression mediated by NK cells and involving the lymphoid-related DC subset FL is known to expand dramatically the number of DCs (Maraskovsky et al., 1996) and to enhance the generation of NK cells (Shaw et al., 1998) in vivo. Since FL has also been reported to have antitumor properties in SCID mice (Lynch et al., 1997), we investigated the potential in vivo regulation of NK activation by DCs and its relevance in antitumor immune responses in FL-treated mice. In MHC class I-negative tumor (mesothelioma AK7)-bearing mice, expansion of all DC subsets following FL (kindly provided by Immunex, Seattle, WA, USA) administration markedly prevented tumor growth not only in immunocompetent mice (Figure 18.1) but also in nude and Rag−/− mice, stressing that T cells were not required for the tumor growth delay. These antitumor effects were mainly mediated by NK cells as demonstrated by the complete

FIGURE 18.1 Tumor growth kinetics following FL administration in day 20 AK7 tumor-bearing B6 mice. Ten micrograms of FL were administered subcutaneously (s.c.) daily for 20 days. Mean tumor sizes are plotted with s.e.m. bars from the start of FL treatment. Asterisks depict significantly smaller tumors in FL-treated animals as compared with the PBS-injected group (P  0.05). Data were reproduced three times with similar results.

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abrogation of FL-mediated antitumor effects in beige mice and following the specific depletion of the NK cells in immunocompetent mice. Interestingly, additive antitumor effects could be achieved using a combination of NK-cell stimulatory factors with FL. While FL treatment or intratumor injection of the recombinant adenoviral vector encoding both IL-12 subunits (AdmIL-12) in established tumors induced tumor growth delay, the combination of both allowed transient complete tumor regression in all treated mice (Figure 18.2A). Similar results were obtained by combining human Chinese hamster ovary (CHO)-derived IL-15 (kindly provided by Immunex) systemic administration with FL (Figure 18.2B). Neutralization of endogenous IL-12 or IFNγ and blocking of the B7 molecules using CTLA-4 failed to inhibit the FL-induced tumor regression. Moreover, FL had significant antitumor effects in type I IFN receptor and Rag2 doubleknockout mice. Therefore, essential NK cell stimulatory cytokines were not involved in vivo. Interestingly, the antitumor effects were detectable following 10 consecutive days of FL therapy, timing corresponding to the peak in the DC expansion. They decreased up to 5–10 days following treatment cessation, corresponding to the decrease of the DC numbers (Maraskovsky et al., 1996). Indeed, the hypothesis whereby DCs could be involved in the FL-induced NK celldependent antitumor effects was confirmed by the fact that a selective depletion of a lymphoidrelated DC subset in tumor-bearing nude mice significantly inhibited the efficiency of the FL therapy (Fernandez et al., 1999a).

DCs promote NK cell cytolytic activity and IFNγ production in vitro To test the possibility that DCs are able to activate resting NK cells, we developed an in vitro system consisting of the co-culture of bone marrow-derived DCs (BM-DCs) or of the growth factor-dependent spleen DC line (D1) (Winzler et al., 1997) with freshly extracted resting NK cells in culture medium not supplemented with any cytokine.

FIGURE 18.2 NK-cell stimulatory factors markedly enhance FL-mediated antitumor effect. Day 20 established AK7 tumor-bearing B6 mice were given 10 µg of FL (s.c.) daily for 20 days. (A). Coadministration of AdmIL-12. 5  108 p.f.u. of AdCO1 (control empty viral vector) or AdmIL-12 were injected intratumorally once at day 10 of FL treatment. (B) Coadministration of IL-15. Fifteen micrograms of human CHO-derived IL-15 were injected intraperitonally daily from day 10 to day 15 of FL treatment. Asterisks represent significantly smaller tumors in FL plus AdmIL-12or plus IL-15-injected groups as compared with FLtreated animals (P  0.05). Mean tumor sizes are plotted with s.e.m. bars from the start of FL treatment. These experiments were performed at least twice with identical results.

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The cytolytic activity of allogeneic and syngeneic resting NK cells against YAC-1 cells was very significantly increased following the coculture with D1 cells either immature or induced to maturation using TNFα (Figure 18.3A and 18.3B). Similar results were obtained by co-cultivating resting NK cells with immature BM-DCs generated in GM-CSF plus IL-4. Once activated by DCs, NK cells are able to lyse not only MHC class I-negative tumor cells such as AK7 (mesothelioma, model used for in vivo studies) or MCA-101 (fibrosarcoma) but also MHC class I-expressing targets such as MCA-205 (fibrosarcoma). The cytokine release function of NK cells is also activated as demonstrated by the presence of IFNγ in the supernatant of DC/NK cell co-culture (Figure 18.3C). Moreover, the NKcell activation is dependent on the number of stimulating DCs as shown in Figure 18.3C. Mature DCs are known to secrete various cytokines capable of activating NK cells (Hart, 1997). However, these soluble factors did not play a major role in the activation of resting NK cells by DCS as shown by the complete abrogation of the DC-induced NK-cell cytolytic activity (Figure 18.4A) and IFNγ secretion (Figure 18.4B) following separation of both populations in transwell experiments. Thus, activation of NK cells by DCs requires intimate cell–cell contact.

FIGURE 18.3 DC directly stimulate NK cell functions in vitro. (A) Freshly isolated splenic NK cells derived from SCID BALB/c (allogeneic) or Rag2/ B6 (syngeneic) mice were co-cultured alone or together with TNFα-conditioned D1 cells. (B) Similarly, freshly isolated splenic NK cells derived from SCID BALB/c mice were co-cultured alone or together with immature or TNFα-conditioned D1 cells. Co-cultures were performed at a ratio of 0.5 NK cell to 1 DC for 18 hours. Viable lymphocytes were tested against YAC-1 cells in a classical chromium release assay. Results are expressed as the percentage of specific lysis at various effector:target ratios and represent the means of triplicate wells. (C) Supernatants from immature D1 cells and SCID mice-derived NK cells cultured alone or together at various ratios were harvested at 40 hours and assayed for the presence of mIFNγ by ELISA. Results are from a single experiment representative of a series of three to five. DENDRITIC CELLS AND INTERACTION WITH OTHER CELLS

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Resting NK cells cannot be triggered by targets or any APCs The activation of resting NK cells by DCs cannot be attributed to the fact that DCs might represent targets for NK cells. First, BM-DCs did not represent targets for resting NK cells (Figure 18.5A) and second, the co-culture of resting NK cells with known targets such as YAC-1 cells and P815-B7.1 (Chambers et al., 1996) did not induce their cytolytic activity (Figure 18.5B). The DCinduced NK-cell activation is not a property of any APCs since macrophages were unable to activate resting NK cells (Figure 18.5C). Moreover, this property seems to be specific to a subset of DCs since BM-DCs generated only in GM-CSF did not induce NK-cell activation (Figure 18.5C). Interestingly, DCs derived from splenocytes using FL, GM-CSF and IL-6 were not able to trigger significantly resting NK cell functions (M. Terme, unpublished results).

Adoptive transfer of DCs at the tumor site significantly impaired tumor progression in therapy

FIGURE 18.4 Cell contact-dependent activation of NK cells by D1 cells. Resting NK cells and TNFαconditioned D1 cells were cultured in a transwell plate together or separated by a porous membrane in the same well at a ratio of 0.5:1 for 18 hours. (A) Viable lymphocytes were tested against YAC-1 cells in a classical cytotoxicity assay. Results are expressed as the percentage of specific lysis at various effector:target ratios and represent the means of triplicate wells. (B) Supernatants were harvested and assayed for the presence of mIFNγ by ELISA. These data represent one out of three independent experiments achieving identical results.

To test directly the role of in vivo crosstalk between DCs and NK cells, tumor-bearing mice were injected at the tumor site with immature D1 cells. Similar results to those obtained following FL treatment were achieved, suggesting that the FL antitumor effects were, at least partly, due to the expansion of DCs. Moreover, demonstration of a major role of NK cells in these D1 cellmediated antitumor effects by abrogation of the effects following selective NK-cell depletion emphasize that DCs directly induce NK-cell activation in vivo.

DISCUSSION These results assign to DCs a novel role since we have shown that these cells are capable of triggering resting NK-cell cytolytic activity against MHC class I-negative and -positive targets and IFNγ secretion independent of exogenous cytokines (Fernandez et al., 1999a).

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FIGURE 18.5 (A) Neither BM-DCs nor D1 cells represent targets for resting NK cells. The cytotoxicity of freshly isolated splenic NK cells derived from SCID mice was tested against YAC-1 cells, BALB/c micederived BM-DCs (GM-CSF  IL-4) or immature D1 cells. (B) Co-culture with YAC-1 or P815-B7.1 cells do not activate resting NK cells. Lytic activity against YAC-1 cells of NK cells co-cultured with either immature BALB/c mice-derived BM-DCs (GM-CSF  IL-4), YAC-1 or P815-B7.1 cells was evaluated after 18 hours. (C) No APCs are able to activate NK cells. The lytic activity of NK cells co-cultured for 18 hours with GM-CSF-propagated BM-DCs or SCID mice-derived macrophages was tested against YAC-1 cells. In all experiments, a classical chromium release assay was performed. Results are expressed as the percentage of specific lysis at various effector:target ratios and represent the means of triplicate wells. Data are representative of one out of three independent experiments.

To our knowledge, very few studies have documented the capacity of discrete cell types, such as B-EBV cell lines or in vitro differentiated macrophages to modulate NK-cell activity

(Chang et al., 1990; Montel et al., 1995). One study reported that HLA-DR positive cells, which could be DCs, are necessary for NK-cell cytotoxicity against virally infected targets (Bandyopadhyay et al., 1986). Monocytes have been shown to downregulate IL-2-mediated augmentation of human NK-cell activity through the secretion of PGE2 (Suzuki et al., 1984). Hanna et al. reported a correlation between the generation of hyporesponsiveness to NK cell activation induced following activation of suppressor macrophages by C. parvum (Hanna, 1983). To our knowledge, no previous report has demonstrated that a cell subset can markedly trigger IFNγ production and cytolytic activity of resting NK cells without additional cytokines. DCs may express critical NK costimulatory receptors and/or cytokines that may act in concert to activate resting NK cells in vitro and potentially in vivo. Indeed, DCs have been shown to produce IL-12 following antigendependent cognate DC–T cell interaction (Macatonia et al., 1995; Winzler et al., 1997), IFNα/β when virally infected (Eloranta et al., 1997) and IL-15 (Jonuleit et al., 1997) to facilitate recruitment and activation of effector cells. However, the constrained ratios between DCs and NK cells necessary for activation and its abrogation following disruption of the intimate cell–cell contact strongly suggest that DC

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A model for the modulation of innate and adaptive immune responses by DCs.

membrane molecules are capable of promoting engagement of NK-cell receptors. It is likely that this membrane crosstalk could be potentiated by soluble factors such as cytokines or chemokines. It is interesting to note that the activation of NK cells can only be induced following coculture with DCs derived under certain culture conditions, such as those generated in GM-CSF plus IL-4 but not those obtained using only GMCSF. These observations suggest that the NK-cell triggering function of DCs might be tightly regulated and that DCs may need to be at a distinct differentiation stage to be capable of activating resting NK cells. Although it has not been shown so far that DCs could directly activate resting NK cells, several authors have demonstrated that IL-2 and/or

IL-12-activated NK cells or LAK cells could recognize B7 or CD40-positive target cells such as authentic DCs, thus overruling class I moleculetriggered inhibitory receptors (Chambers et al., 1996; Carbone et al., 1997; Geldhof et al., 1998). It is noteworthy that after FL completion, the numbers of DC dropped dramatically. It is possible that a negative feed-back of the DC–NK loop could occur to regulate NK activation processes as suggested by Chambers et al., (1998). Recently, there has been a great deal of interest in DC biology and in implementing their attributes for immunotherapy against tumors or infectious diseases. There is accumulating evidence that treatment with in vitro-generated and antigen-pulsed or -transfected DCs can be effective in inducing T cell-mediated immunity

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in mice (Fernandez et al., 1998) and in humans – in normal volunteers (Dhodapkar et al., 1999) as well as in tumor-bearing patients (Nestle et al., 1998). With the description of a novel role for DCs in activating NK cells, the DC-based immunotherapy may not only promote T celldependent antitumor immunity but also the NK cell-mediated counterpart. Recent results show that monocyte-derivedhuman DCs (MD-DCs) can elicit resting human NK-cell cytolytic activity and IFNγ production only following licensing by immortalized fibroblasts or tumor cell supernatants (Fernandez et al., 1999b). The licensing of MD-DCs is medi-ated by soluble factors acting in a crossspecies manner and allows both human and mouse NK cell activation. Interestingly, once NK cells are activated using IL-2, nonlicensed DCs dramatically enhance NK-cell survival and activation in vitro following IL-2 deprivation (Fernandez et al., 1999b). Following Janeway and Medzhitov’s theory, innate immunity controls the development and nature of adaptive immunity (Medzhitov and Janeway Jr, 1997). It is conceivable that tumor homologues of pathogen-associated molecular patterns may be recognized by pattern recognition receptors on DCs that will license DCs to activate NK cells at the tumor site. A second and nonexclusive possibility is that soluble factors secreted by the tumor cell itself, or by the microenvironment, may induce the DCs to acquire the capability to activate NK cells. The interactions between DCs and NK cells in peripheral tissues may consequently result in the triggering of NK-cell cytolytic activity but also of chemokine and cytokine secretion such as IFNγ, that will in turn attract and activate other effector cells of the innate immunity such as macrophages (Figure 18.6). The coordinate interactions of these key players may result in tumor lysis, releasing apoptotic or necrotic bodies that will be taken up, transported and presented by DCs to T cells. We could speculate that the interaction between DCs and NK cells would also result in the ‘activation’ of DCs, that will start to migrate to the draining lymph nodes while maturing. Thus, it can be inferred that DCs

are at the interface between innate and adaptive immune responses.

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Whitmore, J., Miller, R.E. and Schuh, J.C.L. (1997). Nat. Med. 3, 625–631. Macatonia, S.E., Hosken, N.A., Litton, M. et al. (1995). J. Immunol. 154, 5071–5079. Maraskovsky, E., Brasel, K., Teepe, M. et al. (1996). J. Exp. Med. 184, 1953–1962. Medzhitov, R. and Janeway, C.A. Jr., (1997). Curr. Opin. Immunol. 9, 4–9. Montel, A.H., Morse, P.A. and Brahmi, Z. (1995). Cell. Immunol. 160, 101–114. Moretta, A., Bottino, C., Vitale, M. et al. (1996). Annu. Rev. Immunol. 14, 619–648. Moretta, A., Sivori, S., Ponte, M., Mingari, M.C. and Moretta, L. (1998). Curr. Top. Microbiol. Immunol. 230, 15–23. Nestle, F.O., Alijagic, S., Gilliet, M. et al. (1998). Nat. Med. 4, 328–332. Orange, J.S. and Biron, C.A. (1996). J. Immunol. 156, 1138–1142. Pende, D., Parolini, S., Pessino, A. et al. (1999). J. Exp. Med. 190, 1505–1516. Pulendran, B., Lingappa, J., Kennedy, M.K. et al. (1997). J. Immunol. 159, 2222–2231. Raulet, D.H. (1996). Curr. Opin. Immunol. 8, 372–377.

Reis e Sousa, C., Hieny, S., Scharton-Kersten, T. et al. (1997). J. Exp. Med. 186, 1819–1829. Rolstad, B. and Seaman, W.E. (1998). Scand. J. Immunol. 47, 412–425. Salazar-Mather, T.P., Ishikawa, R. and Biron, C.A. (1996). J. Immunol. 157, 3054–3064. Salazar-Mather, T.P., Orange, J.S. and Biron, C.A. (1998). J. Exp. Med. 187, 1–14. Scharton-Kersten, T., Afonso, L.C., Wysocka, M., Trinchieri, G. and Scott, P. (1995). J. Immunol. 154, 5320–5330. Shaw, S.G., Maung, A.A., Steptoe, R.J., Thomson, A.W. and Vujanovic, N.L. (1998). J. Immunol. 161, 2817–2824. Sivori, S., Vitale, M., Morelli, L. et al. (1997). J. Exp. Med. 186, 1129–1136. Suzuki, H., Yamashita, N., Sugiyama, E., Sato, M., Ito, M. and Yano, S. (1984). Anticancer Res. 4, 63–67. Trinchieri, G. (1989). Adv. Immunol. 47, 187–376. Vitale, M., Bottino, C., Sivori, S. et al. (1998). J. Exp. Med. 187, 2065–2072. Winzler, C., Rovere, P., Rescigno, M. et al. (1997). J. Exp. Med. 185, 317–328.

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19 B cells Francine Brière1, Bertrand Dubois2, Jerome Fayette1, Stephanie Vandenabeele3, Christophe Caux1 and Jacques Banchereau4 1 Schering-Plough, Dardilly, France; INSERM U404 ‘Immunité et Vaccination’, Lyon, France; 3 The Walter and Eliza Hall Institute of Medical Research, Post Office Royal Melbourne Hospital, Victoria, Australia; and 4 Baylor Institute for Immunology Research, Dallas, TX, USA 2

It is one of the most beautiful compensations of this life that no man can sincerely try to help another without helping himself. Ralph Waldo Emerson

INTRODUCTION

critical molecules such as CD40L to B lymphocytes. There is now evidence that DCs directly interact with B cells in vitro. Both freshly isolated and in vitro generated DC subsets share the ability to stimulate B cells.

A critical role for dendritic cells (DCs) located within the mucosal epithelium is to capture foreign antigens following tissue injury and subsequently initiate immune responses. Although much is known concerning DCs and their ability to stimulate antigen-specific naïve T cells, it is only recently that the direct contribution of DCs in the regulation of humoral response has been addressed. Several in vitro and in vivo studies have reported the importance of dendritic cells (DC) in the establishment of humoral responses (Inaba et al., 1983; Francotte and Urbain, 1985; Inaba and Steinman, 1985; Spalding and Griffin, 1986; Sornasse et al., 1992; Cebra et al., 1994; Flamand et al., 1994). However, DCs were thought to initiate B cell responses by activation of T cells, which in a second stage provide cytokines and Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

HUMAN DCs ENHANCE GROWTH OF ACTIVATED B CELLS Early B cell activation occurs in the T cell areas of secondary lymphoid tissues (Liu et al., 1991; Toellner et al., 1998). Antigen-activated B cells specifically migrate to T-cell zones (Cyster et al., 1994), where they localized in the proximity of interdigitating dendritic cells (IDC) (Björck et al., 1997). Likewise, in vitro-generated DCs form very tight clusters with allogeneic B cells when cultured in the presence of CD40L-transfected

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cell line, as surrogate activated T cells (Dubois et al., 1997; Fayette et al., 1997). DCs affect the growth of mature B cells at their various stages of differentiation: naïve (Fayette et al., 1997), memory (Dubois et al., 1997) or germinal center B cells (Dubois et al., 1999a). DCs induce IL2-mediated B cell proliferation (Dubois et al., 1997; Fayette et al., 1997). This requires the engagement of DC–CD40, which promotes the release of IL12 and sgp80/IL6R (Dubois et al., 1998).

HUMAN DCs ENHANCE DIFFERENTIATION OF ACTIVATED B CELLS Memory B cell differentiation without cytokines DCs strongly potentiate the differentiation of CD40-activated memory B cells towards essentially IgG secreting cells in the absence of exogenous cytokine. While endogeneous IL12 is not involved, IL6 plays an important role in this differentiation (Dubois et al., 1997). DCs enhance B cell differentiation through the secretion of soluble IL6R α chain, gp80 which complexes to IL6 (Dubois et al., 1997).

Role of IL-12 in DC-induced naïve B cell differentiation into plasma cells During the extrafollicular reaction which occurs in the T-cell rich areas, activated T cells stimulate antigen-specific naïve B cells to proliferate and differentiate into germinal center founder cells or into plasma cells producing mostly IgM (Liu and Arpin, 1997). In vitro, naïve B cells have a low propensity to differentiate into plasma cells (Arpin et al., 1995). However, the further addition of DCs allows activated naïve B cells to differentiate into plasma cells (Fayette et al., 1998). IL-12, secreted by DCs upon CD40 engagement, synergizes with the complex IL6/soluble IL6R α chain produced by DCs for the differentiation of naïve B cells into IgM-secreting plasma cells (Dubois et al., 1998). Addition of IL-10, a com-

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monly used B cell differentiation factor, masks the contribution of endogeneous IL-12 in DCmediated plasma cell differentiation (Fayette et al., 1998). These observations made with in vitro activated B cells have been extended to splenic and lymph node plasmablasts whose survival and differentiation into plasma cells are enhanced by DCs (Garcia de Vinuesa et al., 1999).

HUMAN DCs INDUCE SWITCH RECOMBINATION Naïve B cells undergo IgA isotype switching In the absence of exogenous cytokines, DCs induce surface IgA expression on CD40-activated naïve B cells (Fayette et al., 1997) to a level comparable to that obtained with IL10 and TGFß, the most potent cytokine combination allowing IgA switching (Defrance et al., 1992). Induction of IgA-expressing B cells by DCs is partially mediated by TGFß while neither IL-10 nor IL-12 appear to be involved in the process (Fayette et al., 1997; Brière et al., 1998). DCs provide a critical yet uncharacterized IgA-inducing signal that synergizes with IL-10 and TGFß to induce commitment towards IgA expression of every other naïve B cell (Fayette et al., 1997). Isotype switching is a DNA recombination event which moves the VDJ gene upstream of Cµ gene to the CH gene of the new isotype by deleting the DNA located between the switch regions that lie upstream of each CH gene. The reciprocal recombination products (or switch circles) contain the 3 part of Sµ joined to the 5 part of the S region of the new isotype (Iwasato et al., 1990; von Schwedler et al., 1990). Addition of DCs to cultures of CD40-activated B cells, in the absence of exogeneous cytokines, induces the production of IgA switch circles at levels much higher than those obtained with IL-10 and TGFß. The highest numbers of Sα –Sµ reciprocal junctions is observed upon addition of DCs, IL10 and TGFß (Dubois et al., 1999b). Thus, in a CD40-dependent context, IL-10 is an isotype switch factor for IgA, as well as IgG (Brière et al.,

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1994; Malisan et al., 1996) and in the absence of exogeneous cytokines, DCs promote switch recombination to Cα genes. While DCs alone are able to induce CD40activated naïve B cells to express surface IgA, IL-10 is essential for their differentiation into IgA-secreting cells. Moreover, in the presence of IL-10 and TGFß, DCs skew CD40-activated naïve B cells towards the secretion of IgA. These results extend earlier studies with mouse Peyer’s patch or splenic B cells (Schrader et al., 1990; Schrader and Cebra, 1993; Cebra et al., 1994) as well as pre-B cell lines (Spalding and Griffin, 1986) which were induced to secrete high levels of IgA with a combination of polyclonally activated T cells or TH2 clones and DCs. Importantly, in the presence of DCs, naïve human B cells can be induced to secrete both IgA1 and IgA2 subclasses (Fayette et al., 1997). In contrast to the unique IgA isotype in mice, two IgA subclasses IgA1 and IgA2 exist in humans whose differential in vivo expression suggests distinct regulation (Brandtzaeg, 1995).

DCs and mucosal immune responses In our experimental model, human DCs can trigger (1) an IL-12-dependent systemic type of humoral response which is observed in the presence of IL-2; and (2) an IL-12-independent mucosal type of humoral response in response to cytokines associated to the gut environment such as IL-10 and TGFß (Banchereau et al., 2000). It is tempting to speculate that in vitrogenerated DCs subpopulations share properties with mucosal-associated DCs involved in the regulation of mucosal humoral responses and with DCs present in nonmucosal lymphoid organs that would regulate systemic responses. As DCs have been described in virtually all mucosae, it would be important to determine whether mucosal-associated DCs differentially regulate humoral responses (Hart, 1997). Oral administration of IL-12 in mice, induces secretion of TH2-type cytokines without altering secretory IgA antibody responses while when

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delivered parenterally, IL-12 induces a TH1-shift (Marinaro et al., 1997). Our results demonstrate that in addition to priming T cells towards TH1 development through the production of IL-12, DCs may directly signal naïve B cells during the initiation of the immune response. These findings may have important implications in the outcome of the immune responses obtained with vaccination protocols using a mucosal versus a systemic route.

DC SUBSETS DIFFERENTIALLY REGULATE B CELL FUNCTIONS In vitro-generated DC subsets In vitro and in vivo studies have shown that monocytes are precursors for DCs (Banchereau and Steinman, 1998). In the context of B cell responses, DCs and monocytes are equally able to enhance CD40-activated B cell proliferation. However, DCs are more efficient than monocytes in inducing memory B cells to secrete IgG and IgA in the absence of cytokines (Dubois et al., 1997). Furthermore, both CD34 hematopoietic progenitor- and monocytederived DCs, but not monocytes, induce surface IgA expression on CD40-activated naïve B cells in the absence of cytokines (unpublished observations). CD34-HPC generate two DC subsets in response to GM-CSF and TNFα (Caux et al., 1996, 1998) Langerhans cells and interstitial (dermal) DC (Caux et al., 1996). While both DC subsets are able to enhance the proliferation of CD40-activated B cells and to induce the differentiation of memory B cells, only CD14-derived interstitial DCs can induce naïve B cells to differentiate into IgM secreting cells in response to CD40 ligation and IL-2 (Caux et al., 1990). In addition, ex vivo isolated epidermal Langerhans cells also lack those properties. This suggests that interstitial DCs (e.g. dermal-type) rather than epidermal Langerhans cells could be critical in the launching of primary B cell responses (Caux et al., 1998).

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A role for GCDC in the establishment of germinal center reaction The germinal center is the microenvironment that allows the generation of B cell memory. There, B cells proliferate, undergo somatic mutation, isotype switching, affinity selection and differentiate into either memory B cells or plasmablasts. The germinal center also contains T cells, follicular DCs (FDC) and germinal center DCs (GCDC) (Grouard et al., 1996). These GCDC can be isolated from tonsils as CD4CD11clin cells. When added to cultures of CD40-activated germinal center B cells, they stimulate in an IL-12-dependent manner B cell proliferation in response to IL-2 and drive their differentiation towards plasma cell. In contrast, Langerhans cells isolated from skin are unable to drive those germinal center B cells to proliferate and differentiate. In addition, GCDC induce IL-10independent isotype switching towards IgG1. Thus, GCDC are likely to play a significant role in GC development. In order to generate a humoral immune response, antigen-specific CD4 T cells and antigen-specific B cells must interact. This could be the role of the human germinal center DC (GCDC) population localized within the germinal center (Grouard et al., 1996) and originally described in the mouse as an ‘antigen-transporting cell’ (Tew et al., 1980), which could display the antigen to both T and B cells. Furthermore, such a population could also correspond to the recently identified murine DC subset based on its ability to bind a fusion protein consisting of the cystein-rich portion of the mannose receptor and the Fc portion of human IgG1. In vivo, this latter DC subset was not only able to prime T cells but also to induce production of antigen-specific IgM and IgG1 (Berney et al., 1999). As illustrated in Figure 19.1, DCs could present the antigen concomitantly to both T and B cells. This would imply that DCs may present unprocessed antigen to B cells. Indeed, in vivo studies in the rat have shown that DCs can capture and retain unprocessed antigen which could be transferred to naïve B cells to initiate specific TH2-associated antibody responses (Wykes et al., 1998). DCs would select the rare

FIGURE 19.1 DC serves as ‘a temporal bridge’ between CD4 T helper cells and B cells. In this model, DCs would interact directly with both T cells and B cells. (1) DCs select the rare antigen-specific T cell and this activated T cell, producing cytokines and expressing CD40L, (2) further signal DCs to express high levels of costimulatory molecules. In the last step, (3) fully mature DC could directly interact with B lymphocytes and turn on the humoral response.

antigen-specific T cells and such activated T cells, producing cytokines and expressing CD40L, further signal DCs to express high levels of co-stimulatory molecules. Then, fully mature DCs would directly interact with B lymphocytes and turn on the humoral response. During the extrafollicular reaction, IDCs are also likely to play a role in the induction of an IL-2-dependent IgM plasma cell (Garcia de Vinuesa et al., 1999). Germinal center formation starts with the migration of GC founder cells in the follicles and involves the TH2 type of CD4 T cells (Liu and Arpin, 1997).

MOLECULES INVOLVED IN DC–B CELL INTERACTIONS Several members of the TNF/TNF-R family appear to be involved in the interactions of DCs with B cells. In particular, CD40 activation upregulates OX40L expression on DCs and B cells (Stuber et al., 1995; Ohshima et al., 1997) which can engage T-cell OX40 to promote (1) production of TH2 cytokines (Flynn et al., 1998); (2) migration of CD4 T cells within B cell follicles

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(Brocker et al., 1999) through the upregulation of CXCR5 (Flynn et al., 1998). The importance of OX40–OX40L interaction for B cell responses has been suggested by enhanced proliferation and Ig secretion after cross-linking of OX40L on B cells (Stuber et al., 1995) and by a severely impaired IgG response to a T-dependent antigen after in vivo neutralization of OX40 with polyclonal antibodies (Stuber and Strober, 1996). However, because OX40 and OX40L-deficient mice develop normal humoral responses with appropriate germinal center formation and architecture, the OX40–OX40L pathway may be redundant in vivo for the formation of germinal centers. On the other hand, OX40L on DCs was demonstrated to deliver a critical costimulatory signal for the development of contact hypersensitivity and virus specific CD4 T cells, and OX40 signalling may thus be unique for its ability to generate optimal CD4 T cell responses (Chen et al., 1999; Kopf et al., 1999). Another member of the TNF family may be critical to the development of germinal centers. Identified by several groups, BAFF/Blys/TALL-1 or THANK (Moore et al., 1999; Mukhopadhyay et al., 1999; Schneider et al., 1999; Shu et al., 1999) was found on DCs and T cells. BAFF/Blys binds to a receptor restricted to B cells and induces both proliferation and immunoglobulin secretion by different B cell subsets (Moore et al., 1999; Schneider et al., 1999). In particular, BAFF-transgenic mice develop autoimmune-like manifestations (Mackay et al., 1999). Thus, BAFF/Blys may represent an important costimulator through which DCs regulate B cell proliferation and function. As TNFα and FasL, BAFF/Blys, TWEAK or APRIL are trans-membrane molecules possessing canonical furin cleavage motifs in their stalk region, that are processed for releasing a secreted form by a protease yet to be identified (Chicheportiche et al., 1997; Hahne et al., 1998; Schneider et al., 1999). Decysin, a novel disintegrin-metalloproteinase, isolated from germinal center dendritic cells and specific of mature DCs represents a candidate for the cleavage of molecules of the TNF-family and may thus play an important role in the regulation of T and B cell functions (Mueller et al., 1997).

CONCLUSION Besides activating T cells, DCs can directly activate B cells at their various stages of differentiation. DC subsets may provide B cells with the different cytokines/surface molecules that determine the fate of humoral immune responses, such as peripheral versus mucosal-type of humoral responses. In the context of T cell responses in the human, monocytederived CD11c DCs or virus-activated CD11c DC polarize naïve T cells predominantly towards a TH1 profile, whereas IL3-derived CD11c DC favor TH2 differentiation (Cella et al., 1999; Cella et al., 2000; Kadowaki et al., 2000; Rissoan et al., 1999). IL-12, produced at high levels by monocyte-derived CD11c DCs, but not by CD11c derived DC, plays a fundamental role in recruiting naïve CD4 T cells towards the TH1 pathway. Monocyte-derived CD11c DC subsets, as well as ex vivo purified GCDC, can trigger an IL-12-dependent systemic type of humoral response in the presence of IL-2 and an IL-12independent mucosal type of humoral response

FIGURE 19.2 DCs directly modulate B cell responses. CD40-activated DCs produce IL-X (BAFF/Blys?) which enhances the proliferation of CD40-activated B cells. CD40-activated DCs also secrete IL-12 and sgp80, which binds IL-6 produced by B cells and some DCs. These factors, together with IL-2, induce CD40activated naïve B cells to differentiate into plasma cells secreting IgM and CD40-activated germinal center cells to differentiate into plasma cells secreting IgG1. CD40-activated DCs also provide B cells with the uncharacterized IL-Y, which together with IL-10 and TGFβ permits isotype switching towards IgA1 and IgA2.

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in response to cytokines associated to the gut environment such as IL-10 and TGFß (Figure 19.2). One needs to investigate the effects of CD11c derived DCs on B cell responses as the ability of a DC to produce IL-12 could not only be deciding for the classes of T cell immune responses but also for that of B cells. Furthermore, it will help in refining the DC network complexity on a functional basis.

ACKNOWLEDGEMENTS J.B., from the Baylor Institute for Immunology Research, has been supported by grants from the Baylor Health Care System Foundation, Cap CURE, Ligue Nationale contre le Cancer axe Immunologie and NIH CA78846A. Special thanks to Muriel Vatan and Caroline Alexandre for expert editorial assistance. We would also like to thank Clarisse Barthélémy, Jean-Michel Bridon, Béatrice Vanbervliet, Isabelle Durand and Catherine Massacrier for their continuous support.

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20 Cells of the monocyte/macrophage lineage Thomas C. Manning and Thomas F. Gajewski University of Chicago, Chicago, Illinois, USA

We few, we happy few, we band of brothers. Henry V William Shakespeare

INTRODUCTION

possess potent phagocytic capabilities, there are direct mechanisms whereby these related APCs might interact (i.e. via engulfment of apoptotic or necrotic cell remnants) for antigen presentation.

Dendritic cells (DCs) are thought to be the primary antigen-presenting cell (APC) for initiating the activation of naïve T cells (reviewed in Banchereau and Steinman, 1998). DCs are closely related to cells of the monocyte/macrophage lineage. In fact, DCs and macrophages may represent terminal stages of monocyte differentiation (Caux et al., 1996; Szabolcs et al., 1996; Palucka et al., 1998). In addition to their lineage relationship, DCs and monocyte/macrophages interact in both direct and indirect fashions. Each cell type possesses an ability to present peptide/MHC antigen to T cells and each can secrete multiple soluble factors that alter the immune environment within which participating cell types function. Moreover, T lymphocytes, by virtue of their ability to interact with both DCs and monocyte/macrophage cells, can serve to courier information between these cell types in the absence of direct APC–APC interaction. Finally, since both DCs and macrophages Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

RELATIONSHIP OF DCs TO THE MONOCYTE/ MACROPHAGE LINEAGE Sources of DCs Despite Langerhans’ initial histological description of epidermal interdigitating DCs in 1868, investigations into DC biology were not initiated until 27 years ago (Steinman and Cohn, 1973). DCs can now be isolated directly from tissues such as tonsils, spleen or epidermis, or alternatively can be generated by culturing CD34 hematopoetic precursor cells or CD14 peripheral blood monocytes. DCs are most commonly obtained for clinical use by in vitro

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culture of adherent peripheral blood monocytes; these are termed monocyte-derived dendritic cells (MDDCs). Frequently, the starting cells are depleted of lymphocytes and NK cells by negative selection prior to culturing with cytokines. In practice, immature MDDCs are obtained after a 5 to 7 day culture in the presence of GM-CSF and IL-4. These immature DCs can be induced to mature through a 1 to 3 day culture with various stimuli such as TNFα, bacterial products such as lipopolysaccharide (LPS) or dsRNA, CD40 ligand (CD40L) or monocyteconditioned medium (MCM) (Sallusto and Lanzavecchia, 1994; Bender et al., 1996; Romani et al., 1996). It is not certain if all maturation signals exert the same functional effects.

Myeloid-related DC lineage DCs, Langerhans cells (LCs), and macrophages are all thought to develop from a common CD14 precursor (Inaba et al., 1993; Caux et al., 1996; Szabolcs et al., 1996). From in vitro studies with day 5 CD34 precursor cells cultured in the presence of GM-CSF and TNFα, it has been shown that epidermal LCs arise from CD1a/CD14 precursors, while DCs and macrophages develop from precursors that are CD1a/CD14. Thus at this early time-point, the mutually exclusive expression of CD1a and CD14 appears to separate LC precursors from macrophage/DC progenitors. In addition, it appears that the day 5 CD1a/CD14 DC precursor cells are bipotential in that they can differentiate into either DCs (in the presence of GM-CSF and TNFα) or macrophages (if cultured with M-CSF). Between day 5 and day 12, the DC precursor cells lose expression of CD14 and acquire expression of CD1a (Caux et al., 1996; Palucka et al., 1998). Similar to the observations with CD34 hematopoetic precursor cells, a CD1a/ CD11c progenitor found within the peripheral blood has also been characterized as a Langerhans cell precursor and gives rise to typical Birbeck granule-expressing LCs when cultured with GM-CSF, IL-4 and TGFβ (Ito et al., 1999). It has been reported that CD2 expression by

peripheral blood monocytes is a property of DC precursors (Takamizawa et al., 1997). CD2 monocytes appear to lose CD14 expression and acquire a dendritic phenotype when cultured in the absence of cytokines, while CD2 monocytes retain CD14 and become macrophages under the same conditions (Crawford et al., 1999). During DC development in vitro from both monocytes and CD34 stem cells, several molecules can act to shift differentiation from the DC pathway towards the macrophage pathway. Notably, IL-10 is a potent inhibitor of DC differentiation and favors the macrophage development of DC precursors but not mature DCs (Buelens et al., 1997a, 1997b; Allavena et al., 1998). Vascular endothelial growth factor (VEGF) is another factor that is capable of inhibiting the maturation of DCs (Gabrilovich et al., 1996). Interestingly, VEGF is a frequently secreted product of many tumors, indicating a potential role in immune escape by tumors in vivo. IL-6, a cytokine also secreted by a number of tumors, acts to favor macrophage over DC development (Menetrier-Caux et al., 1998). Both monocytes and immature DCs express the M-CSF receptor, and the addition of M-CSF to adherent monocyte cultures acts to shift development of bipotential precursors toward the macrophage pathway (Menetrier-Caux et al., 1998; Palucka et al., 1998). During the maturation process, DCs lose expression of the M-CSF receptor and become resistant to an M-CSF-induced shift towards macrophage fate. Thus, the balance of cytokines can alter the production of functional mature DCs and shift progenitors towards either a DC or a macrophage fate. Collective evidence indicates that once they have matured, DCs become relatively resistant to phenotype changes. Monocytes have recently been shown to migrate to lymph nodes and differentiate into DCs in vivo following phagocytosis of fluorescent microspheres in subcutaneous tissues (Randolph et al., 1999b). In this system, approximately 25% of phagocytic monocytes differentiated into DCs, while 75% remained in place and acquired the phenotype of tissue macrophages. These findings are consistent with

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the recently reported roles for transendothelial trafficking and Fcγ receptor-mediated endocytosis in promoting DC maturation (Randolph et al., 1998a; Regnault et al., 1999).

Lymphoid-related DCs In considering the relationship of monocyte/ macrophage cells to DC lineage, it is important to point out, that in addition to the better characterized myeloid DC, a separate lymphoidlineage DC may exist (Ardavin et al., 1993; Galy et al., 1995; Saunders et al., 1996; Wu et al., 1996). At least two different cell types have been referred to as lymphoid-related DCs. The first is a mouse CD11cCD8α DC, while the second is a human CD4CD3CD11c plasmacytoid monocyte. In general, these so-called lymphoidrelated DCs express the IL-3 receptor (CD123), and either lack or have very low levels of expression of myeloid-lineage markers such as CD33 (Grouard et al., 1997; Facchetti et al., 1999; Robinson et al., 1999). In addition, while this subset expresses CD8α in the mouse, the human counterpart expresses CD4 (Winkel et al., 1994). There remains a significant degree of uncertainty as to the function of the variously described lymphoid DC populations. Nevertheless, human DCs derived from CD4CD3CD11c cells have been shown to induce TH2 differentiation and the term DC2 has accordingly been applied (Rissoan et al., 1999). It should be noted that this study used human DCs that were CD11c-negative and strongly CD123-positive corresponding to the plasmacytoid monocytes described elsewhere (Grouard et al., 1997; Cella et al., 1999; Facchetti et al., 1999; Robinson et al., 1999). In the mouse, several groups have reported that CD11cCD8a DCs preferentially induce a TH1 response (Maldonado-Lopez et al., 1999; Pulendran et al., 1999; Reis e Sousa et al., 1999; Smith and de St Groth, 1999). It should be noted that these studies were performed using mouse DCs that were in all cases CD11c-positive. Additionally, these CD8 DCs have been shown to express a ligand for Fas and are able to kill CD4 T cells (Suss and Shortman, 1996). Not

surprisingly, given their ability to kill T cells through Fas, a role in maintaining tolerance was suggested but remains to be demonstrated. Thus, it appears that two different populations of DCs have acquired the title of ‘lymphoid-related’. One population in the human is a CD4CD3CD11c plasmacytoid monocyte (Grouard et al., 1997; Cella et al., 1999; Rissoan et al., 1999; Robinson et al., 1999), while the other population described in the mouse is CD11cCD8 (Vremec et al., 1992; Suss and Shortman, 1996; Maldonado-Lopez et al., 1999; Pulendran et al., 1999; Reis e Sousa et al., 1999; Smith and de St Groth, 1999). The potential usefulness of various lymphoid-related DC subsets as immune adjuvants or tolerizing stimuli remains to be thoroughly examined.

DC MATURATION STAGES DCs exist in at least two stages of maturation. In the immature state they are highly potent at acquiring antigen. After receiving a maturation stimulus, their antigen uptake capability decreases in parallel with a dramatic increase in antigen presentation and co-stimulatory molecule upregulation (reviewed in Banchereau and Steinman, 1998). Maturation of immature DCs can be obtained by the addition of factors such as inflammatory cytokines such as TNFα, bacterial products such as LPS, double-stranded RNA or CpG oligonucleotides, monocyteconditioned medium (MCM), TNF-related activation-induced cytokine (TRANCE) or CD40 ligand (CD40L) (Sallusto et al., 1995; Cella et al., 1996; Reddy et al., 1997; Wong et al., 1997; Mackey et al., 1998; Bachmann et al., 1999; Verdijk et al., 1999). Additionally, there is evidence that passage through endothelial surfaces may also serve to induce signals leading to DC maturation (Randolph et al., 1998a). Several TNF family members appear to promote especially important signals for DC maturation. The important role of CD40/CD40L interactions in DC maturation is highlighted by the profound defect in cell-mediated immunity seen in mice deficient in these molecules

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(Campbell et al., 1996; Kamanaka et al., 1996; Soong et al., 1996; Mackey et al., 1998). Nevertheless, a CD40-independent pathway of T-cell activation also appears to be operative in certain circumstances (Borrow et al., 1996; Oxenius et al., 1996; Whitmire et al., 1996; Lu et al., 2000). In this regard, both CD40L and TRANCE have been shown to enhance the survival and adjuvanticity of DCs and to improve T-cell priming (Bennett et al., 1998; Mackey et al., 1998; Ridge et al., 1998; Schoenberger et al., 1998; Bachmann et al., 1999; Josien et al., 2000). TRANCE and CD40L are expressed on the cell surface of activated T cells. While CD40L is preferentially expressed by CD4 cells, TRANCE is expressed on both CD4 and CD8 cells (Josien et al., 1999). Both molecules are also active as soluble recombinant forms that associate as trimeric complexes and thereby act to crosslink TNF-family receptors on DCs. These recombinant soluble TNF-family ligands offer promise for improving the adjuvanticity of DCs used in immunotherapy protocols. Macrophages and monocytes are poised to contribute to DC maturation through various mechanisms. They may do so directly by production of inflammatory cytokines such as IL-1β and TNFα. Additionally, by attracting activated T cells to sites of inflammation through chemokine production, they recruit a cellular source of CD40L (and also TRANCE) for efficient DC activation. Several macrophage products, namely IL-6 and nitric oxide (NO), may also exert negative effects on DC development or maturation (Holt et al., 1993; Menetrier-Caux et al., 1998). Inflammatory monocytes expressing high levels of CD11b but lacking the macrophage marker F4/80 and the DC marker DEC-205 are able to migrate to lymph nodes and themselves differentiate into mature DCs (Randolph et al., 1999b). Thus, arrival of monocytes and macrophages at the scene of inflammation may exert both positive and negative feedback on DC maturation.

MEDIATORS OF DC–MONOCYTE/ MACROPHAGE INTERACTIONS Monocyte-conditioned medium Monocyte-conditioned medium (MCM) represents the DC maturation stimulus currently used most widely in clinical application. In practice, it is obtained by culturing lymphocyte-depleted PBMCs in tissue culture dishes coated with immunoglobulin for 24 hours. Replicating the exact components of this heterogeneous mix of soluble mediators has been problematic, and serves to illustrate the complexity of the interaction between DCs and monocytes that results in full DC maturation (O’Doherty et al., 1993; Bender et al., 1996; Romani et al., 1996; Reddy et al., 1997). In particular, it was observed that monocyte-conditioned medium could not be replaced by the cytokines TNFα, IL-1, IL-6, IL-12 or IL-15. In addition, neutralizing antibodies to IL-1, TNFα, IL-6 or IL-12 did not inhibit the activity of MCM (Bender et al., 1996). Nevertheless, the mean concentrations of TNFα, IL-1β, IL-6 and IFNα present in MCM have been determined, and a cocktail of these four cytokines can act as a partial substitute for MCM (Reddy et al., 1997). Prostaglandins, especially PGE-2, appear to be one important constituent of MCM, and it has been suggested that TNFα, IL-1, IL-6 and PGE-2 in combination is a suitable substitute for MCM (Jonuleit et al., 1997). PGE-2 appears to synergise with TNF in inducing DC maturation but can have differing effects on IL12 production (Rieser et al., 1997; Kalinski et al., 1998). In one study, PGE-2 was shown to preferentially bias toward TH2-inducing mature DCs when used with high concentrations of IL-1β and TNFα (Kalinski et al., 1998). Another group has reported that PGE-2 inhibits IL-12 production in the presence of LPS, but augments IL-12 production in synergy with TNFα (Rieser et al., 1997).

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Chemokines and chemokine receptors Chemokines are small polypeptides of 8 to 10 kDA size that are secreted by various cell types, bind to proteoglycans like heparin, and are able to induce migration of hematopoietic cells along a concentration gradient. Chemokines can be divided into four families based on structural motifs involving critical cysteine amino acid residues (designated CC (or β), CXC (or α), CX3C and C). The cellular response to chemokines is triggered through binding to cell surface chemokine receptors, members of the seven membrane-spanning domain G proteincoupled receptor family. The array of chemokines produced by DCs and to which DCs can respond is vast (reviewed in Dieu-Nosjean et al., 1999; Sozzani et al., 1999). DCs respond to many members of the CC chemokine family including MIP-1α, MIP-1β, MIP-3α, MIP-3β, MIP-5, RANTES, MCP-3, MCP-4, 6Ckine/SLC, MDC, TECK, and also to the CXC chemokine SDF-1. The chemokine receptors mediating DC responses include CCR1, CCR2, CCR5, CCR6, CCR7, CCR9 and CXCR4. Chemokine production by DCs is likewise broad and consists of both inducible and constitutive components. Immature DCs produce MDC and MIP-4/ DC-CK1 constitutively. TECK is produced by thymic DCs. Mature DCs produce chemokines that include MCP-1, IL-8 and MIP-3β, among others. An important advance within the last 2 years has been the demonstration that CCR6 and CCR7 are coordinately expressed on immature and mature DCs, respectively. The expression of these two chemokine receptors appears to play a critical role in directing the migration of immature DCs towards sites of inflammation, and then following antigen capture and maturation triggering, towards the T-cell areas of draining lymph nodes (Dieu et al., 1998; Sallusto et al., 1998; Sozzani et al., 1998). Thus, immature DCs expressing CCR6 migrate towards areas of inflammation in response to MIP-3α. MIP-3α is induced by inflammatory stimuli and is secreted by various cell types including endothelial cells and monocytes (Hieshima et al., 1997; Hromas

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et al., 1997; Rossi et al., 1997). Once immature DCs arrive at the site of inflammation and derive maturation signals, they lose CCR6 expression, upregulate CCR7, and acquire responsiveness to MIP-3β. In contrast to MIP-3α, MIP-3β production is restricted to lymphoid tissue and hematopoietic cells and is particularly highly produced in the periarteriolar T-cell zones of lymph nodes (Yoshida et al., 1997; Dieu et al., 1998; Sallusto et al., 1998). 6Ckine/SLC is a chemokine that is structurally related to MIP-3β, is expressed in lymphoid high endothelial venules and also acts as a ligand for CCR7 (Gunn et al., 1998). The plt strain of mouse exhibits a defect in the expression of SLC as well as defects in lymphocyte and DC homing into lymph nodes (Gunn et al., 1999). It is now thought that the CCR7-dependent migration of maturing DCs into the T-cell areas of lymphoid organs involves both MIP-3β and SLC. In fact, mice deficient in CCR7 show severe disturbances in the migration of B cells, T cells and maturing DCs, along with delayed antibody kinetics and an absence of delayed-type hypersensitivity (DTH) responses (Forster et al., 1999). Additionally, CCR7 has been shown to have a role in localizing the homing of TH1 and TH2 cells to periarteriolar lymphoid sheath and the periphery of the T-cell zones, respectively. This correlates with the expression of CCR7 by naïve and TH1 cells, and its absence on TH2 cells (Randolph et al., 1999a). Recently, a role in DC transendothelial migration has been assigned to the p-glycoprotein molecule (MDR-1) (Randolph et al., 1998b). This protein appears to be important in the migration of maturing DCs away from sites of inflammation and into draining lymphatics. Despite the fact that monocytes and macrophages can secrete numerous chemokines, it is problematic to attribute to these cell types the downstream effects of any one particular chemokine in vivo. Monocytes and macrophages are certainly prominent producers of chemokines (especially CC chemokines) that do have an effect on DCs. The recruitment of activated T cells expressing CD40L is one example of

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how chemokine secretion by monocytes and macrophages can lead to DC activation. Nevertheless, there is redundancy in the production of chemokines and many different cell types have the ability to affect DC function via any one particular chemokine receptor.

Cytokines and cytokine receptors DC maturation can be affected by many of the cytokines secreted by monocytes and macrophages. IL-10, for example, has been shown to block the maturation of DCs (Buelens et al., 1997a, 1997b). IL-18, a product of both DCs and macrophages, has been shown to augment IL-12-dependent IFNγ production by DCs (Fukao et al., 2000). Type I interferons (IFNα and IFNβ) produced by macrophages, T cells and other cells in response to viral inflammation have been shown to inhibit IL-12 production by DCs and thereby prevent DC-dependent TH1 differentiation (McRae et al., 1998). GM-CSF can be produced by macrophages as well as T cells, fibroblasts and endothelium and is an essential cytokine for the in vitro generation of myeloid DCs from either CD34 precursors or peripheral blood monocytes (Caux et al., 1992; Inaba et al., 1992a, 1992b). Nevertheless, GM-CSF does not appear to be critical for DC development in vivo. Although GM-CSF transgenic mice exhibit an increase in the number of DCs isolated from lymphoid organs, GM-CSF or GM-CSF receptor null mice exhibit approximately normal numbers of DCs, suggesting that other cytokines are also important in DC development (Vremec et al., 1997). IL-4 is produced by T cells, mast cells and some bone marrow stromal cells and is used alongside GM-CSF in generating DCs from peripheral blood monocytes. It has been shown to act, at least in part, by suppressing development of DC progenitors along the macrophage pathway (Jansen et al., 1989). FLT-3 ligand is a hematopoetic growth factor that has widespread expression, exists in both membrane bound and soluble forms, and acts on primitive hematopoietic cells. Administration of FLT-3L in vivo leads to large increases in the number of DCs of

both the myeloid and lymphoid-related subsets (Maraskovsky et al., 1996).

Nitric oxide NO is produced by the oxidative metabolism of arginine to citruline via one of at least three isoforms of the enzyme nitric oxide synthase (NOS). Isoforms specific to endothelium (NOS3 or eNOS) and neuronal tissue (NOS1 or nNOS) are expressed constitutively, while an inducible form (NOS2 or iNOS) that is best characterized in macrophages undergoes extensive upregulation in response to certain inflammatory stimuli (reviewed in MacMicking et al., 1997). NOS2 is dramatically upregulated following activation of macrophages with lipopolysaccharide (LPS), TNFα, or IFNγ. NO exerts direct antimicrobial killing effects, but also acts as a soluble immune mediator. It has been shown that NO generated by activated macrophages can inhibit T lymphocyte activation and proliferation (Hoffman et al., 1990; Denham and Rowland, 1992; Langrehr et al., 1992). It appears that NO production by macrophages exerts at least some of this immunomodulatory effect via the downregulation of APC function on neighboring DCs (Holt et al., 1993). NO is also produced by DCs upon their own activation, thus providing a mechanism of negative feedback (Bonham et al., 1996; Lu et al., 1996). Moreover, NO has been shown to induce the apoptosis of DCs. Inhibiting NO generation with the nitric oxide synthase inhibitor NG-monomethyl-L-arginine (NMMA) mitigates this effect (Bonham et al., 1996; Lu et al., 1996). Similarly, in patients with the autoimmune disorder primary biliary cirrhosis, DC stimulatory capacity was found to be impaired relative to healthy controls, and DCs from these patients were observed to produce higher levels of NO. DC stimulatory capacity was restored by the addition of NMMA (Yamamoto et al., 1998). Since the inducible form of nitric oxide synthase (NOS2 or iNOS) is activated by various immunologic stimuli such as LPS, TNFα, or IFNγ, these immunomodulatory effects of NO represent a feedback loop that may serve to regulate DC responsiveness.

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NO generation has also been implicated in regulating the balance between TH1 and TH2 responses. Mice defective in the inducible form of nitric oxide synthase (NOS2 or iNOS) exhibit increased susceptibility to Leishmania major infection yet paradoxically exhibit enhanced TH1 responses (Wei et al., 1995). This enhancement of the TH1 response in NOS2 knockout animals is thought to be at least partly attributable to the ability of NO to inhibit IL-12 production (Huang et al., 1998). However, it now appears that there may be a TH1-inducing effect of NO that selectively occurs at low concentrations in addition to the TH1 suppressive effects seen at higher concentrations (Niedbala et al., 1999). Recent evidence now indicates that in the L. major system NOS2 may act early on to control NK cells and the cytokine response, while later in the response and at higher levels of upregulation, it may work through direct antimicrobial effects (Diefenbach et al., 1998). Since both DCs and macrophages produce NO in response to inflammatory stimuli, this soluble mediator represents a mechanism for DC–macrophage crosstalk.

DIRECT DC–MONOCYTE/ MACROPHAGE INTERACTION Phagocytosis, macropinocytosis and antigen uptake One of the hallmark characteristics of DCs is their tremendous capacity for antigen uptake. DCs accomplish this using several mechanisms. Macropinocytosis is the term used for the ability of immature DCs to take in large amounts of fluid material. It has been estimated that the rate of fluid uptake for a single DC is 1000 to 1500 µm3 per hour, a volume closely corresponding to the volume of a single DC (Sallusto et al., 1995). DCs are additionally capable of phagocytosis of both apoptotic bodies and necrotic cell debris, although their intake capacity in this regard has generally been regarded as less than that of macrophages. Recent evidence indicates, that

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while macrophages engulf apoptotic bodies using various receptor mechanisms including the thrombospondin receptor CD36, the phosphatidylserine receptor and the vitronectin receptor (αvβ3-integrin), DC-mediated phagocytosis may be mediated preferentially through the αvβ3-and αvβ5-integrin molecules (Rubartelli et al., 1997; Albert et al., 1998a). However, full understanding of the receptors involved in DC phagocytosis is lacking. Immature DCs have been shown to secrete exosomes, which are small (50–90 nm) membrane vesicles that contain both MHC class I and II molecules and have been found to induce potent antitumor responses in mice (Zitvogel et al., 1998). Moreover, these secreted vesicles contain molecules such as the heat-shock protein hsc73 and the integrin-binding protein MFG-E8 (which binds to αvβ3 and αvβ5) that suggest mechanisms for transferring immunostimulatory antigens between macrophages and DCs (Thery et al., 1999). Just as there are differences between macrophages and DCs, so there are also differences between DC subtypes with regards to the mechanism of antigen uptake and antigen processing. Epidermal LCs, the first DC subtype described, are characterized by a unique cytoplasmic organelle known as the Birbeck granule, which by electron microscopy bears a striking resemblance to the wooden-style tennis racket (Birbeck et al., 1961). These structures are not identified on any other subtype of mature or immature interstitial DC. Recently, some of the uncertainty surrounding this structure has lifted with the identification of Langerin as an endocytic receptor unique to LCs that induces the formation of Birbeck granules (Valladeau et al., 2000). This receptor is a Ca2-dependent lectin with mannose-binding specificity, and has been suggested to be involved in nonconventional antigen routing (i.e. cross-priming). It has been suggested that Langerin may serve to efficiently internalize mannose-expressing microorganisms and ensure their proper routeing. As a comparison, macrophages and DCs do not express Langerin, but rather express a macrophage mannose-receptor (MMR) that

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mediates internalization into the MHC class II pathway (Sallusto et al., 1995).

Antigen-processing capabilities of DC versus macrophages The processing and loading of exogenously derived antigens on to MHC class I molecules constitutes what has been referred to as the ‘nonclassical’ or ‘exogenous’ pathway and underlies the phenomenon of cross-priming (Bevan, 1976; Carbone and Bevan, 1990; Kurts et al., 1996; reviewed in Rock and Goldberg, 1999). DCs are highly efficient at presenting antigens via the MHC-I exogenous pathway (Svensson et al., 1997; Albert et al., 1998a, 1998b; Rovere et al., 1998; Regnault et al., 1999; Rodriguez et al., 1999). Macrophages appear to be much less efficient at utilizing this exogenous MHC-I pathway, but nevertheless, are also able under certain circumstances to present exogenous antigens via MHC-I (Rock et al., 1993; Norbury et al., 1995; Reis e Sousa and Germain, 1995; Suto and Srivastava, 1995). In particular, DCs may be more potent than macrophages when it comes to presenting antigen derived from apoptotic cells (Albert et al., 1998b). It appears that DCs (and not macrophages) possess a unique pathway that allows exogenously acquired antigen to be exported from the endocytic compartment into the cytosol for access to the proteosome/TAP apparatus (Rodriguez et al., 1999). This export appears to be size-selective, and thus the existence of a DC-specific transporter or pore has been proposed. In addition to the antigen presentation machinery involved in exogenous MHC-I presentation, immature DCs may employ a novel extracellular antigenprocessing capability whereby extracellular proteolysis results in peptide loading on to empty cell surface MHC class II molecules associated with extracellular H-2M/HLA-DM molecules (Santambrogio et al., 1999a, 1999b). Macrophages do not appear to express these empty MHC class II molecules to the same level as immature DCs (Santambrogio et al., 1999b). One interesting point of variation in antigen processing concerns the functional differences

between the so-called normal and ‘immunoproteosome’ proteolytic apparatus (reviewed in Rock and Goldberg, 1999). The immunoproteosome is the version expressed in mature DCs and contains the IFNγ-induced catalytic subunits LMP2, MECL1 and LMP7 in place of the constitutively expressed subunits β1, β2 and β5. Differences in proteolytic ability between the two versions of the proteosome have been demonstrated, and certain peptides may fail to be cleaved equivalently by the two assemblies (Morel et al., 2000).

Apoptotic versus necrotic debris – outcomes There are two main subtypes of cellular debris encountered by phagocytic cells: apoptotic and necrotic. There has been some question as to whether each type of material can be equivalently processed and presented. Some evidence seems to favor the idea that engulfment of necrotic debris will be immunostimulatory, while apoptotic debris may promote tolerance (reviewed in Steinman et al., 2000). DCs are able to engulf bystander apoptotic cells under certain conditions and can thereby derive maturation signals that allow them to efficiently stimulate T cells (Rovere et al., 1998). In particular, it has been shown that DCs can engulf cells of the monocyte/macrophage lineage (Albert et al., 1998b; Yrlid and Wick, 2000). A study using influenza-infected apoptotic monocytes concluded that uptake of apoptotic monocyte debris by DCs led to efficient T-cell stimulation, while uptake of necrotic debris did not (Albert et al., 1998b). More recently, in a different system using fibroblasts rendered necrotic through freeze–thaw cycles or apoptotic via ceramide treatment, the reverse was observed: necrosis rather than apoptosis efficiently activated DCs for T-cell stimulation (Gallucci et al., 1999). Some of the differences observed in these studies may be attributable to the fact that monocytes rendered apoptotic by influenza infection will release IFNα, heat-shock proteins, and other inflammatory mediators indicating that a ‘danger state’ exists (Gallucci et al., 1999). In

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analyzing the ability of DCs to phagocytose and present cellular fragments of dying B cells via the MHC class II pathway, it was observed that both types of debris (apoptotic and necrotic) were extremely well processed (Inaba et al., 1998). Recently, Bhardwaj and colleagues have shed further light on the issue of apoptosis versus necrosis. Using a system of UV-triggered apoptosis and freeze–thaw-induced necrosis, it was shown that phagocytosis of apoptotic bodies by immature DCs provides for optimal cross-presentation of antigen, but that only necrotic debris affects DC maturation and thus the stimulatory properties of the DC presenting the antigen (Sauter et al., 2000). It has recently been observed that a weakly stimulatory subset of DCs from rat intestinal epithelium constitutively transports apoptotic cells to regional lymph nodes in the absence of inflammatory stimuli (Huang et al., 2000). Together, these reports suggest a mechanism whereby the uptake of apoptotic material under noninflammatory conditions might serve to maintain peripheral tolerance to self antigens, while engulfment of apoptotic cells in the presence of necrosis and inflammation might efficiently stimulate a response (Steinman et al., 2000). While it is clear that DCs can efficiently phagocytose and present antigen from infected monocytes and macrophages (Albert et al., 1998b; Yrlid and Wick, 2000), the significance of this is uncertain when considering the relationship of DCs to monocyte/macrophage cells, since DCs can also phagocytose other nonhematopoietic cell types. There is no firm evidence to date that DCs preferentially engulf debris derived from one cell type over another. Nevertheless, the ability of DCs and macrophages to compete for antigens or to transfer antigens via exosome particles provides an interesting potential mechanism for immune regulation.

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DCs VERSUS MONOCYTE/MACROPHAGE CELLS AS IMMUNE ADJUVANTS Because of their demonstrated ability to activate naïve T cells, antigen-pulsed DCs have become the preferred cell type for immunotherapeutic approaches. Early on it was noted that both macrophages and DCs could induce an allogeneic mixed leukocyte reaction (MLR), while only DCs efficiently induced a syngeneic MLR (Guidos et al., 1984). Differences in their antigen-presenting capabilities may also underscore the ability of DCs and macrophages to preferentially induce TH1 or TH2 responses. A study that compared the antibody isotypes induced by immunization with DCs versus peritoneal macrophages revealed that macrophages and DCs both induced specific antibody responses but that DCs preferentially induced TH1 isotypes (IgG2a) while macrophages induced TH2 isotypes (IgE) (De Becker et al., 1994). IL-12 and IL-10 production by the two cell types may be involved in regulating the TH1/TH2 balance. In response to equivalent inflammatory stimuli (IFNγ plus LPS), DCs produce significantly more IL-12 and much less IL10 than do M-CSF-differentiated macrophages (Smith et al., 1998). DCs have shown to be potent immunostimuli in numerous tumor systems in mice, and are being used in a clinical trial of various types of human cancer. Despite their potent efficacy in animal studies, purified DCs are not a requirement for inducing an immune response to a tumor. In mouse studies, it was observed that concomitant administration of IL-12 as an adjuvant allows bulk PBMCs to be substituted for DCs and also enhances the efficacy of DC-based immunotherapy (Fallarino et al., 1999). Moreover, tumor peptide-pulsed PBMCs elicit specific immune responses when used concomitantly with IL-12 in melanoma patients (Gajewski et al., 1999). The efficacy of PBMCs under these conditions is consistent with the recent observation that freshly isolated CD2 peripheral blood monocytes are themselves highly stimulatory without

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a requirement for preculturing (Crawford et al., 1999) and suggests that IL-12 production may underlie the TH1 stimulatory capacity of DCs. Importantly, it appears that strategies to provide additional IL-12 beyond that produced normally by DCs (i.e. IL-12-transfected DCs) can improve the stimulatory properties of DCs (Tuting et al., 1998).

CONCLUSION Although DCs are considered as being derived from cells of the monocyte/macrophage lineage, their function is clearly quite distinct. Within the last decade, the lineage relationship between monocytes, DCs and macrophages has become better understood. Nevertheless, certain lineage issues including the various DC subsets described in both human and mouse remain uncertain. Advances in understanding the array of chemokines and soluble mediators active on these cells have shed important light on the migration pattern of DCs. The large number of chemokines, and the diverse cell types from which they are secreted and on which they act, make their study a complex process. Finally, the various mechanisms through which DCs and macrophages take up and present antigens are beginning to be understood. From this knowledge will follow new approaches to eliciting or tolerizing the immune response.

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21 Endothelium Gwendalyn J. Randolph Institute for Gene Therapy and Molecular Medicine, Mount Sinai School of Medicine, New York, NY, USA

Up to a certain point we can understand living organisms mechanically. But when we look more closely it becomes quite evident that the knowledge we gain hardly touches any fundamental physiologic problem. The way of real advance in biology lies in . . . defining the parts and activities in this whole in terms of implying their existing relationships to other parts and activities. J. S. Haldane Interactions between dendritic cells (DCs) and endothelia occur at key periods in the life history of DCs. Following emigration from the bloodstream by adhesion to and migration across the vascular endothelium, DC precursors develop into immature DCs that survey the subendothelial, peripheral tissue for immunologic danger and for the maintenance of self tolerance. After capturing and processing antigens, the full maturation of DCs into potent stimulators of T lymphocytes proceeds at the same time as their passage across lymphatic endothelium en route to lymph nodes. Evidence is mounting that the simultaneous differentiation of DCs with their migration across endothelium is not merely coincidence, but rather that endothelium, in addition to regulating the trafficking of these antigen-presenting cells, also influences their development.

Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

EMIGRATION OF DC PRECURSORS FROM THE BLOOD TO PERIPHERAL TISSUES The first interactions between endothelium and DC precursors occur when these precursors are recruited to tissues and extravasate across vascular endothelium. Blood-borne precursors of DCs include CD14 monocytes (Randolph et al., 1998a, 1999) and several types of nonmonocytic MHC IICD3CD14CD19CD56 cells (O’Doherty et al., 1993; Thomas et al., 1993; Strunk et al., 1997; Cella et al., 1999; Siegal et al., 1999). Some studies have also characterized the transmigration of CD14 monocytes differentiated to immature DCs in culture with granulocyte–macrophage colony-stimulating factor (GM-CSF) and interleukin (IL)-4 or IL-13 prior to their addition to endothelium (D’ Amico et al., 1998; Lin et al., 1998). These investigations have been valuable to the characterization of DC 275

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maturation and are very relevant to the use of these cells in the clinic. However, studies regarding migration of these cells across vascular endothelium are difficult to place in a physiologic context, since partially matured monocytederived DCs are not found in the circulation. Vascular endothelium regulates the passage of leukocytes from blood to tissues via the coordinated and selective expression of chemokines and the expression of adhesive selectins and ligands for leukocytic integrins. Such regulation results in, for example, the recruitment of neutrophils to tissues only during acute inflammation, and the finely tuned homing of subsets of memory T cells to particular tissues (Butcher and Picker, 1996). In comparison, the trafficking of DCs seems to follow a different pattern in which most types of blood-borne DC precursors so far characterized with respect to interactions with endothelium appear to extravasate into peripheral tissues under steadystate conditions, with augmented recruitment during inflammatory reactions (Figure 21.1). The plasmacytoid monocytic DC precursors are exceptions to this trafficking paradigm, as these cells may exclusively interact with and transmigrate across high endothelial venules (Cella et al., 1999), the specialized vasculature unique to lymph nodes. 

CD14 Monocytes When isolated monocytes are applied to the apical surface of confluent endothelial monolayers grown on a collagenous matrix in vitro, typically at least half of the monocytes migrate across the endothelium to enter the underlying connective tissue matrix within 1–2 hours (Muller and Weigl, 1992; Meerschaert and Furie, 1994). This initial passage of monocytes across cultured endothelium does not require cytokine activation and occurs in response to an endogenous soluble gradient of the chemokine monocyte chemoattractant protein (MCP)-1 (Randolph and Furie, 1995), which is produced constitutively by endothelial cells in vitro and in vivo (Li et al.,1993). MCP-1, then, may participate in the known constitutive recruit-

FIGURE 21.1 Relationship and interactions of DCs with vascular and lymphatic endothelia in the steady state and during immune activation. Blood vessels (schematically identified by the presence of enucleated cells representing red blood cells) contain known precursors of myeloid DCs including CD14CLA monocytes, CD3CD14CD19CD56 lineage markernegative (Lin) CLACD11c nonmonocytic DCs, and CD34CLA precursors of epidermal Langerhans cells (hatched object represents the epidermis). The precursors for interstitial DCs are less well-defined, may vary depending on the tissue to which they home, and may overlap with the CLA precursors just named. In the interstitium, they accumulate around blood vessels. Proposed transendothelial trafficking patterns are shown with solid arrows representing constitutive, steady-state migration events, and dashed arrows representing trafficking under conditions of inflammatory or immune activation. Evidence for these pathways and molecular events that mediate them are discussed in the text. A question mark is placed next to the dashed arrow that suggests that interstitial DCs undergo reverse transmigration into the blood. There is evidence that DCs can traffic across vascular endothelium from tissues to the blood (see text), but which subset of DC migrates in this manner is speculative.

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ment of monocytes to tissues during the steady state. However, differences between the numbers of resident peritoneal macrophages in MCP-1-deficient or CCR2-deficient mice and their wild-type counterparts are negligible, whereas the data clearly demonstrate a key role for this chemokine in recruitment of monocytes during inflammatory states (Boring et al.,1997; Kuziel et al., 1997; Lu et al., 1998). Monocytes and other subsets of DCs express receptors for and chemotactic responses toward several other chemokines (reviewed by Sallusto and Lanzavecchia, 1999; Sozanni et al., 1999), which in addition to MCP-1, probably participate in recruitment of these cells from the blood.

Lin− CD11c cells When unfractionated peripheral blood mononuclear cells are added to unstimulated cultured endothelial monolayers, some CD3 cells, NK cells (Berman et al., 1996), and nonmonocytic blood-derived DC precursors enter the collagen along with monocytes, although CD14 monocytes still represent about 90% of the entering cells (Randolph et al., 1998a). That bloodderived nonmonocytic DC precursors, like monocytes, adhere to and migrate across unstimulated cultured endothelium (Brown et al., 1997) suggests that they may also traffic out of the blood during the steady state. Further supporting this possibility is the observation that these precursors constitutively undergo selectin-mediated rolling in skin microvessels in vivo (Robert et al., 1999). Just as for monocytes, inflammatory conditions probably lead to increased recruitment of these cells (Robert et al., 1999). For both populations of DC precursors, adhesive interactions with the endothelium utilize CD11/CD18 and CD49d integrins (Muller and Weigl, 1992; Meerschaert and Furie, 1994; Brown et al., 1997), and CD31 mediates passage across the endothelial junction via homophilic adhesion (Muller et al., 1993; D’Amico et al., 1998).

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Langerhans cells Keratinocytes and skin vascular endothelium constitutively express the chemokine MIP-3α (Charbonnier et al., 1999). CD1a precursors of Langerhans cells generated in vitro from cord blood-derived CD34 hematopoietic precursors express CCR6, the receptor for MIP-3α, and chemotax in response to this chemokine, in contrast to other DC precursors (Charbonnier et al., 1999). These findings together suggest that MIP3α may recruit Langerhans cells across vascular endothelium and to the epidermis. However, Langerhans cell precursors in the blood, identified as CD34 cutaneous lymphocyte antigen CLA cells (Strunk et al., 1997), do not yet express CD1a. Thus, the hypothesis that MIP3-α mediates trafficking of Langerhans cell precursors from the blood requires analysis of CCR6 expression and chemotactic activity toward MIP-3α of the CLACD34CD1a blood-derived Langerhans cell precursors.

Lin− CD11c− plasmacytoid monocytes A novel population of DC precursors, referred to as ‘plasmacytoid monocytes,’ have been identified in the circulation and are the body’s principle type I interferon-producing cells (Cella et al., 1999; Siegal et al., 1999). Such cells are LinCD11cMHCII cells that are further distinguished by expression of immunoglobulin-like transcripts (ILT) 3 but not ILT1. Both ILT1 and ILT3 are found on myeloid LinCD11c DCs. The interaction of the plasmacytoid monocytes with typical vascular endothelium has not been studied, and whether these cells migrate through peripheral tissues is unknown. However, they appear capable of traversing lymph node high endothelium to enter lymph nodes directly from the blood, a route not utilized by other DC precursors (Kupiec-Weglinski et al., 1988). L-selectin (CD62L) and the CXCR3 chemokine receptor, that mediates chemotaxis to the chemokines IP-10 and Mig, are expressed by these cells (Cella et al., 1999) and may mediate migration across the high endothelium.

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LOCALIZATION OF TISSUE DCs TO THE PERIVASCULATURE After emigrating from the blood, some subsets of DC remain only transiently within the underlying tissue, exiting via lymphatic vessels as veiled cells in less than 3–5 days (Pugh, et al., 1983). The kinetics of monocyte development from bone marrow precursors, extravasation from blood in vivo (Van Furth, 1988), and their differentiation to DCs in the presence of endothelium in vitro (Randolph et al., 1998a) or after administration of latex particles in vivo (Randolph et al., 1999) are all consistent with the idea that monocytes serve as a precursor for the continuously migrating veiled cells (Figure 21.1). Other DCs, such as Langerhans cells, reside within the tissue for more than a week (Katz et al., 1979), unless perturbed by inflammatory stimuli. The migration of Langerhans cell precursors into the epithelium places them in a prime position to encounter and acquire antigens that penetrate protective epithelial barriers. Resident DCs are also abundant in the vascularized tissues that underlie epithelia and, strikingly, they accumulate around blood vessels (Figure 21.1). This steady-state distribution was first noted among the Factor XIII MHC II dermal DCs within normal human skin (Sontheimer, 1989). Using confocal imaging, a three-dimensional reconstruction of the relationship of these cells to endothelial cells in human dental pulp suggests that dendritic processes of the pulpal antigen-presenting cells directly contact the ablumenal surface of the endothelium (Okiji et al., 1997). In the liver, MHC II interstitial DCs are concentrated around vessels of the portal triad, including the bile duct (Prickett et al., 1988). In lymph nodes, the plasmacytoid monocytes localize just beneath the high endothelial venules (Cella et al., 1999). DCs that are recruited to tissues under pathologic conditions including rheumatoid arthritis (Thomas and Quinn, 1996), ulcerated oral tissue (Natah et al., 1997) and inflamed plasmacytoidrich lymph nodes (Cella et al., 1999), also tend to

accumulate around blood vessels. In patients with rheumatoid arthritis, DCs are among the leukocytes recovered in the synovial fluid. In addition, the perivasculature of synovial tissue is enriched in DCs that display various stages of maturation. Some of these cells coexpress monocyte markers along with co-stimulatory molecules that are characteristically enriched on DCs. Thus, this tissue may serve as a site of monocyte differentiation to DCs. In the same perivascular area, cells with a more mature DC phenotype (high expression of co-stimulatory molecules in the absence of monocyte markers) form clusters with T cells (Thomas and Quinn, 1996). What biologic events lead to the predominant accumulation of DCs in the perivascular regions of connective tissue? It is not yet clear whether precursors of DCs are recruited specifically to the perivascular tissue or whether only those precursors that happen to be recruited to areas with abundant blood vessels develop into typical DCs. A study by Fioretti et al. (1998) illustrates the preferential localization of newly recruited DCs to tissue surrounding endothelial cells in situ. These investigators developed a model to explore whether leukocyte recruitment to an implanted tumor would give rise to a protective antitumor immune response in mice. Mastocytoma cells were engineered to express human MCP-3, which activates and/or promotes chemotaxis of human monocytes, T cells, eosinophils and basophils in vitro (Locati and Murphy, 1999). The chemoattractant can also direct the migration of immature DCs (Sozzani et al., 1995) generated in vitro from human monocytes cultured in GM-CSF and IL-4. Interestingly, MCP-3-expressing mastocytomas that were implanted in mice became infiltrated with monocytes, neutrophils, eosinophils and T cells, but mature DEC-205 2A1 DCs were rare within the tumors (Fioretti et al., 1998). In contrast, DEC-205 2A1 DCs accumulated abundantly around blood vessels in peritumoral tissue, even though these sites were not the primary site of chemoattraction by MCP-3. In this model, the perivasculature surrounding the tumor may serve as the most supportive envi-

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ronment for the differentiation of MCP-3recruited cells into DC, or may secrete a chemoattractive signal for DCs that overrides the chemotactic activity of MCP-3. Interestingly, the vasculature within the tumor did not manifest the same capacity to promote accumulation of maturing DC (Fioretti et al., 1998). Indeed, tumors are known to suppress DC maturation, at least in part due to the production of vascular endothelial growth factor (VEGF) (Gabrilovich et al., 1996), which is highly expressed within tumors to support neovasculaturization (Ferrara and Davis-Smith, 1997). Neutralizing mAbs to VEGF reverse the inhibitory activity with respect to DC maturation that is present in human tumor cell supernates (Gabrilovich et al., 1996). Aside from its activity as an endothelial cell growth factor, VEGF mimics the biologic function of macrophage colony-stimulatory factor (M-CSF) and replaces its functions in stimulation of osteoclast development in M-CSFdeficient mice (Niida et al., 1999). Similar toVEGF, production of M-CSF by tumor cells suppresses DC maturation (Menetrier-Caux et al., 1998). As it is becoming increasingly clear that monocytes can differentiate into either macrophages or DCs, one mechanism by which VEGF may inhibit DC development is to promote the maturation of monocytes to macrophages. Besides the possibility that blood vessel endothelium supports the development of DCs, does the accumulation of DCs around blood vessels serve an important function? The answer to this question is unknown. It has been suggested that this cellular distribution might facilitate encounter with T cells as they are recruited to tissues from blood (Thomas and Quinn, 1996). Indeed, memory T cells home selectively to peripheral tissues where relevant antigens may be present (Butcher and Picker, 1996), and possibly these antigens are processed in situ by perivascular DCs. However, the general importance of the lymph node as a site in which antigen-bearing DCs and T cells interact and the relatively immature state of perivascular DCs – more specialized for acquiring and processing antigen than in presenting it – raise questions

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regarding this proposed function of perivascular DCs. Perivascular accumulation of DCs also leaves them in an ideal position to traverse the endothelial barrier in the ablumenal-to-lumenal direction to re-enter the blood (Larsen et al., 1990). That DCs may migrate from tissues to blood is supported by several studies (see below), although this route of migration is probably quantitatively minor in comparison to the exit of DCs from tissues via lymphatic vessels.

TRANSENDOTHELIAL MIGRATION OF DCs FROM PERIPHERAL TISSUES INTO LYMPHATIC VESSELS Clearance of DCs from tissues as they carry processed antigen to lymphoid organs requires migration across an endothelial barrier in the ablumenal-to-lumenal direction. This migratory process has been called reverse transmigration (Randolph and Furie, 1996), a term chosen to distinguish it from the initial passage of DCs from the blood to tissues by migration across vascular endothelium in the lumenal-toablumenal direction.

Reverse transmigration across lymphatic and vascular endothelium in vivo Reverse transmigration across lymphatic endothelium is a key process, as it is thought to be the only means by which macrophages and most DCs gain access to lymph nodes (Barker and Billingham, 1968; Mebius et al., 1991), with the clear exception of plasmacytoid monocytic DC precursors (Cella et al., 1999). DCs comprise a substantial fraction, as much as 20%, of the cellular composition of lymph, even during the steady state (Pugh et al., 1983). The remaining cells in lymph (about 80%) are mainly memory T cells. The enrichment of DCs in lymph is particularly striking considering the rare incidence of nonmonocytic DCs in peripheral blood of less than 1% of leukocytes (O’ Doherty et al., 1993; Thomas et al., 1993). Conversely,

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monocytes enter tissues constitutively, at a rate that may exceed the rate at which tissue macrophages are replenished, but few typical monocytes are observed in lymph, naturally leading to the idea that monocytes may differentiate into DCs during passage through tissue (Randolph et al., 1998a). Secondary lymphoid chemokine (SLC) and macrophage inflammatory protein-3β (MIP-3β, also called ELC) bind to CC chemokine receptor 7 (CCR7) and are potent attractants for T cells and mature DCs. These chemotactic cytokines are highly expressed within lymphoid organs (Dieu et al., 1998; Gunn et al., 1998) and play a critical role in organizing the cellular architecture of lymphoid tissue (Dieu et al., 1998; Förster et al., 1999; Gunn et al., 1999). Lymphatic endothelial cells also display SLC on their lumenal surfaces (Gunn et al., 1998; Saeki et al., 1999) where it may mediate reverse transmigration of DCs into lymph vessels. After application of the contact sensitizer FITC to the skin of either CCR7−/− mice (Förster et al., 1999) or to mice with a naturally occurring plt (paucity of lymphoid T cells) mutation that results in SLC and ELC deficiency (Gunn et al., 1999), there is greatly impaired entry of peripheral DCs into lymph nodes. Moreover, a polyclonal anti-SLC antibody inhibits by approximately half the migration to draining lymph nodes of DCs injected into wild-type mouse footpads (Saeki et al., 1999). However, it is still not completely certain whether SLC mediates entry of DCs into the lymphatic vessels per se or has a more predominant role in promoting the downstream step of entry of DCs into the lymph node from the lymphatics. In comparison to reverse transmigration into lymphatics, reverse transmigration across vascular endothelium is an event that is less widely recognized. However, some studies imply that DCs can move from peripheral tissues to spleen, requiring then that these DCs would have returned to the blood (Larsen et al., 1990). Indeed, the appearance of DCs in one tissue after having been resident in another indicates that these cells re-enter the blood. In the context of transplantation, this phenomenon probably

accounts for the establishment of microchimerism (Starzl et al., 1993; Terakura et al., 1998), which is thought to be key for establishing tolerance to a transplanted tissue. This re-entry into blood would either have had to result from passage through a lymph node into efferent lymph, an event not thought to occur (Steinman, 1991), or the cell must have undergone reverse transmigration across vascular endothelium.

Reverse transmigration in vitro Monocyte-derived cells reverse transmigrate across cultured vascular endothelium in cultures of human umbilical vein endothelial cells (HUVECs) or dermal microvascular endothelium grown on native human connective tissue (Randolph and Furie, 1996), reconstituted collagen gels (Randolph et al., 1998b), or polycarbonate filters (D’ Amico et al., 1998) (Figures 21.2–21.4). Two observations highlight the similarities between these in vitro models and the trafficking of DCs via lymphatic vessels: (1) both reverse transmigration in vitro and the emigration of DCs from human skin explants via authentic lymphatic vessels require functional multidrug resistance 1 (MDR-1) protein (Randolph et al., 1998c), and (2) phenotypic analysis of reverse-transmigrated, monocytederived cells revealed that they differentiate into DCs (Randolph et al., 1998a). Reverse transendothelial migration and differentiation of monocytes to DCs The pattern of differentiation and transmigration of monocytes in the model just described is reminiscent of the trafficking in vivo, in which monocytes constitutively emigrate from the blood, but DCs are much more numerous in lymph than are monocytes. In contrast to the reverse-transmigrated cells, monocyte-derived cells that remain in the subendothelial collagen become macrophages (Randolph et al., 1998a). This model not only substantiated the evidence that monocytes serve as precursors for DCs, as previously

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suggested from in vitro cultures using exogenously added IL-4 and GM-CSF (Sallusto and Lanzavecchia, 1994; Bender et al., 1996; Kiertscher and Roth, 1996; Romani et al., 1996; Zhou and Tedder, 1996), but is also the first report to show that monocytes develop into DCs in response to factors endogenous to the culture system. This observation has been further validated from studies in vivo (Randolph et al., 1999). The key role of endothelial cells in promoting differentiation of monocytes into costimulatory DCs, in the model just described and models used by other investigators (Ferrero et al., 1998; Denton et al., 1999), illuminates a physiologic setting that may promote this differentiation in vivo: during and/or following emigration from

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the bloodstream, monocytes receive signals from endothelial cells that not only direct their trafficking but which also determine their pattern of differentiation (Shortman and Maraskovsky, 1998). In the reverse transmigration model, full maturation of monocytes to DCs is enhanced by the inclusion of phagocytic material in the collagen (Randolph et al., 1998a), suggesting that the action of phagocytosis, at least of phagocytosed ‘foreign’ material may provide a key maturation stimulus in situ. Interestingly, DCs in lymph are frequently seen to contain phagocytosed debris (Kelly et al., 1978; Sokolowski et al., 1978; Pugh et al., 1983; Mayrhofer et al., 1986). Besides mediating the differentation of monocytes into DCs, endothelial cells have also been

FIGURE 21.2 Models to study the influence of endothelial cells on DC maturation and their migration across endothelial monolayers in the forward and reverse directions. (A) Human umbilical vein endothelial cells are grown on reconstituted collagen gels that have been polymerized in tissue culture wells (typically microtiter wells). (B) In another model, endothelial monolayers are cultured on native human amnion that has been stretched across a Teflon ring and held in place with a Viton O-ring. The integrity of these constructs is such that individual cultures can be maneuvered as desired, such as raised on culture stands to permit access to the basal surface (see other configurations in Randolph and Furie, 1995). Both systems allow the investigation of forward and reverse transendothelial migration of DCs and the effect of the endothelium on DC differentiation. DENDRITIC CELLS AND INTERACTION WITH OTHER CELLS

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FIGURE 21.3 Schematic diagram depicting forward and reverse transmigration of monocytes. (I) Incubation of peripheral blood mononuclear cells with endothelium for 1.5 hours results in the transmigration of most monocytes, but few lymphocytes, into the subendothelial collagen. (II) Washing the cultures then leaves an intact endothelial monolayer with monocytes accumulated in the subendothelial matrix, where they engulf the phagocytic particles. (III) During further culture, some of the phagocytic monocyte-derived cells retraverse the same endothelium (t1/2  48 hours) and accumulate in the apical compartment. These reverse-transmigrated monocytes simultaneously differentiate into DCs. The figure shows cultures established with the use of reconstituted collagen. Except for the inclusion of phagocytic particles throughout the extracellular matrix, a similar reverse-transmigration assay can be set-up using the amnion model (Figure 21.2).

found to support long-term proliferation and differentiation of blood-derived CD34 stem cells. In the same study, fibroblasts were unable to substitutefortheendothelium(Moldenhaueretal.,submitted). The mechanisms by which endothelial cells promote differentiation of DCs is not yet clear. Endothelial cells produce GM-CSF (Broudy

et al., 1986), which is probably necessary but unlikely to be sufficient for mediating differentiation of DCs, as recombinant GM-CSF alone does not support generation of DCs from monocytes in culture (Sallusto and Lanzavecchia, 1994). Other known factors that support monocytic differentiation to DCs include the cytokines IL-4 and IL-13 (Chomarat and Banchereau, 1997) and/or engagement of CD40 by its ligand CD154 (Brossart et al., 1998; Grewal and Flavell, 1998). Expression of CD154 has been reported on monocytes and some DCs (Grewal and Flavell, 1998; Salgado et al., 1999), and the highly regulated, transient expression of CD154 appears to limit immuneresponses(Pérez-Melgosaetal.,1999).In the hypothetical scenario in which CD40–CD154 interactions may have a key role in monocyte differentiation to DCs, the role of endothelial cells may be to stabilize the expression of CD154. Interactions between LFA-3 (CD58) on endothelial cells and CD2 on leukocytes stabilizes mRNA for CD154 and increases CD154 levels on cells that express it (Murakami et al., 1999). Endothelial cells are typically cultured on extracellular matrix and can produce their own basement membrane. Thus, the role of extracellular matrix components in participating in the differentiation of DCs must be considered. Some investigations indicate that collagen type I (Brand et al., 1998) and fibronectin (Staquet et al., 1997) promote maturation of DCs. Contamination of these matrix components with lipolysaccharide (LPS) may account for the effects on differentiation (Suri and Austyn, 1998), an explanation that has increased appeal when one considers that the maturation induced by collagen type I did not require strong adhesive interactions of the DCs to the matrix. However, experimental evaluation of this possibility suggested that the maturation-promoting effects of collagen type I could not be completely explained by the presence of LPS (Brand et al., 1998). Subsets of monocytes may preferentially become DCs and reverse transmigrate All of the reports that link endothelial cells with the development of monocytes into DCs include

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steps of transendothelial migration, raising the possibility that the migratory events themselves are especially important to this pathway of differentation. Migration across endothelium may cause this maturation, or alternatively the endothelium may selectively promote the migration and reverse transmigration of subsets of monocytes that can become DCs. Reverse transmigration of monocyte-derived cells in vitro occurs with a half-time of 1–2 days (Randolph and Furie, 1996; Randolph et al., 1998b). Whether or not the endothelial cells are cytokine-stimulated, the kinetics of reverse transmigration are higher than first order (Randolph and Furie, 1996), indicating that the process is regulated by two or more independent rate-limiting events. A straightforward explanation of these kinetics suggests that there are at least two subsets of monocytes that have differing rates of reverse transmigration, and possibly differing potentials for differentiation to DCs. One of these populations appears to reverse transmigrate more rapidly across endothelial monolayers, particularly when the endothelium is pretreated with a proinflammatory cytokine. Just as the systemic administration of proinflammatory mediators, including IL-1,

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promote the clearance of DCs from tissues in vivo (Roake et al., 1995), pretreatment of cultured endothelium with IL-1 greatly enhances the rate of reverse transmigration of at least a subset of monocyte-derived cells in vitro (Randolph and Furie, 1996). There are several molecular markers that may identify subsets of blood cells that preferentially reverse transmigrate and become DCs. A subset of CD16 cells in peripheral blood give rise to DCs (Schäkel et al., 1998). Another group suggests that only those monocytes which express CD2 are DC precursors (Crawford et al., 1999). Still others report that CD14 monocytes that coexpress CD34 preferentially migrate across endothelium and that these cells can be stimulated with GM-CSF to become DCs when recovered from beneath the endothelium (Ferrero et al., 1998). However, these investigators observed a much lower basal transendothelial migration than seen in other models. This may be due to the use of endothelium grown on transwell filters, a system which would minimize the accumulation of soluble factors in the subendothelial space. When endothelium grown on human amniotic connective tissue is washed extensively to remove soluble MCP-1 from the

FIGURE 21.4 Morphological views of reverse-transmigrating cells. HUVEC/collagen cultures were incubated with PBMCs for 1.5 hours, washed to remove cells from the apical compartment, and cultured further for a total of 24 hours. Samples were fixed, embedded in LR White, and processed for electron microscopy. Serial sections of monocyte-derived cells apparently in the process of reverse transmigration (arrows) suggests that they make contact with the apical surface of the endothelium (arrowheads) predominantly via cytoplasmic extensions, with little close contact along the length of the membranes of the two cell types. These observations are consistent with the finding that reverse-transmigrated cells are easily removed from the apical compartment of the cultures by gentle pipeting. The cells appear to contact the endothelial junction, despite no requirement during reverse transmigration for the endothelial junctional protein CD31 that mediates apical-to-basal transmigration. DENDRITIC CELLS AND INTERACTION WITH OTHER CELLS

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collagenous matrix, the basal rate of monocyte migration is greatly reduced (Randolph and Furie, 1995). Alternatively, since the authors were actually measuring the number of monocytes that migrated completely through the transwell filter, it is possible that many more monocytes traversed the endothelium but not the full thickness of the filter. In such cases, their observations would be best explained by the hypothesis that the CD34 subset of monocytes is the most motile monocyte. Indeed, a greater motility of developing DCs relative to undifferentiated monocytes and maturing macrophages might explain the selective reverse transmigration of the former (Randolph et al., 1998a). Molecular mediators of reverse transmigration The molecular events that mediate reverse transmigration are poorly understood. The process is distinguished from apical-to-basal transmigration by the lack of a requirement for MCP-1 (Randolph and Furie, 1996) and CD31 (Randolph et al., 1998b, 1998c). Intercellular adhesion molecule 1 (ICAM-1) mediates the earliest phases of reverse transmigration, but apparently only accounts for the increased kinetics observed under conditions when the endothelium has been stimulated with IL-1 (Randolph and Furie, 1996). It is not clear whether the inability to block reverse transmigration over a longer period with antagonists of ICAM-1 is due to its failure to participate in reverse transmigration under other conditions. Alternatively, during the rather long duration of reverse transmigration, ICAM-1 and an additional adhesive molecule(s) may have redundant roles, such that blocking both ICAM-1 and the other molecule would be necessary to observe an inhibitory effect. ICAM-1 may be needed for interaction of the reversetransmigrating cells with the basal surface of the endothelium. Tissue factor, a transmembrane member of the coagulation cascade (Edgington et al., 1991), is another molecule that may participate in leukocyte–endothelial interactions during reverse transmigration (Randolph et al., 1998b).

Whether ICAM-1 and/or tissue factor have a role in the trafficking of DCs via authentic lymphatic vessels has not been determined. As mentioned earlier, both reverse transmigration in vitro and migration of DCs into dermal lymphatic vessels involves a role for the transmembrane transporter p-glycoprotein (Randolph et al., 1998c), but how this transporter participates in such trafficking is still under investigation. Chemokines, such as SLC, appear to promote the migration of peripheral DCs into lymphatics. How closely reverse transmigration across vascular endothelial cells, which do not express SLC in vivo, mimics this aspect of lymphatic trafficking is also not known. Reverse transmigration is sensitive to inhibition by pertussis toxin (Randolph, unpublished observations), consistent with the possibility that the process is mediated by chemokines and not a result of random migration.

INTEGRATED IN VITRO MODELS TO STUDY THE INFLUENCE OF ENDOTHELIAL CELLS UPON DC TRAFFICKING AND DEVELOPMENT – METHODOLOGY Clearly, much work is still needed to understand the molecular details and biological consequences of the interactions between DCs and endothelium. One limitation to progress in this field is lack of wide-spread use of endothelial culture systems. In addition, there is so far no source of cultured human or murine lymphatic endothelium. The identification of several lymphatic endothelial-restricted molecular markers, including desmoplakin (Schmelz et al., 1994), the VEGF-C receptor Flt-4 (Kaipainen et al., 1995), LYVE-1 (Banerji et al., 1999) and Prox-1 (Wigle and Oliver, 1999), should facilitate the establishment of cultured lymphatic endothelium. Here, somewhat similar, well-characterized methods that use HUVEC monolayers to study reverse transmigration are discussed. A revised model system that incorporates two endothelial

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monolayers, one of lymphatic origin if possible, would be most ideal. Figures 21.2 and 21.3 illustrate schematically two integrated culture systems that permit the study of apical-to-basal transendothelial migration, differentiation, and reverse transmigration back across the same endothelial monolayer. Endothelial cells isolated from umbilical veins (Jaffe et al., 1973) and grown in 20% human serum or fetal bovine serum without additional growth factors are seeded on to human amniotic matrix (Furie et al., 1984; Huang et al., 1988) or a reconstituted collagen type I matrix (Muller et al., 1989) within one to two passages from primary plating (Figure 21.2). Seeding of endothelium on to reconstituted collagen gels is most efficient if the collagen matrix is coated with fibronectin just prior to addition of endothelial cells. In contrast, growth of endothelium on amnion, a native human extracellular matrix composed predominantly of collagens type I and III (Lwebuga-Mukasa et al., 1984) and probably other matrix components, does not require addition of exogenous fibronectin. Thus, one advantage to the latter system is that it is quite straightforward to prepare LPS-free cultures, avoiding the use of commercial sources of extracellular matrix components, in which LPS levels vary from lot to lot. Another feature of the HUVEC/amnion model is that the type solutions, such as chemoattractants (Huang et al., 1988), or volumes of medium (Randolph and Furie, 1995) that bathe the basal surface of the cultures can be controlled or modified at any time as desired, whereas the basal aspect of the cultures using reconstituted collagen is enclosed and relatively inaccessible to manipulation (Figure 21.2). Weighing against these disadvantages, the reconstituted collagen system has a number of advantages, including less laborious set-up and a greater reproducibility between replicates, since the thickness of the amnion can vary somewhat between replicates. The major advantage of the reconstituted collagen gel model is the possibility of incorporating particulates in the matrix (Figure 21.3), since phagocytic particles enhance the differentiation of monocytes to DCs.

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In both model systems, monocytes and other DC precursors that are added to the apical aspect of the cultures migrate across unstimulated endothelium (Figure 21.3). A good physiologically relevant method to determine that endothelium is truly unstimulated and that the system is LPS-free is to add neutrophils to some control wells, as these cells do not migrate across resting endothelial monolayers, but readily migrate across stimulated endothelium (Huang et al., 1988; Furie and McHugh, 1989). To study reverse transmigration (Figure 21.3; see methods in Randolph and Furie, 1996; Randolph et al., 1998a, 1998b), the endothelium should be washed in medium after monocytes have been incubated with the monolayers for a time sufficient to allow maximal transmigration into the underlying matrix (1-2 hours). This washing step removes cells that failed to adhere or adhered loosely to the endothelium during incubation. Essentially all cells that adhere tightly to the endothelium readily migrate into the subendothelial matrix (Muller and Weigl, 1992; Meerschaert and Furie, 1994). Typically, replete culture medium (such as Medium 199) efficiently removes nonadherent cells from the apical compartment, but a more rigorous technique can be used in which the endothelium is washed briefly with Ca2- and Mg2-free buffer containing 1 mmol/L EGTA to disrupt integrin–ligand interactions. Then, complete culture medium is added and the incubation continued for durations sufficient to allow reverse transmigration (Figure 21.4). The two culture systems differ slightly in the half-time of reverse transmigration: between 24 and 36 hours in the amnion cultures (Randolph and Furie, 1996) and about 48 hours in the reconstituted collagen cultures (Randolph et al., 1998b). The recovery of reversetransmigrated cells (by simple pipeting from the apical compartment as detailed in Randolph et al., 1998b) is more efficient in the reconstituted collagen gel model when it is set up in microtiter wells, probably because of a more favorable geometry. If established as described, about 15 000 reverse-transmigrated cells are recovered per microtiter well (Randolph et al., 1998a, 1998b).

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CONCLUSION Inthelastseveralyears,tissueculturemodelscontaining DCs along with T cells or purified DC precursors alone have greatly advanced our understanding of DC maturation and immunologic function.Invitromodelswillnodoubtcontinueto be essential for the identification of key molecular events that can later be tested further for their overall biological and immunological impact using in vivo models.Well-controlled models that investigate DCs in a more physiologic context, such as in the study of their interactions with endothelium, will probably become very valuable tools for gaining still greater insight into the complex and critical ways that these antigen-presenting cells regulate immune responses.

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22 Dendritic cell/dendritic cell interaction Stella C. Knight and Penelope A. Bedford Imperial College School of Medicine, Northwick Park Institute for Medical Research, Harrow, UK

Self-control is the quality that distinguishes the fittest to survive. George Bernard Shaw

INTRODUCTION

or paracrine effect of cytokines produced by DCs. Both of these concepts have recently been introduced into the DC literature and there is only a small amount of published information. Consequently, after discussing this published work, some speculations that arise from these observations will be introduced. The capacity to stimulate proliferation of allogeneic lymphocytes in the mixed leukocyte reaction (MLR) is used as a functional property defining DCs. In this article, we discuss the unique capacity of DCs to transfer antigens, including MHC molecules, to other DCs that can, in turn, stimulate primary T-cell proliferation. One consequence of this functional transfer is that the MLR may, to a large extent, measure the capacity of DCs to transfer antigens to allogeneic DCs rather than their capacity directly to stimulate T cells. Thus, if we adhere to the concept that functional DCs must be able to stimulate a primary T-cell proliferation, only a proportion of the so-called DCs now studied may possess this property. Identifying the relevance

The maturational and functional state of dendritic cells (DCs) and their subpopulations may delineate not only whether an immune response to antigen occurs but also the nature of that response (Banchereau and Steinman, 1998; Kalinski et al., 1999). Following these realisations, the maturation of DCs and the interaction of DCs of different types with lymphocyte populations has been the focus of many studies. However, DC/DC interactions may occur before the DC/lymphocyte contacts; these DC/DC interactions may themselves determine the final signals that are to be delivered to lymphocytes. This early stage of DC intercourse potentially holds the key to whether a response will be stimulated and to establishing the nature of that response. Two aspects of ‘self regulation’ within DC populations will be discussed in this article. The first is the transfer of antigens between DCs and its relevance to the development of primary immune responses. The second is the autocrine Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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and mechanisms involved in antigen sharing by DCs constitutes another layer of understanding of the potential for these cells to regulate immune responses. The cytokine milieu also influences the maturation of DCs. DCs themselves produce cytokines for which they also express receptors (Fisher et al., 1999). A second mechanism by which DCs regulate their own development may be by the autocrine or paracrine effects of cytokines they produce and the modulation of these effects by exposure to different antigens.

RESPONSIVENESS AND NONRESPONSIVENESS INDUCED BY DENDRITIC CELLS Mature DCs are often thought of as nonphagocytic cells. However, Sigbjorn Fossum (Fossum, 1989), when describing DCs from the afferent lymphatics (veiled cells), summed up the position more accurately when he described them as having ‘a phagocytic past . . . and also . . . a phagocytic future’. Thus, Langerhans cells in the skin have phagocytic properties and interdigitating cells within lymphoid organs maintain some specific phagocytic functions; for example, they become packed with apoptosing cells when irradiated spleen cells are injected into mice. Fossum also stated, somewhat prophetically, that ‘the curiously selective taste for other cells suggests that dendritic leukocytes are involved in other activities beyond the presentation of foreign antigens. They could, for example, be key cells in a system for network regulation of T cells’. More recent studies show that some populations of DCs may interact with B cells directly and stimulate them (Wykes et al., 1998). The suggestion that DCs are key cells for network regulation of T cells could, therefore, be extended; they may additionally be involved in network regulation of B cells. DCs are usually defined as the only cells known to be potent at stimulating primary, immune response. Even in neonatal animals, antigen-pulsed DCs can cause stimulation of

immune responses (Ridge et al., 1996). However, DCs can also deliver negative signals to T cells. For example, DCs within the thymus can, under some circumstances, induce nonresponsiveness or negative selection (Inaba et al., 1991). DCs believed to be of a lymphoid-related lineage within the thymus are packed with apoptosing cells and the phenomenon of acquisition of dying cells by DCs has been associated with the development of tolerance. The peculiarity of DCs exposed to an antigen giving, apparently, either positive or negative signalling is also seen in autoimmune disease. DCs expressing autoantigen and injected intravenously can produce autoimmune disease (Knight et al., 1983; Knight et al., 1988; Ludewig et al., 2000). By contrast, DCs exposed to antigen and injected by routes which are considered more immunogenic may produce protection against the development of autoimmune disease (Whitacre et al., 1991; Clare-Salzler et al., 1992). From these various studies, the confusion of DCs initiating either responsiveness or tolerogenesis under different conditions is manifest. Multiple theories on the way in which different signals are delivered have been proposed. Signalling through the T-cell receptor in the absence of a second co-stimulatory signal may produce tolerance, rather than stimulation (Lenschow and Bluestone, 1993). Another idea is that cross-tolerance occurs when antigens are obtained from other cells (Carbone et al., 1998). However, in contradiction of this theory, DCs engulfing cells or cellular fragments that express a foreign antigen can themselves present that antigen to stimulate immunity (Inaba et al., 1998; Albert et al., 1998a; Albert et al., 1998b). The presence of inflammatory stimuli may also convert an otherwise tolerogenic signal into a stimulatory one (Finkelman et al., 1996)

TRANSFER OF ANTIGENS BETWEEN DENDRITIC CELLS A common denominator in the production either of immune stimulation or of tolerance is the efficient acquisition of cellular or other

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antigens by DCs. It has long been known that DCs acquire higher levels of antigens than other cells and employ a variety of mechanisms ranging from phagocytosis to endocytosis and pinocytosis. It has also been established that free MHC molecules can be found in the supernatants of lymphoid cells or even circulating in body fluids (Krangel, 1987; Puppo et al., 1995). Since DCs constitutively express high levels of MHC molecules, it is not surprising that evidence is now accruing that DCs can shed or secrete MHC molecules. Some of the data suggest that such release of MHC may be associated with exosomes shed from DCs (Zitvogel et al., 1998; Thery et al., 1999). DCs also acquire antigens shed from other cells and particularly from other DCs (Knight et al., 1998; Bedford et al., 1999). The capacity of DCs both to shed antigens and to acquire shed antigens provides an effective antigen transfer between DCs, that may be relevant to the role of this cell in the development of either immunity or tolerance. These properties are explored here, firstly in relation to the transfer of MHC molecules and then in the transfer of other antigens including antigenic peptides.

Transfer of MHC molecules between DCs In the development of allograft rejection, the major route of antigen presentation is an indirect one. The evidence is that alloantigens are acquired by DCs of the recipient and are presented syngeneically to produce allograft immunity (Benichou et al., 1997). This observation leads to a rather anomalous situation in that DCs of the graft tissue, as well as those of the recipient, are highly effective in promoting the development of immunity to allografts. The existence of an MLR, in which DCs stimulate proliferation of allogeneic lymphocytes, also indicates that there is something unique about DCs since they are the only cells that are potent at stimulating this reaction (Steinman and Witmer, 1978; Crow and Kunkel, 1982). Since DCs and not other cells cause stimulation of allogeneic lymphocytes, and since enriched T-cell populations respond markedly to stimulation by these allo-

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geneic antigen-presenting cells, it has been accepted that this response is caused by direct stimulation of DCs across an allogeneic barrier. However, high levels of production of MHC molecules by DCs may contribute to their high allostimulating capacity; both in the in vivo and in vitro studies of allostimulation, acquisition of MHC molecules by DCs of the recipient/responder type is a crucial step in the stimulation of alloreactivity. Early studies of in vivo allografting showed large mononuclear cells, that turned out to be DCs, which had acquired alloantigens and were present in graft recipients (Sherwood et al., 1986). More recently, acquisition of MHC molecules by recipient type DCs after injection of allogeneic DCs has been observed (Inaba et al., 1998). Although some of this antigen acquisition may be due to the peculiar appetite of DCs for dying nucleated cells bearing the alloantigen, these observations, nevertheless, raise the question that there may be transfer of MHC molecules between live DCs. Early studies by Dean Mann (Mann and Abelson, 1980) indicated that there was synergy between ‘monocytes’ of stimulator and responder type in the production of an MLR. Accessory cells bearing Ia molecules of responder type have also been implicated in the generation of allospecific cytotoxic T-lymphocyte responses (Weinberger et al., 1982), which is further indirect support for antigen transfer from allogeneic cells to antigen-presenting cells of the responder type. MHC molecules may be transferred freely from DCs to other cell types and much of the evidence purports to show the transfer of antigens directly from DCs to T cells (Arnold and Mannie, 1999). However, there is now direct evidence for the transfer of MHC molecules between allogeneic DCs. Our recent paper shows formally that transfer of MHC molecules between allogeneic DCs is significant (Bedford et al., 1999). The amount of MHC transferred between DCs is far in excess of that seen between other cells of the immune system such as T cells, macrophages, B cells or L cells transfected with MHC molecules or between DCs and these other cell types. DCs can secrete large amounts of MHC molecules. The observation

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that antibodies to MHC class II molecules of the responder type partially block MLR suggests that the MHC molecules themselves may be acquired by DCs, re-processed and presented syngeneically. However, the antigens within the supernatants of DCs may be in the form of exosomes shed from these cells (Thery et al., 1999) providing the possibility that many different materials may be transferred between DCs, perhaps even as ‘lipid rafts’. Other cells such as B cells and tumour cells also produce exosomes that contain MHC molecules (Raposo et al., 1996). The MHC molecules shed from other cells can be acquired by DCs but these DCs do not stimulate primary T-cell responses directly. However, DCs acquiring antigen from other DCs, directly stimulate T-cell responses (Bedford et al., 1999) (Figure 22.1). Evidence that this indirect presentation is a major stimulating pathway in the MLR is that removal of the last remaining DCs in a responder population of lymphocytes can cause severe depletion of the MLR. Very few DCs may be required within the responder population since the physical depletion of DCs

followed by final depletion with specific anti-DC antibodies and anti-class II antibodies with complement are required to deplete the response. Partial reconstitution of an MLR could also be achieved in the DC-depleted responder cells by adding metrizamide gradient-purified DCs from spleen or lymph nodes. The secondary acquisition of MHC class II molecules from allogeneic DCs and their presentation by cells syngeneic with the responder population fits more closely with the rules applying to other forms of antigen-presentation which require syngeneic antigen presenting cells. These observations raise concerns about the interpretation of studies claiming that different populations of DCs can stimulate primary allogeneic T-cell proliferation. The test as studied gives, at least in part, a measure of the capacity of DCs to transfer MHC molecules to DCs that do not already carry those antigens; the capacity of DCs to transfer antigen across this ‘antigen gradient’ provides the stimulus for the primary immune activation. The MLR may not only be a measure of the capacity of DCs to stimulate the T cells directly. In our continued studies, as shown in Table 22.1, if ‘DCs’ from different sources are mixed in culture, all of them are able to produce MHC molecules which are transferred to allogeneic DCs. However, DCs from spleen and lymph nodes acquire the antigen efficiently whereas DCs derived from bone TABLE 22.1 Transfer of MHC Class II molecules between allogeneic DCs Source of DC Bone marrow-derived (GM-CSF and TNFα) Splenic Lymph node

FIGURE 22.1 Stimulation of the MLR. DCs and not other cells are potent stimulators of the MLR. Since DCs stimulate proliferation of partially purified T cells, it was believed that direct stimulation across an allogeneic barrier was the main mode of antigen presentation (top diagram). However, a major component of the MLR is now known to be the stimulation of T cells syngeneically by DCs that acquire MHC molecules from allogeneic DCs (lower diagram).

Donating

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DCs derived from mouse bone marrow stem cells, from spleen or from lymph nodes, when mixed with allogeneic DCs for 24 hours, showed transfer of MHC between the cells as detected using strain-specific antibodies to Ia. All the DC types donated MHC molecules to allogeneic cells but only the splenic and lymph node DCs acquired high quantities of MHC. The results suggest that only mature cells from the spleen and lymph node are specialised to acquire the antigen from other DCs.

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marrow stem cells after culture with maturational cytokines fail to acquire MHC molecules as measured by flow cytometry (Bedford and Knight, unpublished observations). Some of those populations now called ‘dendritic cells’ may not have the capacity directly to stimulate primary T-cell responses.

Transfer of other antigens between DC In secondary immune responses, antigens added into cell cultures containing antigenpresenting cells and sensitised T cells may cause proliferation. However, primary immune responses have been difficult to obtain. A common feature of the studies that have successfully produced primary proliferative responses is the use of a population of stimulator cells that has been pre-pulsed with antigen. Such prepulsed DCs stimulate primary, proliferative and cytotoxic T-cell (CTL) responses efficiently (Macatonia et al., 1989, 1991; Vyakarnam et al., 1991; Williams et al., 1991; Croft et al., 1992; De Bruijn et al., 1992). The number of cells required to produce such primary responses indicated that of the order of one cell per 200–500 CD8+ T cells were responding to a single T-cell epitope of influenza virus (Macatonia et al., 1989); such proportions of specific T-cells are close to the theoretical numbers of cells capable of responding to individual epitopes now detected by using tetramer antibody staining for specific T-cell receptors (Murali-Krishna et al., 1998). Pulsed DCs can thus produce efficient primary responses in vitro. Such responses may be produced by an antigen-handling mechanism similar to that already described for the MLR. Antigens may be transferred syngeneically across an antigen gradient and be ‘re-presented’ to give primary responses. This requirement for antigen transfer between DCs may account for the necessity to use antigen-pulsed DCs to produce primary responses. In vitro transfer The evidence that DCs can produce histocompatibility antigens that are transferred to other

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cell types has been discussed. There is evidence that antigens other than histocompatibility antigensarealsotransferredbetweenDCs.Aswith the transfer of MHC molecules, DCs acquire high quantities of antigens from syngeneic DCs. The amount of antigen acquired by T cells from DCs (Emerson and Cone, 1982; Arnold and Mannie, 1999) is small compared with that acquired by other DCs (Knight et al., 1998). DCs acquiring antigens from other DCs are also powerful simulators of primary T-cell proliferation, whereas DCs acquiring antigen directly do not stimulate primary responses in highly purified, DC-depleted T-cells. Thus, using antigen-pulsed DCs, responses were only obtained when DCs not directly exposed to antigen were present additionally in the responder population. If all the DCs expressed the same antigen, there was no primary T-cell stimulation (Knight et al., 1998) (Figure 22.2). This synergy between DCs initially with and without antigen accounts for the requirement for pre-pulsed DCs to stimulate primary responses. It is of interest to identify the mechanisms involved in this type of DC interaction. It is not clear whether the antigen is transferred between DCs together with the MHC molecules to which they may be bound. Mature lymph node DCs express high levels of MHC class II at the cell surface although synthesis may be reduced (Kampgen et al., 1991) and some class II molecules at the cell surface may be acquired from other DCs. The internalisation of different MHC class II molecules from the cell surface into different types of vesicles (Knight et al., 1987, 1989) may, in part, reflect the processing of secondarily acquired MHC molecules. Current theories to explain the requirement for DCs with secondarily acquired antigens to induce primary T-cell stimulation include the idea that antigen that is ‘self processed’ by DCs provides a negative signal to the T cells, while processed antigen acquired from other DCs signals positively. Thus, a commonly distributed antigen would, on balance, provide negative signalling to the T cells. By contrast, an antigen, present only on a proportion of DCs, would be acquired by some DCs that had not already processed this antigen. DCs acquiring antigen

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FIGURE 22.2 Stimulation of primary T cell responses to antigen. Antigen that is processed directly by DCs fails to stimulate T-cell proliferation. The antigen may also be transferred between DCs but, without an antigen gradient between these DCs, neither the directly acquired nor the transferred antigen stimulates T cells (top diagram). However, antigen acquired from other DCs is stimulatory in cells that do not already express that antigen (lower diagram). The antigen transfer between DCs may be in the form of ‘dendrosomes’ (exosomes with the capacity to stimulate primary immune responses on acquisition by DCs), which can also contain factors such as heatshock proteins that may potentiate the immune response. The outcome of these effects is that DCs do not stimulate responses to widely distributed environmental antigens. They will initiate responses to antigens not directly in their own environment, particularly if they are damaging.

from other DCs, i.e. antigen that was not selfprocessed would cause positive signalling. Self/nonself discrimination could equate to the balance between self processed and nonself processed antigen in the DC. A major impetus to the development of immune responses could, therefore, be the distribution of antigen on DCs. If an antigen at either high or low concentration was universally expressed, then ‘tolerance’ would be maintained. This ubiquity of antigens and DCs could account for the tolerogenic effects of intravenous exposure to antigen (Miller et al., 1979). However, an antigen not widely expressed but arriving asymmetrically would be presented. One can speculate that if the first DC was also

damaged, then the production of materials such as heat-shock protein or some change in other chaperone proteins would be associated with the development of positive signalling on transfer of processed antigen to the recipient DC. Such a system incorporates not only the popular view that damage promotes the maturation of DCs (Matzinger, 1994) but also provides a mechanism whereby common environmental antigens are tolerogenic but less widely distributed and damaging antigens are immunogenic. There are some clues as to the nature of the material that is transferred between DCs providing support for this hypothesis. DCs shed exosomes that contain not only MHC molecules but also heatshock proteins and integrin molecules (Thery et al., 1999). DCs may also shed functional mannose receptor (Jordens et al., 1999). Other cell types produce exosomes, but the studies on primary T-cell stimulation in vitro, like those with the MLR, indicate that only the exosomes from DCs, that have been efficiently acquired by other DCs, produce primary T-cell stimulation directly (Knight et al., 1998). We suggest the term ‘dendrosomes’ for the primary stimulatory material shed from DCs. In vivo effects The previous sections have described the production of antigen-specific materials by DCs that can stimulate primary T-cell responses even across an allogeneic barrier. Such materials, if they can be obtained in quantity and stabilised, could provide ‘the Holy Grail’ for vaccination – an antigen-specific, primary stimulatory factor that can be effective even across an allogeneic boundary. Dendrosomes shed from DC-bearing tumour antigens have already been used to produce protective tumour immunity (Zitvogel et al., 1998). From the in vitro studies described earlier, it is most likely that such materials require DCs of the recipient for their presentation and stimulation. It is also likely from in vitro studies that cellular or other antigenic materials released from DCs may need to be acquired by DCs that have not already been exposed to that antigen in order to produce primary immune

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responses. The effectiveness of hybrids between tumour cells and DCs in stimulating immunity (Scott-Taylor et al., 2000) may induce protection via a similar mechanism. One formal set of experiments directed at this question of antigen transfer in vivo indicates that antigen can be transferred between different types of DC in order to produce an immune response. Thus, CD8α+ DCs exposed to antigen and injected into animals remained at the local intradermal site of injection and yet responses were produced. CD8α+ cells may have transferred antigen to CD8α- migratory DCs travelling to the draining lymph nodes and stimulating primary immune response (Smith and de St Groth, 1999). Other formal experiments investigating antigen transfer between DCs in vivo in development of primary immune responses are lacking. Since there is a lack of formal evidence on this point, in the last part of this section we will speculate on the importance of antigen transfer by quoting some examples where a mechanism of this type could be operating. In neonatal animals, when large quantities of antigen are injected, tolerance can result. Antigens that are present around the thymus capsule are very rapidly acquired and distributed throughout DCs within the thymus and may, therefore, be expected to cause negative signalling by their wide distribution. A prediction would be that, if antigen was injected only on DCs pulsed with antigen, an antigen gradient would be created with positive rather than negative signalling. Experiments have shown this to be the case (Ridge et al., 1996). Another prediction can be made in the case of chronic infection; further exposure to widely distributed antigens is unlikely to produce positive signalling and immunity. However, the use of cryptic antigens, which are not major antigens of viruses, for example, might be expected to give positive signalling. The use of cryptic rather than commonly immunogenic antigens from CMV virus, produced protective immunity in chronic infection (van der Most et al., 1996). In early studies on tumour immunity, mice bearing large tumours showed tumour regression when DCs not exposed to tumour antigens were injected;

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DC-bearing tumour antigens were ineffective (Knight et al., 1985). Such studies provide support for the idea that antigen transfer may be occurring; one interpretation of such data is that tumour immunity was initiated by the creation of an antigen gradient between DCs with and without tumour antigen. Protection against tumours given DCs pulsed with tumour antigen may occur efficiently only when small tumours are present (Knight et al., 1985). If self antigens or nonself antigens that are widely distributed are signals for DCs to produce negative rather than positive signalling, then these effects are relevant, not only to the development of protective immunity, but also to the production of autoimmunity. A major cause of autoimmunity is believed to be the development of immune responses to self antigens initiated when they are delivered as part of organisms which possess epitopes that are cross reactive with self proteins. However, incorporation of antigens of the host could equally be a stratagem evolved by the organism for evading the host immune response. It is striking that the production of autoimmunity by DCs exposed to autoantigens is achieved by intravenous injection, (Knight et al., 1983, 1988) a route associated with nonresponsiveness. By contrast, the use of more immunogenic routes of immunisation results in protection against the development of autoimmunity (Clare-Salzler et al., 1992). The inflammation associated with DCs exposed to autoantigens could thus be associated with nonresponsiveness to antigen. DCs with autoantigen, or processing organisms with antigens crossreactive with self antigens, when reaching the target tissue, would encounter other cells processing the same antigens, there would be no antigen gradient and negative signals would result. Since migration of DCs to the lymph nodes is associated with maturation of the capacity to stimulate T cells this situation might result in lack of DC migration and accumulation of DCs within the target tissue. Accumulation of DCs within the target tissues in autoimmune disease is a common, early feature. Inappropriate accumulation of DCs, perhaps some of them having been stimulated by ‘innate

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mechanisms’, may cause nonspecific inflammation. One example providing support for such a hypothesis is the development of reactive arthritis following genital tract infection with Chlamydia trachomatis, an organism that bears epitopes crossreacting with collagen. C. trachomatis induces production of IL-12 in DCs. Accumulation of DC-bearing chlamydial antigens and nonspecific accumulation of lymphocytes occurs in the joints in postchlamydial arthritis (Stagg et al., 1996). Nonresponsiveness to foreign organisms because of the crossreactive ‘self-like’ components may occur and could be operating at the level of DC/DC interactions. The observations on antigen transfer and the crosstalk between DCs, are controversial but they lead to novel views of the role of DCs in the development of immunity. At the very least, the efficiency with which DCs can transfer antigens, both acquired and constitutive, to other DCs is a mechanism for increasing the sensitivity of responses in rare populations of naïve lymphocytes. However, the current evidence is that this transfer of antigens may not just enhance the development of primary immune response but may also be a fundamental part of the control of those immune responses.

AUTOCRINE PATHWAYS IN DCS The cytokines produced by DCs and their effects on the immune system are discussed in detail elsewhere in this volume. However, in this chapter, the principal of autocrine or paracrine effects of the DC cytokines that promote the development of DCs polarised to produce different immune responses, will be highlighted briefly with reference to just two of the major cytokines. The two cytokines concerned are IL12 and IL-4, which play seminal roles in the development of TH1 and TH2 T-cell responses. Early in the immune response, DCs are the major cells producing IL-12 (Macatonia et al., 1995). However, DCs and their progenitors express IL-12 receptors (Fisher et al., 1999).

IL-12 can act on these receptors, stimulating, via an NFκB pathway, the production of further IL-12 (Grohmann et al., 1998). Exposure to IL-12 in developing DCs also causes the upregulation of CD80 on the surface of these DCs, which then stimulate primary T-cell responses more robustly (Kelleher and Knight, 1998). Some viruses, bacteria and parasites can promote the production of IL-12 by DC (Reis e Sousa et al., 1997; Gorak et al., 1998; Hessle et al., 1999; Konecny et al., 1999). The promotion of IL-12producing DCs by the presence of IL-12 may be a long-lived effect in vivo that acts through an effect of IL-12 on the stem cells giving rise to DCs (Kelleher et al., 1999a). The long-term autocrine or paracrine effect of the IL-12 on the in vivo developing DC system may provide one mechanism by which that system is ‘educated’ by early antigenic experience. Other viruses and bacteria are known to switch the immune system to produce less cell-mediated immunity and more antibodymediated effects. One such virus, Rauscher leukaemia virus, was recently shown to switch the production of cytokines in DCs from IL-12 to IL-4 (Kelleher et al., 1999b). DCs also express IL-4 receptors and IL-4, therefore, is likely to cause effects in DCs signalled via this receptor. Some published evidence indicates that there is an autocrine loop for IL-4 in DCs. This information shows that high doses of IL-4 will initiate the apoptotic death of DCs (Rissoan et al., 1999). Our own studies have now shown that IL-4 itself can act on DCs to initiate the production of IL-4 in those cells (Maroof and Knight, in preparation). Thus, a similar autocrine pathway to that described for IL-12 may exist. As suggested in Figure 22.3, it seems likely that exposure of developing DCs to some organisms, or the presence of IL-4 itself, in the milieu of the developing cells will initiate or promote the production of IL-4 within DCs. In the presence of excess IL-4, however, the cells will undergo apoptosis suggesting an autocrine or paracrine feedback mechanism when there is overproduction. These two examples of autocrine pathways in DCs may indicate important control mechanisms in developing DC populations which,

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FIGURE 22.3 Autocrine/paracrine effects of IL-4. Exposure of DCs to IL-4 during their development causes the initiation of IL-4 production in the DCs providing a possible autocrine or paracrine loop polarising the cytokine profile within DCs and influencing the type of immune response generated. Higher concentrations of IL-4 lead to death of the DCs and may constitute a feedback mechanism for removing DCs when cytokine production reaches a certain level.

again, may operate within these populations themselves and precede the later two-way multiple interaction with cells of the lymphoid system.

CONCLUSIONS In conclusion, developing DCs may interact at multiple levels during their development and maturation. Such interactions will include the transfer of antigens, both endogenously and exogenously acquired by DCs, to other DCs. When these other DCs are exposed directly to the same antigen there is no stimulation of Tcells. If the DCs are not themselves directly exposed to the antigen, and the transfer occurs across an antigen gradient, then the DCs with secondarily acquired, processed antigen may be potent stimulators of primary T-cell activation; DCs may not, therefore, stimulate responses to antigens in their own immediate environment. A second self regulatory mechanism for DCs involves the autocrine or paracrine effect produced by different DC populations; such effects may occur with production in DCs of the

cytokines IL-12 and IL-4 associated with TH1 and TH2 responses. Production of these cytokines in DCs may act on DCs themselves, further polarising the cytokine production in these cells. Knowledge of DC/DC interactions is in its infancy. The role which different subpopulations of DC, and their different states of maturation, play in these multiple DC/DC interactions are beginning to show the ways in which DCs exposed to antigen can regulate their own development – before they influence the development of immune responses within the lymphocyte populations.

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Inaba, K., Turley, S., Yamaide, F. et al. (1998). J. Exp. Med. 188, 2163–2173. Inaba, M., Inaba, K., Hosono, M. et al. (1991). J. Exp. Med. 173, 549–559. Jordens, R., Thompson, A., Amons, R. and Koning, F. (1999). Int. Immunol. 11, 1775–1780. Kalinski, P., Hilkens, C.M., Wierenga, E.A. and Kapsenberg, M.L. (1999). Immunol. Today 20, 561–567. Kampgen, E., Koch, N., Koch, F. et al. (1991). Proc. Natl Acad. Sci. USA 88, 3014–3018. Kelleher, P. and Knight, S.C. (1998). Int. Immunol. 10, 749–755. Kelleher, P., Williams, N.J. and Knight, S.C. (1999a). Immunol. Lett. 65, 51–54. Kelleher, P., Maroof, A. and Knight, S.C. (1999b). Eur. J. Immunol. 29, 2309–2318. Knight, S.C., Farrant, J., Chan, J., Bryant, A.E., Bedford, P.A. and Bateman, C. (1988). Clin. Immunol. Immunopathol. 48, 277–289. Knight, S.C., Fryer, P., Griffiths, S., Harding, B., Dixey, J. and Ansell, B.M. (1989). Clin. Exp. Immunol. 78, 19–25. Knight, S.C., Fryer, P.R., Griffiths, S. and Harding, B. (1987). Immunology 61, 21–27. Knight, S.C., Hunt, R., Dore, C. and Medawar, P.B. (1985). Proc. Natl Acad. Sci. USA 82, 4495–4497. Knight, S.C., Iqball, S., Roberts, M.S., Macatonia, S. and Bedford, P. (1998). Eur. J. Immunol. 28, 1636–1644. Knight, S.C., Mertin, J., Stackpoole, A. and Clarke, J.B. (1983). Proc. Natl Acad. Sci. USA 80, 6032–6035. Konecny, P., Stagg, A.J., Jebbari, H., English, N., Davidson, R.N. and Knight, S.C. (1999). Eur. J. Immunol. 29, 1803–1811. Krangel, M.S. (1987). Hum. Immunol. 20, 155–165. Lenschow, D.J. and Bluestone, J.A. (1993). Curr. Opin. Immunol. 5, 747–752. Ludewig, B., Ochsenbein, A.F., Odermatt, B., Paulin, D., Hengartner, H. and Zinkernagel, R.M. (2000). J. Exp. Med. 191, 795–804. Macatonia, S.E., Hosken, N.A., Litton, M. et al. (1995). J. Immunol. 154, 5071–5079. Macatonia, S.E., Patterson, S. and Knight, S.C. (1991). Immunology 74, 399–406.

Macatonia, S.E., Taylor, P.M., Knight, S.C. and Askonas, B.A. (1989). J. Exp. Med. 169, 1255–1264. Mann, D.L. and Abelson, L. (1980). Cellular Immunology 56, 357–364. Matzinger, P. (1994). Annu. Rev. Immunol. 12, 991–1045. Miller, S.D., Wetzig, R.P. and Claman, H.N. (1979). J. Exp. Med. 149, 758–773. Murali-Krishna, K., Altman, J.D., Suresh, M. et al. (1998). Immunity. 8, 177–187. Puppo, F., Scudeletti, M., Indiveri, F. and Ferrone, S. (1995). Immunol. Today 16, 124–127. Raposo, G., Nijman, H.W., Stoorvogel, W. et al. (1996). J. Exp. Med. 183, 1161–1172. Reis e Sousa, C., Hieny, S., Scharton-Kersten, T. et al. (1997). J. Exp. Med. 186, 1819–1829. Ridge, J.P., Fuchs, E.J. and Matzinger, P. (1996). Science 271, 1723–1726. Rissoan, M.C., Soumelis, V., Kadowaki, N. et al. (1999). Science 283, 1183–1186. Scott-Taylor, T.H., Pettengell, R., Clarke, I. et al. (2000). Biochim. Biophys. Acta 1500, 265–279. Sherwood, R.A., Brent, L. and Rayfield, L.S. (1986). Eur. J. Immunol. 16, 569–574. Smith, A.L. and de St Groth, B.F. (1999). J. Exp. Med. 189, 593–598. Stagg, A.J., Hughes, R.A., Keat, A.C.S. and Knight, S.C. (1996). B. J. Rheumatol. 35, 1082–1090. Steinman, R.M. and Witmer, M.D. (1978). Proc. Natl Acad. Sci. USA 75, 5132–5136. Thery, C., Regnault, A., Garin, J. et al. (1999). J. Cell Biol. 147, 599–610. van der Most, R.G., Sette, A., Oseroff, C. et al. (1996). J. Immunol. 157, 5543–5554. Vyakarnam, A., Matear, P.M., Cranenburg, C. et al. (1991). Int. Immunol. 3, 939–947. Weinberger, O., Germain, R.N., Springer, T. and Burakoff, S.J. (1982). J. Immunol. 129, 694–697. Whitacre, C.C., Gienapp, I.E., Orosz, C.G. and Bitar, D.M. (1991). J. Immunol. 147, 2155–2163. Williams, N.A., Hill, T.J. and Hooper, D.C. (1991). Immunology 72, 34–39. Wykes, M., Pombo, A., Jenkins, C. and Macpherson, G.G. (1998). J. Immunol. 161, 1313–1319. Zitvogel, L., Regnault, A., Lozier, A. et al. (1998). Nat. Med. 4, 594–600.

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23 Dendritic cells in the context of skin immunity Adriana T. Larregina and Louis D. Falo Jr University of Pittsburgh School of Medicine, Pittsburgh, PA, USA

You never really understand a person until you consider things from his point of view . . . ’til you climb inside his skin and walk around in it. Atticus Finch, in To Kill a Mockingbird Harper Lee

INTRODUCTION

homeostasis of the integument, have led to consideration of the skin as an immune-competent organ. Nominations such as skin-associated immune system (SALT) (Streilein, 1983), skin immune system (SIS) (Bos and Kapsemberg, 1986, 1993; Bos, 1997), dermal microvascular unit (Sontheimer, 1989) dermal immune system (DIS) (Nickoloff, 1993) or pilosebaceous unit (Böhm and Luger, 1998) are examples of efforts to classify cellular and humoral elements with immune-related functions that are either resident or attracted to the skin during inflammatory or immune responses. Skin cells and structures that are associated with immune function are listed in Table 23.1. We will focus on morphological aspects, phenotype and functions of the population of dendritic cells (DCs) resident in the skin. Individual descriptions of each member of the skin immune system exceeds the scope of the

Throughout evolution higher organisms have developed nonspecific and specific defense mechanisms to protect themselves against foreign aggressors. Nonspecific mechanisms include a first line of defense represented by mechanical barriers and products of secretion. When epithelial barriers are breached, nonspecific inflammatory and adaptive immune responses are activated. The characteristics of adaptive immune response are specificity, memory and discrimination of self from nonself antigens. Externally, the skin and internally, the mucous membranes constitute these first mechanical barriers fully equipped to trigger nonspecific inflammatory and specific immune responses. The coordinated capacity of cutaneous immune cells to initiate and regulate the intensity of skin immune responses in order to maintain the

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TABLE 23.1 Skin cells and structures with immuneassociated function Epidermis

Dermis

Keratinocytes

Dendrocytes

Langerhans cells αβ: CD8 and CD44

Mast cells Vascular endothelial cells

T lymphocytes γδ: (mouse) Cutaneous nerves

Lymphoid endothelial cells Fibroblasts Tissue macrophages T lymphocytes αβ CD8 AND CD4 Pilosebaceous unit Dermal microvascular unit

Modified from Bos et al., 1993, 1997; Nickoloff, 1993; Böhm and Luger, 1998.

present chapter, and their function will be mentioned only in the context of their association with skin DCs.

LANGERHANS CELLS General aspects and cytological characteristics Skin Langerhans cells (LCs) were first described as intraepithelial neurons by Paul Langerhans in 1868. In 1876, Arstein and in 1896, Herxeimer considered LCs for the first time, as epidermal passenger leukocytes. During the following years, different origins and functions were given to LCs. It was not until 1976, and after the description of the allostimulatory function of spleen DCs (Steinman and Cohn, 1973), that epidermal LCs were considered as leukocytes from the immune system, involved in the development of hypersensitivity reactions (Silberberg et al., 1976). It is now very well established that epidermal LCs are antigen processing and presenting cells resident in the epidermis capable of taking up antigens and migrating to draining lymph nodes where they present processed antigens to naïve or memory T cells (Banchereau and Steinman, 1998). Situated in the basal and suprabasal layers of the epidermis, LCs constitute an intraepithelial net with a sentinel function capable of capturing

and processing nonself or self-antigens. They represent 2–4% of the total epidermal cell population and their density varies between approximately 200 and 1000 cells/mm2 of the epidermal surface depending on the anatomical area (more abundant in face and neck skin) (Berman et al., 1983). LCs are very difficult to visualize in sections routinely stained with H&E, however, they can be identified using metal impregnation techniques, enzymatic detection of adenosine triphosphatase (ATPase), or immunohistochemistry (reviewed by Schuler in 1991). Ultrastructurally, the hallmark of LCs is the Birbeck granule (BG), a rod-shaped trilaminar membrane structure initially described by Birbeck in 1961. In 1966 BGs were found in cells of histiocytosis X, a neoplasm of LCs (Basset and Nezellof, 1966). Although not well-determined, the BG has been postulated to be an organelle with antigen uptake function (Strobel et al., 1996). Besides BGs, LCs contain abundant mitochondria, lysosomes, Golgi regions, intermediate filaments of vimentin, and a dark indented nucleus (Schuler et al., 1991).

PHENOTYPE OF LANGERHANS CELLS AND DERMAL DENDRITIC CELLS The origin and functional stage of LCs determine their complex phenotype. LCs and dermal dendritic cells (DDCs) are leukocytes, originated from bone marrow progenitors that colonize the skin after leaving the peripheral blood circulation. Upon entering the skin, resident epidermal LCs and DDCs remain as resting cell populations until a ‘danger signal’ induces their activation and mobilization to draining lymph nodes. The phenotype of LCs, and probably DDCs, differs according to their functional state, which is determined by molecular changes in the microenvironment such as secretion of cytokines, chemokines or other proinflammatory factors (Romani et al., 1988). Differences in the phenotype and function of skin LCs at different maturation stages are listed in Table 23.2.

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TABLE 23.2

Characteristics of resting and mature Langerhans cells

Characteristic Cytology Dendritic processes Birbeck granules Acidic organelles Phenotype Langerin/lag Antigen presenting molecules MHC-class I MHC-class II (surface) CD1a CD1c Fc receptors CD32 FCεRI Adhesion molecules β1 integrins CD44 CD54 CD58 Costimulatory molecules CD40 CD80 CD86 Activation markers CD83 CD25 CD69 Cytokine/chemokine receptors GM-CSF-Rα (CD116) Common β Chain (CD131) TNFRII (CD 120bb) IL-IRI (CD121a) IL-IRRII (CD121b) IFNγ R (CD119) IL-6Rα (CD126) Gp-130 (CD130) C5aR (CD88) CCR1 CCR4 CCR5 CCR6 CCR7 CXCR4 Platelet activating factor Cytokine/Chemokines secretion IL-1β IL-6 IL-12 IL-15 IL-18 TNFα IFNγ SCF MIP-1α MIP-2 Antigen uptake/processing Antigen presenting

Resting LC

Mature LC

  

 / 



/

   

   /

 

/ 

  (S) / 

α4↑/α6↓ v3v6v9 ↑  

/  

  

  

  

  (subpopulation)        (subpopulation)       

               

  /  /

    

     /

    / 

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Molecules commonly used to identify Langerhans cells and dermal dendritic cells Because LCs and DCs in general lack specific lineage markers, they were defined as leukocyte populations expressing the CD45 molecule and the myeloid markers CD13 and CD33, and are negative for other lineage-specific markers: CD3, CD19, CD20, CD14, CD16 and CD56. Although not restricted to DC lineage, the molecules CD1a, and CD1c expressed by human LCs, and MHC class II, CD11c, or DEC-205 in mouse, have been broadly used to identify LCs. Recently, the presence of the langerin and the intracytoplasmic lag antigen, has also been used as specific markers for LCs (Kashihara et al., 1986; Valladeau et al., 2000). It has been demonstrated that anti-lag antibody recognizes an intracellular epitope of the langerin molecule. Langerin is a Ca2-dependent type II transmembrane protein with a single carbohydrate recognition domain, which is specifically expressed on the membrane of the LCs. Ligation of langerin by the specific mAb antibody DCGM4, or mannan, resulted in the internalization of the langerin and in the generation of BGs. The expression of langerin is downmodulated after LC maturation consistent with the diminished number of intracellular BGs (Valladeau et al., 2000). According to their HLA-DR expression, two different LC populations can be observed in the epidermis: DRlow (60–75%) and DRhigh (25–10%). The cause of this increased expression and its significance has not been determined (Dezutter-Dambuyant et al., 1984). Probably these cells correspond to a subpopulation of large, activated LCs that can be easily seen in freshly isolated epidermal sheets (Romani et al, 1991). Dermal dendrocytes (DDCs) share most of the phenotype of mature LCs, however they express the intracytoplasmic transglutaminase clotting factor XIIIa (FXIIIa) (Headington, 1986; Cerio et al., 1989), lower levels of CD1a and CD1c than LCs, and they express CD1b (Nestle et al., 1993; Lenz et al., 1993). In addition, DDCs can be dis-

tinguished from immature LCs by the absence of E-cadherin, or lag. According to the presence of FXIIIa, CD1a and CD14, three different populations of DCs migrate from dermal explants: (1) FXIIIa, CD1a, CD14; (2) FXIIIa, CD1a, CD14; (3) FXIIIa, CD1a, CD14. In T-cell allogeneic proliferation assays, the first and second populations induced strong stimulatory responses, however, the third population (CD1a CD14) induced a level of T-cell proliferation similar to that of circulating monocytes (Nestle et al., 1993; Nestle and Nickoloff, 1995). Other studies suggested that the stimulatory function of DDCs was only achieved by a trace cell population of dermal CD1 LC-like cells (Meunier et al., 1993; Sepulveda-Merril et al., 1994). However, it is interesting that these last studies used DDCs obtained after enzymatic dermal disruption and not after migration. Thus, it could be assumed that most of the DDCs isolated correspond to a resting DC population and, like freshly isolated epidermal LCs, they are very weak stimulators of T-cell responses. In mice, two populations of DDCs had been described. CD11b and CD11b and cells expressing the marker NLDC-145 (DEC-205) can be found in the dermal perivascular unit (Duraiswamy et al., 1994). Upon maturation, LCs downregulate CD1a, E-cadherin, langerin and lack BGs, acquiring a similar phenotype of the migrated population of DDCs, however mature LCs do not express FXIIIa (review by Nestle and Nickoloff, 1995).

Molecules involved in antigen capture and antigen presentation function As professional antigen presenting cells (APCs) LCs express class I (HLA-A, B and C), (Bronstein et al., 1993), and class II (HLA-DR, DP, and DQ) molecules from the major histocompatibility complex (MHC) (Sontheimer et al., 1986). They also present the invariant chain CD74 that is related with transportation assembly and posterior membrane expression of α and β subunits of the class II molecules (Austyn and Wood, 1993). Related to their antigen capture function, LCs

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constitute the only epidermal cell population that express Fc-IgG receptor type II (CD32) (Romani et al., 1991; de la Salle et al., 1992). They also exhibit high and low affinity receptors for the Fc portion of IgE (Fcε-RI and Fcε-RII) and IgE-binding protein (EBP), all structures implicated in allergen uptake (Bierber et al., 1992; Rieger et al., 1992). Murine LCs and DCs express DEC-205 multilectin receptor. Human LCs do not express the macrophage mannose receptor multilectin, related to the internalization of mannosylated antigenic proteins (Mommaas et al., 1999), however the presence of langerin, with a mannose-specific binding site, in the membrane of immature human LCs, has been related to the internalization and processing of mannose molecules (Valladeau et al., 2000). Resting LCs do not express or they express only very low levels of the co-stimulatory molecules CD80 (B7.1) or CD86 (B7.2) (Symington et al., 1993; Wolf et al., 1994). After maturation, they express de novo, or increase the expression of these three molecules. Binding the co-receptor CD28 and CTLA4 (CD154) on the membrane of T cells, CD80 and CD86 participate in the generation of the second signal to activate a T cellspecific immune response during the DC/T cell dialogue (Azuma et al., 1993). LCs also express low levels of the CD40 molecule. The crosslinking of the CD40 molecule induces DC activation (Caux et al., 1994), and prevents DCs from undergoing apoptosis ‘in vitro’ (Ludewing et al., 1995).

Receptors for complement, chemokines, cytokines and growth factors Langerhans cells are capable of responding to nonspecific inflammatory stimuli and mediators such as pro-inflammatory cytokines, complement and growth factors are implicated in their entrance, homing, activation and migration from the skin. They express functional receptors for CR3bi (CD11b–CD18) (De Panfilis et al., 1990), C5aR (CD88) (Morelli et al., 1996), and for the chemokine receptors CCR1, CCR4, CCR5, CCR6, and CXCR4. After maturation they

305

downregulate CCR6 and upregulate CCR7, a molecule that is involved in lymph node chemotaxis and homing. Importantly, CXCR4 and CCR5 are receptors for the T-tropic and Mtropic strains of HIV-1, and their relation with productive HIV-1infection of DCs and its transmission is being investigated (reviewed by Sozzani et al., 1999). Immature LCs also express GM-CSFαR (CD116), IL-1R type I and type II (CD121a and b), TNF-RI (55 kDa) (CD 120a, only in mouse DCs), TNF-RII (75 kDa) (CD120b, mouse and human LCs), IL 6R α chain and low β chain (CD126 and CD130, respectively), and IFNγ R (CD119). After maturation in short-term cultures, they upregulate the common β chain of GM-CSF, IL-3 and IL-5 (CD131), IL-1R type I and type II, they express de novo IL-2Rα (CD25) and IL-6Rβ (CD131), and they downmodulate TNFαRII (Larregina et al., 1996). Dermal dendrocytes migrated from human dermal explants present a similar pattern of cytokine receptors to that of cultured LCs. In addition, they express IL3Rα (CD123), IL-4Rα (CD124), IL-7Rα (CD127), and the common γ chain of IL-2R, IL-4R, IL-7R, IL-9R, and IL-15R (CD132) (Larregina et al., 1997).

Molecules involved in the migratory cell cycle and homing Langerhans cells and DDCs constitute migratory cell populations. They arrive in the dermis possibly as an immature precursor from peripheral blood and reside as immature cells in the epidermis or in dermis until a stimulatory signal is able to induce their migration. During this migratory cycle, LCs differentially modulate adhesion molecules that allow their interaction with other cells or with extracellular matrix proteins. Skin DCs, their precursors, and 15–20% of the circulating T cells that colonize the skin, express in their membranes the molecule CD15s, also known as Siayl Lewis X or cutaneous lymphocyte-associated protein (CLA) (Ross et al., 1994). CD15s is a co-receptor for E-selectin (CD62 E) expressed on the surface of activated endothelial

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cells. The presence of CD15s allows the attachment and transendothelial migration of circulating cells to the dermis (Koszick et al., 1994). Keratinocyte/LC association is mediated through E-cadherin, a membrane glycoprotein that mediates Ca2-dependent unions. During their activation, LCs decrease the level of Ecadherin expression and are ‘released’ from neighboring epidermal cells (Tang et al., 1993). Resident LCs express β1-integrins (α4, and α6) (Le Varlet et al., 1991), and the molecule CD44s. After activation with TNFα, LCs downregulate α6 integrins (Le Varlet et al., 1992; Price et al., 1997) and they increase the expression of α4 integrin and CD44v3, v6 and v9 (HaegelKronenberger et al., 1998). β1-integrins and CD44 isoforms mediate the adherence of LCs to extracellular proteins such as fibronectin and hyaluronic acid, respectively. β2 integrins (CD11a, CD11c and CD18) (Simon et al., 1993), as well as adhesion molecules from the immunoglobulin superfamily (CD50, CD54, CD58 and CD102) (Hart and Pricket, 1993; Acevedo et al., 1993; Teunissen et al., 1994) are expressed preferentially after LC maturation and mediate adherence to activated endothelial cells and to T cells during antigen presentation. Curiously, LCs also express high levels of CD43, a glycoprotein rich in sialic acid residues, that confers negative charges and can act as an anti-adhesion molecule. The blockade of CD43 enhances the clustering of DCs with T cells and improves the consequent T-cell responses (Fanales-Belasio et al., 1997)

Other markers expressed by skin dendritic cells All DCs express in their nuclei and cytoplasm the protein S-100 (Cochin et al., 1981) that is also seen in other cells with dendritic shape, like melanocytes, Schwan cells and neurons. In humans, LCs and DCs derived from peripheral blood precursors express the molecule CD4, a receptor for the human immunodeficiency virus-1 (HIV-1) (Rappersberger et al., 1988). Cytokine-stimulated human LCs and mature

DCs express the membrane molecule CD83 identified for the first time in a population of mature peripheral blood DCs (Zhou and Tedder, 1995). Although its functional importance in DCs is unknown, it is being used as a maturation marker for human DCs. LCs also have enzymatic markers such as ATPase, nonspecific esterase and endogenous peroxidase. They do not express lysozyme, α1 antitrypsin or α1 antiquimotrypsin, enzymes produced by macrophages (Romani et al., 1991).

ONTOGENY OF SKIN DENDRITIC CELLS As we discussed previously, DCs present in the skin are CD45 leukocytes expressing myeloid lineage-restricted markers such as CD13 and CD33, together with the monocyte/macrophagerelated molecules CD14 (only DDCs and LCs in some skin pathologies), and CD68 (only intracytoplasmic in LCs) (Romani et al., 1991). In 1986, Murphy et al. observed that the repopulation of LCs in human skin from patients undergoing bone marrow transplants (thus devoid of LCs) was dependent on cells bearing a monocyte/macrophage phenotype and containing abundant melanosomes. After colonizing the epidermis, these cells acquired expression of CD1a and BGs, while still expressing monocyte/macrophage markers and having melanosomes. In a third step, and 5 weeks after transplantation, the epidermis of these patients was populated with CD1 LCs with abundant BGs and devoid of melanosomes and macrophage markers. They concluded that LC repopulation of the epidermis was related to the infiltration of the epidermis with cells from the macrophage/phagocytic lineage coming from the underlying dermis. Most of the knowledge that we have now concerning LC/DDC lineage is the result of in vitro studies performed during the last years. In 1992, for the first time two different groups demonstrated that DCs could be generated in vitro from human cord blood CD34 progenitors in the presence of GM-CSF and TNFα

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(Santiago-Schwartz et al., 1992; Caux et al., 1992). Importantly, with this combination of cytokines, DCs differentiated along two different pathways (Caux et al., 1996a). One of these populations, derived from a CD1a, E-cadherin, BGs, cell, resembled LCs in phenotype and function. The other more related to the DDC population derived from a CD14 precursor, expressed CD1a, CD14, the clotting factor XIIIa, CD11b, and CD68, but lacked BGs and E-cadherin. Besides the use of GM-CSF, human myeloid LCs can be obtained from CD34 progenitors in the presence of IL-3 and TNFα (Caux et al., 1996b). The basis of this redundant function of GM-CSF and IL-3 could be explained since both cytokines share the same β-chain of their receptors. In this regard, the myelopoietic effect observed with the use of TNFα could be due to a specific upregulation of the common β chain for the GM-CSF/IL-3 receptor in the CD34 precursor (Caux et al., 1996b). Other growth factors such as stem cell factor (SCF) or Flt3 ligand (Flt3L) have been shown to dramatically increase the yield of DCs obtained in culture, however without inducing any preferential differentiation towards LCs or DDCs (Maurer and Stingl, 1999). Nonetheless, the presence of transforming growth factor β1 (TGFβ1) has been demonstrated to be crucial for the development and homing of LCs. This is confirmed by the observation that LCs can be obtained in vitro from human cord blood CD34 progenitors in the presence of TGFβ1 under serum-free condition (Strobel et al., 1996, 1997), and more recently, in combination with GM-CSF and IL-4, from human peripheral monocytes (Geissmann et al., 1998). The observation that TGFβ/1 mice are devoid of LCs, together with the demonstration that the absence of LCs in these animals is not due to a defect at the progenitor level, clearly demonstrates an important role of TGFβ1 in differentiation and homing of LCs (Borkowski et al., 1996, 1997). It is now clear that LCs can be obtained from CD34 circulating progenitors and that the expression of the skin-homing molecule CLA is necessary to originate cells with LC phenotype

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(Strunk et al., 1996, 1997). Recently a more immediate precursor (CD1 CD11c) present in peripheral blood was also seen to differentiate into LCs when cultured in the presence of GMCSF plus IL-4 and TGFβ1 (Ito et al., 1999). However, the intrinsic mechanisms that induce transendothelial migration of these progenitors or precursors from peripheral blood into the skin remain unknown. In the last few years, DCs have been also derived in vitro from an early lymphoid T cell lineage precursor. This DC of lymphoid origin can be identified by the expression of the surface molecule CD8α (Saunders et al., 1996). In mice, the cytokine Flt3-L, has been shown to increase DCs of both lymphoid and myeloid origin (Maraskosky et al., 1996). However, the presence of CD8α cells has not yet been formally demonstrated in mouse or human skin under physiologic conditions. In this regard it is interesting that the Ikaros mice, lacking all cell populations derived from lymphoid lineage, has a normal number of epidermal LCs (Georgopoulos et al., 1994).

Skin entrance, homing and migration of dendritic cells It is well-established that DCs arrive at peripheral organs and reside in them as immature cell populations, developing a sentinel function. After antigen capture they emigrate towards secondary lymphatic organs to present the processed antigens to specific T lymphocytes (reviewed by Bancherau and Steinman, 1998). During their migration, DCs interact with different cell types, as well as with extracellular proteins. This process, which includes directional migration and modulation of the adhesion molecule expression, is at least partially coordinated by different cytokines. Of particular interest, the chemokines molecules, a subfamily of cytokines with chemotactic effect on leukocytes (Premarck and Schall, 1996), have emerged in the last few years as an important network of molecules related to the directional migration and homing of DCs (reviewed by Sozzani et al., 1999).

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To migrate from peripheral blood into the skin, DCs or their precursors need to attach to endothelial cells and then migrate through the endothelial wall. Some adhesion molecules such as ICAM-2 are expressed constitutively by endothelial cells, but most of the molecules related to leukocyte migration are upregulated by proinflammatory cytokines, such as TNFα and IL-1 (D’Amico et al., 1998). Differential expression of adhesion molecules by skin endothelial cells is also important to control leukocyte migration into the skin. Examples of specialization by skin endothelial cells are the molecules NCAM (CD56) (Mizutani et al., 1994), the receptor for thrombospondin (CD36) (Swerlick et al., 1992) and E-selectin (CD62e) (Picker et al., 1991). E-selectin, which is preferentially expressed by endothelial cells from inflamed skin, is the co-ligand for the molecule CLA present in the membrane of leukocytes that are homing in the skin (Picker et al., 1991). β2 integrins expressed by DCs are involved in the processes of endothelial attachment and also transendothelial migration. The platelet endothelial cell adhesion molecule (PECAM-1) seems to be important during transendothelial migration since blockade of its endothelial coligand CD31, partially inhibits this process (D’Amico et al., 1998). Concerning DC directional migration, the role of the chemokines is of crucial significance. Recent studies demonstrated that the CC chemokines, macrophage inflammatory protein 1α (MIP-1α), MIP-1β and RANTES, that bind CCR1 and CCR5, were able to increase the migration of human DCs in in vitro chemotactic assays up to three-fold (Sozzani et al., 1995, 1997). After entering the skin, DCs will populate the epidermis, as resting LCs, or the dermis as dermal dendrocytes. The specific mechanisms leading to this preferential homing have not been elucidated. As we already described, the cytokine TGFβ1 seems to be of primary importance in determining the presence of epidermal LCs, since the TGFβ/1 mice completely lack epidermal LCs, but not DCs in other locations. Resident LCs remain as a resting immature

cell population until an adequate stimulus induces environmental changes leading to LC activation and migration. Diverse stimuli such as activation of complement factors (Morelli et al., 1996), lipopolysaccharide (LPS) (Roake et al., 1995), sensitizers (Macantonia et al., 1987), ultraviolet B light irradiation (UVB) (Moodycliffe et al., 1992) allogeneic transplantation and skin organ explants (Austyn and Larsen, 1990; Larsen et al., 1990; Morelli et al., 1995), and more recently, genetic immunization, are able to induce LC maturation and migration (Condon et al., 1996; Progador et al., 1998). The proinflammatory cytokines IL-1β and TNFα play a central role in the induction of LC migration and it has been proposed that LCs require both signals for their mobilization. IL-1β binds both IL-1RI and IL-1RII receptors expressed by immature LCs, but only IL-1R1 is functional while IL-1RII acts as a decoy receptor (Cumberbach et al., 1997). TNFα exerts its function by binding to TNFαRII, also expressed by immature LCs (Wang et al.,, 1996). The main source of IL-1α and TNFα is epidermal keratinocytes, while IL-1β is mainly produced by LCs (Heufler et al., 1992). IL-1β is the first cytokine secreted in response to topical allergens and IL-1β mRNA can be detected in LCs as early as 15 min after exposure to sensitizers (Enk et al., 1993). In vivo experiments have shown that local injection of IL-1β induces LC maturation and migration that can be inhibited by general administration of antagonist antibodies (Cumberbach and Kimber, 1995; Cumberbach et al., 1994, 1997). TNFα also has an important role in inducing LC migration. Different studies have demonstrated that intradermal injection of TNFα stimulates epidermal LC migration and their consequent accumulation in afferent lymph nodes (Cumberbach et al., 1994; Cumberbach and Kimber, 1995) and that the administration of TNFα blocking antibodies results in inhibition of the migration induced by LPS, UVB, or contact sensitizers. TNFα also induces a rearrangement at the cytoskeleton level of DCs by inducing the polymerization of F-actin and a lack of vinculin-containing adhesion structures, changes that end in the acquisi-

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tion of high cell motility (Winzler et al., 1997). On the other hand, TNFα, IL-1α and IL-1β induce downregulation of E-cadherin by activated LCs, and as a consequence LCs can be released from neighboring keratinocytes (Schwarzenberger and Udey, 1996). Different mechanisms inducing skin inflammatory responses might produce the same stimuli for LC emigration. LCs released from the epidermis still have to interact with extracellular matrix proteins such as laminin, fibronectin and collagen, present in the epidermal dermal junction or in the underlying dermis. The molecules involved in cell/extracellular matrix interactions are β1 integrins (Ruoslahti and Pierchbacher, 1987). LCs express α4, α5 and α6β1 integrins (LeVarlet et al., 1991), and in vitro studies have demonstrated that LCs attached firmly to laminin and fibronectin, but do not adhere to collagen. The binding of LCs to laminin is mediated by α6 integrins while the interaction with fibronectin depends on the presence of α4 and α5 integrins (LeVarlet et al., 1992). Interestingly the administration of blocking antibodies to α6 integrins completely inhibits the accumulation of migrating DCs in the draining lymph nodes, while no effect was observed with the blockade of α4 integrins (Price et al., 1997). In addition, after exposure of the epidermis to skin sensitizers, LCs produce and secrete matrix metalloproteinase-9 (MMP-9), a type IV collagenase that degrades gelatine, collagen type IV and type V (Kobayashi, 1997). Another molecule that is implicated in extracellular matrix attachment and migration of LCs is CD44, which mediates adhesion to immobilized hyaluronidate. As previously mentioned, immature LCs express the standard form of CD44, and after maturation they upregulate the isoforms v3, v6 and v9. In addition to its adhesion molecule function, CD44s ligation induces monocyte-derived DCs to secrete IL-10, IL-6, and IL-12p70 and the ligation of the CD44 v6 or v9 induces the release of TNFα, IL-8 and IL-1β (Haegel-Kronenberg et al., 1998). After a short trip, human and mouse LCs enter dermal afferent lymph vessels and migrate towards draining lymph nodes (Lukas et al.,

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1996; Weinlich et al., 1998). During their migration, DCs have been seen to acquire some maturation molecules such as CD86, while still conserving some markers of their immature stage like the invariant chain (Welinch et al., 1998). This observation might confirm that DCs start migrating before completing their maturation, and they can undergo a further maturation during their interaction with different molecules such as extracellular proteins, lymphatic endothelial cells or T cells. The entrance and homing of DCs in the paracortical area of local lymph nodes seems to be directed by the chemokines MIP3β and secondary lymphoid tissue chemokine (SLC). Both chemo-attractants are expressed in T cell-rich lymphoid areas of secondary lymphatic organs, and bind to the CCR7 receptor that is upregulated during DC maturation (Dieu et al., 1998; Saeki et al., 1999).

FUNCTIONAL ASPECTS OF LANGERHANS CELLS AND DERMAL DENDRITIC CELLS Maturation and immunostimulatory function Resident LCs and probably resident DDCs have a very efficient antigen uptake function, but they are weak antigen presenters and they are not very efficient in inducing T cell allostimulation (Steinman, 1991). After maturation, both LCs and DDCs become potent immunostimulatory cells able to prime naïve T cells. Several studies have shown that: (1) LCs and DDCs can capture and process protein antigens and present antigenic peptides to naïve T cells (Nestle et al., 1998); (2) after migrating from skin organ explants both cell populations are potent stimulators of allogeneic naïve T cells (Lenz et al., 1993); and (3) LCs and DDCs can efficiently sensitize T cells to haptens in vivo and initiate hypersensitivity reactions (Kurimoto et al., 1994), and LCs can also induce antigen-specific cytotoxic T lymphocyte (CTL) responses (Celluzzi and Falo, 1997).

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Resting LCs accumulate class II molecules into cytoplasmic vesicles, while expressing low membrane levels of them (Mommaas et al., 1995; Cella et al., 1997). After stimulation with proinflammatory cytokines or after the ligation of CD40, an active synthesis of class II molecules begins, and they accumulate in vesicular or multilamellar compartments where they can be loaded with antigenic peptides, and are then efficiently transported to the cell membrane as stable dimers. The upregulation of class II molecules is followed by a downregulation of their synthesis, thus converting the now mature DCs into efficient antigen-presenting, but weak antigen-processing cells (Sallusto et al., 1995). The activation of mouse and human DCs seems to be regulated by the expression and intranuclear translocation of the inducible transcription factor relB, a gene product member of the nuclear factor (NF)-κB family of transcription factors (Rescigno et al., 1998). NF-κB factors play an important role regulating the expression of a number of genes encoding proteins with immune and inflammatory functions. This includes expression of genes encoding cytokines (IL-1α/β, IL-2, IL-3, IL-6, IL-8, IL-12, TNFα), growth factors (GM-CSF, M-CSF and G-CSF), cytokine receptors (CD25), adhesion molecules (ICAM-1, VCAM-1, E-selectin and Mad-CaM), stress proteins, complement factors and immunoregulatory molecules, such as MHCclass I, class II and β1 microglobulin. Potent inducers of NF-κB are bacterial cell membranes, LPS, virus, proapoptotic and necrotic stimuli such as the release of oxygen free radicals, IL-1, TNFα and UVB and γ irradiation (reviewed by May and Gosh, 1998). Immature LCs synthesize neither relB protein nor mRNA, however both can be found in both mouse and human mature DCs (Carrasco et al., 1993; Clark et al., 1999). In mice, the disruption of the relB gene results in an impaired antigenpresenting cell function and absence of DCs in secondary lymphoid organs, but a normal number of epidermal LCs (Burkly et al., 1995). Mature immune competent DCs express high levels of MHC-class I and class II molecules, as well as adhesion and co-stimulatory molecules

such as CD54, CD58, CD80, CD86 and CD40 necessary for DC/TC dialogue and they produce and secrete both subunits of IL-12, (p35 and p40) ILβ-1, and IL-6. As a result specific populations of CD4 T helper cells, and/or CTLs will be induced. Consequently, the delivery of antigens to DCs in situ is a rational approach for the development of immunization approaches. The unique stimulatory function of LCs or DDCs and their ideal situation as cellular elements of the skin immune system, make the skin a suitable organ for the development of immunomodulatory strategies for the prevention or treatment of tumors or infectious diseases (reviewed by Tüting et al., 1998). During the last few years, genetic immunization using naked DNA or viral vectors has emerged as an important tool to engineer skin dendritic cells in situ. Topical application of naked DNA (Fan et al., 1999) or adenoviral vectors (Tang et al., 1997; Yu et al., 1999), as well as biolistic delivery (Figure 23.1), (Condon et al., 1996; Progador et al., 1998) or intradermal injections of naked DNA (Torres et al., 1997; Akbari et al., 1999) encoding viral or tumor proteins have resulted in the activation and induction of protective immune responses specific to the encoded antigen. Moreover the presence of bacterial stimulatory sequences present in plasmid backbones, such as CpG motifs, provides the adjuvant effect necessary to induce an effective immune response (Pisetsky, et al., 1996; Tighe et al., 1998; Jakob et al., 1998; Hartman et al., 1999).

Skin dendritic cells and downregulation of the immune response While the immune stimulatory function of skinderived DCs has been broadly investigated, we know less regarding their role in mechanisms leading to the downregulation of the immune response. In the skin, the phenotype and state of activation of resident LCs and DDCs are determined at least in part by different cytokines, growth factors, hormones and peptides,

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FIGURE 23.1 Electron microscopy of human skin dendritic cells after biolistic delivery of naked DNA. (A) A human dendritic cell from the dermal microvascular unit, containing 1 µm gold particle near the nucleus (arrow). (B) Higher magnification of the dermal dendritic cells shown in A. (C) A dendritic cell migrated from human epidermal dermal explants after biolistic gene delivery, also shows an intracytoplasmic gold particle (arrow). DC, dendritic cell; EC, endothelial cell; MC, mast cell; M, macrophage.

released by neighboring cells and nerves under different conditions. Some factors can induce the activation of resting DCs, while others can maintain their immature state or inhibit their migration. Amongst the latter, TGF-β1 that is constitutively released by keratinocytes and immature LCs have been shown to prevent the maturation of human LCs induced by LPS, TNFα, or IL-1β (Geissman et al., 1999). Low amounts of IL-10 are spontaneously produced by keratinocytes in normal skin and its secretion is dramatically augmented after a UVB irradiation dose that can abrogate the response to skin sensitizers (Niizeki and Streiling, 1997). Moreover IL-10 is produced by some progressive melanomas, and it has been postulated that this can be a mechanism of melanoma immune evasion (Enk et al., 1997). IL-4, another TH2

cytokine inhibits in vitro the migration of human epidermal LCs, through the downregulation of TNFR II (Takayama et al., 1999). Other factors and peptides that are related to negative regulation of the immune response are released by terminal skin nerves. These include calcitonin gene-related peptide (CGRP) and proopiomelanocortin (POMC), a precursor that can be cleaved into several bioactive neuropeptides. Two POMC-derived peptides are α melanocyte stimulating hormone (α-MSH) and adrenocorticotropic hormone (ACTH) (Schauer et al., 1994). Nerve fibers expressing CGRP are situated very close to LCs and purified CGRP inhibits the antigen-presenting function of cultured LCs (Hosoi et al., 1993). α-MSH that is produced by cebocytes of the pilosebaceous unit (Böhm and

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Luger, 1998) antagonizes the pro-inflammatory effect of IL-1, and TNFα inhibits the production of IFNγ by human lymphocytes and induces the secretion of IL-10 (Bhardwaj et al., 1996). α-MSH, is able to transform immature DCs into tolerizing DCs capable of inducing regulatory T cells (Grabbe et al., 1996; Groux et al., 1997). The engineering of DCs capable of inducing anergy or tolerance instead of stimulation will be of significant relevance for the prevention or treatment of transplant rejection and autoimmune diseases.

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24 Dendritic cells in the respiratory tract Laurent P. Nicod and L. Cochand Hôpitaux Universitaires de Genève, Switzerland

You might as well question why we breathe. If we stop breathing, we’ll die. If we stop fighting our enemies, the world will die. Victor Laszlo, in Casablanca

INTRODUCTION

mediators potentially able to either decrease inflammation or to activate immature DCs in the vicinity and favor a TH1 or TH2 response.

The airways and the alveoli constitute a large surface continuously exposed to organic and inorganic particles, as well as to viruses or bacteria. The mucociliary clearance allows the removal of many potential pathogenic antigens impacted along the airways. If the epithelial surfaces are, however, disrupted, a local network of dendritic cells extending from the basement membrane up to the tight junctions between the apical side of the epithelial cells, is ready to take up antigens and to migrate towards the lymph nodes to trigger T cells in the central immune system. Dendritic cells are also found around the vessels and in small numbers in the alveolar walls. In contrast, the alveoli contain mostly alveolar macrophages differentiated in phagocytic cells usually devoid of surface markers required for giving second signals during antigen presentation. They can, however, depending on the dangers encountered, release either anti-inflammatory cytokines or inflammatory Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

DISTRIBUTION OF PULMONARY DENDRITIC CELLS When we first stained normal human lung for HLA-DR-positive cells, alveolar macrophages, unlike murine macrophages were brightly stained. HLA-DR-positive cells were also seen in the interstitium of the lung. Many had a mature macrophage appearance, but others displayed elongated processes similar to those demonstrated by DCs in lymphoid organs. The majority of these DCs were found in the bronchial mucosa either above or below the basement membrane (Plate 24.1) or deep in the lamina propria of the bronchia. In the mucosa below the epithelium, occasional aggregates of lymphocytes of the bronchus-associated

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lymphoid tissue (BALT) were visualized and were associated with DCs. Many DCs were found in the perivascular loose connective tissue and in the pleura (Nicod et al., 1986) (Plate 24.2). These findings were confirmed not only in humans, but also in mice (Sertl et al., 1986). Shortly afterwards, DCs were shown to constitute a tight network around the airways. This was well shown by Patrick Holt’s group, as they performed tangential sections of the airways. They thus demonstrated that DC processes formed a network like Langerhans cells do in the skin (Shon-Hegrad et al., 1991), but the majority of these cells do not express CD1a (OKT6) as do Langerhans cells.

IMMATURE PHENOTYPE OF LUNG DENDRITIC CELLS DCs are widely distributed in the blood, the skin and in most tissues, especially in lymphoid organs, where they display an important phenotypic diversity, the significance of which remains unclear (Hart, 1997). The difficulty in obtaining a large number of DCs from human tissues has been palliated by the recent development of methods to generate DCs in vitro. The methods used differ in two aspects: (1) the use of cord blood precursors or adult blood monocytes (Mo); and (2) the use of different cytokines. Romani and associates (Romani et al., 1996) have derived DCs from human Mo. Mo were cultured for 6 to 7 days in GM-CSF and IL-4 to obtain ‘immature’ DCs. Immature DCs were characterized by a high endocytic activity that can be measured by fluorescein isothiocyanate dextran incorporation (Sallusto et al., 1995), but a low capacity to stimulate T cells. When challenged by inflammatory stimuli such as TNFα, lipopolysaccharide (LPS), or monocyte-conditioned medium (Romani et al., 1996; Reddy et al., 1997), these cells become mature DCs. They lose their capacity to incorporate FITC-dextran, but become strong T-cell stimulators while expressing high levels of CD80, CD40, CD86 and CD83. This latter (CD83) is a specific cell-surface marker of mature DCs whose function is unknown (Zhou and Tedder, 1995).

DCs isolated from human lung have a phenotype similar to freshly isolated blood DCs (Scheinecker et al., 1998). They express high levels of LFA3 and MHC class II molecules (Nicod and El Habre, 1992) and low levels of CD14 molecules. The α chain of the β2 integrins (CD11c) is known to be expressed on mouse spleen DCs more than on any leukocytes, however in human lung, CD11c is expressed to the same extent as it is on monocytes. The same appears true for human blood DCs as compared with monocytes (Freudenthal and Steinman, 1990). Lung DCs have a weak CD83 expression (Cochand et al., 1999), like immature DCs (Reddy et al., 1997). CD1a, a characteristic cellsurface molecule present on epidermal LCs (Péguet-Navarro et al., 1995), was shown to be present on blood-derived DCs as reported by others (Scheinecker et al., 1998). However, CD1a was consistently absent on lung DCs (Cochand et al., 1999). Lung DCs, like immature DCs, show only limited expression of CD40, CD80 and CD86 (Cochand et al., 1999), whereas human epidermal DCs (Rattis et al., 1996) and murine spleen or skin DCs (Inaba et al., 1994) do express significant levels of CD86, but no CD80. Cryostat sections of mice tissues (Inaba et al., 1994) confirm the low expression of CD80 and CD86. Although the expression of B7 molecules can be upregulated by 1 to 3 days in vitro culture without intentional stimulation (Inaba et al., 1994), our results and others show that lung DCs are in a so-called immature phenotype, (Table 24.1). The capacity of DCs to uptake and process antigens is highly dependent on the stage of differentiation of DCs. Immature DCs have the capacity to phagocytose particles and microbes (Henderson et al., 1997). To do so they express receptors that mediate absorptive endocytosis in particular the mannose as well as Fcγ and Fcε receptors (Sallusto et al., 1995; Fanger et al., 1996; Engering et al., 1997). Inflammatory stimuli result in loss of capturing machinery and in an increase of T-cell stimulatory function. Lung DCs have an ability to endocytose dextran, which is inbetween immature and mature

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TABLE 24.1 Cell surface phenotype of monocytes (Mo), immature DC (iDC), mature DC (mDC) or lung DC Markers

Mo

CD14  CD83  CD40  CD80  CD86  CCR1  CCR5  MHC class I/II  Mannose R 

iDC

mDC

Lung DC

        

    

        

 

blood-derived DCs. Lung DCs uptake dextran and the number of molecules internalized as judged by the mean fluorescence intensity, is rather high. Lung DCs, like immature blood-derived DCs express both CCR1 and CCR5 (Cochand et al., 1999), whereas mature DCs migrating towards lymphoid structure no longer express these two receptors, but are likely to acquire the CCR7 described in other systems (Sallusto et al., 1998).

LUNG DENDRITIC ORIGIN AND FATE Evidence accumulates in favor of DCs being derived from peripheral blood monocytes or partially differentiated blood-derived dendritic cells, recruited into tissues, via yet undefined mechanisms. DC populations in peripheral tissues such as skin (Chen et al., 1986) and muscle (Leszcynski et al., 1985) turn over relatively slowly, exhibiting a half-life of 30 days. Respiratory tract DCs could have a more rapid turnover at the mucosal surfaces where they are in close contact with inhaled particles or organisms. The origin and turnover of respiratory tract DCs have been studied in normal rats employing a radiation chimera model (Holt et al., 1994). In this model, the supply of precursor DCs from bone marrow is interrupted by X-irradiation of animals while shielding the thoracic cavity to

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prevent local tissue damage, followed by transplantation of congenic bone marrow; host/ donor DCs can be discriminated in frozen tissue sections via appropriate pairs of monoclonal antibodies. Given the rate of decline of the host DCs post irradiation and pre-engraftment of congenic bone marrow, and the subsequent rate of replenishment of the tissue population with DCs, a close approximation of population halflife was estimated at 2–3 days for intraepithelial DC, 7–10 days for parenchymal DCs and 21 days for skin DCs (Holt et al., 1994). The high turnover rate of bronchial DCs would then be close to those of the gut, the half-life of which has been estimated at 3 days (Fossum, 1989). The most striking responses have been documented following challenge of rats with aerosols containing bacterial lipopolysaccharides (ShonHegrad et al., 1991), or heat-killed bacteria (McWilliam et al., 1994). In these models, DC influx occurred within 20 minutes of the end of challenge, and peaked within 2 hours, at which time local airway intra-epithelial DC numbers are up to 2.5 times baseline levels. The enhanced density of DCs remains for a further 24– 48 hours, prior to migration to regional lymph nodes (McWilliam et al., 1994). Similar DC recruitment has been observed following live virus infection of the tracheal epithelium, commencing concomitantly with the first expression of viral nucleoproteins within infected airway epithelial cells, and also within 2–6 hours of challenge of immune animals with an aerosol containing a soluble recall antigen (McWilliam et al., 1997). After antigen uptake, airway DCs migrate to the paracortical T-cell zone of the draining lymph nodes of the lung, where they interact with naïve T cells (Xia et al., 1995). When labelling DCs with green fluorescent protein (GFP), GFP-labelled DCs injected in the trachea of mice, were traced in the draining mediastinal lymph nodes 24 hours after injection, but not in non-draining lymph nodes or spleen (Havenith et al., 1993). By 48 hours after injection, labelled DCs had disappeared from the draining lymph nodes, in agreement with data from subcutaneous injection of DCs (Ingulli et al., 1997). Their

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disappearance being perhaps the result of apoptosis after interaction with antigen-specific T cells. Activated dividing T cells can be found after a few days at distance from the hilar nodes (B. Lambrecht et al., 2000). The migration of activated DCs and later of memory T and B cells is shown in Figure 24.3.

DENDRITIC CELLS AND T CELL ACTIVATION DCs have the unique capacity to induce primary immune responses. DCs are potent stimulators of a primary MLR in both experimental animals and in humans (Steinman et al., 1983). They are very effective in stimulating T cells to help B cells develop into plasma cells (Inaba and Steinman, 1985). DCs also appear to be the most potent stimulators of CD8 cytotoxic T lymphocytes (CTLs) (Inaba et al., 1987). Both antigen-specific CTLs, as well as antigen nonspecific natural killer cells can be elicited by DCs (Young and Steinman, 1990). DCs can prime an immune

response in vivo even in the absence of adjuvants (Inaba et al., 1990). However studies on antigen presentation by lung DCs are still scarce. DCs purified from human lung support T-cell response to both exogenous antigens (Nicod et al., 1989) and alloantigen (Nicod and El Habre, 1992; Van Haarst et al., 1996). When tested in parallel with endogenous lung macrophages, human lung DCs are considerably more potent for T-cell activation (Cochand et al., 1999). The crucial role of lung DCs, in vivo, for the presentation of inhaled antigen to previously activated or memory T cells has been shown lately. B.N. Lambrecht et al. after selective depletion of lung DCs showed that dendritic cells are required for the development of chronic eosinophilic airway inflammation in response to ovalbumine (Lambrecht et al., 1998). The priming of T cells by nasal or lung DCs in vivo is being better understood (Vermaelen et al., 2001). It is likely that airway macrophages and the ‘alveolar milieu’ profoundly modulate the priming of T cells by lung DCs.

FIGURE 24.3 Migration of bronchial dendritic cells (DCs). DCs migrate to interbronchial lymph nodes to initiate an immune response. If most alveolar macrophages are eliminated along the airways, some may also migrate to lymph nodes. Memory T cells primed from the BALT or GALT can recirculate to the mucosal surfaces to be activated locally. DENDRITIC CELLS IN THE PERIPHERY

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ADHESION MOLECULES AND DC – T CELLS INTERACTION

ADHESION MOLECULES AND DC–T CELLS INTERACTION When T cells are cultured in the presence of lung DCs, typical cell aggregates or clusters become apparent (Flechner et al., 1988; Nicod et al., 1990). At this critical stage, the physical association of the two cells seems to be due to an antigen-independent mechanism (Inaba et al., 1989). Subsequently, the specific regulation by the TCR of MHC molecules displaying antigenic peptides is facilitated. Studies of the molecular basis of the interaction between T cells and DCs have shown that adhesion-dependent interactions are partially mediated through binding of integrins. The β1integrins family includes receptors that bind to extracellular matrix. Antibodies to β1-integrins reduce DC-induced alloreactions to the same extent as monoclonal antibodies against β2integrins do, with a synergistic effect when antibodies to both integrins are combined (Nicod and El Habre, 1992). Elevated levels of VLA-5 are found on both DCs and monocytes, and the mean fluorescence intensity is only slightly increased on DCs compared with monocytes. VLA-4 is only weakly present on lung DCs and its ligand, vascular cell adhesion molecule (VCAM), is not detected (Nicod and El Habre, 1992). Among the β2-integrins, LFA-1 (CD11a/ CD18) is decreased on human lung DCs as compared with blood monocytes, however, its ligand, ICAM-1, is slightly more expressed on lung and blood DCs as compared with monocytes (Freudenthal and Steinman, 1990). In blocking experiments, antibodies against LFA-1 or ICAM-1 only partially decrease T-cell proliferation (Nicod and El Habre, 1992). The low efficacy of these antibodies in blocking an alloreaction could be partly related to the presence of ICAM-2 and/or ICAM-3 on DCs. However, the β2 integrins are important in DC–T cell interactions, because monoclonal antibodies against the common β2 chain or the α chain of CD11c (p150, 95) markedly decreases Tcell proliferation. The α chain of CD11c is more expressed on DCs than are CD11a or CD11b. In mouse spleen, DCs express the highest levels of

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CD11c (Steinman, 1991) of any leukocyte. In human lung, however, CD11c is expressed to the same extent as it is on monocytes. The same appears true for human blood DCs as compared with monocytes (Freudenthal and Steinman, 1990). LFA-3 is highly expressed on peripheral blood and lung DCs compared with monocytes (Freudenthal and Steinman, 1990). CD2 is absent from DCs. LFA-3 which binds to T cell CD2 participates in the binding of T cells and DCs, stabilizing cell clusters (King and Katz, 1989; Hauss et al., 1995) and participates in T cell activation via an interaction with the TCR–CD3 complex (Breitmeyer et al., 1987). A profound inhibition is observed when antibodies against either LFA-3 or CD2 are used during DC–T cell interactions. The maturational program in DCs is accompanied by increased expression of the co-stimulatory B7-1/CD80 and B7-2/CD86 molecules interacting with CD28 on T cells. In the absence of B7-CD28 crosslinking of the APC, MHC and the T cell receptor lead to T cell anergy (Doherty, 1995). Both CD28 and the closely related CTLA-4 molecule bind members of the B7 family. Interaction of CTLA-4 may mediate a negative signal to dampen the immune response (Gribben et al., 1995). Although lung DCs have an immature phenotype with low CD80 and CD86 expression, even after the isolation procedure, lung DCs shared a strong capacity, like mature DCs to stimulate T-cell proliferation. The role of CD80 and CD86 on lung DCs was demonstrated by using blocking antibodies (Cochand et al., 1999). These results are in agreement with Scheinecker and associates (Scheinecker et al., 1998). However, the only partial inhibition obtained when using the potent inhibitor CTLA-4, which blocked both CD80 and CD86 in our experiments, could suggest the presence of other co-stimulatory signals. In vitro, the ligation of CD40–CD40L can increase the expression of CD80 and CD86 and also the production of IL-12. These effects lead to a maturation of immature DCs and to a stronger T-cell stimulation (Cella et al., 1996). A

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similar function of CD40 has been shown on human epidermal cell (Sallusto et al., 1998) and on mouse lung DC (Masten et al., 1997). Surprisingly, the same CD40mAb clone inhibiting the MLR induced by human epidermal DC (Sallusto et al., 1998) had no effect on the MLR induced by human lung DCs. This could imply that enough co-stimulatory signals were already present when the experiments were performed, which might be induced by the culture conditions (Holt et al., 1993).

THE ROLE OF CYTOKINE RELEASED BY DCs IN ACTIVATING T CELLS Cytokines such as IL-1α or IL-1β (Dinarello, 1988), IL-6 (Wong and Clark, 1988), or TNFα (Yokota et al., 1988) have all been described as providing T cell co-stimulatory signals. Although powerful APCs, lung DCs like blood-derived DCs produce little or no IL-1 (McKenzie et al., 1989; Nicod et al., 1990). Using a sensitive enzyme immunoassay for IL-1α and IL-1β, lung DCs demonstrated far less surface-bound and secreted IL-1 than monocytes. Furthermore, IL-1 was produced in relatively small quantities during a DC-induced MLR as compared with monocytes. In addition, blocking antibodies to either IL-1α or IL-1β did not interfere with human lung DC-induced T cell proliferation. However, in contrast to the findings with DCs, the MLR induced by lung AMs was inhibited by blocking antibodies to IL-1 (Nicod et al., 1990). TNF-α was also shown to enhance anti-CD3 activated T-cell proliferation, as well as the response of T cells to antigen-bearing paraformaldehyde-fixed APCs (Plaetinck et al., 1990). The receptor involved in murine as well as human thymocyte stimulation appeared to be the TNFsR-75 (Scheurich et al., 1987; Tartaglia et al., 1991). TNFα induces IL-2 receptor and gene regulatory factors, such as c-fos or NFκB in T lymphocytes (Tartaglia et al., 1993; Grilli et al., 1993). For human lung DCs, TNFα production is unlikely completely to explain why these cells are such potent APCs, because

LPS-stimulated DCs produce less TNFα than do AMs (Nicod et al., 1990). However, using high doses of a TNF inhibitor, a fusion protein recombinant TNFsR-p55Hγ3, we were able to reduce T-cell proliferation and TH1 and TH2 cytokine production induced by DCs, monocytes or antiCD3 antibodies. This study supports the importance of TNFα as a co-stimulatory factor in all of these situations (Nicod et al., 1996). Human peripheral blood DCs appear to be poor producers of IL-6 (Vakkila et al., 1990). Blocking antibodies against IL-6 does not decrease a DC-induced MLR, suggesting that IL-6 plays little role during alloreaction (Nicod et al., 1990). IL-6 has been claimed to contribute to the early generation of TH2 cells by an IL-4dependent mechanism (Rincon et al., 1997). However IL-6-deficient mice have normal TH2 development. The role of IL-6 on T helper determination in lung has still to be revealed. IL-12 is the principal TH1-inducing cytokine and acts by inducing signal transduction and activator of transcription 4 (STAT 4) in undifferentiated TH0 cells. Dendritic cells constitutively produce IL-12 (Macatonia et al., 1995) and its production is further enhanced by microbial stimulation. Activated macrophages and NK cells can also be an early source of IL-12 (Hsieh et al., 1993; Isler and Nicod, 2000). Recently, IL-18 (interferon gamma inducing factor, IGIF) has been cloned and shown to cooperate with IL-12 to induce TH1 development by maintaining the expression of the IL-12Rβ2 chain on developing TH1 cells (Xu et al., 1998). Human DCs produce constitutively pro-IL-18 (personal data) which needs to be cleaved by caspase 1 to become active. The release of active IL-18 by murine or human DCs has been shown recently (Stoll et al., 1998). PGE2 and IL-10 are produced in response to a variety of stimuli by epithelial resident cells or alveolar macrophages, and can reduce the production of biologically active IL-12 in DCs (Koch et al., 1996; Kalinski et al., 1997; Isler et al., 1998). DCs grown in the presence of these factors induce differentiation of naïve T helper cells into the TH2 phenotype. Thus the microenvironment in which the DCs reside during antigen

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uptake, might influence the production of IL-12 and the prolongation of T helper responses induced by DCs.

ALVEOLAR MACROPHAGES AND DENDRITIC CELLS AMs constitute 85% of the cells in the alveoli, whereas DCs account for no more than 1% of the cells in the lung compartment. AMs phagocytose and neutralize foreign organisms and particles that reach distal airways and produce oxygen radicals, cytokines, lyzozyme, complement and many other substances. If they are good phagocytic cells, they are poor APCs and may even suppress T-cell proliferation. A defect in accessory molecules such as CD80 and CD86 has been shown in normal subjects (Chelen et al., 1995). The release of inhibitory substances like IL-10 (Isler et al., 1998) or PGE2 has been demonstrated (Metzger et al., 1980). In contrast

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when AM are exposed to bacterial products such as lipopolysaccharides, AM release higher levels of TNFα compared to peripheral monocytes, whereas monocytes released higher levels of IL-10 compared to AM (Boehringer et al., 1999). This demonstrates the higher inflammatory properties of AM which may influence theoretically the phenotype and maturation stage of DCs in their vicinity. The type of pathogens or stimuli encountered by AM may be important for the ensuing immunity. Indeed IL10 and TNFα production can be dissociated depending on the signalling pathways triggered, such as tyrosine kinases or phosphatases (Boehringer et al., 1999). This is of particular interest since the production of IL-12 by AM themselves was found to be tightly regulated by the autocrine production of IL-10 (Isler et al., 1998). In Figure 24.4, we show how AM could influence neighboring DC activation and how TH1/TH2 differentiation could be influenced when memory T cells migrate to the vicinity of

FIGURE 24.4 Interaction of macrophages (MØ) and dendritic cells (DC). Macrophages which are the major population in the alveoli can release mediators when encountering foreign particles (innate immunity), which can modulate the specific immunity by releasing cytokines, which can activate immature DC or influence the T helper pattern of reactivated memory cells. Interaction of macrophages with epithelial or mesenchymal cells are omitDENDRITIC CELLS IN THE PERIPHERY

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interstitial DCs while macrophages release either IL-10 or IL-12. The role of epithelial or mesenchimal cells is omitted for the sake of clarity. In some pathological states such as sarcoidosis, AM express high levels of CD86, CD40 and CD30L like mature DCs would do (Nicod and Isler, 1997). These changes do occur under yet quite unknown stimulatory factors. Indeed we have been unable to stimulate these changes with a variety of cytokines in vitro. However it appears clear that AM can express markers thought to be rather specific for dendritic cells, despite their so-called end-stage differentiation state. Similar changes have been found in mice when AM had been exposed to bleomycin (Tager et al., 1999). In histiocytosis X there is an accumulation of mature dendritic cells in the lung with features of Langerhans cells. Cytokines such as GM-CSF, TNFα and IL-1α could play a major role in the local environment where these Langerhans cells migrate or proliferate (Tazi et al., 1999). These macrophages having acquired DC phenotypes or DC/Langerhans cells attracted by the local conditions may profoundly influence local T cell immunity in pathological states, interfering with gas exchanges, the prime function of the lungs. In normal states, AM being poor APC or having even an immunosuppressive role (Metzger et al., 1980), protect the air spaces from deleterious inflammatory processes.

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PLATE 24.1 Dendritic cells above and below the basement membrane of the human airways. The basement membrane below the mucosa (arrows) and above the lamina propria is shown by a star in the frozen section seen by transmission electro-microscopy (A) The HLA-DR stained DC with their elongated processes are pointed out on the same section seen by fluorescence.

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PLATE 24.2 Dendritic cells around human lung vessels. HLA-DR DCs are stained by immunoperoxidase and found in the loose connective tissue around the vessels. DCs are shown with arrowheads and the lumen of the vessel with the star.

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25 Intestinal dendritic cells G. Gordon MacPherson, Fang-Ping Huang and Li Ming Liu Sir William Dunn School of Pathology, University of Oxford, UK

What was in his mind, I don’t know. I only know what was in his stomach. In Search of the Castaways Jacques Paqarel

INTRODUCTION: DENDRITIC CELLS AND INTESTINAL IMMUNE RESPONSES

systemic responses to foreign antigen however, interaction of the intestinal (or other mucosal) specific immune system with antigen may lead to active tolerance. Intestinal immune responses may even be dichotomous, in that oral antigen can induce a local secretory IgA response, but animals (and perhaps humans) may be hyporesponsive to the same antigen when challenged systemically following oral administration (oral tolerance). The cellular and molecular mechanisms underlying oral tolerance are complex and controversial (reviewed in Kagnoff, 1996; Garside and Mowat, 1997; Strobel and Mowat, 1998) but there is increasing evidence that dendritic cells (DCs) may play a central role (Viney et al., 1998), (see below). Immune responses to antigen in the intestinal lumen are initiated at two or possibly three sites, Peyer’s patches (PP), mesenteric lymph nodes (MLN) and possibly the lamina propria of intestinal villi (LP) (see Abreu Martin and Targan, 1996; Mayer, 1997; Mowat and Viney, 1997; Brandtzaeg, 1998; Curr. Topics Microbiol.

The gastro-intestinal tract faces a continual immunological onslaught from antigens (Ag) in food, commensal bacteria, and on occasion, potentially pathogenic bacteria, viruses, parasites and perhaps prions. Yet, for the most part, we are totally unaware of the immune responses that are occurring all the time in the gut. In other sites, such responses would inevitably lead to local inflammation, but in the gut, this outcome is happily a rare event. It does, of course, occur at times and food allergies and many chronic inflammatory bowel diseases represent an inappropriate response to foreign or self antigen. Intestinal immune responses, as with responses elsewhere, may be active but silent (subclinical), as evidenced by the frequent clinical infections with ‘non-pathogenic’ microorganisms seen in patients and animals with immunodeficiencies. In contrast to most Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Immunol. vol. 236, 1999, for reviews of intestinal immunity). The initial event in activation of a naïve T cell is presentation of antigen by DC (Banchereau and Steinman, 1998), but it now seems probable that DCs have more subtle regulatory roles in T cell activation and that they may give non-Agspecific signals that affect or determine the differentiation of the T cell. Thus, in designing mucosal vaccines, in developing immunotherapy for tumours or inflammatory/autoimmune diseases, a clear understanding of intestinal DC properties and functions is essential. We are however, only at the beginning of such understanding. Study of intestinal DCs is very limited, largely due to the difficulty of extracting DCs from tissues, and, in humans, the difficulty of obtaining intestinal tissues other than the colon and rectum. Considerably more is known about the properties of DCs in PP and MLN than of those in LP, and in humans, information is only available in any detail about colonic DCs. In this chapter we will consider DCs present in PP, LP and MLN in terms of their phenotypes, properties and functions in the initiation and regulation of immune responses to enteric antigen. Inevitably, we will raise more questions than we have answers, but perhaps we will stimulate increased efforts to unravel the functions of DCs in these critical sites.

IN SITU IDENTIFICATION OF INTESTINAL DENDRITIC CELLS It is crucial to the understanding of many intestinal disorders to be able to identify DCs by immunohistological criteria in normal and pathological specimens There are however, major difficulties with the identification of DCs by these criteria alone. In situ, DCs can be tentatively identified by their irregular morphology in conjunction with immunostaining for MHC class II and other surface markers that are more or less specific for DCs (it should be noted that there do not seem to be any monoclonal

Abs (mAb) that recognise all DCs and only DCs). Expression of MHC class II and an irregular morphology are not however sufficient for DC identification as many macrophages, particularly in inflamed tissues, may express MHC class II. Other markers such as CD83 (Zhou and Tedder, 1995) and fascin (Mosialos et al., 1996), and those identified by the Hart group (Hock et al., 1994; Green et al., 1998) have not been used widely as yet. Given these provisos, DCs appear to be relatively abundant in the gut. The majority of DCs lie in the LP underlying the epithelium, but in the small intestine of the rat, lined by columnar epithelium, there is evidence for a population of DCs that resides above the basement membrane, and there is also much evidence that DCs below the basement membrane can extend processes up between epithelial cells (Maric et al., 1996). It is not clear from in situ studies whether phenotypic or functional differences exist between DCs in these different sites. PP are the major portals by which antigen in the intestinal lumen is made available to the cells of the immune system. Antigen is transported across the epithelium by specialised epithelial cells, M cells, that possess relatively few microvilli and are capable of transcytosis of soluble molecules and particles. A number of pathogens have ‘hijacked’ this route to bypass the epithelial barrier (see Wolf and Bye, 1984; Gebert et al. 1996; Semin. Immunol. vol. 11, 1999 for reviews of M cells). DCs have been described at two sites in PP, the T cell areas, in which DCs are thought to correspond to IDC in lymph nodes, and in the subepithelial area underlying the dome. This is the region into which antigen is delivered by M cells, and it is an attractive hypothesis that DCs in this area capture antigen and transport it to T areas for presentation to recirculating T cells. In the mouse, DCs in the two sites differ in their expression of surface markers. Subdome DCs are negative for DEC-205 but express CD11c, whereas T area DCs express both markers (Kelsall and Strober, 1996; reviewed in Iwasaki and Kelsall, 1999b). Recently Iwasaki and Kelsall (personal communication) have shown by

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immunofluorescence that subepithelial dome DCs express markers typical of myeloid DCs whereas those in T cell areas express CD8α/α, and are negative for CD11b typical of lymphoid DCs. They have also identified a third population of DCs, CD11c, CD11b, DEC-205, present in both the subdome and T cell areas.

LIFE HISTORY AND MIGRATORY PROPERTIES OF GUT DCs Steady state Recruitment Little is known about the recruitment of DC precursors to tissues. However, it is clear that expression of chemokine receptors by DCs is a regulated phenomenon. Caux’s group (Dieu et al., 1998) have shown that immature CD34 precursor-derived human DCs respond via CCR6 to MIP-3α, but following maturation, lose responsiveness to MIP-3α, and downregulate CCR6, while become responsive to MIP-3β. In the tonsil, MIP-3α, is expressed in follicle-associated epithelium, whereas MIP-3β is expressed in T-cell areas. This suggests that DC maturation is associated with modulations of chemokine receptor expression that would direct immature DCs to peripheral areas, but on receipt of maturation signals they would be targeted to T cell areas. Kelsall’s group however, (Iwasaki and Kelsall, personal communication) suggest that in murine PP, the epithelial subdome DCs and T-cell area DCs represent distinct myeloid and lymphoid lineages, respectively. They find that isolated subdome DCs express CCR6 and migrate towards MIP-3α, whereas all PP DCs express CCR7 and respond to MIP-3β and SLC. Interestingly, they find that splenic myeloid DCs, although expressing CCR6 do not migrate towards MIP-3α. They suggest that precursors of both PP DC subsets enter PP following recognition of SLC secreted by high endothelial venules and that myeloid DC responsiveness to MIP-3α is induced by TGF-β, present at high levels in PP.

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Exit The ability of DCs to act as APC depends on their migrating to T-cell areas of secondary lymphoid tissues (draining lymph nodes or PP). We have studied the migration and turnover of DCs in the normal rat small intestine (Pugh et al., 1983). Dividing DC precursors in the bone marrow can be labelled by i.v. tritiated thymidine (3H-TdR) or bromodeoxyuridine (BrdU). As there is no evidence for significant DC division in the intestine, the minimal time taken for labelled DCs to appear in pseudo-afferent intestinal lymph represents the time from the last division in the marrow and includes the time taken to traverse the intestine. Labelled DCs appear in lymph within 48 hours, with peak numbers arriving at 3–4 days. Thus DCs spend a minimum of 48 hours and a modal time of 3–4 days in the intestine before migrating to the nodes. We cannot accurately estimate the maximum time in the intestine because the rate of decline of labelled DC appearance in lymph is affected both by input into the gut of cells that were labelled early in their differentiation in marrow and those that have spent longer periods in the gut. These data show that intestinal DCs turn over much more rapidly than Langerhans cells, but with similar kinetics to murine splenic DCs (Steinman et al., 1974) and rat lymph node DC (Fossum, 1989). In the respiratory tract of rats, a different approach, measuring the kinetics of DC reconstitution after depletion by corticosteroids or irradiation, suggests that the average half-life of DCs is about 2 days (Holt et al., 1994). Thus it appears that in the absence of any known stimulation, DCs spend only short times in mucosae before migrating to draining nodes. Models for DC populations in the intestine are shown in Figure 25.1.

Stimulated migration Recruitment Inflammatory stimuli cause a rapid influx of DCs and/or DC precursors into some mucosal sites. Thus Holt’s group has shown that large numbers

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FIGURE 25.1 tissues.

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Diagram of dendritic cell populations identified in rodent intestine and associated lymphoid

of DC precursors enter respiratory epithelium within hours of challenge with bacterial, viral or protein antigen. These cells spend up to 48 hours in the epithelium and then migrate to the draining nodes (McWilliam et al., 1996). Interestingly, similar stimuli did not result in DC accumulation in the peritoneal cavity or epidermis. It is not known whether similar recruitment occurs in intestinal mucosae. Exit A variety of nonspecific stimuli can affect DC migration dramatically, possibly acting via a final common pathway involving TNF-α and IL-1. I.v. endotoxin induces a rapid increase in the numbers of DCs migrating in lymph from

the intestine. This effect occurs within 6 hours, peaks at 12–18 hours and is over by 48 hours and results in an approximately 10-fold increase in the numbers of DCs that can be collected over that period. The source of the migrating DCs appears to be the LP as at 24 hours, the numbers of OX62 cells in the LP were much reduced. It seems unlikely that the LPS was acting directly on DCs, because in contrast to bone marrowderived DCs, where LPS induces nitric oxide synthase induction and NO secretion (Powell and MacPherson, in preparation) we cannot detect any effects of LPS on isolated lymph DC. TNF-α, possibly macrophage-derived, is involved because an anti-TNF Ab markedly inhibited the effects of LPS. The DCs released following LPS administration do not differ from

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steady-state DCs in MLR stimulation, but other properties were not investigated (MacPherson et al., 1995). In other models (murine heart and kidney (Roake et al., 1995)), IL-1 is also involved in stimulating DC migration. Recent studies in the rat (Matsuno et al., 1996; Kudo et al., 1997) have identified a novel migratory pathway for DCs. Following i.v. particle administration, DCs endocytose particles within the blood (probably in hepatic sinusoids), and then translocate into hepatic lymph, ending up in the hepatic/coeliac nodes. This route has important implications for the understanding of the immune response to blood-borne pathogens.

ISOLATION OF DCs FROM THE GUT DCs are a rare cell type in all tissues and their isolation is fraught with difficulties. Most techniques involve mincing the tissue, enzymatic digestion, differential adherence and negative or positive selection. The yields are low (we routinely start with six or more whole intestines to prepare LP or PP DCs (Liu and MacPherson, 1995a)) and the proportion of DCs that is recovered is unknown. The procedures used to isolate DCs are associated with two major problems – they may be selective for subpopulations of DCs and they may induce changes in the isolated DCs that do not represent normal in vivo events. This is particularly true of procedures that involve overnight incubations at 37C, as many studies have shown that such incubation induces changes analogous to maturation or differentiation, especially if cytokines are present in the incubation mixture. An alternative approach to the study of intestinal DCs has been in use in our laboratory for some years. Lymph DCs (L-DC) are normally removed from the lymph in the first node they reach. Mesenteric lymphadenectomy in the rat, sheep and mouse (Pugh et al., 1983; Rhodes, 1985; Mayrhofer et al., 1986; Bujdoso et al., 1989) has been used to remove this blockage to migration. After a period of weeks, the afferent and

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efferent lymphatics of the removed nodes join up, with the result that DCs can be collected by cannulation of central lymphatics, in the rat the thoracic duct. Such DCs have recently left the intestine physiologically, can be collected in the cold and can be concentrated by simple density gradient centrifugation with or without negative or positive selection. By combining Nycoprep sedimentation with magnetic bead (MACS) separation, we can routinely obtain up to 90–95% DC purity. These DCs are as close to a physiological population as can be acquired at present. Such L-DCs are, however, difficult to collect in large numbers and represent DCs at just one stage in their life history. In addition we do not know whether these DCs derive from the LP, PP or both. Their study has however given a number of important insights into DC properties and functions.

FUNCTIONAL PROPERTIES OF ISOLATED GUT DCs Tissue DCs Relatively few studies have examined the phenotypic and functional properties of DCs isolated from the gut LP or PP. Pavli et al., (1990) isolated DCs from murine PP and LP. They found that these cells resembled splenic DCs in phenotype and function. We used similar approaches to isolate DCs from rat LP and PP (Liu and MacPherson, 1995a). Yields were small and DCs could not be enriched to more than 30–40% purity. DCs freshly isolated expressed high levels of MHC class II, and in contrast to freshly isolated LCs (Schuler and Steinman, 1985) and heart or kidney DCs (Austyn et al., 1994), gave intermediate levels of stimulation in a MLR when compared to lymph DCs. After overnight culture with GM-CSF, LP DCs became as potent MLR stimulators as lymph DCs (Liu and MacPherson, 1995a). APC have been isolated from human colon (Mahida et al., 1988). These cells gave good stimulation in a MLR, but the authors considered that they had both macrophage and DC

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characteristics. In contrast, another study (Pavli et al., 1993) showed that when macrophages and DCs were separately isolated from human colon, most MLR stimulation could be attributed to DCs. PP DCs have been studied more intensively in mice. Ruedl et al., (1996) have shown that freshly isolated CD11c PP DCs did not express DEC-205, were functionally immature in terms of T-cell activation, but were actively endocytic and could phagocytose latex particles. After culture with GM-CSF, TNF-α, or anti-CD40 mAb, the DCs expressed CD11c and DEC-205, lost the ability to process native antigen, downregulated MHC class II and invariant chain synthesis, upregulated B7 and became potent stimulators of resting T cells, thus acquiring the properties of mature DCs. It is suggested that the subdome DCs migrate to T-cell areas, but there is no direct proof for this. This differentiation is similar to that described for LCs in culture, but it is not known whether it represents normal maturation in the absence of inflammatory stimuli. Kelsall’s group has also isolated and characterised PP DCs, and has shown that they have important immunomodulatory properties (see below). Ardavin’s group has recently made an intensive phenotypic study of DCs isolated from lymphoid organs (Anjuere et al., 1999). They find that in contrast to spleen and lymph node, PP DCs belong mainly to the lymphoid subgroup (CD8, DEC-205hi). DCs have also been isolated from porcine PP (Makala et al., 1998), they expressed high levels of MHC class II and were potent stimulators of an allogeneic MLR.

Lymph DCs DCs isolated from the thoracic duct lymph of mesenteric-lymphadenectomised rats have recently left the small intestine and represent DCs actively involved in antigen transport (Pugh et al., 1983; Liu and MacPherson, 1991, 1993). These cells appear to differ functionally from DCs isolated either from peripheral tissues or secondary lymphoid tissues. Thus they are fully mature in terms of MLR stimulation and their potency does not change for at least 72

hours in culture, but they retain the ability to process native antigen for the same period (Liu and MacPherson, 1995b). A partial explanation for these observations comes from recent experiments (Liu et al., 1998). We have shown that in rat intestinal lymph, DC subpopulations can be distinguished by their expression of CD4 and OX41, a SIRP family member (Robinson et al., 1986; Adams et al., 1998). CD4/OX41 DCs are more effective APC for naïve and sensitised T cells, survive better in culture but do not lose the ability to process antigen in culture. CD4/OX41 DCs are weak APC, contain phagolysosomes but do not express detectable FcR, survive poorly in culture, and in culture completely lose the ability to process native antigen whilst becoming as strong stimulators of a MLR as the CD4 cells. This DC subpopulation is very strongly reactive for nonspecific esterase, and recently we have shown that this activity derives from apoptotic intestinal epithelial cells (Huang et al., 2000). Thus functionally CD4 DCs resemble LCs, whereas the CD4 DCs do not. We do not know how these two populations relate to each other, but in vivo kinetic studies show that their life spans and turnover times are similar, suggesting that the CD4 DCs are not precursors of CD4 cells.

UPTAKE OF ANTIGEN BY GUT DCs In order to stimulate an immune response, antigen needs to gain access to DCs. The presence of DCs and their processes within intestinal epithelia (Maric et al., 1996) may facilitate this interaction, but at present it is not known if these DCs can migrate to central lymphoid tissues. Soluble macromolecules can enter subepithelial tissues via M cells in PP (reviewed in Curr. Topics Microbiol. Immunol. vol 236, 1999; Semin. Immunol. vol. 11, 1999) but can also gain access via intact epithelial surfaces (reviewed in Sanderson and Walker, 1993). Particulate antigens can similarly cross the epithelial barrier and can be found in PP and MLN. However, the relative efficiency of uptake at the two sites,

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and their relative importance in the induction of immune responses is unclear (see Oral Tolerance, Ann. N.Y. Acad. Sci., 1995; Thomas et al., 1996, for reviews). Although mucosal DCs are strategically placed to acquire antigen via mucosal surfaces, there is very little concrete evidence that they can do this, and even less that they do so with pathogenic organisms. We have shown that soluble antigen, given by gavage or intra-intestinal injection, is acquired by DCs in the intestinal wall, and that antigen-bearing DCs appear in intestinal lymph within 6 hours (Liu and MacPherson, 1991, 1993). Recently we have shown that 8–12 hours after injecting FITClabelled HSA into the small intestine, 4–5% of lymph DCs contain detectable FITC (Huang and MacPherson, unpublished). DCs from antigenfed rats can stimulate sensitised T cells and more importantly, can sensitise naïve T cells following subcutaneous injection. It was important in these experiments to show that the injected DCs were presenting antigen directly and this was done by injecting parental strain antigen-bearing DCs into an F1 recipient and showing that T cells were only sensitised to the MHC of the injected DCs (Liu and MacPherson, 1993). We could not determine from these experiments the origin of the antigen-bearing DCs, they could have arisen from PP, LP or both. Kelsall and Strober (1996) have shown that CD11c DCs isolated from murine PP after feeding OVA are able to activate OVA-specific naïve T cells in vitro. Thus as expected, PP are clearly a route for delivery of antigen to mucosal DCs, but a route via LP cannot be excluded. In contrast to studies with soluble antigen, very little is known about the interaction of mucosal DCs with particulate antigen or pathogens in situ. Mayrhofer et al., (1986) showed that in rats infected with S. typhimurium, cells with DC characteristics in pseudoafferent intestinal lymph contained Salmonella antigen. A preliminary report has shown that following infection of mice with S. typhimurium expressing green fluorescent protein, bacteria co-localise in PP with cells expressing DC characteristics (N418 expression, lack of macrophage

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markers) (Hopkins and Kraehenbuhl, 1997). A porcine virus localises in PP DCs after intranasal inoculation (Cheon and Chae, 1999). Recently, microcapsules have been detected in intestinal DCs after oral inoculation (Lomotan et al., 1997). A recent report (Vazquez-Torres et al., 1999) is of considerable interest. They have shown that intestinal Salmonella are rapidly (15 min) translocated into the blood stream and thence the spleen and that in the blood stream the bacteria are contained within CD18 cells, possibly macrophages or DCs. The cells were not characterised further but the significance of this report is that it suggests a migratory route from the intestine to blood that cannot involve lymph and passage through the mesenteric nodes, as it is so fast and because essentially all DC/ macrophages in lymph are filtered out in the node.

REGULATION OF IMMUNE RESPONSES BY INTESTINAL DC Self and oral tolerance All studies using isolated gut DCs have shown that they are capable of activating naïve or memory T cells. We have shown that oral antigen is acquired by DCs in the intestinal wall and transported to mesenteric lymph nodes, and that these DCs can activate naïve T cells following adoptive transfer (Liu and MacPherson, 1993). Normally, however, oral antigen is tolerogenic and evidence suggests that even antigen targeted directly to DCs may be tolerogenic (Finkelman et al., 1996). It is an attractive hypothesis that under noninflamed ‘nondangerous’ (Matzinger, 1994) conditions, DCs transport antigen to lymph nodes where it may actively tolerise naïve T cells, and that this may form part of normal immune regulation. Recently, experiments using antigen expressed only on peripheral tissues have shown clearly that a bone marrow-derived APC is required for tolerance to this surrogate self antigen (Adler et al., 1998) and DCs are strong candidates for this

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role (Heath et al., 1998). In contrast to some current dogma, much evidence suggests that DCs are continually trafficking from peripheral tissues to lymph nodes in the absence of inflammatory or other ‘danger’ stimuli (Smith et al., 1970; Pugh et al., 1983). We have recently shown that the weakly immunostimulatory, CD4/ OX41 lymph DC transport apoptotic intestinal epithelial cells to T-cell areas of Peyer’s patches and mesenteric nodes, and that this traffic exists in gnotobiotic rats (Huang et al., 2000). We suggest that this traffic serves to tolerise newly formed T cells to peripheral self antigen. As these same DCs can acquire intraintestinal antigen, this may represent one mechanism involved in oral tolerance.

The switch from tolerance to immunogenicity How this steady state, where tolerance is the primary outcome, is changed to one of active immune induction, is at present unknown. It is an attractive hypothesis that changes in the local cytokine environment, possibly manifested in epithelial cells, modulate DC recruitment and gene expression, and that subsequently these DCs are able to induce active immunity in T cells (reviewed in Strober, 1998). Recently, Viney’s group (Williamson et al., 1999) have shown that in mice whose intestinal DC populations have been expanded by Flt3-ligand treatment, tolerogenic responses to oral antigen can be changed to immunogenic by co-administration of antigen and cholera toxin or IL-1, and that these changes are accompanied upregulation of CD80 and CD86 on intestinal DCs.

Regulation of T-cell differentiation There is currently much interest in the regulation of immune responses in the gut. Apart from the potential for immunotherapy of autoimmune, inflammatory and hypersensitivity diseases, there is the need to base vaccine design on an understanding of the mechanisms by which intestinal and other mucosal responses are regulated. Some principles are becoming

apparent, but there is clearly a long way still to go. That DCs have a role in this regulation is apparent, but details of how their activity is regulated, and how they modulate the differentiation of lymphocytes are far from clear. Immune responses initiated in PP are characterized by activation of B cells to secrete IgA. Although it is clear that T cells are finally responsible for inducing B cells to switch from IgM to IgA or IgG, the local signals that determine the differentiation patterns of those T cells are still unclear, although roles for IL-4, IL-5, IL-10 and TGF-β have been suggested (reviewed in McIntyre and Strober,1999). There is accumulating evidence that DCs have a role in inducing the differentiation of T cells to a mucosal phenotype. In early studies (Spalding et al., 1983) it was shown that cells with similar properties to those of splenic DCs could be isolated from murine PP and in further studies it was suggested that these DCs could have important roles in determining the quality of immune responses initiated in PP. Thus, when mixtures of PP DCs and T cells were isolated and added to sIgM splenic B cells, polyclonal activation of the B cells led to IgA synthesis, whereas the use of splenic DCs led only to IgM synthesis. It was suggested that it was the DCs and not the T cells that were important in stimulating this isotype switch. Cebra’s group (Schrader et al., 1990) have shown that PP DCs and splenic DCs could both support IgA secretion by appropriately primed B cells. Murine lamina propria, PP and splenic APC have been compared in terms of their ability to activate naïve T cells in vitro (Harper et al., 1996). Different patterns of cytokine secretion were induced by the different APC, with splenic cells inducing both TH1 and TH2 cytokines, PP APC inducing IgA cytokines, but LP cells inducing only IFN-γ and TGF-β. However, as the APC populations contained only 1–2% DCs but large numbers of T and B cells, it is difficult to be certain that the differences seen were due to the influence of the APC. Everson (Everson et al., 1998) has evidence that suggests that cytokine secretion by mucosal DCs may influence T cell activation. He showed that activation of T cells by mucosal DCs

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induced a TH2 pattern of cytokine secretion, whereas splenic DCs induced TH1 cytokines. Again, the mechanisms by which DCs influence cytokine secretion by T cells are unknown. Recently, Kelsall’s group (Iwasaki and Kelsall, 1999a, 1999b) have also investigated the properties of PP DCs. Importantly, they isolated DCs in a way that would minimise stimulation of the DCs. They found that if PP DCs were used to stimulate TCR-transgenic T cells in vitro, on restimulation with anti-CD3 and anti-CD28 the T cells secreted high levels of IL-4 and IL-10, but low levels of IFN-γ. In contrast, splenic DCs induced mainly IFN-γ secretion. If cultures were set up with PP DCs in the presence of anti-IL-10 or anti-TGF-β, IFN-γ secretion was enhanced, whereas these Abs had no effect if splenic DCs were used. They also showed that if PP DCs were incubated with anti-CD40 Ab, IL-10 secretion was induced, whereas anti-CD40 had no effect on splenic DCs. Thus PP DCs may have a major role in determining the differentiation fate of naïve T cells. Recently, much interest has centred on a third class of CD4 T cell, regulatory (Tr1) cells. These cells have critical roles in the regulation of inflammation in the intestine (Powrie et al., 1994, 1996; Groux et al., 1997; Read et al., 1998; Seder et al., 1998), reviewed in (Powrie, 1995; Powrie et al., 1997; Mason and Powrie, 1998). IL-10 and TGF-β are critical factors for the induction of Tr1 cells (Powrie et al., 1996; Asseman and Powrie, 1998) and it is an attractive hypothesis that DCs are responsible for secretion of these cytokines, although production of TGF-β by DCs has apparently not yet been documented. If PP DCs and splenic DCs are so different, it remains to be determined how these differences are brought about, but Kelsall has suggested that TGF-β may be involved.

B cell activation It has become clear that DCs also interact directly with B cells and can modulate B-cell differentiation. We have shown that DCs can interact directly with naïve B cells and that they can retain native antigen and make it available for

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recognition by B cells. In addition they provide signals that are necessary for the initiation of isotype switching (Kushnir et al., 1998; Wykes et al., 1998), reviewed in Immunol. Rev. (1999). Recent work from Bancherau’s group supports the concept that DCs may have a direct role in isotype switching in mucosal tissues. They showed that polyclonal activation of ‘naïve’, sIgM human tonsillar B cells in the presence of CD40 ligand and blood-derived DCs led to a skewing of the response towards IgA (Fayette et al. 1997). This area of research is worthy of further exploration; the roles of DC and/or stromal cells in the generation of IgA switch and helper T cells, and in isotype switching in B cells are quite obscure.

INTESTINAL DCs AND RETROVIRAL INFECTION HIV is most often acquired through infection of mucosal surfaces and DCs are strategically placed to encounter the virus, transport it to lymphoid tissues and infect T cells (reviewed in Schneider et al., 1996; Knight and Patterson, 1997; Pope, 1999; Rowland-Jones, 1999; Klagge and Schneider Schaulies, 1999). It is difficult to investigate the role of DCs as HIV transporters in humans, but some important evidence has come from studies of SIV in rhesus macaques (Spira et al., 1996). After vaginal inoculation of SIV, viral DNA was detected by in situ PCR. Viral DNA was first detected in lamina propria cells with the characteristics of DCs in both the vagina and the cervix. Within 2 days, infected cells were detected in the subcapsular sinus and paracortex (T cell area) of the draining nodes. Interestingly, no infected cells were seen in vaginal or cervical epithelium, even though many LCs are present in vaginal epithelium. It is possible that LCs are binding or endocytosing virus without it replicating, this would prevent detection by PCR. This study provides suggestive evidence that DCs are the primary target of SIV (and HIV ) in mucosal infection. Studies in human skin, have however suggested that although DCs may be the first cells to acquire

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the virus, DC–T-cell clusters are the major sites of HIV replication (Pope et al., 1994, 1995). Pope’s group have now (Pope, 1998) isolated similar clusters from mucosal surfaces of SIVinfected macaques and shown that the DC–Tcell conjugates support active viral replication. HIV-infected DC cell syncytia have been observed at the lympho-epithelial surface of the tonsil (Frankel et al., 1996, 1997). Two more recent studies have suggested that local CD4 T-cell infection occurs rapidly after mucosal infection of macaques (Hu et al., 1998; Stahl Hennig et al., 1999). Infection was intravaginally or via non-traumatic application to the tonsil, respectively. After vaginal infection, most of the infected cells extracted from the epithelial tissues were CD4 T cells. There is also evidence that HIV infection can decrease numbers of DCs in mucosal tissues. Thus Lim et al., (1993) showed a decrease in cells with DC phenotype in the duodena and Spinillo et al. (1993) in the cervical epithelium of HIVinfected patients. This might lead to increased susceptibility to intercurrent infections.

CONCLUSIONS AND FUTURE DIRECTIONS The understanding of intestinal DCs is in its infancy. Many studies are still descriptive, relying, particularly in humans, on morphological and immunocytochemical identification of DCs. We know a moderate amount about the distribution of DCs in the gut and are starting to realise that DCs represent complex populations of cells whose lineage and functional relationships to each other are largely unknown. It is becoming clear that they have important roles in determining the different outcomes of T-cell recognition of antigen. We do not understand the roles of DCs in nonperturbed states, and know very little about how the changes in DC properties and functions are induced by inflammation or other ‘danger’ stimuli. The molecular basis of DC function is even more poorly understood. We know very little about the signals that inform DCs to position

themselves at particular sites or to leave those sites, or about the regulation of those signals by environmental stimuli. We do not know how far we can translate the properties of DCs grown in vitro to those that exist in tissues. HIV is probably the most studied mucosal infection in relation to DCs. It is, however, one of many clinically important intestinal infections, and the roles of DCs in immune responses to these infections remain to be defined, yet the development of effective vaccines to these infections must be based on understanding the nature of protective immunity and the roles of DCs in inducing the appropriate response.

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Robinson, A.P., White, T.M. and Mason, D.W. (1986). Immunology 57, 239–247. Rowland Jones, S.L. (1999). Curr. Biol. 9, R248–R250. Ruedl, C., Rieser, C., Bock, G., Wick, G. and Wolf, H. (1996). Eur. J. Immunol. 26, 1801–1806. Sanderson, I.R. and Walker, W.A. (1993). Gastroenterology 104, 622–639. Schneider, T., Ullrich, R. and Zeitz, M. (1996). Semin. Gastrointest. Dis. 7, 19–29. Schrader, C., George, A., Kerlin, R. and Cebra, J. (1990). Int. Immunol. 2, 563–570. Schuler, G. and Steinman, R.M. (1985). J. Exp. Med. 161, 526–546. Seder, R.A., Marth, T., Sieve, M.C. et al. (1998). J. Immunol. 160, 5719–5728. Smith, J.B., McIntosh, G.H. and Morris, B. (1970). J. Anat. 107, 87–100. Spalding, D.M., Koopman,W.J., Eldridge, J.H., McGhee, J.R. and Steinman, R.M. (1983). J. Exp. Med. 157, 1646–1659. Spinillo, A., Tenti, P., Zappatore, R., De Seta, F., Silini, E. and Guaschino, S. (1993). Gynecol. Oncol. 48, 210–213.

Spira, A.I., Marx, P.A., Patterson, B.K. et al. (1996). J. Exp. Med. 183, 215–225. Stahl Hennig, C., Steinman, R.M., Tenner Racz, K. et al. (1999). Science 285, 1261–1265. Steinman, R.M., Lustig, D.S. and Cohn, Z.A. (1974). J. Exp. Med. 139, 1431–1435. Strobel, S. and Mowat, A.M. (1998). Immunol. Today 19, 173–181. Strober, W. (1998). Ann. N Y Acad. Sci. 859, 37–45. Vasquez-Torres, A., Jones Carson, J., Baumler, A.J. et al. (1999). Nature 401, 804–808. Viney, J.L., Mowat, A.M., O’Malley, J.M., Williamson, E. and Fanger, N.A. (1998). J. Immunol. 160, 5815–5825. Wolf, J.L. and Bye, W.A. (1984). Ann. Rev. Med. 35, 95–112. Williamson, E., Westrich, G.M. and Viney, J.L. (1999). J. Immunol. 163, 3668–3675. Wykes, M., Pombo, A., Jenkins, C. and MacPherson, G.G. (1998). J. Immunol. 161, 1313–1319. Zhou, L.J. and Tedder, T.F. (1995). J. Immunol. 154, 3821–3835.

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26 Dendritic cells in the liver, kidney, heart and pancreas Raymond J. Steptoe 1, Peta J. O’Connell 2 and Angus W. Thomson2,3 1 Walter and Eliza Hall Institute of Medical Research, Victoria, Australia; Thomas E. Starzl Transplantation Institute and Departments of Surgery and 3 Molecular Genetics and Biochemistry, University of Pittsburgh, Pittsburgh, PA, USA 2

That swift as quicksilver it courses The natural gates and alleys of the body Hamlet William Shakespeare

INTRODUCTION

monocytic precursors (as discussed elsewhere in this volume), is that a great deal of our recently acquired understanding of DC biology has derived from cells generated in this fashion. While DCs in the vascularized nonlymphoid organs are clearly important in tissue and immune homeostasis, it is a lament of those seeking to gain an understanding of the role of DCs in these processes that these DCs are becoming increasingly less well understood relative to their in vitro-derived counterparts. This is inevitably a consequence of the relative difficulty of dealing with DC from nonlymphoid tissues due to their existence as a ‘trace’ population in these tissues. It is, however, important that the gains in our understanding achieved through the use of in vitro-generated cells are applied to, and validated in, organs such as the liver, heart, kidney and pancreas.

Much of the impetus for the study of DCs in the organs reviewed in this chapter had originally arisen from a desire to understand the immunological processes underlying rejection of these organs following allogeneic transplantation. Study of DCs in these organs ultimately led to the concept that the nature of (donor) DCs in these organs may influence, in a dichotomous fashion, the nature of allograftinduced responses vis-à-vis acceptance or rejection (tolerance or immunity). Similarly, interest has been shown in the potential role that DC in nonlymphoid tissues may play in maintaining peripheral self-tolerance. A consequence of the advent of in vitro propagation of DCs from either hematopoietic progenitor cells or cytokine-induced differentiation of

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LIVER Histological location and phenotype While Langerhans cells described in epithelial sites, such as the oral mucosa (Hutchens et al., 1971) and epidermis (Silberberg-Sinakin et al., 1978; Stingl et al., 1978), had been shown to be related to DCs, Hart and Fabre (1981a) provided a ground-breaking immunohistological demonstration of the presence of presumptive DCs in interstitial connective tissue of many nonlymphoid solid organs, including the liver. Previously, Forsum et al. (1979) had demonstrated sparsely distributed individual major histocompatibility complex (MHC) class II cells in the liver of the guinea pig, however, staining had been attributed to Kupffer cells, the resident macrophage population of the liver. Immunostaining of tissue sections localizes MHC class II cells primarily around the periportal area and central veins, with a few cells scattered throughout the parenchyma of rat (Hart and Fabre, 1981a; Steiniger et al., 1984; Spencer and Fabre, 1990), mouse (Witmer-Pack et al., 1993; Woo et al., 1994) (see Figure 26.1) and human (Prickett et al., 1988; Ballardini et al., 1989) livers. The constitutive expression of MHC

II antigens (Ags), and the dendriform morphology of these cells, led to their initial classification as DCs. While MHC class II Ags are also expressed constitutively by human (Hart and Fabre, 1981a; Prickett et al., 1988), but not rodent (Steiniger et al., 1984) Kupffer cells, the restricted localization of DCs allows them to be readily distinguished from Kupffer cells, which are distributed uniformly throughout the parenchyma (Hart and Fabre, 1981a; Spencer and Fabre, 1990; Witmer-Pack et al., 1993). The presence of MHC class II presumptive DCs, has also been reported in the liver capsule (Prickett et al., 1988). Unlike Kupffer cells, liver DCs do not express nonspecific esterase or α-napthylacetate esterase activity (Hart and Fabre, 1981a; Lautenschlager et al., 1988) facilitating distinction of these cell types. In situ examination of liver following intravenous (i.v.) administration of colloidal carbon (Hart and Fabre, 1981a; Steiniger et al., 1984; Witmer-Pack et al., 1993) or sheep red blood cells (SRBC) (Lautenschlager et al., 1988), has led to the conclusion that, unlike Kupffer cells, liver DCs do not phagocytose these particles in vivo. However, cellmediated transfer of colloidal carbon to the celiac (liver-draining) lymph nodes has been observed following i.v. injection of this agent.

FIGURE 26.1 Identification of DC in mouse (C57BL/10J) liver (portal tract), heart, kidney (cortex) and pancreas (islet) by immunohistochemical staining for MHC class II (IAb) (400). Micrograph courtesy of Dr V.M. DENDRITIC CELLS IN THE PERIPHERY

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While this finding was previously attributed to migrating Kupffer cells (Hardonk et al., 1986; Matsuno et al., 1990) more recent studies suggest it may be due to phagocytic DC progenitors translocating from blood to celiac lymph within the liver (Matsuno et al., 1996, 1997) (vide infra). This is supported by observations of ED1ED2OX-6 bead-bearing DCs in the hepatic sinusoids 1 hour following i.v. injection of latex spheres (Sato et al., 1998). Phenotypic TABLE 26.1 Human liver dendritic and Kupffer cell antigen expression determined by immunomorphology mAb specificity

Liver interstitial DC

Kupffer cells

MHC class I





MHC class II HLA-DP HLA-DQ HLA-DR

  

  

LCA (CD45)





FcR CD16 CD32 FcRI

  

 / /

Adhesins/ Complement receptors CD11a CD11b CD11c CD18

   

   

Other CD4 CD14 CD39 CD40

   

   /

Modified after Prickett et al. (1988). TABLE 26.2

and ultrastructural analyses suggest that these DCs represent immature, ‘Ag-processing’ DC (vide infra) (Sato et al., 1998). In vitro generated DC progenitors propagated from stem cells present in adult liver are able to phagocytose opsonised SRBC, however, this activity is lost following provision of ‘maturation’ signals by the extracellular matrix protein type-1 collagen (Lu et al., 1994) that is spatially associated with liver DC in vivo (vide infra). Thus, differences in the location, morphology, phenotype and other characteristics underline the distinct and separate nature of DC and Kupffer cells in this organ (Table 26.1). Recent immunomorphological studies in this laboratory have indicated that while presumptive MHC class II DCs are sparsely distributed in the liver, and the density of cells per unit volume may be low compared with other solid organs (Table 26.2), the total DC content of murine livers is high, up to 5- to 10-fold that of other mouse parenchymal organs, such as heart and kidney (Table 26.3) (Steptoe et al., 2000). In mice, the β2 integrin p150,90 (CD11c) is expressed by DCs in a wide variety of lymphoid and nonlymphoid sites (Metlay et al., 1990) and is widely used as a marker of DCs in this species. The total number of hepatic CD11c cells (assessed by flow cytometry) per liver is roughly one sixth that of the spleen (approximately 0.7  106 versus approximately 4  106 CD11c cells/organ, respectively) (Shaw et al., 1998). Both CD8α (CD11bhi, CD11c) and CD8α (CD11blo, CD11c) DCs may be identified in mouse livers (O’Connell et al., 2000). CD8α and CD8α DCs each constitute ≤1.0% of freshly

Relative MHC II cell density in parenchymal organs Cell density (cells/cm3  106)a

Strain

Heart

C57BL/10

6.64  3.4b,d

a

Kidney

Pancreas

Liver

5.04  0.55c,d

4.45  1.6b,d

1.08  0.25

b

Sections were stained with anti-IA mAb visualised using biotinylated anti-rat Ig and ABC/DAB. Immunopositive cells were enumerated using the ‘disector’ principle. b Significantly different from liver (P  0.05). c Significantly different from liver (P  0.005). d No significant difference between other groups. From Steptoe et al. (2000). Copyright  2000 of Elsevier Science B.V. DENDRITIC CELLS IN THE PERIPHERY

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Total MHC II cell number per organ MHC class II cell number (mean  1 S.D.  106)a (% of total CD45 cells)b

Strain

Pancreas

Heart

Kidney

Liver

C3H

0.34  0.01c (52.9%)

0.11  0.01c (40.3%)

0.78  0.04c (55.8%)

1.17  0.10c (21.2%)

B10.BR

0.31  0.05d (51.5%)

0.44  0.07e (38.2%)

0.63  0.04f (47.1%)

3.04  0.18 (25.5%)

C57BL/10

0.44  0.01 (60.6%)

0.43  0.03 (50.1%)

0.46  0.08 (56.5%)

2.87  0.26g (14.5%)

a

Sections were stained with anti-I-Ak (C3H) or anti-I-Ab (C57BL/10 and B10.BR) mAb visualized using biotinylated anti-rat Ig and ABC/DAB. Immunopositive cells were enumerated using the ‘fractionator’ principle. b Calculated percentage of CD45 cells co-expressing MHC class II. c Significantly different from all other C3H organs analyzed (P  0.001). d Significantly different from B10.BR liver and kidney (P  0.001). e Significantly different from B10.BR liver (P  0.001) and kidney (P  0.05). f Significantly different from all other B10.BR organs analyzed (P  0.001). g Significantly different from C57BL/10 heart, pancreas (P  0.001) and kidney (P  0.01). From Steptoe et al. (2000). Copyright  2000 of Elsevier Science B.V.

isolated liver NPC, however, both populations can be enriched to 10–15% by overnight culture and metrizamide density centrifugation (Figure 26. 2). The total number of hepatic DCs (both CD8α and CD8d) may be greatly augmented by systemic administration of the hemopoietic growth factor Flt3 Ligand (L) (Steptoe et al., 1997; O’Connell et al., 2000) (Figure 26.2). Flt3L mobilizes stem and progenitor cells from bone marrow and results in accumulations of large numbers of functional DCs in lymphoid and nonlymphoid tissues. Following Flt3L administration (10 µg for 10 days) the number of lowbuoyant density CD11c DCs recovered from the livers of treated mice is strikingly elevated by up to 650-fold (Steptoe et al., 1997; O’Connell et al., 2000). Maximal increases in DC numbers are observed after 10 days of continuous treatment (Shaw et al., 1998). Flt3L potently induces equal expansion of both CD8α and CD8α subsets in vivo (O’Connell et al., 2000). Histologically, clusters of infiltrating MHC class II CD11c cells can be observed, not only in the portal areas (where DCs are normally located), but also throughout the parenchyma of Flt3Ltreated animals (Shurin et al., 1997). Extensive characterization of the in situ

phenotype of rodent liver DCs has not been performed and thus few reports are available detailing the location of cells immunoreactive with mAbs recognizing DC-associated Ags within the liver. In the rat, MHC class II and/or OX62 cells are localized predominantly in portal areas, with only low numbers of cells visualized in the sinusoids and around the central vein (Brenan and Puklavec, 1992; Matsuno et al., 1996; Sato et al., 1998). Double immunostaining for ED1 (antimonocyte/ macrophage), ED2 (antimacrophage/Kupffer cells) and OX6, identifies two distinct phenotypes of hepatic DC, ED1ED2OX6, and ED1ED2OX6 (Sato et al., 1998). Interestingly, rat hepatic MHC class II DCs appear to express variable levels of CD4 (Steiniger et al., 1984; Stein-Oakley et al., 1991), in common with DCs in other peripheral tissues of this species (Steiniger et al., 1984; Darden et al., 1990; Schon-Hegrad et al., 1991). In the mouse, CD11c (Witmer-Pack et al., 1993) and DEC-205 cells (Shurin et al., 1997) are predominantly located periportally, as reported for MHC II cells. The in situ phenotype of human liver DCs has been examined more extensively and is summarized in Table 26.1. DCs in the normal mouse liver, exist in an ‘immature’ or ‘Ag-processing’ phenotype, as

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10

4

10

10

4

LIVER

0.3% 3

10

3

10

10

4

2

10

1 0

10

0

3

10

0

10

10

1

10

2

10

3

10

4

0

10

10

2.0%

1

10

2

10

3

10

10

4

10

4

10

4

10

10

10

4

10

4

10

2

4

1

10

10

0

0.1% 0

10

1

10

2

1.1%

10

10

1

0.3% 3

3

10

10

3

13.8% 10

Day 0

10

2

10

3

0.6%

12.6%

2

-10

2

-10

-10

2

Day 0-DC

3

1 0 0

1

10

2

10

3

10

4

0

10

10

1

10

2

10

3

10

3 2

10

CD11b

1

10

0

10

CD8α

0

10

4

10

3

10

10

3 2

10

0

10

2

10

27.2%

29.6%

10

3

10

2

10

1

10 10

0

1.7%

1

10

10

10

10

4

10

1.3%

0.8%

Day 10

10

0 4

10

4

10

4

2

10

1

1

10

10

0

10

10

10

0

0.7%

10

1

10

10

1

Enriched

0

10

1

10

2

10

3

10

4

10

0

10

1

10

2

10

3

10

CD11c

Isotypes

FIGURE 26.2 Detection of CD8α and CD8α DC in livers from both normal and Flt3L-treated (10 µg; 10 days) B10 mice. DCs were enriched from normal liver by overnight culture of nonparenchymal cells (NPCs) followed by metrizamide density centrifugation. Freshly isolated or DC-enriched NPCs were double-stained with FITCconjugated anti-CD8α or anti-CD11b mAb, together with anti-CD11c PE mAb, and then analyzed by flow cytometry. The results are representative of two separate experiments.

described for DCs in other nonlymphoid tissue sites, rather than the ‘Ag-presenting’ phenotype reported for DCs in lymphoid tissue (reviewed in Steinman, 1991). Immunostaining for the costimulatory molecules CD40, CD80 (B7-1) and CD86 (B7-2) (Inaba et al., 1994; Morelli et al., 2000) in sections of normal mouse liver indicates that liver DCs do not constitutively express detectable amounts of these molecules in situ, a

phenotype characteristic of DCs in other peripheral tissues (Inaba et al., 1994). Interestingly, CD86 expression is evident on Kupffer cells in normal mouse liver (Inaba et al., 1994). The predominant localization of immature, ‘Agprocessing’ DCs within portal tracts is consistent with the notion that DCs are found in locations where immune surveillance may be necessary. In the case of the liver, the portal system draining

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DENDRITIC CELLS IN THE LIVER , KIDNEY, HEART AND PANCREAS

the mucosal surface of the gut, is exposed to both foodborne immunogens and pathogens and to symbiotic eukaryotic organisms. Interestingly, the development of tolerance to antigen administered either orally or via the portal venous tract may be dependent on liver DCs (Yang et al., 1994; Gorczynski et al., 1996). The immature in situ phenotype of liver DCs (Plate 26.3) is consistent with observations of DC isolated by collagenase digestion of the liver, followed by metrizamide density centrifugation to obtain the low buoyant density nonparenchymal cells (NPCs), from both normal and Flt3L-treated mice (Woo et al., 1994; O’Connell et al., 2000). Freshly isolated CD11c DCs, including both CD8α and CD8α, are MHC class IIlo, CD40/lo, CD80lo and CD86lo. However, consistent with DCs from lymphoid tissues, mouse liver DC exhibit upregulation of MHC class II and costimulatory molecule expression (Figure 26.4) together with concomitant downregulation of CD16/32 (FcγRII) and F4/80 following overnight culture (Woo et al., 1994; O’Connell et al., 2000).

Population dynamics Demonstration of the replacement of donorderived DCs in liver allografts with those of host origin confirms that this pool of cells is maintained, at least in part, from progenitor cells derived from the host’s hemopoietic system (Valdivia et al., 1993). While data are not available to indicate the population dynamics of these cells in humans, isolation of DCs from lymph draining the liver following bromodeoxyuridine feeding indicates that in rats, the DC migration rate from the liver is approximately 1  105 DCs/h (Matsuno et al., 1995, 1996). Furthermore, approximately half the DCs leaving the liver via lymph have arisen by division within the previous 5.5 days (Matsuno et al., 1996). Additionally, DCs bearing ingested latex particles appear quickly (within 1 hour) in lymph draining the liver of rats administered latex particles i.v. (Matsuno et al., 1996; Sato et al., 1998). It has been proposed that these particle-laden DCs may not be derived from DCs resident within the liver, rather that they develop from a marginated pool in the circulation which rapidly translocates

0

10

1

10

2

10

3

10

4

10

0

10

1

2

10

10

3

10

4

10

0

11.4%

0

32.3%

0

10.3%

0

48.8%

16

16

30

16

Immature (freshly-isolated) DC

0

10

1

10

2

10

3

10

4

10

0

10

1

10

2

10

3

10

4

10

71.0%

0

10

1

10

2

10

3

10

MHC class II

4

10

0

10

1

10

2

10

CD40

3

10

4

10

0

71.6%

0

0

70.4%

0

69.3%

16

16

30

16

Mature (overnight-cultured) DC

0

10

1

10

2

10

3

10

4

10

CD80

0

10

1

10

2

10

3

10

4

10

CD86

FIGURE 26.4 Flow cytometric analysis of liver CD8α DC from B10 mice after 10 days Flt3L administration. NPC were triple immunolabeled with (i) anti-CD11c PE, (ii) anti-CD8α CyC, and (iii) anti-MHC class II (IAb), anti-CD40, anti-CD80 or anti-CD86-FITC. Histograms show the expression of MHC class II, CD40, CD80 or CD86 for freshly isolated (immature) and overnight-cultured (mature) CD8α CD11c DCs. The data demonstrate that culture of freshly isolated, immature liver CD8α results in phenotypic maturation. Similar results were obtained for CD8α CD11c DCs. Open profiles denote isotype-matched controls. The results are representative of four separate experiments. DENDRITIC CELLS IN THE PERIPHERY

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LIVER

via hepatic sinusoids to hepatic lymph vessels (Matsuno et al., 1996; Kudo et al., 1997). The liver may therefore represent an important site in which circulating DCs that have phagocytosed particulates can gain access to lymph draining to celiac lymph nodes. Adoptively transferred allogeneic DCs that migrate from blood to celiac lymph nodes (apparently via this pathway) are able to induce proliferation in alloreactive T cells in the paracortical regions of the celiac nodes (Kudo et al., 1997). These results raise the possibility that celiac lymph nodes may represent an important site for induction of immune responses to bloodborne pathogens, particularly as the rate of DC migration via this route appears to increase following i.v. administration of particulates (Austyn, 1996). Liver DC normally traffic, via lymph, to the celiac lymph nodes (Matsuno et al., 1995, 1996) and possibly to the spleen via blood (although this has not been demonstrated). However, if the lymphatic vessels are disrupted, as in the case of liver transplantation, larger numbers of donor-derived MHC class II DCs are detected in both the spleen and celiac lymph nodes (Sun et al., 1995; Bishop et al., 1996; Steptoe et al., 1999).

Ontogenic development Few reports have detailed the ontogenic development of DC populations in the liver. In humans, isolated HLA-DR cells (of undefined histological type) have been reported in the liver of 8-week-old embryos. At this time, the only other HLA-DR cells that exist are DCs in the thymus (Natali et al., 1984). Development of an adult-equivalent distribution of HLA-DR DClike cells is attained in most human fetal tissues, including the liver, by 26 weeks of gestation (Natali et al., 1984). In the rat, van Rees et al. (1988) demonstrated an absence of MHC class II DC in the liver until after birth. Cell populations expressing the macrophage-associated Ags recognized by the mAbs ED1, ED2 and ED3 however, developed prenatally. A similar absence of MHC class II cells in the liver has also been reported prenatally in mice (Natali et al., 1981).

Function DC isolated from liver tissue The functional capacity of liver DCs has not been examined extensively, although studies indicate that these cells are able to acquire the potent allostimulatory capacity attributed to DCs isolated from other tissue sites. Klinkert et al. (1982) provided evidence that the ability to act as accessory cells in periodate-induced mitogenesis was associated with DC-enriched populations from rat liver. Lautenschlager et al. (1988) demonstrated that rat liver DCs, isolated as overnight-cultured, nonadherent lowbuoyant density cells (LBDC), when adoptively transferred, primed naïve allogeneic recipients for accelerated allograft rejection. Interestingly, in these experiments, the DC-enriched preparations from liver contained approximately 80% DCs, but primed for accelerated allograft rejection only as efficiently as crude splenocytes which contained 5% DCs (Lautenschlager et al., 1988). This raises the question whether liver DCs are as functionally competent as those from spleen. DCs enriched from mouse liver using a similar technique of harvesting LBDC after overnight culture, either in the presence or absence of granulocyte–macrophage colonystimulating factor (GM-CSF), effectively stimulate allogeneic mixed leukocyte responses (MLR) in vitro (Woo et al., 1994; Steptoe et al., 1997). Human DCs that migrate overnight from thin pieces of liver tissue exhibit a phenotype of intermediate maturity similar to LC, and activate naïve cord blood lymphocytes with equivalent efficiency at low stimulator:responder ratios (Goddard et al., 2000). However, it is unclear from these studies how the relative allostimulatory ability of liver DCs that undergo spontaneous maturation in vitro (overnight culture), compares with that of ‘classical’ immunostimulatory DCs isolated from lymphoid tissues. The stimulatory capacity of freshly isolated unmanipulated liver DCs has not been assessed, although the absence of CD80 and CD86 expression by these cells in situ (Inaba et al., 1994) (see also Plate 26.3) suggests that, in common with DCs from

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other nonlymphoid tissue sites, these cells would exhibit weak stimulatory capacity. Phenotypically, CD11c CD8α or CD8α DC freshly isolated from livers of Flt3L-treated mice are immature, and undergo spontaneous maturation following overnight incubation (Figure 26.4) (O’Connell et al., 2000). It has been reported that overnight-cultured hepatic DCs from Flt3L-treated mice exhibit similar levels of MHC class II, but elevated levels of CD80 and CD86 compared with controls (Steptoe et al., 1997). There is also evidence that the allostimulatory activity of overnight-cultured, DC-enriched liver NPCs is higher when these cells are obtained from Flt3L-treated animals compared with controls (Steptoe et al., 1997). However, this may represent the elevated number of DCs in the former preparations due to Flt3L administration, and is consistent with observations that Flt3L treatment markedly increases the number of in vitro generated liver-derived ‘DC progenitors’ (vide infra) that can be propagated from mouse livers (Drakes et al., 1997b; Steptoe et al., 1997). As reported for immature, liver-derived DC progenitors generated from normal mice (Lu et al, 1994; Thomson et al, 1995), mature overnight-cultured Flt3L-induced liver DCs, including CD8α DCs, home in vivo to secondary lymphoid tissues following their local subcutaneous (s.c.) injection (Figure 26.5) (Morelli et al., 2000; O’Connell et al., 2000). In vitro analysis has shown that mature CD8α and CD8α hepatic DCs are strong and equally efficient stimulators of allogeneic TH1 (mainly) and TH2 cell proliferation in primary MLR (Figure 26.6) (Morelli et al., 2000; O’Connell et al., 2000). Following s.c. injection of bulk Flt3L-induced DC, clusters of IFNγ-producing cells, and a few IL-4-producing cells, were detected in splenic T-cell areas 2 days later. Double immunostaining for cytokines and the allogeneic MHC class II of the donor DC, demonstrated that at least some of the cytokineproducing areas were spatially associated with donor DCs (Morelli et al., 2000). Recently, both stimulatory and regulatory activity has been ascribed to mononuclear NPCs from Flt3Ltreated mice. Portal vein immunization with

FIGURE 26.5 Tissue ‘homing’ of overnight-cultured, sorted B10 (H2b) liver CD8α DC to allogeneic C3H (H2k) recipient draining popliteal lymph node, 2 days after s.c. administration into the hind footpad. Trafficking of CD8α DC was analyzed by (A) PCR detection of donor-specific DNA (MHC class II; IAb) in the draining popliteal lymph node (LN). Amplification of β-actin was used as the internal positive control. (B) Additionally, two-color immunohistochemistry, using anti-IAb mAb followed by ABCAP (black), and anti-CD3ε mAb followed by ABC-Px (grey) identified CD8α DC (arrows) in the subcapsular and paracapsular areas. Only a few of the CD8α DCs were identified in the paracortical T cell areas (grey) (100). From O’Connell et al. (2000) Copyright  2000 The American Association of Immunologists. Staining and micrograph (B) courtesy of Dr A.E. Morelli.

Flt3L-induced DCs in mice, has been associated with TH2 polarization of host T cells and prolongation of allograft graft survival (Gorczynski et al., 1999). A DEC-205 OX-2 hepatic mononuclear cell (presumptive DC) from both normal and Flt3L-treated mice, was found to inhibit T cell proliferation induced in vitro, by allogeneic DCs. Cells derived from these MLR cultures were associated with extended graft survival upon transfer to kidney transplant recipients. IL-2 and IFN-γ production was inhibited in vitro, and IL-4 and IL-10 production was enhanced (Gorczynski et al., 1999). Interestingly, human

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2

10

3

10

4

10

4

10

4

3

5.9%

1

10

2

10

3

10

4

10

0

10

1

2

10

3

10

4

10

4

10

2.5% 2

10

0

10

1

10

2 1

10

0

10

0

10

0

10 10

4

10

3

3

10

10

2

10

IL-10-PE

10

1

10

2

10

3 2

10

1

10

1

10

10

2

10

1

10 10

1

10

0

10

18.5%

0

10 4

4

10

10

3

10

3

2

10

IL-4-PE

0

10

1

10

0

10

0

10 0

10

4.1%

10

3

4

10

2

10

3

0.1%

0.1% 0

10 4

4

10

10

3

10

3

2

10

10

1

10

IFNg-PE

10

1

10

2

10

3

10

2

10

1

10

0

10 0

10

10

Isotype control-PE

8.6%

14.2%

1

B

10

4

0.2%

10

A

10

10

4

LIVER

0

10

1

10

2

10

3

10

4

10

0

10

1

10

2

10

3

10

4

10

CD4-FITC

FIGURE 26.6 Detection of Th1 (IFNγ) and Th2 cytokines (IL-4 and IL-10) in C3H CD3 CD4 T cells after in vitro stimulation with overnight-cultured, γ-irradiated allogeneic (B10) CD8α DC (A), or CD8α DC (B) from Flt3Ltreated liver. After 3 day MLR, the T cells were harvested and restimulated with anti-CD3ε plus anti-CD28 mAbs in the presence of Brefeldin A for 5 hours at 37C. Thereafter, the cells were triple immunolabeled with (i) anti-CD3 CyC, (ii) anti-CD4-FITC and (iii) anti-IFNγ, IL-4 or IL-10-PE. Only CD3 T cells are illustrated in the Figure; APC and other minor contaminating populations in the MLR were gated out according to their lack of expression of CD3. The data are representative of three separate experiments.

liver DC, unlike epidermal LC, prodice IL-10 following overnight culture in GM-CSF (Goddard et al., 2000). Liver-derived DCs collected from the thoracic duct of celiac lymphadenectomized rats possess allostimulatory activity in MLR assays, equivalent to that of DCs of intestinal origin collected in a similar manner from mesenteric lymphadectomized rats (Matsuno et al., 1995, 1996). DCs of intestinal origin collected from mesenteric lymph have been characterized previously, and while able to stimulate MLR responses they retain the capacity to ingest and process antigen (Liu and MacPherson, 1995a, 1995b). This suggests that these cells may be intermediate between ‘Ag-processing’ and ‘Ag-presenting’ stages. Whether liver-derived DCs in celiac lymph are in a similar stage of development is unclear. In the studies of Kudo et al. (1997) however, DCs bearing latex beads that were adoptively transferred i.v. and subsequently isolated from celiac lymph were unable to phagocytose further particulates in vitro. Therefore, the studies of Matsuno et al. (1995, 1996) suggest that the DCs that migrate from the liver via the lymphatics are either already ‘mature’ immunostimulatory cells, or that they rapidly mature upon leaving the liver. The evidence provided by in situ studies of costimulatory molecule expression (Inaba et al., 1994) (see also plate 26.3)

suggests the latter alternative. This situation may, how-ever, be altered when DC-bearing particulates translocate from blood to celiac lymph as discussed above (Kudo et al., 1997). Donor (or so-called ‘passenger’) leukocytes, resident in organ allografts at the time of transplantation, may remain in the graft or traffic to lymphoid tissues in hosts that may or may not be treated with immunosuppressive therapy. It has been postulated that these immature DCs could then constitute a population of cells that might interact with alloreactive T cells in a potentially ‘tolerogenic’ fashion (Steinman et al., 1993). Such cells could be important in contributing to the relatively favorable outcome observed with liver allografts compared with other commonly transplanted organs. Interestingly, augmentation of not only the number of DCs, but also the functional capacity of these cells in murine donor livers by pretreatment with Flt3L (vide supra) results in reversal of the spontaneous acceptance of allogeneic livers normally observed in this species, and results in rapid graft rejection (Steptoe et al., 1997). It was concluded from these studies that, under normal circumstances, donor-derived DCs do not induce an effective T cell-mediated anti-allograft response in liver recipients, but that this may be altered by increasing the number and functional capacity of these cells in the donor organ.

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DC progenitors propagated in vitro from adult liver hemopoietic stem/progenitor cells As with other tissue sites, one of the major difficulties of studying liver DCs is the inability to readily obtain large numbers of these cells. In an effort to overcome this problem, Lu et al. (1994) developed a procedure whereby DCs could be generated from stem/progenitor cells present within the mouse liver NPC population using a modification of the procedure described for GM-CSF-mediated propagation of DCs from mouse blood or bone marrow (Inaba et al., 1992a, 1992b). DC propagated in this manner expressed high levels of CD45, CD11b, CD24 (HSA), and CD44, but only low to moderate levels of the macrophage marker F4/80, the antigen uptake-associated receptor CD16/32 (FcγRII), ICAM-1 (CD54) and the DC-restricted markers DEC-205, CD11c and 33D1 (Lu et al., 1994). These cells exhibit the classic veiled morphology of DCs, yet exhibit only low levels of CD40 and the CD28/CTLA4 ligands CD80 and CD86 (Lu et al., 1994; Drakes et al., 1997b). Thus, these cells were termed liver-derived ‘DC progenitors’. Extended periods of culture or exposure to typical DC maturation-inducing stimuli, including tumor necrosis factor-α (TNF-α), interferon-γ (IFN-γ) or bacterial lipopolysaccharide (LPS) did not induce high levels of MHC class II antigen expression or the acquisition of fully ‘mature’ DC characteristics (Lu et al., 1994). However, culture of these liver MDC progenitors in the presence of collagen type-1, fibronectin or laminin – extracellular matrix proteins that DC are spatially associated within liver, resulted in ‘maturation’ (Lu et al., 1994; Drakes et al., 1997a,). For example, liver-derived DC progenitors were unable to effectively stimulate MLR responses, in contrast to those cultured in the presence of type-1 collagen for 3 days (Lu et al., 1994). Likewise, phagocytic activity was downregulated (Lu et al., 1994). It is therefore interesting to speculate that contact with these extracellular matrix proteins during migration may provide signals which induce ‘maturation’ of hepatic DCs. When the migratory potential of liver DC pro-

genitors was examined by i.v. injection into allogeneic recipients, they were found to home to the periarteriolar lymphoid sheath (T-cell area) of the spleen, and to express moderate to high levels of donor MHC class II (Lu et al., 1994; Thomson et al., 1995). Recent studies from this laboratory demonstrate that mouse liverderived DC progenitors and mature liver myeloid DCs exhibit similar mRNA expression profiles for CC and CXC chemokines and their receptors (Thomson et al., 1999; Drakes et al., 2000). RANTES (regulated upon activation, normal T cell expressed and secreted) and MIP (macrophage inflammatory protein)-1α were the most prominently expressed chemokines, as was their common receptor, CCR1 (Drakes et al., 2000). MIP-1α production by liver-derived DCs was induced both by LPS and coculture with allogeneic T cells. MIP-1α was also found to be chemotactic for liver-derived DCs, inducing their in vitro migration (Drakes et al., 2000). The reduced capacity of GM-CSF-stimulated liver-derived DC progenitors to induce allogeneic T-cell proliferative responses led to speculation that they may act as ‘tolerogenic’ or anergyinducing APCs. This was examined by pretreatment of pancreatic islet allograft recipients with liver-derived DC progenitors of donor origin, 7 days prior to islet cell transplantation. The result was prolongation of islet allograft survival (Rastellini et al., 1995). More recent studies indicate that the comparative immune privilege of hepatic allografts may be due, in part, to the induction of TH2 cytokine responses by liverderived DC progenitors (Khanna et al., 2000). Thus, liver-derived myeloid DC progenitors induced only minimal T-cell proliferation and weak cytotoxic T responses in vitro, while clusters of IL-10- and IL-4-secreting mononuclear cells could be detected in draining lymphoid tissues following s.c. injection into allogeneic hosts. By contrast, comparatively high numbers of IFNγ, but no IL-10- or IL-4-secreting cells, were induced by bone marrow-derived myeloid DCs (Khanna et al., 2000). Recently, a novel population of normal adult mouse liver-derived DCs, propagated in response to IL-3 and CD40 ligation has been described (Lu et al., 2000).

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These DCs, that appear to induce T regulatory cells in vitro, prolong organ allograft survival, and could conceivably play a role in liver transplant tolerance. The potential therapeutic role of these putative tolerogenic liver-derived DC populations is discussed in detail elsewhere in this volume. Whether DCs propagated from liver NPCs are developmentally identical to DCs freshly isolated from liver tissue remains unclear, although the limited comparative studies to date indicate similar characteristics for all features examined.

Disease In viral hepatitis, interdigitating DCs (IDCs) have been identified by ultrastructural analysis (Bardadin and Desmet, 1984) and by immunohistochemical techniques (van den Oord et al., 1990) in areas of piecemeal necrosis in human liver. IDCs are often observed closely associated with CD8 T cells in the periphery of necrotic areas. Large numbers of HLA-DR presumptive DCs have also been observed in areas of ‘spotty’ inflammation (van den Oord et al., 1990). Additionally, HLA-DR DCs have been noted amongst CD4 T cells in severely inflamed portal tracts (van den Oord et al., 1990). It has been suggested that within the central area of portal tracts, CD4 T cells are activated by DCs, whereas the periportal and centrilobular parenchyma appears to be the site where CD8 cells are mediating cytotoxic activity (van den Oord et al., 1990). Mature, activated CD83 DC may be critical for the recruitment and survival of activated tumor-specific lymphocytes during carcinogenesis. However, whilst the frequency of activated (CD83) DCs in the peripheral blood of patients with hepatocellular carcinoma is similar to those with liver cirrhosis, their number is significantly lower in liver tissue, and they have not been identified infiltrating cancer nodules (Chen et al., 2000). Follicular DC neoplasms have also been identified in liver (Arber et al., 1998; Shek et al., 1998). These rare, and generally lymphoid tissue-associated tumors harbor the Epstein–Barr virus (EBV), which is suggested to

play a role in disease pathogenesis. This has important implications for organ transplantation, as EBV in spindle cell tumors is usually associated with immunosuppressed children and adults. The antitumor role of DC has recently been highlighted in a murine hepatic tumor model. Flt3L administered alone, or in combination with IL-12, significantly reduced the number of hepatic metastases (Peron et al., 1998). Flt3L treatment was associated with significant infiltration of tumor borders by both lymphocytes and DCs, concomitant with an increased number of apoptotic bodies (Peron et al., 1998). The number of DCs was enhanced, both in the liver parenchyma and the tumor metastases, supporting the proposal that DCs may be directly involved in the antitumor effect of Flt3L (Peron et al., 1998). DCs may play a role in the development of primary biliary cirrhosis (PBC). They have been observed frequently within the bile duct in the early phases of PBC, but are much less common in advanced disease where they may be more often located periductally (Ballardini et al., 1989; Demetris et al., 1989; Kaji et al., 1997). The numbers of IDCs are significantly higher compared with CD83, activated DCs (Tanimoto et al., 1999), suggesting a role for DCs of different activation states in disease pathogenesis. In a transplant setting, Demetris et al. (1991) provided immunohistochemical evidence that donorderived DCs within rat liver allografts were clustering with, and possibly activating, infiltrating CD4 and CD8 T cells. Likewise, DCs are found within the portal inflammatory infiltrate in obliterative arteriopathy (chronic allograft rejection) (Demetris et al., 1989), and they have been observed interacting with T cells present within the cellular infiltrate (Oguma et al., 1988, 1989).

KIDNEY Histological location and phenotype Early reports showed DCs to be distributed throughout the urinary tract (Hart and Fabre, 1981a, 1981b). Subsequent reports have provided

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further details of the presence of DCs in the kidney of rat (Steiniger et al., 1984; Leszczynski et al., 1985a; Gieseler et al., 1997), mouse (Austyn et al., 1994; Steptoe et al., 2000), musk shrew (Kerschbaum et al., 1995), and human (Hart et al., 1981; Daar et al., 1983; Hancock and Atkins, 1984). Morphometric analysis has indicated that normal mouse kidneys contain approximately 0.5–0.8  106 DCs (Steptoe et al., 2000) (Table 26.3), readily detected in the cortex and medulla and equivalent to ~25–30% of the DC content of mouse spleen. Human and rat glomeruli contain few MHC class II DCs (between one and three per glomerulus in the mouse and rat), whereas intertubular areas are rich in these cells, both in the cortex and medulla (Hart et al., 1981; Daar et al., 1983; Hancock and Atkins, 1984; Steiniger et al., 1984; Leszczynski et al., 1985a; Gieseler et al., 1997). In the mouse and rat, DCs exhibit the greatest density in the medullary region (Steiniger et al., 1984; Steptoe et al., 2000). Additionally, a cortico-medullary gradient of MHC class II expression has been reported in rat, with cortical DCs exhibiting the highest expression levels (Steiniger et al., 1984). Interestingly, in the bladder (Hart and Fabre, 1981b), MHC class II DCs are located beneath the epithelial layer, as in other epithelial surfaces. Despite expressing large quantities of MHC class II in situ, rat kidney DCs, like those in heart and pancreas, reportedly express no detectable quantities of invariant chain (Ii), which is in contrast to the high expression of Ii by DCs in lymphoid tissues (Saleem et al., 1997). As Ii expression is required for expression of antigen-bearing MHC class II molecules, this may be an important mechanism regulating the expression of self-antigens by DCs in peripheral nonlymphoid tissues. Immunohistochemical studies have indicated that kidney DCs are phenotypically similar to those in other nonlymphoid sites, and lack markers considered characteristic of macrophages (Daar et al., 1983; Hancock and Atkins, 1984; Kaisling and Le Hir, 1994; Gieseler et al., 1997). Actin-binding protein (p55), a marker of mature DCs, is expressed by few cells in normal human kidney (Sonderbye et al., 1998), thus

indicating the immature nature of DCs in this organ. Ultrastructurally, rat renal DCs resemble those in other nonlymphoid tissue sites, and are readily distinguished from co-existing macrophages and monocytes (Gaudecker et al., 1993), which they may outnumber (Kaisling and Le Hir, 1994). Austyn et al. (1994) reported that CD16/32 (FcγRII), CD11b and the macrophage antigen detected by the mAb F4/80 were present on freshly isolated MHC class II presumptive DCs from mouse kidney. However, following overnight culture, expression of these markers was reduced (Austyn et al., 1994). In contrast to DCs from lymphoid tissues, the DC-associated markers DEC-205 and 33D1 were not expressed by these cells when freshly isolated. Likewise, IL-2Rα (CD25) expression was absent (Austyn et al., 1994), as with other nonlymphoid tissue DCs. Kidney DCs do not phagocytose colloidal carbon in situ (Hart and Fabre, 1981a). They do phagocytose zymosan particles in vitro when freshly isolated, but, with less avidity than freshly isolated mouse cardiac DC (Austyn et al., 1994). The location of DC lining the bladder, ureter and in the vicinity of tubules within the kidney suggests a role for these cells in surveillance for invading pathogenic organisms. However, this hypothesis has not been examined in experimental models of urinary tract infection.

Population dynamics The population dynamics of renal DCs are similar to those of DCs in other nonlymphoid tissue sites. Irradiation and reconstitution studies have provided an estimate of population turnover in the order of 7–10 days in the rat (Leszczynski et al., 1985a). Systemic administration of IFN-γ is reported to result in an increase in MHC class II DCs in the kidneys of mice, with a 2–3-fold increase observed after 3 days of treatment. A slow reduction to near normal levels occurs within 2–3 weeks of cessation of treatment (Skoskiewicz et al., 1985). A similar pattern has been reported in the rat, however the extent of DC increase was not as dramatic (Leszczynski et al., 1986). Conversely, treatment with IFN-α and

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IFN-β results in a reduction of the density of DCs within the mouse renal cortex (Maguire et al., 1990). It is unclear whether these effects are mediated by altered DC trafficking patterns. Systemic LPS administration mobilizes DCs from the kidney, and also results in subsequent recruitment of MHC class II DC progenitors into this organ (Roake et al., 1995a, 1995b). These effects of LPS administration were attributed to mediation of DC migration by the cytokines TNF-α and IL-1α (Roake et al., 1995b). The ‘injury response’ associated with organ harvest and transplantation has been shown to induce the accumulation of DCs within kidney grafts subsequent to transplantation into syngeneic hosts (Penfield et al., 1999). No reports have detailed the specific role of chemokines in modulating DCs traffic through the kidney.

Ontogenic development Fewdetailsareavailableoftheontogenicdevelopment of DC populations in the kidney. In the human, MHC class II DCs first appear between 8 and 13 weeks gestational age, and increase in number between 12 and 21 weeks of gestation, at a rate similar to that in other nonlymphoid sites (heart,lung,pancreas)(Hofmanetal.,1984;Natali et al., 1984). Little is known of the ontogenic development of DCs in the kidney of rodents other than mice, in which MHC class II cells are absent until after birth (Natali et al., 1981).

Function An extensive and demanding study of the functional capacity of murine kidney DCs was performed by Austyn and colleagues (1994). The results indicated that freshly isolated renal DCs possessed little capacity to stimulate resting allogeneic T cells in MLR assays, or to provide accessory cell activity in oxidative mitogenesis. As with DCs from other sites, stimulatory capacity was acquired following overnight culture (Austyn et al., 1994). Similar observations have been reported for DCs isolated from rat glomeruli (Gieseler et al., 1997). Depletion of passenger leukocytes by ‘parking’

in allogeneic hosts (Lechler and Batchelor, 1982) or a protocol of cyclophosphamide and irradiation (McKenzie et al., 1984a) prior to kidney allografting has indicated that renal DCs are capable of providing a strong allogeneic stimulus in vivo. Velasco and Hegre (1989) have provided similar data implying that the absence of DCs from fetal rat kidneys renders these organs less immunogenic than those of adults. These studies provide in vivo evidence complementary to that derived from in vitro studies indicating the fully functional nature of kidney DCs when mature. The role of DCs in kidney allograft rejection has been utilized as a model system to demonstrate that ex vivo PUVA treatment (ultraviolet A irradiation of the donor organ in the presence of the photoactive substance 8-methoxypsoralen) may reduce the functional capacity of DCs within the grafts (Gaudecker et al., 1993). Efforts to deplete kidney DCs (e.g. use of anti-CD45 mAbs) prior to human organ transplantation have not, however, led to significant improvement in graft survival, suggesting that ‘indirect’ presentation of alloantigens by infiltrating host-derived DCs may be an important mechanism underlying elicitation of rejection.

HEART Histological location and phenotype Cardiac DCs are closely associated with the endocardial blood vessels and connective tissue and are generally aligned parallel to cardiac myocytes with their processes interdigitating between the myocytes (Hart and Fabre, 1981a; Steiniger et al., 1984) (see also Figure 26.1). DC density in normal ACI rat heart has been estimated as 55–60 DC/mm2 (Forbes et al., 1986; Darden et al., 1990) with a density range of approximately 10–75 DC/mm2 between strains (Darden et al., 1990). Total cardiac DC number in mice is low, approximately 0.4  106 cells per heart (Steptoe et al., 2000). In addition to DCs, a distinct population of resident tissue macrophages can be distinguished by their expression of the macrophagerestricted Ags recognised by the mAbs BMAC-5

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and ED2 in the rat (Spencer and Fabre, 1990) and also by the presence of nonspecific esterase activity (Hart and Fabre, 1981a). Rat cardiac DCs express variable levels of CD4 and the CD68-like antigen recognised by the mAb ED1, but not CD8 or the resident tissue macrophage antigen recognised by the mAb ED2 (Darden et al., 1990; Zhang et al., 1993; Suzuki, 1995). They are thus similar phenotypically to those in other peripheral tissues. Like DCs in rat kidney (vide supra), rat cardiac DCs do not express detectable quantities of invariant chain (Saleem et al., 1997). In mice, cardiac DCs express variable amounts of CD16/32 (FcγRII), CD11b and the panmacrophage marker recognised by the mAb F4/80, but not IL-2Rα (CD25) when freshly isolated, with little change observed in the phenotype of these cells following overnight culture (Austyn et al., 1994). DEC-205 and 33D1 are reported to be absent from freshly isolated cardiac DC from normal mice (Austyn et al., 1994), although CD11c has been reported on freshly isolated cells (Steptoe et al., 1997). In situ immunohistochemistry has also revealed the expression of CD11a, CD44 and macrosialin on murine heart DC (Austyn et al., 1994). Little phenotypic information is available for cardiac DCs in humans, but no differences were noted between these DCs and those in other nonlymphoid sites (Daar et al., 1983; Hancock and Atkins, 1984). Ultrastructural analysis indicates that Birbeck granules present in Langerhans cells of the epidermis are absent from cardiac DCs . While rat cardiac DCs do not phagocytose colloidal carbon in vivo (Spencer and Fabre, 1990), freshly isolated mouse cardiac DCs phagocytose zymosan particles in vitro with an avidity greater than that exhibited by MHC class II DCs freshly isolated from the kidney (Austyn et al., 1994).

Population dynamics Population turnover is rapid in rat heart, with a speedy decline in numbers seen following lethal irradiation, such that DCs are virtually absent after 5 days (Hart and Fabre, 1981a). Consistent

with this rapid turnover are observations that the MHC class II DC population rapidly reconstitutes following bone marrow transplantation (Hart and Fabre, 1981a; Leszczynski et al., 1985b). The prolonged presence of donor MHC class II cells in heart allografts parked in athymic nude or normal ACI rat recipients (~10% at 37 weeks) indicates that, in a similar fashion to Langerhans cells (Czernielewski et al., 1985; Chen et al., 1986; Czernielewski et al., 1987), a long-lived or immobile population of DCs may also be present in the heart (Burris et al., 1989). The accumulation of donor-derived DCs in the spleens of heart allograft recipients (Larsen et al., 1990) demonstrated the ability of mouse cardiac DCs to traffic to lymphoid tissue, providing evidence of the role of DCs in central priming of anti-allograft immunity. Mobilization of DCs from heart tissue is very rapid, with donorderived DCs detected in the spleens of recipient mice as early as 24 hours after heart transplantation. DC accumulation was maximal 2 to 4 days post-transplant (Larsen et al., 1990). The rapid reduction in the number of donor-derived DCs in recipient spleens after 4 days post-transplant indicated that donor DCs are required only to initiate the allograft rejection response and not to maintain the anti-allograft response, and is consistent with recent reports of rapid killing of DCs following interaction with T cells. Administration of the hematopoietic growth factor Flt3L has been shown to result in a 1.5-fold increase in the number of DCs in mouse cardiac tissue (Steptoe et al., 1997). As in the kidney, systemic administration of LPS to mice results in the mobilization of DCs from cardiac tissue and a subsequent accumulation, by recruitment, of MHC class II DC progenitors (Roake et al., 1995a, 1995b). It appears that this effect is mediated, at least in part, by TNF-α and IL-1α (Roake et al., 1995a) as reported for DCs in other tissue sites.

Ontogenic development In the human, MHC class II leukocyte populations appear to first develop in the heart

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between 13 and 17 weeks of gestation, relatively late compared to other nonlymphoid human tissues examined (Hofman et al., 1984).

Function Freshly isolated mouse heart DCs exhibit only a weak capacity to stimulate resting allogeneic T cells in MLR assays or to costimulate oxidative mitogenesis, but this activity is markedly increased following overnight culture (Austyn et al., 1994). Depletion of DCs from heart tissue allografts by anti-MHC class II mAb (Faustman et al., 1982) or from vascularized heart allografts by treatment with cyclophosphamide and total body irradiation (McKenzie et al., 1984b) prior to transplantation extends allograft survival. Conversely, augmentation of the number of cardiac DCs by Flt3L pretreatment of donors exacerbates heart allograft rejection (Steptoe et al., 1997). These results provide in vivo evidence of the potentially strong T-cell stimulating capacity of these nonlymphoid tissue DCs. A key issue that has intrigued transplant immunologists is whether DCs can prime T-cell alloreactivity peripherally within allografts in addition to within central lymphoid tissues. Using a rat vascularized heart allograft model, Forbes et al. (1986) demonstrated intragraft cell–cell interaction of donor MHC class II DC and graft-infiltrating T cells. As the CD4CD8 T cell ratio at day 4 post-transplant was approximately 2 :1, it is likely this represented productive DC–T-cell interactions rather than cytotoxic T cell-mediated killing of DCs. While further analysis is required to determine if these interactions lead to T-cell activation, this observation may indicate that DCs possess the ability to prime T-cell activation and proliferation peripherally (in nonlymphoid sites), at least when provided with the appropriate activation signals subsequent to transplantation.

Disease Experimentally induced myocardial infarction in rats results in accumulation of DCs in the

‘border zones’ of infarct sites (796 DC/mm2 versus 82/mm2 in normal hearts) 7 days postinfarct where they exhibit prominent clustering with CD4 T cells (Zhang et al., 1993). DC density then returns to relatively normal levels 21 days following the infarction event. Interestingly, within the center of the infarct lesion, only relatively minor changes in DC density occur. In an experimental model of actively induced experimental autoimmune myocarditis in rats, DCs expressing CD80 and CD86 are the first leukocytes observed infiltrating around small cardiac blood vessels following immunization with cardiac myosin (Suzuki, 1995; Dimitrijevic et al., 1998). Primary lesions are reported to subsequently form at these sites (Suzuki, 1995). At later timepoints, DCs may become less prominent as APC due to the influx of MHC class II macrophages (Dimitrijevic et al., 1998). It was suggested that in this model, DC mediated the initial antimyocardial destructive events and that the subsequent macrophage infiltration was a response to the localized tissue necrosis induced by DCs (Suzuki, 1995). Likewise, cardiac DC density has been observed to be markedly increased in the acute and subacute phases of human myocarditis (Yokoyama et al., 2000).

PANCREAS Histological location and phenotype MHC class II presumptive DCs are distributed throughout both the exocrine and endocrine portions of the pancreas. In humans, individual islets of Langerhans contain 0–6 intra-islet DCs (Leprini et al., 1987) which comprise approximately 1% of all islet cells (Lu et al., 1996). Overall, DCs located within islets constitute only a small proportion (approximately 4%) of the total MHC class II interstitial cells within normal human pancreata, the remainder located in the exocrine pancreas (Leprini et al., 1987). Human intra-islet DCs are restricted predominantly to the periphery of the islets, where these cells exhibit a more extensive dendriform morphology than those present in the

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exocrine pancreas (Leprini et al., 1987). In other species examined, for example the dog and pig (Shienvold et al., 1986), rat (Steiniger et al., 1984; Lloyd et al., 1987) and mouse (Steptoe et al., 2000), the distribution of DCs appears similar (Figure 26.1), although some predilection for blood vessels has been reported in the rat (Steiniger et al., 1984). Total pancreatic DC number in normal mice is approximately 0.3–0.4  106 per pancreas (Steptoe et al., 2000). The total number of DCs per islet appears similar between species, at least in human, rat, mouse and dog (Gebel et al., 1983; Shienvold et al., 1986; Leprini et al., 1987). In the rat, pancreatic DCs express variable levels of CD4, with a population of CD4/MHC II cells similar to those reported in heart (Steiniger et al., 1984; Darden et al., 1990) (vide supra). Human pancreatic DCs may be comprised of two phenotypically distinct subpopulations. The majority (approximately 70% of all MHC class II cells) are negative for surface and cytochemical markers of macrophages, exhibit a markedly dendriform morphology, while the remainder express some markers usually associated with macrophages and display an ovoid morphology (Leprini et al., 1987; Lu et al., 1996). The majority of human pancreatic DC express CD68 (Lu et al., 1996), while a proportion of intra-islet DCs express substantial levels of CD57 (Leprini et al., 1987)

Population dynamics There have been no reports on the population kinetics of pancreatic DCs.

Ontogenic development Examination of the development of MHC class II expression in the pancreata of rodents indicates that, as in many other nonlymphoid tissues, MHC class II leukocytes are absent until the time of parturition, but that adult-equivalent populations of DCs develop by 1 month postpartum (Fujiya et al., 1985). In larger animals, DC development in the pancreas occurs at an earlier time-point. In pigs, rare MHC class II

leukocytes have been observed in the exocrine portion as early as the 48th day of gestation (Fujiya et al., 1985). In humans, the presence of MHC class II presumptive DCs has been widely reported in human fetal pancreas between 14 and 20 weeks of gestation (Danilvos et al., 1982; Hofman et al., 1984; Oliver et al., 1988), but may develop as early as 8 gestational weeks (Koo Seen Lin et al., 1991). The rate of DC development reportedly undergoes rapid acceleration between 13 and 16 weeks of gestation (Hofman et al., 1984; Fujiya et al., 1985). The distinct populations of macrophages and DCs described in adults have also been observed in fetal pancreata (Koo Seen Lin et al., 1991; Jansen et al., 1993).

Function The functional activity of pancreatic DCs has not been examined directly in vitro and it is unclear whether these cells express T-cell stimulatory activity or maturational patterns equivalent to DCs from other nonlymphoid tissue sites. The strongest evidence of the T-cell stimulatory activity of pancreatic DCs has been provided by transplantation studies. Depletion of MHC class II presumptive DCs from islets by anti-MHC class II (Faustman et al., 1981, 1982; Lloyd et al., 1987) or anti-DC mAb pretreatment (Faustman et al., 1984, 1985) prior to allotransplantation extends the subsequent survival time of these pancreatic islet grafts. Likewise, when pancreatic islets were treated with UV irradiation (which inhibits the allostimulatory capacity of DCs) prior to allotransplantation, the survival time of the grafts was extended (Lau et al., 1984a, 1984b). DCs isolated from pancreatic lymph nodes (PLN) of prediabetic nonobese diabetic (NOD) mice exhibit extensive clustering with T cells derived from NOD PLN, but not with T cells from nonpancreatic LN of NOD mice or PLN from nondiabetic mouse strains (Clare-Salzler and Mullen, 1992). This apparently indicates the carriage of pancreatic islet-derived antigen by pancreas-emigrating DCs that subsequently interact with Ag-specific T cells. Further evidence of transport of islet-derived antigen is

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REFERENCES

provided by studies in which subcutaneous injection of PLN-derived DCs protects NOD mice from subsequent diabetes development via the induction of regulatory T cells (ClareSalzler et al., 1992). It is currently unclear whether pancreas-derived DCs are responsible for the antigen transport that results in crosstolerance as described by Heath and co-workers (reviewed in Heath et al., 1998), or whether another APC/cell type or passive transfer is responsible.

Disease DCs are a prominent cell type in the leukocytic infiltrate observed in pancreatic islets of rodent models of diabetes. In prediabetic BB rats, MHC class II dendriform cells accumulate at the margins of islets of Langerhans in elevated numbers prior to the infiltration of islets by T lymphocytes and macrophages (Voorbij et al., 1989; Ziegler et al., 1992). In NOD mice, MHC class II DC are the first hematopoietic cells to accumulate around and infiltrate into pancreatic islets during the early stages of insulitis (Jansen et al., 1994) and have been identified, along with macrophages, as the earliest source of TNF-α in NOD islets (Dahlen et al., 1998). It appears that islet-infiltrating DEC-205 DCs exist as a network of cells expressing VCAM-1 and ICAM-1 around which infiltrating lymphocytes are organized (Lo et al., 1993). Infiltration of islets by DCs in NOD.SCID mice is somewhat reduced, suggesting a degree of dependency on T cells for maximal DC infiltration (Dahlen et al., 1998). Interestingly, in nonobese diabetesresistant (NOR) mice, that are MHC-identical to NOD mice, islet infiltration exhibits a protracted block at the earliest stages of infiltration, without subsequent T-cell infiltration or T cellderived inflammatory cytokine production (Fox and Danska, 1998). Transgenic expression of TNF-α in the islets of neonatal NOD mice exacerbates diabetes development, apparently via enhanced (DC-mediated) presentation of isletderived antigens (Green et al., 1998). It is therefore interesting to speculate that activation of intra-islet DCs via interaction with T cells (or T

cell-derived cytokines) or other inflammatory cytokines, such as TNF-α, may be crucial to the eventual development of invasive insulitis and diabetes in susceptible rodents, or conversely, tolerance in nondiabetes prone animals.

Acknowledgements The authors’ work is supported by National Institutes of Health grants DK 57228-01 (RJS), DK 49745 and AI 41011 (AWT) and by the Roche Organ Transplantation Research Foundation (AWT). We thank Drs. A.E. Morelli and S. Goddard for valued input and discussion.

REFERENCES Arber, D.A., Weiss, L.M. and Chang, K.L. (1998). Semin. Diagn. Pathol. 15, 155–160. Austyn, J.M. (1996). J. Exp. Med. 183, 1287–1292. Austyn, J.M., Hankins, D.F., Larsen, C.P., Morris, P.J., Rao, A.S. and Roake, J.A. (1994). J. Immunol. 152, 2401–2410. Ballardini, G., Fallani, M., Bianchi, F.B. and Pisi, E. (1989). Autoimmunity 3, 135–144. Bardadin, K.A. and Desmet, V.J. (1984) Histopathology 8, 657–667. Bishop, G.A., Sun, J., DeCruz, D.J. et al. (1996). J. Immunol. 156, 4925–4931. Brenan, M. and Puklavec, M. (1992). J. Exp. Med. 175, 1457–1465. Burris, D.E., Gruel, S.M. and Rao, U.K. (1989). Transplantation 47, 1085–1087. Chen, H.-D., Chenglin, M., Yuan, J.-T., Wang, Y.-K. and Silvers, W. (1986). J. Invest. Dermatol. 86, 630–633. Chen, S., Akbar, S.M., Tanimoto, K. et al. (2000). Cancer Lett. 148, 49–57. Clare-Salzler, M. and Mullen, Y. (1992). Immunology 76, 478–484. Clare-Salzler, M.J., Brooks, J., Chai, A., Van Herle, K. and Anderson, C. (1992). J. Clin. Invest. 90, 741–748. Czernielewski, J., Vaigot, P. and Prunieras, M. (1985). J. Invest. Dermatol. 84, 424–426. Czernielewski, J.M. and Demarchez, M. (1987). J. Invest. Dermatol. 88, 17–20. Daar, A.S., Fuggle, S.V., Hart, D.N.J. et al. (1983). Transplant. Proc. 15, 311–315. Dahlen, E., Dawe, K., Ohlsson, L. and Hedlund, G. (1998). J. Immunol. 160, 3585–3593.

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Danilvos, J.A., Hofman, F.M., Taylor, C.R. and Brown, J. (1982). Diabetes 31 (Suppl), 23–28. Darden, A.G., Forbes, R.D., Darden, P.M. and Guttmann, R.D. (1990). Cell. Immunol. 126, 322–330. Demetris, A.J., Qian, S., Sun, H. et al. (1991). Am. J. Pathol. 138, 609–618. Demetris, A.J., Sever, C., Kakizoe, S., Oguma, S., Starzl, T.E. and Jaffe, R. (1989). Am. J. Pathol. 134, 741–747. Dimitrijevic, M., Milenkovic, P., Milosavljevic, P. and Colic, M. (1998). Transplant. Proc. 30, 4149–4150. Drakes, M.L., Lu, L., McKenna, H.J. and Thomson, A.W. (1997a). Adv. Exp. Med. Biol. 417, 115–120. Drakes, M.L., Lu, L., Subbotin, V.M. and Thomson, A.W. (1997b). J. Immunol. 159, 4268–4278. Drakes, M.L., Zahorchak, A.F., Takayama, T., Lu, L. and Thomson, A.W. (2000). Transplant. Immunol. 8, 17–29. Faustman, D.L., Hauptfeld, V., Lacy, P. and Davie, J. (1981). Proc. Natl Acad. Sci. USA 78, 3864–3868. Faustman, D., Kraus, P., Lacy, P.E., Finke, E.H. and Davie, J.M. (1982). Transplantation 34, 302–305. Faustman, D.L., Steinman, R.M., Gebel, H.M., Hauptfeld, V., Davie, J.M. and Lacy, P.E. (1984). Proc. Natl Acad. Sci. USA 81, 3864–3868. Faustman, D.L., Steinman, R.M., Gebel, H.M., Hauptfeld, V., Davie, J.M. and Lacy, P.E. (1985). Transplant. Proc. 17, 420–422. Forbes, R.D.C., Parfrey, N.A., Gomersall, M., Darden, A.G. and Guttmann, R.D. (1986). J. Exp. Med. 164, 1239–1258. Forsum, U., Klareskog, L. and Peterson, P.A. (1979). Scand. J. Immunol. 9, 343–349. Fox, C.J. and Danska, J.S. (1998). Diabetes 47, 331–338. Fujiya, H., Danilovs, J., Brown, J. and Mullen, Y. (1985). Transplant. Proc. 17, 414–416. Gebel, H.M., Yasunami, Y., Diekgraefe, B., Davie, J.M. and Lacy, P.E. (1983). Transplantation 36, 1592–1600. Gieseler, R., Hoffmann, P.R., Kuhn, R. et al. (1997). Scand. J. Immunol. 46, 587–596. Goddard, S., Hubscher, S.G., Lane, P., and Adams, D.H. (2000). Transplantation 69, (Suppl.) S250 (Abstract 534). Gorczynski, R.M., Cohen, Z., Fu, X.M., Hua, Z., Sun, Y. and Chen, Z. (1996). Transplantation 62, 1592–1600. Gorczynski, L., Chen, Z., Hu, J. et al. (1999). J. Immunol. 162, 774–781. Green, E.A., Eynon, E.E. and Flavell, R.A. (1998). Immunity 9, 733–743. Hancock, W.W. and Atkins, R.C. (1984). Transplant. Proc. 16, 963–967. Hardonk, M.J., Dijkhuis, F.W., Grond, J., Koudstaal, J. and Poppema, S. (1986). Virch. Arch. B Cell. Pathol. Incl. Mol. Pathol. 51, 429–442.

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PLATE 26.3 Distribution of DCs, and their expression of costimulatory molecules in normal mouse liver (A and C) and in Flt3L-treated liver (B and D). In A and B, tissue sections were double immunolabeled with anti-CD11c PE (DC in red) plus anti-CD40 FITC (double positive cells in yellow: red  green). In C and D, sections were double immunolabeled with anti-CD11c PE (DC in red) plus anti-CD86 FITC. (A) In control livers, DCs are evident as single cells in the perivenular region (asterisk) and the portal space, and do not express CD40. (B) After 10 days Flt3L treatment, the number of DC increased dramatically, large cell aggregates were associated with the central vein (asterisk) and the portal spaces. Very few DCs expressed CD40 (double positive cells in yellow, indicated by arrows). (C) In control liver, DCs did not express CD86. (D) After Flt3L treatment, DC remained as immature APC lacking expression of CD86 (no double positive cells in yellow were detected). Cell nuclei were counterstained with DAPI (deep blue). Double immunofluorescence, 200. Staining and micrograph courtesy of Dr A.E. Morelli, from Morelli et al. (2000). Copyright  2000 Lippincott Williams & Wilkins.

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27 Dendritic cells in the spleen and lymph nodes Bali Pulendran, Karolina Palucka and Jacques Banchereau Baylor Institute for Immunology Research, Dallas, TX, USA

The known is finite, the unknown infinite; intellectually we stand on an islet in the midst of an illimitable ocean of inexplicability. Our business in every generation is to reclaim a little more land. T.H. Huxley

INTRODUCTION

give rise to DC precursors that circulate in the blood, lymphatics and lymphoid tissues. DC precursors that home to the tissues reside there as immature DCs, which are efficient at capturing antigens through several mechanisms including phagocytosis and endocytosis. Infection, and the subsequent recognition of pathogen products by these immature DCs, can trigger a maturation program, where they migrate to the secondary lymphoid microenvironments, harboring antigens of the pathogen. When they reach the secondary lymphoid organs, they are now mature DCs and so can present their antigens to T cells, to initiate T-cell immunity. In the present review, we will focus on DCs in secondary lymphoid organs. Much of our present understanding has come from in situ studies in rodents, so these studies represent the focus of this review. However, we will allude to several emerging studies in the human system to illustrate similarities between the various systems.

Secondary lymphoid microenvironments are immunological crucibles in which adaptive immunity is initiated. It is there that dendritic cells (DCs), present to the naïve T cells, antigens they have acquired elsewhere (Ingulli et al., 1997). This antigen-presentation triggers a cascade of events that culminates in the clonal expansion of antigen-specific T and B cells, and ultimately the establishment of immunological memory. Given this pivotal role in the adaptive immune response, DCs occupy a center stage of modern immunological research. Today we know that DCs represent a complex system of cells with multiple subsets, which may represent cells of distinct lineages, as well as cells at different maturation stages within a single lineage (Banchereau and Steinman, 1998). Our present view of DC development can be summarized as follows: DCs develop from committed progenitors in the bone marrow, which

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DEVELOPMENT OF DENDRITIC CELLS Ontogenic heterogeneity DC development in vivo is a complex interplay between distinct DC lineages and specific microenvironmental signals (Banchereau and Steinman, 1998). One of the most striking aspects of DC biology is the plethora of DC phenotypes that have been described both in vitro and in vivo, in murine and human studies. Despite much effort, understanding DC development in vivo has been difficult for several reasons: (1) DCs occur at very low frequencies in vivo making it difficult to obtain sufficient numbers for detailed analysis. Therefore intense efforts have focussed on studying various DC subsets grown in vitro. (2) It is not clear which of the in vitro generated DC subsets actually occur in vivo. This is especially true in the case of in vitro generated human DC subsets, because the corresponding in vivo experiments in human lymphoid tissues are difficult to perform. (3) Subtle changes in a particular in vitro culture system can yield DCs with slightly distinct phenotypes. In such cases, it is never clear whether to consider these DCs to be the same or different. Despite these problems, at least two distinct ontogenic pathways for DC development have been proposed. Thus, DCs have been postulated to arise from either myeloid or lymphoid-committed hematopoeitic precursors (Hart 1997; Banchereau and Steinman, 1998; Shortman et al, 1998;) and these developmental pathways are discussed below (see Chapters on myeloid DC and thymic DCs). Recent studies, however, have challenged the notion of lineage restriction of DC subsets (Traver et al., 2000).

A model for DC development: postulated myeloid and lymphoid lineages DCs from postulated myeloid lineage Hematopoietic progenitor cells (HPC) which give rise to cells of myeloid lineages, including

granulocytes and macrophages, can also give rise to Langerhans cells and myeloid-related DCs in secondary lymphoid tissue (Hart, 1997; Banchereau and Steinman, 1998). Evidence for the myeloid origins of DCs comes predominantly from in vitro studies, in which myeloid DCs can be generated in the presence of GM-CSF and TNFα, from either bone marrow or cord blood progenitors, or from blood precursors (Caux et al, 1992; Inaba et al., 1992a, 1992b; Romani et al., 1994; Sallusto and Lanzavecchia, 1994; Young et al., 1995; Caux and Banchereau, 1996; Steinman et al., 1998). In mice, injections of a polyethylene glycol modified form of GMCSF has been shown to expand DCs which resemble myeloid DCs in vivo (Pulendran et al., 1999; Daro et al., 2000). However, mice deficient in GM-CSF or GM-CSF-Rβ do not have reduced numbers of DCs in lymphoid tissues, suggesting that GM-CSF is not essential for DC development in vivo (Vremec et al, 1997a). In mice and in humans, the myeloid DC lineage has been subdivided into the Langerhans and interstitial DC pathways (reviewed by Banchereau and Steinman, 1998). In vitro cultures of human CD34HPCs yield precursors of two DC populations: Langerhans cells (located in vivo, in epidermis and epithelia of skin and mucosal tissues) and interstitial DCs (located in vivo in the dermal layer of skin and in the tissue interstices) (Caux and Banchereau, 1996; Caux et al., 1996). The addition of maturation signals such as CD40-L, induces further maturation of these precursors into mature DCs. In vivo, the precursors of Langerhans and interstitial DCs are likely to be the CD11c HLA-DR lineage-ve precursors, found in the peripheral blood of humans (Ito et al., 1999; Palucka et al., – in preparation). When cultured in vitro with GMCSF  TNFα, these CD11c precursors give rise to mature, myeloid DCs (Ito et al., 1999; Cella et al., 1999; Pulendran et al., 2000). When cultured with GM-CSF  IL-4 in the presence of TGFβ, they give rise to Langerhans cells (Ito et al., 1999). Gene targeting studies in mice suggest that the developmental pathways of Langerhans cells and interstitial ‘myeloid’ DCs are regulated by

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different genes. Thus, mice deficient in the NFκB like transcription factor Rel B, have reduced numbers of CD8α myeloid DCs (Wu et al., 1998; Crowley and Lo, 1999). In contrast, TGFβ/ mice have reduced numbers of Langerhans cells (Borkowski et al., 1996). However, some recent studies challenge the notion that mouse Langerhans cells are of myeloid origin, since they acquire CD8α expression (a lymphoid marker) upon maturation in vivo (Anjuere et al., 2000; Merad et al. 2000). DCs from postulated lymphoid precursor populations DCs have also been postulated to develop from lymphoid-related precursor populations. In vivo studies in mice demonstrate that transfer of purified, thymic precursors into irradiated hosts, results in the development of T cells, B cells, NK cells and CD8α DCs, but not cells of the myeloid lineage (Vremec et al., 1992; Ardavin et al., 1993; Wu et al., 1996; Shortman et al., 1998). However, there is no clonal evidence to suggest that the same precursor cell that gives rise to lymphoid cells, also yields CD8α DCs. Therefore, these cells have been termed the ‘putative-lymphoid related DCs’ (Shortman et al., 1998). The early thymic precursors can also generate thymic DCs in vitro, when cultured with a combination of cytokines such as IL-1β, IL-3, IL-7, TNF, Flt3-Ligand (FL), c-kit ligand (KL) and CD40 ligand (CD40-L) (Saunders et al., 1996). The conspicuous absence of GMCSF, a cytokine which is essential for the in vitro development of myeloid precursor-derived DCs suggests that GM-CSF is not required for this pathway of DC development (Saunders et al., 1996). In contrast to these findings, a recent study suggests that CD8α DCs can develop from a common myeloid progenitor in vivo (Traver et al. 2000). Thus the lineage origins of DCs is currently a matter of intense debate, and the findings by Traver et al. may reflect a plasticity in the developmental fate of DC precursors. In the spleens and lymph nodes of mice, DC subsets with a similar surface phenotype to CD8α thymic DCs exist, but the lineage origins

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of these is not known. In this review, we will simply refer to these DCs as ‘CD8α DCs’. In humans, a subpopulation of HPCs expressing CD10 and committed to lymphoid lineage, can yield DCs in vitro suggesting the existence of human lymphoid-related DCs (Galy et al., 1995). Again, clonal evidence is lacking. Similarly, DCs have also been cultured from human thymic precursors (Res et al., 1996). However, these putative human lymphoid-related DCs are phenotypically indistinguishable from their myeloid DC counterparts, based on the expression of classical lineage-specific markers such as CD2, CD4 and CD33. Furthermore, unlike murine DCs, human thymic DCs do not express high levels of surface CD8α, but rather CD4 (Sotzik et al., 1994; Winkel et al., 1994). Further speculation about the lymphoid origins of DCs in humans, comes from the recently characterized plasmacytoid CD11c DC precursors in the blood originally described as plasmacytoid T cells or plasmacytoid monocytes (MullerHermelink et al., 1983; Prasthofer et al., 1985; Facchetti et al., 1989). These cells die rapidly after isolation, and are critically dependent of IL-3 for survival and CD40-L for maturation (Grourad et al., 1996a). The resulting DCs do not express CD11b, CD13 and CD33, surface markers classically considered as myeloid (Grourad et al., 1996a). A phenotypically similar population of adult blood cells, have been shown to express the pre-TCR and to contain precursors of mature CD4TCRα/β T cells (Bruno et al., 1997), hence the presumption of lymphoid origin of CD11c DC. Whether the T cells and the CD11c DCs actually develop from the very same precursor cell is not known.

Cytokines that regulate DC development in vitro While the understanding of DC biology has progressed through the analysis of DC subsets generated in vitro, the analysis of DC subsets in vivo has been hampered by the lack of specific growth factors in vivo. However, the demonstration that cytokines such as Flt-3 ligand, GM-CSF and G-CSF can expand various DC subsets

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in vivo, has created new opportunities for investigating DC functions in vivo (Maraskovsky et al., 1996; Pulendran et al., 1997, 1999, 2000; Shurin et al., 1997; Arpinati et al., 2000; Maraskovsky et al., 2000; Daro et al., 2000). The roles played by these cytokines are considered below: Flt-3 ligand Administration of the hematopoietic growth factor Flt-3 ligand (FL) (McKenna et al., 1995; Brasel et al., 1996; Lyman and Jacobson, 1998) to mice, results in a dramatic expansion of both CD8α and CD8α DCs at multiple sites including spleen, lymph nodes, thymus, liver, lung, bone marrow, peripheral blood, gut-associated lymphoid tissues and peritoneal cavity (Maraskovsky et al., 1996; Pulendran et al., 1997; Shurin et al., 1997). FL appears to mediate its effects by targeting primitive hematopoietic progenitors in the bone marrow and inducing their expansion and differentiation under the influence of additional molecular interactions. Both CD8α and CD8α DCs are expanded, but the relative representation of these DC subsets are preserved in FL-treated mice (Maraskovsky et al., 1996; Pulendran et al., 1997). Consistently, mice deficient in FL have reduced numbers of both CD8α and CD8α DCs in various organs (McKenna et al., 2000). The properties of these subsets are discussed below, under the heading: ‘DCs in distinct microenvironments’. In mice injected with FL, antigen-specific T- and B-cell responses (Pulendran et al., 1998), as well as antitumor responses (Lynch et al., 1997) are dramatically enhanced. Similarly, recent studies in humans demonstrate that FL expands both DC precursors and immature DCs in blood (Maraskovsky et al., 2000; Pulendran et al., 2000). In particular, the numbers of CD11c immature DCs, as well as the plasmacytoid CD11c DC precursors are dramatically increased. It now remains to be seen whether this enormous expansion of DCs causes enhanced immune responses to vaccine antigens, as observed in FL-treated mice (Pulendran et al., 1998).

GM-CSF Initial studies of injecting GM-CSF into mice did not reveal any effects on DC development (Maraskovsky et al., 1996); however subsequent studies with a chemically stabilized form of GM-CSF (pegylated) has revealed its potent effects in expanding CD8α DCs in the spleen (Pulendran et al., 1999; Daro et al., 2000). Mice treated with pegylated GM-CSF display dramatically enhanced antibody responses to soluble antigens (Pulendran et al., 1999). Functionally and phenotypically, the CD8α DCs expanded by GM-CSF appear similar to those expanded by FL in many respects (Pulendran et al., 1999; Daro et al., 2000). These studies suggest that GM-CSF can play a role in CD8α DC development in vivo. However, as mentioned above, GM-CSF does not appear to be obligatory for DC development in vivo, as mice deficient in GM-CSF or GM-CSFRβ have normal numbers of DCs (Vremec and Shortman, 1997). Furthermore, overexpression of a GM-CSF transgene in mice does not significantly increase DC numbers in peripheral lymphoid organs, although DC numbers in the lung and peritoneal cavity are increased (Metcalf et al., 1996). The effect of GM-CSF in expanding DCs in humans, has not yet been investigated in detail. Nevertheless, GM-CSF is well established as a factor which mobilizes CD34HPCs (known to contain DC progenitors) (Fischmeister et al., 1999), and systemic GM-CSF administration leads to increased numbers of blood monocytes (DC precursors) (Avigan et al., 1999). It remains to be demonstrated whether its potent effects as a vaccine adjuvant, in eliciting antibody and CTL responses against peptides and proteins in humans, (e.g. Jager et al., 1996) are mediated via DCs.

G-CSF: a novel growth factor for DCs in humans Similarly to initial studies with GM-CSF, administration of G-CSF into mice was not observed to expand DCs in the spleen (Maraskovsky et al., 1996). Other organs were not examined in this study. As is the case with pegylated GM-CSF

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(Pulendran et al., 1999; Daro et al., 2000), it is possible that a stabilized form of G-CSF might indeed expand DC subsets. Recent work suggests that G-CSF injections into healthy donors, results in a dramatic increase in the numbers of plasmacytoid CD11c DC precursors in the peripheral blood (Arpinati et al., 2000; Pulendran et al., 2000). Phenotypically and functionally, the DCs derived from these precursors appear similar to those generated by FL injection. However, unlike the case with FL-treated donors, the numbers of CD11c precursors were not increased in G-CSF donors. Thus FL and G-CSF appear to expand distinct DC subsets, or DC precursors, in the peripheral blood of humans. Overall, these studies suggest that different cytokines can expand distinct DC subsets or DC precursors in vivo, in mice and in humans. As will be discussed below, in mice these cytokines appear to regulate immune responses differently. Whether and how this regulation applies to human remains to be determined.

DENDRITIC CELLS IN LYMPHOID MICROENVIRONMENTS The heterogeneity of DCs and macrophages in lymphoid tissues, has long been recognized by pathologists and histologists, but we are only beginning to isolate these various subsets of antigen presenting cells, to characterize their functional properties. In rodents, rapid progress is being made in this regard, but in humans a thorough investigation of the various DC subsets in the spleen, lymph nodes and the thymus are eagerly awaited. Our current knowledge is summarized below.

Spleen and lymph nodes Mice (Plate 27.1 and Table 27.1) In mice, at least four subsets of DCs can be identified in the spleen and lymph nodes. These include: Subset 1: CD11c CD8α DEC-205

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CD11b/dull (in spleen and lymph nodes), termed ‘CD8α DCs’. Subsets 2 and 3: CD11c CD8α DEC205 CD11bbright, which can be subdivided into a 33D1 CD4 subset (subset 2), and a 33D1ve CD4-ve subset (subset 3), (in spleen and lymph nodes), termed collectively as ‘CD8α DCs’. Subset 4: CD11c CD8α/ DEC205 CD11bbright, termed as ‘Langerhans cells DCs’. Most authors classify subsets 2 and 3 together as a single myeloid-related DC subset (Vremec et al., 1997; Pulendran et al., 1997; Reis e Sousa et al., 1997; Maldanado et al., 1999; Ohteki et al., 1999). Subset 1 may be related to the CD8α ‘putative-lymphoid-related DCs’ in the thymus (Ardavin et al., 1993), and so has often been operationally defined as ‘lymphoid DCs’. Langerhans cells DCs (subset 4) can only be identified in the lymph nodes (Salomon et al., 1998; Anjuere et al., 1999). These various DC subsets differ with respect to their phenotype, function and localization in distinct microenvironments. The CD8α DC subsets are localized deep in the T-cell rich areas of the periarteriolar lymphatic sheaths (PALS) (De Smedt et al., 1997; Pulendran et al., 1997). In contrast, the CD8α DC subsets are localized in the marginal zone areas of the spleen (De Smedt et al., 1996; Pulendran et al., 1997). Activation of the CD8α DC subsets, by injection of LPS or toxoplasma extracts, causes their migration from the marginal zones to the T-cell rich PALS (De Smedt et al., 1997; Reis e Sousa et al., 1997). In lymph nodes, the CD8α subset likely resides in the subcapsular sinus, and also in the PALS region, neighboring the B-cell follicles. As demonstrated above, phenotypically these four subsets exhibit striking differences, but also express similar levels of molecules involved in antigen presentation such as MHC class II, CD40, CD80, CD86 (Table 27.1). Interestingly, the only known marker that can distinguish splenic CD8α DCs in the spleen from thymic DCs is a BP-1 glutamyl aminopeptidase. The function of all these subsets have not yet been explored completely, but the current status of our knowledge is described below. Most studies have focussed on the CD8α and CD8α subsets in the spleen and, to our knowledge, no

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DC subsets in the secondary lymphoids organs of mice CD8α DCs (putative lymphoid)

CD8α DCs (33D1 and 33D1-subsets) (putative myeloid)

Langerhans cell derived DCs

Phenotype

CD11C CD8α DEC205 CD11bdull/ 33D1 CD86 CD40 Class II MHC Birbeck granule Lag

CD11c CD8α DEC205 CD11b 33D1(subset) CD86 CD40 Class II MHC Birbeck granule Lag

CD11c CD8α (CD8α?)* DEC205 CD11bdull 33D1 CD86 CD40 Class II MHC Birbeck granule Lag

Localization

PALS of spleen lymph nodes and Peyer’s patch Thymic cortex

Marginal zones of spleen LPS induces migration from marginal zones to PALS Subepithelial dome of Peyer’s patch Subcapsular sinus of lymph nodes T cell areas of lymph nodes

Langerhans cell derived DCs in T cell areas of lymph nodes Immature Langerhans cells in epithelia of skin and mucosa

Function IL-12 production IFNγ production Ag capture Ag processing CD4-T cell priming: (a) in vitro (b) in vivo CD8-T cell priming (a) in vitro (b) in vivo



/







?

/ 

 

Immature LC (); LC-DC (?) 

  (Th1)

  (Th2)

 

 

  ? ?

* Some recent studies by Anjuere et al. (2000) and Herad et al. (2000) suggest that Langerhans cells acquire CD8α expression during maturation.

study has examined the function of the 33D1 CD4 and 33D1 CD4 subsets separately. Antigen uptake and processing Both the CD8α and CD8α DC subsets from the spleen can take up soluble ovalbumin and dextran equally efficiently (Pulendran et al., 1997; Daro et al., 2000). However, the CD8α DC subset is significantly more efficient at

taking up particulate antigens, such as zymosan (Pulendran et al., 1997; Leenen et al., 1998). Both the CD8α and CD8α DC subsets from the spleen process soluble antigens equally efficiently (Daro et al., 2000). Costimulatory molecules Both DC subsets express similar levels of class II MHC, CD40 and CD86, even in their resting

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state (Pulendran et al., 1997; Daro et al., 2000). These DCs do not express CD80, unless they are activated (Pulendran et al., 1997; Daro et al., 2000). Cytokine production The CD8α DC subset, but not the CD8α DC, can be induced to secrete high levels of biologically active IL-12, by Staphylococcus aureus Cowans (Pulendran et al., 1997; Maldanado et al., 1999); Toxoplasma extract (Reis e Sousa et al., 1997), Listeria (Otekhi et al., 1999), and LPS (Pulendran et al. submitted). Furthermore, IL-12 production by the CD8α DCs elicits IFNγ production in that DC subset (Ohteki et al., 1999). Priming of T cells (1) In vitro: Both the CD8α and CD8α DC subsets from the spleen can prime allogeneic CD4 and CD8 T cells in vitro (Süss and Shortman 1996; Kronin et al., 1996; Daro et al., 2000). In some studies, the CD8α DC subset appears to prime the allogeneic CD4 and CD8 T cells less efficiently than the CD8α DCs, in vitro (Süss and Shortman 1996; Kronin et al., 1996). In these studies, the CD8α DCs were observed to induce apoptosis in a fraction of the CD4 T cells (Süss and Shortman 1996), and to be unable to support cytokine production in CD8 T cells (Kronin et al., 1996). Both subsets can prime naïve, antigen-specific CD4 T cells equally efficiently (Daro et al., 2000). (2) In vivo: Both the CD8α and CD8α DC subsets from the spleen, pulsed with soluble peptides or proteins, can prime antigen-specific CD4 T cells in vivo (Pulendran et al., 1999; Maldanado et al., 1999). After in vivo priming, the CD8α lymphoid DCs elicit Th1 responses, in naïve, antigen-specific CD4 T cells, while the CD8α myeloid DCs elicit Th2 or Th0 responses (Pulendran et al., 1999; Maldanado et al., 1999). Both subsets can prime CD8 T cells in vitro (Daro et al., 2000) and in vivo (Ruedl and Bachmann, 1999). Recently, a novel subset of dendritic-like cells was shown to enter the B-cell follicles during the first few days of a primary immune response,

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and prime antigen-specific B and T cells (Berney et al., 1999). Phenotypically, this population does not resemble any of the populations described above. However, its localization in B-cell follicles makes it a likely counterpart of the germinal center dendritic cells (GCDCs), which we will discuss below. Humans Immunohistochemical studies of DCs in the spleens and lymph nodes of humans have revealed an exquisite diversity of DC subsets in the red-pulp, marginal zones and PALS of the spleen (Wood et al., 1985; Buckley et al., 1987; Takahashi et al., 1998). As with mice, different subpopulations of macrophages and dendritic cells seem to occupy discrete microenvironments. DC-like cells can be observed in the T-cell rich areas of the PALS, B-cell rich follicles and in the follicular mantle. However, it is not clear which of these subsets are actually dendritic cells, and the correlation between these subsets and those described in vitro, or in the peripheral blood of humans, is under investigation. In humans, DC subpopulations have been characterized best in tonsils, with at least three subsets: two in the PALS, and one in the B-cell rich follicles. These are: (1) CD40CD86CD83CD11c interdigitating DC (IDC) within the T-cell rich areas (Bjorck et al., 1997). IDCs at different stages of maturation, as determined by CD83 expression, can be found in tonsils, lymph nodes and spleen. (2) CD4CD3CD11c DCs which are present in T-cell rich areas and locate around and within high endothelial venules of reactive lymph nodes or tonsils (Grouard et al., 1996a; Cella et al., 1999). The precursors of these CD4 CD3 CD11c DCs are also found in the blood (O’Doherty et al., 1994; Grouard et al., 1996a; Rissoan et al., 1999; Cella et al., 1999), and are otherwise known as plasmacytoid T cells or plasmacytoid monocytes (Muller-Hermelink et al., 1983; Prasthofer et al., 1985; Facchetti et al., 1989). These CD11c precursor cells represent a unique subset, shown recently to release large

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as the proliferation of CD40-activated B cells, in the presence of IL-12 (Dubois et al., 1997, 1999). Thus, the compartmentalization of these DC subsets in discrete human lymphoid microenvironments suggests distinct functions. These DC subsets are discussed in detail elsewhere in this volume (see Chapters on myeloid DCs, lymphoid DCs and thymic DCs). Here we would like to summarize briefly our current view of differentiation pathways of blood DC precursors and their mature progeny (Figure 27.2). Recent data show that monocytes constitute immediate precursors of CD11c DC (Palucka et al. submitted). These CD11c DCs can give rise to Langerhans cells, interstitial DC or macrophages, depending on cytokine environment, in vitro (Ito et al., 1999). The mature progeny of LCs and intDCs may home to different microdomains in secondary lymphoid organs, which however the presently available tools do not permit to conclude as the origin of IDCs and GCDCs.

amounts of IFNα, when exposed to viruses (Siegal et al., 1999). Furthermore, these DCs are able to induce T cells to secrete IL-4 (type 2 responses) (Rissoan et al., 1999), and high levels of IL-10, suggesting a possible regulatory function (Rissoan et al., 1999; Pulendran et al., 2000). (3) CD4CD3CD11c germinal center DC (GCDC) can be identified in the germinal centers of human tonsils (Grouard et al., 1996b). They represent less than 1% of germinal center cells, and express Fcγ receptors such as CD16, CD32 and CD64, as well as complement receptors (CR1, CR2 and CR3). This phenotype may explain their efficient binding of immune complexes. GCDCs may represent the mature progeny of blood CD11c precursors, as both uniquely express the metalloproteinase decysin, which is involved in the cleavage of TNF-family molecules (Mueller et al., 1997). In vitro, GCDCs can interact with both T cells and B cells. Thus, they can induce naïve T cell proliferation, as well

CD14 CD11c

Cytokine

CD14 CD11c

M-CSF

GM-CSF TNF/IL-4

Macrophage

••••••••

Interstitial DC

CD123 CD11c IL-3

TGFβ

••••••••

Langerhans DC

••••••••

Plasmacytoid DC

FIGURE 27.2 Dendritic cell subsets in humans. Two subsets of dendritic cell precursors circulate in the blood, CD11c DC precursors and CD11c CD123 DC precursors. CD11c precursors originate from monocytes, under the influence of variety of cytokines, e.g. IL-3 (Palucka et al. – unpublished). These cells can give rise to three types of antigen presenting cells: Langerhans cells, interstitial DCs and macrophages. It is thought that interdigitating DCs (IDCs, which are found in the PALS) and germinal center DCs (GCDCs) may represent a mature progeny of Langerhans cells and interstitial DCs, respectively. Nevertheless, present tools do not permit any conclusions about the origin of mature DCs found in secondary lymphoid organs. CD11c CD123 DC precursors, originally described as plasmacytoid T cells or plasmacytoid monocytes, are thought to represent human lymphoid-related DCs and are critically dependent on IL-3 for survival and on CD40L for maturation in vitro. These CD11c DCs are also observed in the PALS. DENDRITIC CELLS IN THE PERIPHERY

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Peyer’s patch In mice Peyer’s patches, two subsets of DCs have been identified (Kelsall and Strober, 1996; Ruedl and Hubele, 1997; Anjuere et al., 1999): (1) CD11c DEC-205 CD8α CD11bdull/subset; and (2) CD11c DEC205 CD8α CD11bbright subset. Phenotypically, these appear similar to the CD8α and CD8α DC subsets in the spleen and lymph nodes, respectively. The CD11c DEC205 population is restricted to the T-cell areas, and the CD11c DEC-205 population is present below the follicle associated epithelium of the subepithelial dome (SED) area, a strategic location that allows access to luminal antigens that have been transcytosed via M cells. Because of their phenotypic similarities to the DC subsets in the spleen and lymph nodes, it is tempting to speculate that these two subsets correspond to CD8α and CD8α DC subsets in the spleen, which are localized in the T-cell zones and marginal zones, respectively. Whether the SED DCs are immature DCs that migrate to the T-cell zones of the Peyer’s patch to become the interdigitating DCs, or whether these two DC subsets have entirely separate lineages is presently unknown. Recent studies in mice suggest that total DC populations from Peyer’s patch (which include both of the DC subsets described above) produce IL-10 and induce the differentiation of TH2 cells (Iwaskai and Kelsall, 1999). In contrast, total spleen DCs (which include the lymphoid and myeloid DC subsets), prime naïve T cells towards the TH1 pathway (Iwasaki and Kelsall, 1999). Similar findings have also been observed with DCs from the respiratory tracts of rats (Stumbles et al., 1998). However, fractionated CD8α and CD8α DCs in the Peyer’s patch appear to elicit TH1 and TH2 responses (Iwasaki and Kelsall, submitted), similar to what has been observed in the spleens (Pulendran et al., 1999; Maldanado et al., 1999). The CD8α DCs from Peyer’s patch appear to induce stronger TH2 responses than the corresponding subset in the spleen. This may account for the differences in responses elicited by total spleen DCs versus total Peyer’s patch DCs.

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Little is known about DC subsets in the human Peyer’s patch, although elegant immunohistochemical studies have revealed striking heterogeneity and microenvironmental compartmentalization of antigen-presenting cells (e.g. Mahida et al., 1989).

Thymus DCs in the thymus are reviewed elsewhere in this volume, but here they are discussed only briefly to compare them with DCs in the spleen, lymph nodes and Peyer’s patches. In mice and in humans, thymic DCs are mainly located in the medulla and at the cortico-medullary junction (reviewed by Sprent and Webb, 1995). Thymic DCs are intimately associated with T cells, as observed with the interdigitating DCs of the spleen and lymph nodes. In mice, thymic DCs have the phenotype CD11c class II MHC DEC205 CD8α CD11bdull/ CD86 CD40 HSA. This phenotype is very similar to that of the CD8α DCs found in the spleen. One marker that is not expressed by spleen or lymph node CD8α DCs, but which is present on thymic DCs is BP-1, a glutamyl aminopeptidase (Vremec and Shortman, 1997). In mice, thymic DCs have been shown to play a role in the negative, but not positive selection, of T cells by several studies (reviewed by Sprent and Webb, 1995; see Chapter on thymic DCs). In humans, 3 distinct populations of thymic DCs are identified. The major population is CD11b CD11c CD45R0low, does not express myeloid markers and can be induced to secrete bioactive IL-12. One minor population is CD11b CD11chigh and CD45R0high, expresses decysin PNA and is a weak producer of IL-12. A second minor population is the CD11c CD123 plasmaytoid DCs, which do not secrete IL-12 (Vandenbalee et al.)

Migration of DCs to lymphoid microenvironments DC migration is reviewed extensively elsewhere in this volume, so we will discuss it only briefly

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here. Once precursors of DCs leave the bone marrow, they have the formidable task of navigating to the tissues, and then to the secondary lymphoid organs. What are the signals that direct DC migration? This question has provoked a flurry of recent studies that implicate chemokines and chemokine receptors in DC migration. Allogeneic skin transplantation models and injection of labeled DCs into mice demonstrate that DCs migrate to the T-cell rich areas of lymphoid organs (reviewed by Austyn, 1999). Pathogen products, such as LPS can trigger the release of TNFα or IL-1, which mediate DC maturation and migration from the peripheral tissues to the secondary lymphoid organs. These inflammatory mediators inhibit the responsiveness of immature DCs in the tissues to MIP3α, by downregulating CCR6, which is a receptor for MIP3α. Concomitantly, the DCs upregulate CCR7, which confers responsiveness to MIP-3β and to 6Ckine. Thus maturing DCs migrate via the afferent lymph and enter the lymph nodes. The expression of MIP-3β and 6Ckine in the stromal cells of the T-cell rich areas and the expression of 6Ckine by the high endothelial venules guide the migration of these cells to the PALS (see reviews Cyster, 1999; Melchers et al., 1999). The role of MIP-3β and 6Ckine in DC migration to the T-cell zones is confirmed by the phenotypes of mice deficient in these molecules. Plt mice, that lack 6Ckine (Gunn et al., 1999), and CCR7/ mice (Forster et al., 1999) have a deficiency in DC and T cell migration to the lymph nodes.

REGULATION OF IMMUNE RESPONSE BY DENDRITIC CELLS TH1/TH2 regulation Emerging evidence suggests that distinct DC subsets are involved in directing different classes of immune responses in vivo (Pulendran et al., 1999; Maldanado et al., 1999), and in vitro (Rissoan et al., 1999). In mice, the CD11c DEC205 CD8α DC subset in the spleen prime TH1

responses in vivo, while the CD11c DEC205 CD8α DC subset primes TH2 or TH0 responses (Pulendran et al., 1999; Maldanado et al., 1999). Both subsets appear to prime naïve CD4 T cells equally efficiently (Pulendran et al., 1999; Maldanado et al., 1999). Consistent with this TH1/TH2 skewing, GM-CSF which preferentially mobilizes CD8α DCs in mice, elicits mainly IgG1 antibodies in response to soluble antigens, whereas Flt-3 ligand which mobilizes both CD8α and CD8α DC subsets, also elicits IgG2a antibodies (Pulendran et al., 1999), a TH1 signature. DCs from IL-12/ mice fail to induce TH1 responses, suggesting a critical role for IL-12 in CD8α DC-induced TH1 responses (Maldanado et al., 1999). CD8α, but not CD8α DCs can be induced to make large amounts of IL-12 (Pulendran et al., 1997; Reis e Sousa et al., 1998; Maldanado et al., 1999; Ohteki et al., 1999), and IFNγ. The mechanism by which CD8α DCs induce TH2 cytokines is not established, although IL-13, IL-10, IL-6, and OX40-L are good candidates. In humans, monocyte derived CD11c DCs polarize naïve CD4 T cells towards the TH1 pathway, while the CD11c DCs polarize towards a TH2/TH0 pathway (Rissoan et al., 1999). The extent of T-cell polarization by CD11c DCs may be related to their differentiation/maturation stages (Cella et al., 1999). The precursors of these CD11c DCs were recently demonstrated to be capable of producing large amounts of IFNα, upon stimulation with herpes simplex virus or influenza virus (Siegal et al., 1999; Cella et al., 1999). The ability of these DCs to elicit TH2 differentiation needs to be reconciled with their IFNα−producing potential. It is likely DCs exhibit considerable plasticity, such that under certain conditions (e.g. viral infections), the CD11c DCs promote TH1 responses, rather than TH2 responses. This concept is explored more thoroughly in the section entitled: ‘Plasticity of DC phenotype and function.’

Tolerance and immunity As mentioned above, and elsewhere in this volume, thymic DCs are involved in the negative

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selection of T cells. In the periphery, a role for DCs in establishing T-cell tolerance has not been formally demonstrated. In fact, the available evidence suggests that DCs can abrogate peripheral T-cell tolerance against soluble antigens (Pulendran et al., 1998), viral antigens (Shimizu et al., 1998) tumors (Gong et al., 1998) and transplant antigens (Steptoe et al., 1997), and in the neonates (Ridge et al., 1996). However, in vitro work from Shortman’s group suggests that, in mice, both CD8α and CD8α DCs can stimulate T cells, but that the CD8α DC subset can limit the proliferation of T cells (Süss and Shortman, 1996; Kronin et al., 1996). For CD4 T cells, the CD8α DCs kill a proportion of the T cells they activate (Süss and Shontman, 1996). In the case of CD8 T cells, they limit cytokine production (Kronin et al., 1996). These observations should now be reconciled with the known ability of these DCs to skew towards the TH1 and TH2 pathways of differentiation (Maldanado et al., 1999; Pulendran et al., 1999).

B-cell priming This is dealt with extensively, elsewhere in this volume. Here, it is suffice to say that DCs can enhance the differentiation of CD40-activated memory B cells to IgG secreting plasma cells (Dubois et al., 1997). This mechanism is likely to be important in germinal centers, where Bcell affinity maturation and differentiation occur. Consistent with this, germinal center dendritic cells (GCDCs) can stimulate CD40 activated germinal center B cells and induce their differentiation to plasma cells (Dubois et al., 1997, 1999).

CONCLUSIONS: NOVEL CONCEPTS IN DC BIOLOGY Although, research on the functional heterogeneity and microenvironmental localization of the various DC subsets in the body is only beginning, a number of emerging themes are already evident. These are discussed below.

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Functional and phenotypic heterogeneity Much of the current research in DC biology until now has been aimed at discovering the striking heterogeneity of DC phenotypes and function in vivo. Immunohistochemical analyses of lymphoid organs in mice and in humans is revealing a rich diversity of DC subpopulations in discrete microenvironments. This has prompted intense efforts to determine whether these subsets have distinct functions. Already, there are hints that distinct DC subpopulations exert different functions. Two broad principles seem to be emerging: (1) DCs in different lymphoid organs may have distinct functions. Thus, Peyer’s patch DCs (Iwasaki and Kelsall, 1999), or respiratory tract DCs (Stumbles et al., 1998) may skew towards TH2 responses. In contrast total DCs from spleens may skew towards TH1 (Iwasaki and Kelsall, 1999), or TH0 (De Smedt et al., 1996) responses. Given that both the spleen and Peyer’s patch have phenotypically similar subsets of DCs (Maraskovsky et al., 1996; Iwasaki and Kelsall, 1999; Anjuere et al., 1999), these observations suggest that the function of DCs can be altered by the local environment. This plasticity of DC function is discussed in greater length below. (2) Even within a particular lymphoid organ, distinct DC subsets are localized in discrete microenvironments. These distinct DC subsets may have distinct functions. For example, in the spleen, the marginal zone (CD8α) DCs direct TH2 responses, while the T-cell zone (CD8α) DCs elicit TH1 responses (Maldanado et al., 1999; Pulendran et al., 1999). Thus, a relevant question is whether TH1 and TH2 responses are initiated in discrete microenvironments. If indeed this were the case, then it suggests that lymphoid organs may be organized into functionally distinct microdomains. Even within a seemingly homogenous area, such as the Tcell zone of the spleen or lymph nodes, it is possible that there are discrete cellular compartments within which distinct phases of a Tcell response occur. Thus, T-cell responses may be initiated in the outer PALS, near the B-cell

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rich follicles, but the clonal expansion of T cells may be downregulated by DCs, deep in the PALS.

Plasticity of DC phenotypes and function The concept that distinct DC subsets exert distinct immune functions should be viewed in the context that DC subsets display considerable plasticity, in their function and phenotype. For example, DCs that normally induce TH1 profiles can be induced to skew towards the TH2 pathway, when treated with anti-inflammatory cytokines such as IL-10 (Caux et al., 1994; De Smedt et al., 1996; Liu et al., 1998) and TGFβ (Τakeuchi et al., 1997), or with steroids or PGE-2 (Kalinski et al., 1998). These observations are consistent with the fact that DC subsets isolated from distinct organs elicit different T-helper responses (Iwasaki and Kelsall, 1999). It is thus likely that DC function is modified by the exposure to a particular cytokine or pathogen product in a given microenvironment.

Summary Unique features such as functional heterogeneity, motility and developmental plasticity bestow the DC system of cells with great adaptability, to confront a large array of potential pathogens. We have already begun to study their functional heterogeneity and to harness this in several clinical trials. The challenge now lies in understanding how the dendritic cells play a role in establishing a specific pattern of immunity, during a particular infection, and to tap this enormous potential to intelligently manipulate immune responses in our own bodies.

ACKNOWLEDGEMENTS This work was supported by grants from NIH DK57665-01; AI48638-01; AI-99-011 and Baylor Health Care Foundation (to B.P.).

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PLATE 27.1 Dendritic cell subsets in secondary lymphoid microenvironments of mice. In the spleen, two major subsets of DCs can be identified: the CD8α (putative lymphoid-related) DCs subset, and the CD8α(putative myeloid-related) DC subset. The latter can be further subdivided, based on CD4 and 33D1 expression. CD8α DC subsets are localized in the PALS, while CD8α DC subsets are in the marginal zones. Certain microbial products (LPS, toxoplasma extracts), can induce the rapid migration of CD8α marginal zone DCs to the PALS. In the lymph nodes, three major subsets can be identified: CD8α DCs, CD8α DCs, and the Langerhans cell derived DCs (CD11c CD8α/ CD11bbright DEC205). The CD8α DCs and the Langerhans DCs are likely localized in the T-cell zones. The CD8α DCs are likely in the subcapsular sinus, and in the T-cell zones bordering the B-cell follicles. In the Peyer’s patch, two major DC subsets can be identified: a CD8α DC subset and a CD8α DC subset. As in the spleen, and lymph nodes, the CD8α DC subset is localized in the T-cell zones, while the CD8α DC subset lies beneath the subepithelial dome.

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28 Dendritic cells in the central nervous system Francesca Aloisi 1, Barbara Serafini 1, Sandra Columba-Cabezas1, and Luciano Adorini 2 1

Istituto Superiore di Sanità, Roma; and 2

Roche Milano Ricerche, Milano, Italy

The difficulty lies, not in the new ideas, but in escaping the old ones, which ramify, for those brought up as most of us have been, into every corner of our minds. John M. Keynes

INTRODUCTION

above-mentioned authors, this chapter will deal exclusively with research on DCs in the CNS. We shall briefly review the unique structural and functional features that enable the CNS to maintain immune privilege, while not totally escaping immune surveillance. In addition, we shall describe studies dealing with the identification of DCs in the diseased CNS and discuss the potential contribution of DCs to CNS immunopathology. The research reviewed will be presented with the perspective that understanding the role of DCs in CNS immune reactivity could lead to more rational approaches in the therapy of neuroinflammatory disorders, such as the putative autoimmune disease multiple sclerosis (MS).

Dendritic cells (DCs), the most potent inducers of immune responses, are present in most body tissues, but are excluded from the neural environment. This has been traditionally viewed as a main feature contributing to the immune-privileged status of the eye and central nervous system (CNS). Despite the validity of this tenet, recent evidence indicates that DCs are present in defined eye and CNS-associated compartments and are recruited to neural tissues under inflammatory conditions, suggesting a role for DCs in the regulation of intracerebral immune reactivity. In an article published in the first edition of this book, McMenamin and Forrester (1999) discussed issues of DC phenotype and localization in the normal eye and CNS, along with a general description of the structural features, immune-privileged status and immune surveillance mechanisms of these tissues. Because of the exhaustive review of DC involvement in ocular pathophysiology provided by the Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

THE CNS IMMUNE PRIVILEGE The CNS parenchyma is composed of a very complex and heterogeneous network of neuronal cells surrounded by different types of glial

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cells. The latter comprise astrocytes, with a primary role in the control of neuronal survival and activity; oligodendrocytes, enwrapping axons with their myelin sheaths and ensuring optimal signal conduction; and microglia, a type of downregulated tissue macrophage, often referred to as the CNS intrinsic immune system. Neuronal cells and oligodendrocytes are highly susceptible to ‘bystander damage’ and edema that accompany inflammatory or cell-mediated immune responses. Given the poor regenerative capacity of the neural tissue, inflammatory episodes in the CNS often lead to permanent neurological impairment and may have consequences for the survival of the organism. To preserve its structural and functional integrity, the CNS tissue has evolved unique mechanisms that strictly regulate local immune and inflammatory responses. These include: (1) the presence of a blood–brain barrier (BBB), restricting access of immune molecules and cells to the CNS tissue; (2) the lack of a conventional lymphatic system for drainage of antigens and immune cells to secondary lymphoid organs; (3) an immunosuppressive microenvironment that inhibits activation of immune cells, either resident in the CNS or recruited from the periphery; and (4) the absence of DCs within the CNS parenchyma, which reduces the probability to present intracerebral antigens to peripheral T cells and initiate CNS-directed immune responses. In the following sections we shall describe the basic features of CNS immune privilege and discuss evidence indicating continuous, albeit discreet, immune surveillance of the CNS, despite strict regulation of immune reactivity.

The blood–brain barrier The blood–brain barrier (BBB) is the brain/ spinal cord protective shield. Its primary role is to maintain CNS homeostasis by regulating exchange of ions, nutrients, hormones and metabolites between the blood and the CNS parenchyma. Structurally, the BBB is composed of specialized non-fenestrated microvascular endothelial cells connected by tight junctions

and surrounded by a basal lamina (Figure 28.1). The endothelial basal lamina borders the perivascular space that contains the pericytes, contractile cells with macrophage-like activities (Thomas, 1999). A distinct population of perivascular macrophages/microglia is present on either side of the basal lamina (Perry et al., 1993). Perivascular cells are a dynamic bone marrow-derived population able to rapidly sense immunological signals in the blood and transduce them to the neural parenchyma. Owing to their scavenger and phagocytic functions, and to constitutive expression of major histocompatibility complex (MHC) class II molecules, perivascular cells are thought to have a key role as first-line sentinels and antigen-presenting cells (APCs) of the CNS (Hickey and Kimura, 1988; Perry et al., 1993). Essential elements of the BBB are also the astrocytic endfeet which surround the abluminal surface of CNS endothelial cells, outside the basal lamina, and influence both the tightness and trafficking role of the barrier (Engelhardt, 1997). The BBB is present throughout the CNS, except in the choroid plexuses and a set of small midline structures bordering the cerebral ventricles, collectively denominated circumventricular organs (CVOs). The choroid plexuses are leaflike, highly vascular structures that project into the lateral, third and fourth cerebral ventricles. They comprise fenestrated capillaries embedded in a loose connective tissue stroma and are covered by secretory epithelial cells, joined by tight junctions that act as a barrier between the blood and the cerebrospinal fluid (CSF) (Figure 28.1). Choroid plexuses are responsible for the continuous secretion of the CSF that fills the ventricular system and the subarachnoid space at the CNS surface. CVOs are considered to be points of communication among blood, CSF and brain parenchyma, and have been implicated in a myriad of functions, including regulation of body fluid homeostasis, behavior and endocrine responses (Johnson and Gross, 1993). The meninges (dura mater, arachnoid and pia mater) are fibrocellular layers lining the brain/spinal cord surfaces and the blood vessels entering or leaving the CNS tissue. While the

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dura contains permeable fenestrated vessels, the arachnoid membrane situated below acts as a blood–CSF barrier, limiting entry of blood components into the subarachnoid space (Figure 28.1). Traffic of immune cells through the BBB The intact BBB is thought to have an active role in limiting the passage of immune molecules and cells from the blood into the neural parenchyma. Its ability to act as a physical barrier for immune cells is, however, not absolute. Cells of the monocyte/macrophage lineage are constantly recruited from the blood circulation to replenish the pool of CNS perivascular macrophages. Although at a very slow rate, they also enter the CNS parenchyma to differentiate into microglia (Hickey et al., 1992; Lawson et al., 1992). Under certain physiological conditions (e.g. postpartum or after pairing), mast cells migrate into the CNS tissue (Silverman et al., 2000). Moreover, activated T cells, independently of their antigenic specificity, can cross the intact BBB, suggesting constant patrolling of the CNS by the immune system (Wekerle et al., 1986, Hickey et al., 1991). The signals regulating immune cell traffic through the intact BBB are still unknown. BBB disruption and perivascular infiltration of inflammatory cells (macrophages, activated T cells and B cells) into the CNS represent the pathological hallmarks of MS and its animal model experimental autoimmune encephalomyelitis (EAE), as well as of viral, bacterial or parasitic encephalopaties (Engelhardt, 1997; Hickey, 1999). Under inflammatory conditions, activated leukocytes penetrate the BBB according to the multistep model of leukocyte– endothelial cell recognition (Butcher, 1991; Springer, 1994). This involves sequential interactions of adhesion molecules with their counterreceptors on activated endothelial cells and leukocytes, as well as of chemokines with their receptors. Inflammatory mediators, like interferon-γ (IFN-γ), interleukin-1 (IL-1) and tumor necrosis factor-α (TNF-α), induce expression of adhesion molecules (Engelhardt, 1997; Lee and

FIGURE 28.1 Schematic representation of the blood–brain and blood–cerebrospinal fluid (CSF) barriers with possible pathways for CNS antigens or APCs carrying CNS antigens to reach lymphoid organs. Antigens present in the interstitial fluids of the CNS parenchyma can drain directly into the CSF or along perivascular spaces (1). Antigens or APCs draining with the CSF along the ventricles and the subarachnoid space can reach the venous blood through the arachnoid villi in the meninges (2) or drain to cervical lymph nodes via direct connections of the subarachnoid space and periarterial spaces with nasal lymphatics and cervical lymph nodes (3). Both macrophages and DCs are optimally located in the meninges and choroid plexuses to survey fluxes of pathogens and other potentially noxious substances from the blood into the CNS tissue, as well as all fluids and materials leaving the CNS. It is still unknown whether choroidal and meningeal DCs have the ability to migrate into the lumen of blood vessels or if they can enter the subarachnoid space or ventricles and drain with the CSF to cervical lymph nodes. At the blood–brain barrier, pericytes and cells with macrophagic properties are located in the perivascular spaces. The ability of perivascular cells to reverse transmigrate across the blood–brain barrier endothelium and reach the spleen via the blood stream remains to be demonstrated.

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Benveniste, 1999) and production of chemokines (Glabinski and Ransohoff, 1999) in cerebral endothelial cells and astrocytes, and have been implicated in BBB disruption. In MS and EAE, interactions between CD54 (intercellular adhesion molecule-1) and CD106 (vascular cell adhesion molecule-1) on activated brain endothelia and their counter-receptors (LFA-1 and VLA-4, respectively) on activated encephalitogenic T cells are thought to play a crucial role in the migration of the latter through the BBB (Yednok et al., 1992; Engelhardt, 1997). In both diseases, disruption of the BBB is an early event during the development of CNS lesions and can rapidly lead to vasogenic edema with its clinical consequences. Matrix metalloproteases produced by T lymphocytes and macrophages have been shown to play a key role in the disruption of the BBB basal lamina, thereby facilitating penetration of inflammatory cells into the CNS parenchyma (Leppert et al., 1996).

The CNS lymphatic drainage Although lacking conventional lymphatic vessels, the CNS contains fluid compartments thought to represent a modified lymphatic system (reviewed by Cserr and Knopf, 1997). These include the interstitial fluids surrounding neurons and glia, the perivascular spaces lined by the subendothelial basal lamina, and the CSF filling the ventricles and the subarachnoid space comprised between the arachnoid and pia mater. The CSF acts both as a fluid cushion and as a ‘sink’ removing products of the cerebral metabolism into the venous blood. CNS fluids and intracerebrally injected antigens also drain to cervical lymph nodes (Cserr and Knopf, 1997). Viruses infecting the CNS have been shown to induce humoral and cell-mediated immune responses in cervical lymph nodes (Stevenson et al., 1997). Soluble antigens within the CSF drain along the ventricular system and subarachnoid space. From this compartment, they can either enter the dural venous sinuses through arachnoid villi and reach the blood and spleen, or drain to cervical lymph nodes (Figure 28.1). Antigens present in interstitial fluids can drain

into the CSF or reach the cervical lymph nodes along periarterial pathways in the brain and meninges. Studies in rodents have demonstrated that the subarachnoid space and the perivascular space of the nasal-olfactory artery connect, via the cribriform plate, with nasal lymphatics and cervical lymph nodes (Cserr and Knopf, 1997). Migration of APCs carrying CNS antigens along these draining routes is, however, still to be demonstrated.

The CNS immunosuppressive environment Although physical barriers between the CNS and the general circulation represent effective obstacles, the CNS itself actively contributes to its immune privilege. Inhibitory influences from the neural environment have long been proposed to contribute to the downregulated phenotype of microglia as compared to macrophages present in perivascular spaces, meninges or choroid plexuses (Perry et al., 1993). A number of molecules with immunosuppressive activity are synthesized in the CNS. These include: cytokines (transforming growth factor-β), neuropeptides (vasoactive intestinal peptide, α melanocyte-stimulating hormone), βadrenergic receptor agonists and lipids (gangliosides). Evidence is also emerging that neurons themselves have an important role in downregulating the intracerebral expression of proinflammatory molecules and in controlling microglia phenotype. Electrically active neurons and factors released during normal neuronal activity, like neurotrophins, inhibit IFN-γ-induced expression of MHC class II and costimulatory molecules on microglia (Neumann and Wekerle, 1998; Wei and Jonakait et al., 1999). Moreover, neurons constitutively express the integral membrane protein OX-2 which has been reported to play a key role in regulating macrophage/microglia and DC activation (Hoek et al., 2000). Neuronal degeneration has been suggested to impair this negative control, thereby leading to exaggerated intracerebral inflammatory responses (Neumann and Wekerle, 1998). Both reactive

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astrocytes and microglia have been identified as sources of immunosuppressive molecules (e.g. TGF-β, prostaglandin E2) inhibiting macrophage/microglia and T lymphocyte activation (reviewed by Aloisi et al., 2000). Regulation of inflammatory reactions by the CNS parenchyma may also occur through active elimination of CNS-infiltrating immune cells. In EAE, apoptosis of encephalitogenic T cells, via Fas (Suvannavejh et al., 2000) or TNF receptor 1 signaling (Bachmann et al., 1999), is thought to represent the main mechanism for the termination of intracerebral immune responses.

Absence of DCs in the normal CNS parenchyma Initiation of an immune response requires capture of antigens at the site of entry, transport to local lymph nodes and presentation to naïve T lymphocytes. DCs have turned out to play the key role in this process as the most efficient antigen-presenting cell type (Steinman, 1991; Banchereau and Steinman, 1998). It has long been recognized that antigens sequestered behind the BBB are not recognized by the immune system and that certain viruses can persist in the brain for prolonged periods of time without sensitizing the immune system. However, immune responses to intracerebral particulate antigens (e.g. allogeneic grafts, myelin proteins, or pathogens) are readily mounted following peripheral immunization with the same antigens, indicating that TABLE 28.1

recognition of CNS-associated antigens does occur once these are seen by the immune system, outside the CNS (reviewed by Perry, 1998; Matyszak, 1998). All available experimental evidence is consistent with the observation, in several species, that the CNS is devoid of resident, intraparenchymal DCs. Hart and Fabre (1981) first described the presence of Ia DCs in all rat tissues analyzed with the exception of brain. Further immunohistochemical analyses have shown that intraparenchymal expression of MHC class II antigens is confined to microglia (reviewed by Sedgwick and Hickey, 1997). Lack of DCs in the normal CNS parenchyma has been confirmed by studies using monoclonal antibodies (mAbs) against DC-restricted markers (Witmer-Pack et al., 1995; Matyszak and Perry, 1996; McMenamin, 1999; Serafini et al., 1999, 2000) (Table 28.1). In the mouse, these include mAbs N418 (specific for the β2 integrin CD11c) (Metlay et al., 1990), NLDC-145 (reacting with DEC-205, an integral membrane protein involved in antigen processing) (Witmer-Pack et al., 1995), and MIDC-8 (reacting with a still unidentified intracellular antigen expressed by interdigitating DCs of mouse secondary lymphoid organs and mature DCs in vitro) (Inaba et al., 1997). In the rat, absence of DCs within the CNS is indicated by lack of staining with the OX62 mAb, specific for an α-like integrin subunit expressed on rat DCs and γ/δ T cells (Brenan and Puklavec, 1992). Interstitial folliculostellate cells in the anterior pituitary have been

DC compartmentalization in the normal central nervous system

Identification criteria Mouse DCs DEC-205 (NLDC-145 mAb) CD11c (N418 mAb) MIDC-8 mAb Rat DCs OX-62 mAb Human DCs Ultrastructure

Parenchyma

Choroid plexuses

Meninges

References

  

  

  

1, 2 1 1







3, 4





N.D.

5, 6

, present;  absent; N.D., not determined. References: 1, Serafini et al., 1999, 2000; 2, Witmer-Pack et al., 1995; 3, Matyszak and Perry, 1996; 4, McMenamin, 1999; 5, Serot et al., 1997; 6, Hanly and Petito, 1998. DENDRITIC CELLS IN THE PERIPHERY

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reported to exhibit some DC-like morphological and immunopheno typical features but the precise identity of these cells remains to be clarified (Allaerts et al., 1997). The presence of DCs and DC precursors in all non-neural body tissues raises interesting questions on the mechanisms underlying the exclusion of DCs from the CNS parenchyma. Does the BBB prevent the migration of DCs or DC progenitors into the CNS or is the CNS microenvironment not conducive to DC differentiation? DCs residing in nonlymphoid tissues share a common myeloid progenitor with macrophages (and thus very likely also with microglia), and cells of the myeloid lineage enter the CNS to differentiate into perivascular macrophages and microglia, suggesting that the neural environment may prevent DC differentiation. Neurons themselves could have a major role in inhibiting the emergence of both macrophage (as discussed above) and DC phenotypes. Cells expressing DC markers (CD11c, DEC-205, 33D1, MHC class II, CD86) have been identified in granulocyte–macrophage colonystimulating factor (GM-CSF)-supplemented cultures from the neonatal mouse CNS, indicating that at least in vitro CNS-derived myeloid progenitors can differentiate into DCs (Fischer and Bielinsky, 1999). We have observed that an enriched population of CD11b microglia isolated from the adult mouse CNS contains a minor fraction of myeloid progenitors that proliferate in vitro in the presence of GM-CSF and are induced to express features of mature DCs when TNF-α or lipopolysaccharide is added to the cultures (our unpublished observations). It remains to be determined whether myeloid progenitors reside within the CNS parenchyma proper, perivascular spaces or BBB-free areas, and retain the potential to differentiate into DCs following CNS injury or infection.

Localization of DCs in the meninges and choroid plexuses of the normal CNS Over the last few years, several studies have described the presence of DCs in the meninges

and choroid plexuses (Figure 28.1). DCs were identified by morphological criteria, high MHC class II expression and absence of macrophage markers, within the stroma and between the epithelial cells of human choroid plexuses (Serot et al., 1997, Hanly and Petito, 1998). In rats and mice, meningeal and choroid plexus DCs were mainly characterized using DC-specific markers (Matyszak and Perry, 1996; McMenamin, 1999; Serafini et al., 1999, 2000) (Table 28.1). Although numerous macrophages but only scattered DCs were initially identified in these compartments, a recent study using whole-mount preparations of rat meninges and choroid plexuses has revealed that both sites are populated by a surprisingly rich network of DCs (McMenamin, 1999). Numerous OX-62, MHC class II DCs, with dendritic or pleiomorphic morphology and negative for macrophage markers (ED1, ED2, complement receptor 3), have been detected in all three meningeal layers (dura mater, arachnoid and pia mater) and in the stroma of the choroid plexuses. Both the lack of a BBB and the presence of extensive networks of MHC class II DCs in the choroid plexuses and meninges are consistent with the massive accumulation of inflammatory cells at these sites during infectious and autoimmune CNS diseases. Meningeal and choroid plexus DCs resemble the immature DCs residing in peripheral tissues, specialized for antigen capture (Banchereau and Steinman, 1998). Under nonpathological conditions, they do not express molecules typical of mature, lymphostimulatory DCs, like CD40, CD80 and CD86 (Serot et al., 2000; our unpublished observations). We have found that MIDC8, a marker for mature murine DCs (Inaba et al., 1997), is also not expressed in the meninges and choroid plexuses (Serafini et al., 2000). Due to direct contact of the pia mater and arachnoid with the CSF draining along the subarachnoid space, and to the close apposition of the pia mater with the basal lamina and the underlying subpial astrocytic endfeet, meningeal DCs appear to be in a key position for sensing the internal neural environment. As the choroid plexuses also pump metabolites from

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the CSF into the blood, choroid plexus DCs may endocytose and process CSF-derived antigens. Interestingly, cytoplasmic labeling for the antiinflammatory cytokine IL-10 was recently shown in most intraepithelial DCs of human choroid plexuses, suggesting that these cells may have an immunosuppressive role and contribute to maintain immune tolerance to CNS proteins (Serot et al., 2000). Meningeal and/or choroid plexus DCs could also initiate immune responses to pathogenic antigens as shown by specific immune responses to grafts or infectious agents placed in the cerebral ventricles or subarachnoid space (Mason et al., 1986; Stevenson et al., 1997; Perry, 1998). At present, it is not known whether DCs can cross the choroid plexus epithelium and migrate into the CSF or if they are present within the CSF under inflammatory conditions. However, the preferential accumulation of inflammatory cells in the periventricular white matter of MS patients (Raine, 1994) suggests that DCs entering the CNS from the ventricles may facilitate immune cell recruitment and activation. Choroid plexus and meningeal DCs could reach secondary lymphoid organs through the blood circulation or drain with CSF to the nasal lymphatics and cervical lymph nodes (Figure 28.1). To our knowledge, no studies have attempted to localize DCs in the CVOs. As these BBB-free areas contain blood-derived macrophages and activated microglia (Pedersen et al., 1997), it would be interesting to know whether they also contain DCs. In conclusion, the evidence reviewed in this section indicates that DCs, although excluded from the CNS parenchyma proper, are in key positions to contact potential pathogens or autoantigens entering or leaving the CNS. This would enable DCs to participate in the immune surveillance of the CNS tissue, by acting as APCs capable of inhibiting or inducing T-cell activation. As shown for DCs in peripheral tissues, the immunostimulatory or tolerogenic function of CNS-associated DCs would depend on a variety of factors, with a strong influence of the inflammatory or noninflammatory context (Banchereau and Steinman, 1998).

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Microglia as the sentinels of the CNS tissue While DCs have a central role as sentinels of the immune system, microglia are key elements in the surveillance of the CNS microenvironment. As with tissue DCs, microglia are myeloidlineage related cells and exhibit a ramified morphology (Perry et al., 1993). In normal conditions, microglia can be regarded as a form of downregulated tissue macrophage. They exhibit a very slow turnover rate, no phagocytic or endocytic activity, and low expression of CD45, MHC class II molecules, complement receptor 3 and Fc receptors (Perry et al., 1993; McMenamin and Forrester, 1999). As discussed above, inhibitory signals from the CNS microenvironment and an intact BBB must be determinant in maintaining microglia in a quiescent state. The precise role of the extensive network of resident microglia in the normal CNS tissue remains elusive, although the presence of numerous receptors for CNS signalling molecules (ATP, neuropeptides and neurotransmitters) and unique membrane ionic channels suggests that microglia continuously monitor the physiological integrity of their microenvironment. The main function of microglia in the adult brain is to respond very rapidly to any disruption of brain’s homeostasis, such as direct damage to neurons, neurodegeneration, infection or autoimmune attack (Streit and Kincaid-Colton, 1995). In analogy with the long and thin processes of DCs which are important for sensing the environment and interacting with T and B cells, the ramified, crenellated processes emerging from microglial cell bodies are essential for sensing the CNS microenvironment and reacting to even very subtle alterations of its composition. Upon CNS injury and depending on the extent of neuronal damage, microglia progressively lose their ramified morphology and acquire properties of full-blown macrophages (Perry et al., 1993; Streit and Kincaid-Colton, 1995; Kreutzberg, 1996). These include phagocytic activity, production of cytotoxic molecules (free oxygen intermediates, nitric oxide), proteases,

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prostanoids, pro- and anti-inflammatory cytokines, chemokines, and growth factors. Microglia activation is indispensable for protecting the CNS parenchyma from microbial infections, removing damaged neurons or myelin, and inducing repair processes. Migration and proliferation of microglia (or possibly their direct progenitors) within CNS lesions has also been described (Kreutzberg, 1996). Comparison between the antigen presenting function of DCs and microglia Over the past 15 years there has been considerable interest in understanding whether the CNS harbors potential APCs involved in sustaining cell-mediated immune responses against neurotropic infectious agents or CNS autoantigens. Studies in radiation bone-marrow chimeras and EAE models have provided evidence for a major role of CNS perivascular and meningeal macrophages in presenting myelin antigens to encephalitogenic T cells (Hickey and Kimura, 1988; Ford et al., 1996; Sedgwick and Hickey, 1997). So far, there is very little evidence that cells of the monocyte/macrophage lineage can leave the CNS perivascular space and reach lymphoid tissues, possibly carrying with them antigens from behind the BBB (Broadwell et al., 1994). Among intraparenchymal cells, astrocytes, the most abundant CNS glial population, are thought to have only a marginal role in antigen-specific T cell responses (reviewed by Sedgwick and Hickey, 1997; Aloisi et al., 2000). In inflammatory conditions, reactive astrocytes express MHC class II and adhesion/costimulatory molecules very rarely, if at all (Kreutzberg, 1996). In vitro studies have revealed that astrocytes fail to process native proteins and to prime T cells, but can restimulate T cells (mainly TH2 cells) in the presence of antigenic peptides (Aloisi et al., 1998, 1999b). Owing to their ability to suppress macrophage/ microglia activation and TH1 cells via inhibition of IL-12 production by microglia (Aloisi et al., 1997), astrocytes could have an important function in limiting CNS inflammation. Most available evidence points to microglia as the principal intracerebral cell type with APC

function. Because microglia upregulate expression of MHC class II molecules in essentially all known CNS pathologies (Kreutzberg, 1996), a key function of activated microglia is thought to be the MHC class II-restricted presentation of processed antigenic peptides to CD4 helper T cells. Antigen-presenting microglia can display immunostimulatory or inhibitory activities. In autoimmune CNS diseases, such as MS and EAE, microglia upregulate a number of adhesion/ costimulatory molecules (CD40, CD54, CD80/ CD86) and are thought to contribute to the restimulation of CNS-infiltrating encephalitogenic TH1 cells (Aloisi et al., 1997, 1998; Krakowski and Owens, 1997). Similar to what has been observed with macrophages, signals from TH1 cells, particularly IFN-γ, are essential for promoting microglia APC function. In vitro studies using long-term cultured or acutely isolated microglia from the rodent or human CNS generally agree that microglia, once activated, are good stimulators of T-cell effector functions (Becher and Antel, 1996; Ford et al., 1996; Aloisi et al., 1998, 1999b). Similarly to DCs and macrophages, microglia produce the TH1-inducing cytokine IL-12 and restimulate TH1 cells with high efficiency (Aloisi et al., 1998, 1999b; Krakowski and Owens, 1997). Bacterial components as well as interactions between CD154 on activated TH1 cells and CD40 on microglia have a crucial role in the induction of microglial IL-12 production (Aloisi et al., 1999a), possibly contributing to intracerebral skewing of TH1 responses during infectious and autoimmune CNS pathologies. Although comparable to DCs in the restimulation of primed CD4 T cells, both TH1 and TH2, microglia are less efficient than DCs in the priming of naïve T cells (Aloisi et al., 1999b). Microglia can also inhibit CNS immune responses. Ford et al. (1996) have shown that activated microglia isolated from the CNS of rats undergoing graft-versus-host disease fail to induce T cell proliferation and cause some degree of T-cell apoptosis, suggesting a regulatory role for microglia in limiting spreading of immune responses. Anti-inflammatory mediators (IL-10, TGF-β and prostaglandin E2) secreted by activated microglia are also likely to

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contribute to the downregulation of intracerebral immune reactivity (Aloisi et al., 2000).

DCs IN CNS PATHOLOGY DCs in CNS autoimmune diseases DCs have been implicated in the initiation and maintenance of autoimmune diseases, such as insulin-dependent diabetes, autoimmune thyroiditis and rheumatoid arthritis (Thomas and Lipsky, 1996). DCs capturing self-antigens in the target organ and migrating to regional lymph nodes may initiate the activation of autoreactive T cells and support chronic inflammation by sustaining successive waves of priming of naïve T cells. A recent study in a transgenic model of diabetes suggests that DCs recruited to the target tissue are also involved in the maintenance of the inflammatory milieu by local activation of T cells and formation of organized lymphoid structures (Ludewig et al., 1998). An important issue to clarify is the contribution of DCs to CNS autoimmune pathologies. MS is a chronic inflammatory disease involving the white matter of the CNS and leading to loss of neurological function. Although the exact cause of the disease remains unknown, MS presents the main features of an autoimmune disease (Conlon et al., 1999). Histopathological hallmarks of MS include perivascular T cell and macrophage infiltrates extending into the white matter, demyelination and axonal injury (Raine, 1994). Some forms of the disease are thought to be sustained by activation of B cells and autoantibody production (Lucchinetti et al., 1996; Genain et al., 1999). Our current understanding of the pathogenesis of MS invokes both genetic factors underlying disease susceptibility and environmental factors triggering the disease. Viral infections are thought to play a role in the initiation of CNS autoimmune reactivity, either because a virus shares amino acid sequences or conformations with CNS proteins (molecular mimicry) or because a viral infection creates the inflammatory conditions leading to the activation of autoreactive T cells (Conlon et al., 1999).

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Despite extensive analysis of the inflammatory infiltrates in post-mortem MS brains, identification of DCs in MS lesions is still uncertain. By using an anti-CD1a mAb recognizing a β2-microglobulin-associated polypeptide on human dendritic cells (Longley et al., 1989), Battistini et al. (1996) described the presence of rare CD1a cells in the perivascular immune infiltrates of chronic-active MS lesions. We have shown that perivascular process-bearing cells expressing the costimulatory molecule CD80 persist in chronic-inactive MS lesions (De Simone et al., 1995). These could represent either perivascular macrophages or residual DCs entering the CNS during the active phases of the disease. Further investigations are needed to establish if and when in the development of MS lesions DCs are recruited to the CNS. Bartholome et al. (1999) have recently proposed that abnormal over-production of IL-12 by circulating DCs may be related to MS pathogenesis. DCs in EAE Experimental autoimmune encephalomyelitis (EAE) is an animal model of CNS inflammation that has been extensively used to study the pathogenesis of MS (Bradl and Linington, 1996). It is characterized by multifocal perivascular, meningeal and intraparenchymal CNS inflammatory infiltrates, comprising mainly T cells and macrophages. EAE is induced either by immunization with CNS antigens (mainly myelin proteins or their immunodominant epitopes) or by adoptive transfer of T cells primed in vitro with myelin antigens. It is mediated by TH1 cells, which, after migration across the BBB, recognize the target antigen on local CNS APCs and secrete pro-inflammatory cytokines (IFN-γ, IL-2, TNF-β) leading to massive T cell and macrophage recruitment to the CNS, neurological deficits and myelin destruction (Steinman, 1996). DCs endocytosing myelin antigens at the site of subcutaneous immunization and migrating to the draining lymph nodes are essential for EAE initiation. This is supported by the finding that DCs efficiently activate encephalitogenic

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T cells for transferring EAE (Gautam and Glynn, 1989). Moreover, a recent study has shown that DCs pulsed with an encephalitogenic myelin basic protein peptide (Ac1–11) interact with Ac1–11specific TCR transgenic naïve T cells in the peripheral lymph nodes of recipient mice, leading to induction of EAE (Dittel et al., 1999). The identity of the APCs restimulating encephalitogenic T cells in the CNS is less clear. Although most of the above-reviewed studies implicate CNS perivascular macrophages and intraparenchymal microglia in the presentation of neural antigens, evidence is emerging that DCs might also play a role in intracerebral T-cell activation. In a rat model of acute EAE, Matyszak and Perry (1996) revealed the presence of small numbers of OX62, MHC class II cells in CNS perivascular infiltrates, but very rarely within the adjacent spinal cord parenchyma. Suter et al. (2000) have detected the presence of transcripts specific for the class II transactivator (CIITA) form I (specific for dendritic cells) and form IV (IFN-γ-inducible) in the brain and spinal cord of mice developing acute EAE. These authors have also shown that CD11c DCs are present in the CNS immediately before the appearance of clinical signs but after the recruitment of CD11b macrophages, suggesting that macrophages may provide chemotactic signals for DCs. Absence of CIITA and MHC class II expression in early infiltrating macrophages has led to the hypothesis that intracerebrally recruited DCs may have a key role in presenting CNS antigen to encephalitogenic CD4 T cells. To obtain more information on temporal appearance, persistence, and functional maturation of DCs in the inflamed CNS, we have recently performed a detailed immunohistochemical analysis of the CNS in SJL mice at different stages of EAE induced by the proteolipid protein (PLP) peptide 139–151 (Serafini et al., 1999, 2000). These mice developed an acute form of EAE and, less frequently, a chronic or relapsing disease. At variance with the findings of Suter et al. (2000), we have found that DCs are apparently the first leukocytes to be recruited into the CNS following EAE induction. Just before the appearance of CNS inflammatory

infiltrates and EAE clinical signs, process-bearing cells expressing DEC-205, CD11c and MHC class II molecules, but not the mature DC marker MIDC-8, were identified in the spinal cord white matter, immediately beneath the pial surface (Plate 28.2A). This distribution suggests that DCs normally residing in the meninges may cross the basal lamina, which separates the pia mater from the glia limitans, and migrate into the CNS parenchyma. The mechanisms underlying this early DC migration are still unknown. During acute, chronic and relapsing phases of PLP 139–151-induced EAE, DEC-205 and MIDC-8 DCs accumulated within the inflammatory infiltrates in the meninges, submeningeal white matter and around intraparenchymal blood vessels in the spinal cord, brain stem and cerebellum (Plate 28.2 B, C). The most pronounced CNS infiltration by DCs was observed during EAE relapses, when numerous DCs, often colocalizing with CD4 T cells, were found in the spinal cord white matter (Plate 28.2 D). While DCs present in CNS perivascular infiltrates are likely to originate from blood-derived precursors,DCsscatteredintheCNSparenchyma could have differentiated from endogenous precursors under the influence of protracted inflammatory stimuli. DCs infiltrating the CNS of EAE-affected mice appeared to be phenotypically similar to mature DCs as they expressed CD40 and CD86 molecules. Intracerebral DC maturation could be induced by TNF-α and IL-1 synthesized within the inflamed CNS (Tanuma et al., 1997), interaction with CD154 (CD40 ligand) expressed on encephalitogenic TH1 cells (Gerritse et al., 1996), and transendothelial DC migration across the activated BBB endothelium (Randolph et al., 1998). Recruitment and maturation of DCs into the CNS have been confirmed in further studies using Biozzi AB/H mice immunized with whole spinal cord homogenate, a model of chronic-relapsing EAE (Mattner et al., 2000). During the chronic-relapsing phases of the disease, we observed the presence of huge inflammatory infiltrates in the CNS parenchyma with considerable numbers of DCs (Plate 28.2E and our unpublished data). Early recruitment and persistence of DCs

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within the CNS of EAE-affected mice strongly suggest that intracerebral DCs participate both in the onset and maintenance of CNS autoimmunity. They could efficiently restimulate encephalitogenic T cells crossing the BBB and produce chemokines that attract additional DCs, macrophages and CD4 T cells, as well as CD8 T cells and B cells to the CNS (Sallusto et al., 1999; Tang and Cyster, 1999). Consistent with this view, a recent study by Carson et al. (1999) has shown that intracerebrally injected, mature DCs cause disproportionate recruitment of CD8 T cells to the CNS. DCs capturing CNS antigens generated during tissue injury and migrating to cervical lymph nodes or spleen may also be involved in the stimulation of naïve T cells specific for CNS autoantigens. This scenario is supported by the finding that intracerebrally injected mature DCs home to cervical lymph nodes (Carson et al., 1999) and could explain epitope spreading, the appearance of T cell specificities for myelin determinants distinct from the initiating ones, that accompanies progression to chronic EAE (Lehmann et al., 1992; McRae et al., 1995). Because accumulation of B cells in CNS perivascular spaces and development of humoral autoimmunity also occur during EAE chronicization (Linington and Lassmann, 1987; Lucchinetti et al., 1996), another possible mechanism through which DCs may sustain CNS autoimmune reactivity is by supporting intracerebral B-cell growth and differentiation (Dubois et al., 1999). The establishment of DC-T and -B cell interactions within the inflamed CNS could stimulate the production of antimyelin autoantibodies detected in EAE and MS lesions (Genain et al., 1999), and the intrathecal B-cell clonal expansion associated with MS (Tourtellotte et al., 1984). Intracerebrally recruited DCs could also contribute to the formation of the organized lymphoid structures that have been described in the CNS lesions of mice with adoptively transferred EAE (Raine et al., 1984) and in MS plaques (Prineas, 1979). As proposed for other chronic inflammatory disorders (Ludewig et al., 1998; Ruddle, 1999), lymphoid neogenesis in the CNS may represent a key event main-

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taining the autoimmune response against neural antigens. Two distinct subsets of DCs, myeloid- and lymphoid-related, have been recently identified and proposed to differentially affect T-cell responses (reviewed by Reid et al., 2000). In the mouse, lymphoid DCs induce TH1 and myeloid DCs TH2 cells. No information is as yet available on the possible involvement of different DC subsets in the acute and chronic phases of EAE.

DCs in CNS infectious diseases Only a few studies have addressed DC localization in the infected CNS. Hanly and Petito (1998) examined autoptic brain tissue from individuals with acquired immunodeficiency syndrome (AIDS) and found that human immunodeficiency virus (HIV )-infected cells in the choroid plexuses had the morphological and immunophenotypical appearance of DCs. DCs could have a key role in systemic HIV infection, being the first cells infected by HIV and representing a reservoir of the virus during the period of clinical latency that follows the early viremic phase of systemic infection. Infection of the CNS by HIV is thought to be mediated by virusinfected mononuclear phagocytes passing through the BBB, but the above findings suggest that DCs, rather than monocytes/macrophages, carry HIV into the CNS and act as a CNS reservoir of infection. The fate of HIV-infected choroidal DCs is unknown. They could migrate from the choroid plexus into the CSF and play a role in the pathogenesis of HIV encephalitis. HIV-infected DCs draining with CSF or leaving the choroid plexuses through the choroidal fenestrated blood vessels could also reach cervical lymph nodes or spleen, contributing to HIV spreading outside the CNS. Cells expressing the DC markers CD11c and 33D1 were localized within the perivascular and parenchymal inflammatory foci in the CNS of mice chronically infected with Toxoplasma gondii (Fischer et al., 2000). CD11c cells isolated form T. gondii-infected brains showed the phenotype of myeloid DCs (DEC-205lo, F4/80, CD8α) and expressed MHC class II, CD40,

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CD54, CD80 and CD86 molecules. These mature, brain-derived DCs appeared very efficient in triggering antigen-specific and allogeneic T-cell responses and secreted IL-12. DC-like cells generated from GM-CSF-supplemented primary mouse brain cell cultures expressed CD86 and secreted IL-12 following in vitro challenge with T. gondii. Based on these findings, Fischer et al. (2000) have proposed that parasite-induced expansion and functional maturation of DCs into the CNS, possibly from a pool of resident myeloid precursor cells, may represent a mechanism contributing to persistent intracerebral inflammation. Matyszak and Perry (1996) have developed a model of chronic CNS inflammation and demyelination based on injection of bacillus Calmette-Guérin (BCG) into the rat CNS parenchyma and subsequent peripheral immunization with the same pathogen. This protocol elicits a strong and persistent delayed-type hypersensitivity response with T cell and macrophage recruitment to the CNS, disruption of the BBB, myelin breakdown and axonal damage. The CNS inflammatory infiltrates include numerous OX62, MHC class II DCs. It has been proposed that the BCG-induced immune response provokes CNS tissue damage with release of cerebral antigens and activation of CNS autoreactive T cells. In this scenario, DCs recruited to the CNS would process CNS soluble antigens, or antigens associated with injured or apoptotic cells, and transport them to lymph nodes to prime successive waves of autoreactive T cells (Matyszak, 1998).

DCs in other CNS pathologies Little information is available on the possible involvement of DCs in nonimmune CNS pathologies. To our knowledge, only one study has examined the presence of CD1a cells in contused human brain tissue (Holmin et al., 1998).The acute inflammatory response detected in the CNS a few days after trauma consisted of numerous monocytes/macrophages, reactive microglia, polymorphonuclear cells and T lymphocytes. The human DC marker CD1a was

detected on a small number of cells with a dendritic morphology, although these cells were referred to as microglia. Langerhans’ cell histiocytosis (LCH) is a pathology which affects most body tissues, but rarely the CNS. Primary cerebral forms of LCH involve predominantly the hypothalamus leading to hypothalamic dysfunction with diabetes insipidus. By ultrastructural (presence of Birbeck granules) and immunocytochemical (CD1a expression) criteria, Langerhans cells have been identified in brain lesions, although being less numerous than in lesions developing in peripheral tissues (Eriksen et al., 1988; Mazal et al., 1996). Daniel et al. (1985) have described the presence of numerous cells containing Birbeck granules within a primary intracranial histiocytic lymphoma, with predominant meningeal localization. The origin of intracranial Langerhans’ cells is still unknown, as they could either migrate into the CNS or represent a resident CNS population. These cells may proliferate abnormally as in Langerhans’ cell histiocytosis or more aggressively give rise to a form of malignant histiocytic lymphoma.

DC RECRUITMENT TO THE INFLAMED CNS Leukocyte recruitment to distinct anatomical sites is a complex process mediated by chemotactic cytokines, the chemokines (Baggiolini, 1998). Chemokines are small secreted proteins produced by leukocytes themselves and by endothelial and parenchymal tissue cells, particularly under inflammatory conditions. Based on the relative position of the first two cystein residues, chemokines have been classified into four groups (CXC, CC, C and CX3C). All the chemokine receptors identified so far are seventransmembrane-spanning, G-protein-coupled receptors. Most chemokines act on distinct leukocyte populations and each leukocyte population has receptors for and responds to many chemokines, indicating a complex and partially redundant network (Mantovani, 1999). DCs also require chemokines for recruitment

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to lymphoid and non-lymphoid tissues. DCs themselves produce chemokines and, depending on their maturation stage, express different subsets of chemokine receptors (Sallusto et al., 1998, 1999; Sozzani et al., 1999; Dieu-Nosjean et al., 1999). For example, immature DCs express a number of receptors (CCR1, CCR2, CCR5 and CCR6) that bind inflammatory chemokines (MIP-1α, MIP-1β, MCP-1, RANTES and MIP-3α) able to recruit DCs to sites of tissue damage or infection. Upon maturation, DCs upregulate another set of receptors (CCR7 and CXCR4) and gain responsiveness to chemokines (ELC/MIP-3β or TCA4/SLC and TABLE 28.2

SDF-1, respectively) that attract DCs to T-cell areas within lymph nodes. DC recruitment to the inflamed CNS may also be regulated by chemokines produced intracerebrally. Induction of chemokine mRNAs or proteins has been demonstrated in a variety of CNS diseases, including trauma, ischemia, Alzheimer’s disease, EAE, MS and AIDS dementia (Glabinski and Ransohoff, 1999). Considerable attention has been given to chemokines produced in EAE and MS, in the attempt to understand their possible role in the recruitment of immune cells, particularly T cells and macrophages, during disease progression (Table

Expression of chemokines and chemokine receptors in the inflamed CNS in EAE and MS

Chemokine CXC or α family GROα/CXCL1 (murine KC)a IP-10/CXCL10 Mig/CXCL9 CC or β family I-309/CCL1 MCP-1/CCL2 MCP-2/CCL8 MCP-3/CCL7 MIP-1α/CCL3 MIP-1β/CCL4 RANTES/CCL5 MIP-3α/CCL20 MDC/CCL22 C10d/CCL6 CX3C or δ family Neurotactin/Fractalkine/ CX3CL1

Chemokine receptor

CNS disease

Chemokine cellular target

References

EAE EAE, MS MS EAE

Neu T T mDC, Mo, T, B

1, 2 1–5 4 6

CCR2 CCR3 CCR1, 2, 3 CCR1, 5 CCR5 CCR1, 3, 5 CCR6 CCR4

EAE EAE, MS MS EAE, MS EAE, MS MS, EAE MS, EAE EAE EAE EAE

Mo, T iDC, Mo, T, NK Mo, T, NK iDC, Eo, Ba, T, NK iDC, Mo, T iDC, Mo, T iDC, Eo, Ba, Mo, T iDC, T, B iDC, mDC, T, NK Mo

1 1, 2, 3, 6–11 9–11 1, 6, 9, 10, 11 1, 2, 4–6, 8, 10–12 1, 2, 4–6, 8, 10, 11 1, 2, 4–6, 8, 10, 11 13 14 15

CX3CR1

EAE

Mo, T, NK

6, 16

CXCR3b CXCR3 CXCR4c

The Table lists chemokines and/or chemokine receptors detected in the CNS parenchyma or CSF of EAE-affected rodents (mice or rats) and individuals with MS. Chemokines are also named according to Zlotnik and Yoshie, 2000. a GROα binds to CXCR2; expression of this receptor in MS and EAE is still unknown. b Detected in MS brains only. c CXCR4 binds the α chemokine SDF-1, whose presence in EAE or MS has not yet been described. DC, dendritic cell; iDC, immature DC; mDC, mature DC; Ba, basophils; Eo eosinophils; Neu, neutrophils; Mo, monocytes; GRO, growth-regulated oncogene; IP-10, interferon-inducible protein 10; Mig, monokine induced by interferon-γ; MCP, monocyte chemotactic protein; MIP, macrophage inflammatory protein; MDC, macrophagederived chemokine; RANTES, regulated on activation normal T cell expressed and secreted. d The receptor for C10 is unknown. References: 1, Godiska et al., 1995; 2, Glabinski et al., 1997; 3, Ransohoff et al., 1993; 4, Sørensen et al., 1999; 5, Balashov et al., 1999; 6, Jiang et al., 1998; 7, Hulkover et al., 1993; 8, Simpson et al., 1998; 9, McManus et al., 1998; 10, Simpson et al., 2000; 11, Rajan et al., 2000; 12, Miyagishi et al., 1995; 13, Serafini et al., 1999, 2000; 14, our unpublished data; 15, Asensio et al., 1999; 16, Pan et al., 1997. DENDRITIC CELLS IN THE PERIPHERY

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28.2). In EAE, mRNA levels for KC, IP-10, MIP-1α, RANTES, MCP-1, MCP-3, and fractalkine/neurotactin increase before or at the onset of clinical signs and, in some cases, remain elevated during the course of disease (Godiska et al., 1995; Glabinski et al., 1997; Pan et al., 1997). MIP-1α production was shown to correlate with acute EAE onset whereas MCP-1 production correlated with relapses (Karpus and Kennedy, 1997). By immunohistochemistry, in situ hybridization or CSF analysis, increased levels of MIP-1α, MIP-1β, IP-10, RANTES, MCP-1, MCP-2 and MCP-3 have been demonstrated in MS brains (Miyagishi et al., 1995; McManus et al., 1998; Simpson et al., 1998; Balashov et al., 1999; Sørensen et al., 1999). In the inflamed CNS, chemokine receptors have been shown to be expressed on different leukocyte subsets and to be upregulated on resident glial cells (Balashov et al., 1999; Glabinski and Ransohoff, 1999; Sørensen et al., 1999). CNSinfiltrating leukocytes, endothelial, neuronal and glial cells have all been demonstrated to contribute to intracerebral chemokine production (Glabinski and Ransohoff, 1999).The specific role of each chemokine in EAE or MS is still unclear, as most chemokines probably act in concert to recruit different leukocyte subsets into the CNS. Among the chemokines upregulated in MS and EAE (Table 28.2), many are known to attract immature DCs to inflammatory sites (Dieu-Nosjean et al., 1999; Sozzani et al., 1999). Using immunohistochemical and RT-PCR techniques, we have shown that MIP-3α, an inflammatory chemokine active on immature DCs, T and B cells (Varona et al., 1998; Dieu-Nosjean et al., 1999 ), is not expressed in the normal CNS but is upregulated during the acute, chronic and relapsing phases of PLP 139151-induced mouse EAE (Serafini et al., 1999, 2000). Intracerebral expression of CCR6, the only known receptor for MIP-3α, was also increased in this model. MIP-3α was mainly localized in inflammatory cell infiltrates during acute EAE, whereas numerous astrocytes in the spinal cord produced MIP-3α during chronic and relapsing disease phases. This switch in the production of MIP-3α, and possibly of other relevant chemokines, could be critical for the sustained

recruitment of DCs, T and B cells into the CNS and for EAE progression. More recently, we have shown that MDC, a chemokine active on immature as well as mature DCs (Sozzani et al., 1999) and its receptor CCR4 are also strongly upregulated in the CNS during acute and chronic/ relapsing phases of EAE (our unpublished observations). It remains to be determined if chemokines regulating migration of mature DCs and naïve T cells to secondary lymphoid organs, such as ELC/MIP-3β and TCA4/SLC (Cyster, 1999), are produced in the inflamed CNS. This could lead to intracerebral T-cell priming as well as formation of ectopic lymphoid tissue (Fan et al., 2000). Although these findings suggest that DC recruitment to the CNS may occur under inflammatory conditions, the critical chemokines remain to be identified. EAE induction in chemokine or chemokine receptor-deficient mice and the administration of chemokine-neutralizing antibodies or chemokine receptor antagonists should help to clarify this issue.

CONCLUSIONS Research reviewed herein strongly implicates DCs in CNS immune surveillance and in the pathogenesis of CNS autoimmunity. While exclusion of DCs from the CNS parenchyma proper makes it less susceptible to potentially damaging cell-mediated immune responses, DC localization at anatomical sites that are constantly exposed to antigens leaving or entering the CNS ensures that this is not totally ignored by the immune system. It would be interesting to know whether CNS-associated DCs sense the specialized neural microenvironment and play a role in the maintenance of CNS immune privilege. Many questions remain to be answered with respect to the contribution of DCs to CNS autoimmune responses. Although studies in the EAE model indicate that DCs have a role in the development and maintenance of CNS autoimmunity, direct evidence for DC involvement in human CNS autoimmune diseases, like MS, is still lacking. The role of different DC subsets in the regulation of CNS autoimmune responses

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and the precise phenotype of DCs recruited to the inflamed CNS also remain to be defined. Another important issue to clarify is the identity of the signals enabling trafficking and maturation of DCs within the CNS. Clarification of these issues will be essential to elucidate the contribution of DCs to CNS autoimmunity. This, in turn, will be instrumental to design therapeutic manipulation of DC function or blockade of DC migration to the CNS. It remains to be seen if DC targeting could be more effective than current immunomodulatory therapies in the treatment of CNS autoimmune disorders. Given the power of DCs in the regulation of the immune response, this may prove to be a worthwhile effort.

ACKNOWLEDGEMENTS The authors thank C.M. Curianò for graphical work. The authors’ work described in this article was supported by Research Project on Multiple Sclerosis and Project ‘Inflammatory, oxidative and autoimmune mechanisms in CNS diseases’ of the Istituto Superiore di Sanità/Italian Ministry of Health.

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PLATE 28.2 Immunohistochemical detection of DCs in the CNS at different stages of EAE development. (A–D) Immunostainings were performed on 10-µm cryostat spinal cord sections of SJL mice immunized with PLP peptide 139–151 emulsified in complete Freund’s adjuvant. (A) Preclinical EAE stage. Staining with anti-CD11c mAb reveals the presence of scattered process-bearing CD11c DCs in the subpial spinal cord white matter; (B, C) Acute EAE. Serial sections stained with MIDC-8 and anti-CD4 mAbs (B and C, respectively) showing colocalization of MIDC-8 DCs and CD4 T cells in perivascular infiltrates. (D) Relapsing EAE. Widespread infiltration of the spinal cord white matter by DEC-205 DCs. (E) Post-relapse phase in a Biozzi AB/H mouse with chronic-relapsing EAE induced by immunization with whole spinal cord homogenate in complete Freund’s adjuvant. Numerous DEC205 DCs are present within a massive inflammatory infiltrate invading most of the spinal cord. Original magnifications, 500 in A, D and E; 1000 in B and C.

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29 Dendritic cells in the eye Paul G. McMenamin 1 and John V. Forrester 2 1

University of Western Australia, Nedlands (Perth), Western Australia; and 2

University of Aberdeen, Aberdeen, UK

Look in mine eye-balls, there thy beauty lies ‘Venus and Adonis’ William Shakespeare

INTRODUCTION

DC function in the eye will precede a discussion of why there appear to be a lack of DCs in the neural retina and how this fits with current concepts on immune responses and inflammation in the eye.

In this review, we will summarise the existing state of knowledge of dendritic cells (DCs) and other potential antigen presenting cells (APCs) within the eye. In this context we will consider the retina and uveal tract of the eye (Figure 29.1), but not the external surface of the eye and lids which more or less resemble skin and mucosal surfaces with respect to DC distribution and function. The first part of the review describes the organisation of the eye in relation to how immune responses are mediated via the retinal vasculature and uveal tract. A discussion of the likely candidates for APCs in the eye will first consider the evidence for whether microglia are specialised macrophages or represent a type of DC. Other macrophage populations, such as perivascular cells and uveal tract macrophages, will be briefly discussed with relevance to their possible role as APCs. The identification and location of DCs in the uveal tract of the eye will then be addressed. The factors which regulate Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

The general layout of the retina and uveal tract of the eye The eye consists essentially of three layers: the retina is the innermost layer surrounded by the uveal tract (choroid posteriorly and ciliary bodyiris anteriorly) which are both protected by the outer fibrous envelope, consisting of the cornea and sclera (Figure 29.1). The retina consists of two primary layers, an inner neurosensory retina and an outer simple epithelium, the retinal pigment epithelium (RPE). Between the neural retina and RPE is a potential space, the subretinal space, across which the two layers form a loose adhesive contact. The retina consists of several cell types of which neural cells, including photoreceptors,

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bipolar, horizontal, amacrine and ganglion cells form the major components. However, other cell types are also present such as glial cells, vascular endothelium, pericytes and microglia. The choroid consists of vascular pigmented connective tissue with larger vessels externally and a rich bed of large diameter fenestrated capillaries, the choriocapillaris, lying directly beneath the RPE. The ciliary body consists of a smooth pars plana which extends from the anterior limit of the retina to the pars plicata anteriorly. The latter consists of 120 circumfer-

FIGURE 29.1

entially arranged radial ridges, known as ciliary processes, which surround the lens (Figure 29.1). The ciliary processes secrete aqueous humour (AqH) (2–4 µl/min) which circulates around the lens before it passes through the pupil into the anterior chamber from where it leaves via the aqueous outflow pathways. The ciliary processes consist of a connective tissue stroma, rich in fenestrated capillaries covered by a double layer of neuroepithelium which forms an important part of the blood–aqueous barrier (Figure 29.1). The ciliary body is frequently the site of early

Schematic diagram summarizing the anatomy of the eye and the sites of the blood–ocular barriers. DENDRITIC CELLS IN THE PERIPHERY

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inflammatory changes in both clinical anterior and posterior uveitis and the experimental models of these conditions (Nussenblatt, 1991; Forrester et al., 1995). The iris is the most anterior portion of the uveal tract and forms a perforated diaphragm between the anterior and posterior chambers and serves to regulate, via changes in pupil diameter, the amount of light entering the eye. It consists of an incomplete cellular anterior border layer, a highly vascularised and richly innervated pigmented connective tissue stroma and posteriorly a thin layer of smooth muscle, the dilator pupillae, lies approximated to the posterior iris pigment epithelium. A circumferential band of smooth muscle, the sphincter pupillae, lies close to the pupil.

The ‘immune privileged’ status of the eye The interior compartments of the eye have been recognised for decades as possessing unusual immune status or ‘privilege’ (Barker and Billingham, 1977). It has been postulated that ocular tissues, vital for survival but without regenerative capacity, have evolved mechanisms to regulate the nature and magnitude of local immune responses unlike other tissues where ‘bystander’ damage and oedema that accompany acute inflammatory or cell-mediated immune responses may have less significant functional consequences. Mechanisms proposed to underlie this immune privilege include. (1) The blood–ocular barrier (BOB), comprising of the blood–retinal barrier and the blood– aqueous barrier (Figure 29.1); (2) the absence of specialised lymphatic drainage in the eye; (3) the absence or low expression of major histocompatibility complex (MHC class I and II) molecules; and (4) the absence of DC in the neural parenchyma. (1) The BOB functions to regulate the passage of macromolecules (drugs, hormones, high molecular weight proteins), microbial pathogens and intravascular leukocytes from the lumen of

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vessels into the extravascular tissue compartment. The restricted entry of leukocytes into the neural retina and aqueous humour was considered a central tenet of the concept of ‘immune privilege’, namely the neural environment and the tissues of the anterior segment of the eye were protected from surveillance by immune cells. The BOB primarily serves to regulate for an optimal extracellular environment facilitative of neural transmission (Rowland, 1985). (2) The small volume of extracellular space in the retina reduces the requirement for a lymphatic system to remove excess extracellular fluid, although the AqH in the eye is widely regarded as a form of ‘lymphatic system’. The absence of a lymphatic drainage within the eye (and brain) was thought to permit sequestration of antigens in these organs thereby blocking the afferent limb of the immune response thus partly explaining the phenomenon of ‘immune privilege’ (Medawar, 1948). Aqueous humour drains via the iridocorneal angle into Schlemm’s canal (or sinus venosus sclerae in most nonprimates), and thence into collector channels and episcleral veins. Antigens (Ag), either soluble or perhaps associated with ocular DC (see below), leaving the anterior chamber via this route are thought to travel to the spleen where they elicit antigen-specific immune deviant responses (see reviews, Streilein, 1993, 1999). However, there is recent evidence from studies utilising congenic mice that antigens placed in the anterior chamber stimulate antigen-specific T-cell clones in the submandibular lymph nodes (Egan et al., 1996), thus suggesting communication with lymphatic vessels. It is also possible that antigen may leave via the nonconventional AqH outlflow pathways, passing posteriorly into the suprachoroidal space from where fluids may escape via the loose connective tissue surrounding nerves and vessels piercing the sclera. Interestingly there have also been rare reports of lymph channels in the mammalian choroid (Krebs et al., 1988) which seems plausible as there are extensive lacunae in the avian choroid which have the characteristics of lymphatic vessels (De Stefano and Mugnaini, 1997). Indeed the choroid could be regarded as the

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lymphovascular organ of the eye (Forrester and McMenamin, 1999). The phenomenon of anterior chamberassociated immune deviation (ACAID) which results following experimental instillation of antigen into the anterior chamber of the eye in a range of species (see reviews, Streilein 1993, 1999), has been proposed as a paradigm of immune deviation which underlies the immunological tolerance to antigens placed in the anterior chamber. It is proposed that following antigen administration, an antigen-specific ACAIDinducing signal leaves the eye and traffics to the spleen, where regulatory T lymphocyte responses are induced (Ferguson et al., 1987). In mice receiving intracameral soluble antigen (viz. injected directly into the anterior chamber) this signal is reported to be cell-associated and depletion studies suggest the relevant antigenic signal was carried by F4/80 cells (presumptive macrophages) (Wilbanks et al., 1991). However, based on existing knowledge of macrophage function and new data on the APC capacity of iris-derived macrophages (Steptoe et al., 2000) it is more likely that ocular DC act as the sentinel cells carrying antigen-specific signals from the eye. (3) Until recently the apparent paucity of MHC class I and class II expression in the retina and supporting tissues, including the tissues lining the anterior chamber, appeared to fit with an earlier proposed mechanism of immune privilege in the eye (and brain) which stated that T cells entering the neural environment would be unable to recognise endogenous or exogenous antigens and thus be prevented from entering a state of activation (for review see Sedgwick, 1997). Despite the apparent specialisations in the eye described above, which would seem to impose limitations on the afferent and efferent limbs of the immune response, ocular inflammatory responses do occur to both endogenous autoantigens (S-antigen, rhodopsin, interphotoreceptor retinol binding protein, recoverin, phosducin, lens proteins) and exogenous antigens derived from infectious agents (viral, bacterial and protozoan). Thus antigen

sampling and processing clearly takes place somewhere within the eye. The precise identity of the APCs in the eye, which transport antigen to the secondary lymphoid tissues, has been the subject of much interest in ocular immunology. (4) It has been proposed that DCs play little or no role in immune responses within the CNS (Steinman, 1991). This stands in marked contrast to the pivotal role of these cells in regulating immune responses in other tissues. The view that the eye (and brain) was devoid of MHC class II DCs derives from earlier immunohistochemical investigations of class II expression in conventional histological or frozen tissue sections in a variety of species in which only rare scattered or isolated cells were identified (see reviews McMenamin 1994, 1997; McMenamin and Forrester, 1999). However, as we will discuss presently, there are numerous potential APCs within the tissues which comprise the eye (vide infra). Thus it is probably more accurate to state that this ‘deficiency’ in the CNS and eye is relative and that routes of interaction with the immune system do indeed exist but are more tightly regulated than in other organs.

Factors that regulate entry of immune cells into the eye While the concepts of the blood–retinal barrier and blood–aqueous barrier have been fashioned on the basis of exclusion of transport of large molecules from the blood stream into the parenchymal tissue of the retina and aqueous humour (Greenwood, 1992), the assumption has been that these barriers naturally extend to cells. Inflammatory processes frequently occur in the brain and retina, thus it would appear that these barriers are readily broken down. Furthermore, contrary to earlier notions there is recent evidence that normal CNS tissue is ‘patrolled’ by lymphocytes (Sedgwick, 1997) although the low incidence of extravasated lymphocytes in the normal CNS parenchyma means that they are rarely likely to be encountered in histological sections and it is likely that the infiltrating T cells are activated or blast cells since resting T cells do not normally enter the tissues (Sedgwick et al.,

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1998). How such cells gain access to the normal brain and retina is not clear. During inflammation or pathological conditions affecting the CNS alterations in the adhesion properties of the endothelium occur which facilitate leukocyte extravasation. There is good in vitro evidence that cells close to the CNS vessels, such as astrocytes and perivascular macrophages (Figure 29.2), when activated are capable of synthesising cytokines, such as TNF, that may cause upregulation of adhesion molecules such as ICAM1 and E-selectin (Joo, 1994). The fact that in autoimmune conditions, such as MS or uveitis, autoreactive T cells enter the CNS, has lent support to the concept of regular T-cell trafficking in the brain (see reviews, Sedgwick, 1995, 1997). During and indeed prior to overt inflammation in the eye several processes are entrained which ensure that there is access not only of lymphocytes, but several other subsets of inflammatory cells such as monocytes, neutrophils, NK cells and DCs. Inflammatory cells are attracted to the site of damage by chemokines

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(e.g. MIP-1α) and other molecules released by tissue cells such as astrocytes, microglia and retinal pigment epithelial cells (Miyagishi et al., 1997; Gourmala et al., 1997; Crane et al., 1999). Endothelial cells also participate in this process by releasing chemokines but more importantly by altering their adhesiveness for circulating leukocytes by expressing specific adhesion molecules for leukocyte integrin receptors such as ICAMs-1,2 and 3, VCAM and E-selection (Mesri et al., 1994). Morphological changes occur during this process which have been compared with the appearance of high endothelial venules (HEVs) in lymph nodes (McMenamin et al., 1992, 1993). Considerable debate has centred on whether leukocyte emigration in CNS/retinal endothelium occurs trans-cellularly or para-cellularly at the site of the disengaged tight junctions (Greenwood, 1992). As stated above, under normal circumstances a few lymphocytes, albeit activated, appear to have access to the CNS tissue which enables them to ‘patrol’ the tissue for organisms, such as

FIGURE 29.2 Schematic diagram of a blood vessel in the brain parenchyma illustrating the components of the vessel wall which contribute to the blood–brain barrier. From the lumen outwards these are the vascular endothelial cells (EC), basal lamina (shaded zone) and the glia limitans composed of astrocyte (A) foot processes. Note the relative position of perivascular microglia (MG) and perivascular macrophages (PVM). DENDRITIC CELLS IN THE PERIPHERY

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viruses and bacteria. It is assumed that a low grade patrol mechanism also operates in the normal retina. We have recently shown that low numbers of interferon-γ-positive T lymphocytes appear to infiltrate the retina prior to the onset of inflammation in a model of experimental autoimmune uveoretinitis (EAU) induced by a preparation of mixed retinal antigens. This is followed by strong expression of chemokines on the endothelial cells at the sites of inflammation (MIP-1α and MCP-1) (Crane et al., 2000) (Plate 29.3). Monocytes/macrophages also appear to be capable of locating extravascularly in the perivascular spaces in the brain and the retina (Dick et al., 1995). However, not all cells appear to have similar access to the CNS. In particular DCs appear to be excluded from the retina, but clearly migrate freely into other non-neural tissues in the eye including the uveal tract (vide infra). The reason for their exclusion from the neural retina is not clear but may relate to their relative lack of integrin receptor expression in the resting state (Ruedl and Hubele, 1997). The extent of T-cell trafficking in the uveal tract with its less developed vascular barriers (including a ‘leaky’ choriocapillaris) would be expected to be greater than in neural tissues. However, lymphocytes were originally reported to be absent or rare in the normal uveal tract of rodents and in human iris biopsies studied by conventional histological methods. However, recent immunohistochemical staining of ocular tissue wholemounts has revealed a consistent but low density of T cells (4 cells/mm2 in the rat iris, 20 cells/mm2 in the normal rat choroid) (McMenamin and Crewe, 1995; Butler and McMenamin, 1996). Newly emerging evidence has shown that many cells and tissues in the intraocular environment (retina, cornea) are pro-apoptotic which may act as a powerful mechanism of regulating infiltration of T cells and other immune cells into the intraocular tissues and compartments (Griffith et al., 1995; D’Orazio et al., 1999). There is good evidence that when Fas+ lymphocytes infiltrate the eye and come into contact with Fas ligand positive tissues lining the anterior chamber they undergo apoptosis in situ. This is also true for other

FasL positive cells in the eye such as the retinal pigment epithelium. When immune cell-mediated damage occurs in the eye, there is clear evidence of T cell activation, but the role of putative resident APCs is largely unknown. Candidate APCs include parenchymal cells such as astrocytes, oligodendrocytes and endothelium and nonparenchymal haematogenous-derived immune cells including microglia, perivascular macrophages and other macrophage populations. Only the nonparenchymal immune cells will be dealt with in detail in this review. The potential role of parenchymal cells as APCs in the context of the CNS has been extensively discussed by Sedgwick and Hickey (1997). Furthermore there is recent evidence that astrocytes rather than acting as APCs may have a role in downregulating the immune response (Aloisi et al., 2000).

MICROGLIA Current concepts on retinal microglia (phenotype, distribution, ontogeny) Recent advances in our understanding of microglial cells have been derived largely from data in the brain parenchyma and retina. The retina, being an extracerebral portion of the brain and being flat, is comparatively accessible and lends itself to forms of experimental manipulation not possible with the brain. For example, retinal flatmount preparations aid in the display of the regular array of microglia (Figure 29.4). Scholarly reviews of the history of the discovery, characterisation, origin and nature of microglia have been recently published (Thanos et al., 1996; Becher et al., 2000). Microglia are a stable population of highly ramified or dendriform cells of bone marrow origin. They have nonoverlapping territories with neighbouring microglia. Microglia, along with perivascular cells, are believed to represent the resident macrophages of the neural parenchyma (Flaris et al., 1993; Cuzner, 1997). It is estimated that the mouse brain contains 3.5  106 microglial cells, a figure comparable to the

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FIGURE 29.4

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‘Plan’ view of microglia in retinal wholemount stained with the mAb Ox42.

number of macrophages in the liver. Within the parenchyma of the retina (and brain) microglia express the complement receptor 3 antigens (CD11b in rat and CD11c in human) and are CD45low in the rat. There has been some controversy over whether microglia express the resident macrophage marker ED2. However, it would appear that the consensus opinion is that they are ED2-negative (McMenamin et al., 1992b; Dick et al., 1995; Zhang et al., 1997; Yang et al., 1999) and that the retina lacks significant populations of ED2-expressing macrophages apart from those in the perivascular space (see above). The extent of expression of MHC class II by microglia is also controversial (see McMenamin and Forrester, 1999). Studies on retinal microglia indicate that they are generally MHC class II negative at least in mice and most rat strains (McMenamin et al., 1992b; Choudhury et al., 1994; Zhang et al., 1997), but in humans retinal parenchymal and perivascular microglia appear to express some MHC class II. This may in part relate to post-mortem changes which are of necessity unavoidable in most situations. In summary, it appears that subpopulations of microglia, especially perivascular microglia, express MHC class II, albeit at low levels, in some species/strains. In addition, microglia have been reported to express accessory molecules B7/B7.1 (Williams et al.,

1994) although there is counter-evidence that they lack these co-stimulatory molecules and B7.2 (Inaba et al., 1995). Matsubara et al. (1999) have recently shown that freshly isolated microglia in culture express ICAM-1 and B7-2 but require stimulation with IFN-γ before they express B7-1 and MHC class II. However, in similar studies we have found the opposite effect, i.e. that human retinal microglia, cultured in the presence of IFN-g downregulate their expression of MHC class II, as well as other costimulatory molecules required for antigen presentation such as CD40 and CD86 (Clark et al., 1985; Broderick et al. 2000). In addition, cultured human microglia appear to elaborate significant quantities of IL-10 in vitro, supporting an anti-inflammatory rather than a proinflammatory role for these cells. More recently, we have also shown that human retinal microglia express the neuronal– leukocyte ligand OX2 (Dick et al., 2001). OX2 is an integral membrane protein which is expressed on a variety of cells including follicular dendritic cells, B cells, endothelium and neurones (Clark et al., 1985). It is a member of the immunoglobulin superfamily of trans-membrane proteins with an Ig domain which has homology to similar costimulatory molecules such as CD2, CD150, 2B4, CD80/86 (Borriello et al., 1997). The OX2 receptor is exclusively expressed on cells of the myeloid

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lineage especially monocytes/macrophages (Preston et al., 1997) and it has been suggested that the role of OX2 in the CNS is to regulate macrophage-mediated damage. Thus the expression of OX2 on retinal microglia may indicate a function in down-regulating immune responses.

The function of microglia It has been suggested that in the resting state microglia have a constitutive role in ‘cleansing’ extracellular fluid in the CNS, for example by degradation of neurotransmitters, and maintaining a state of ‘vigilance’ by monitoring changes in their extracellular milieu (Kreutzberg, 1996). Indeed pinocytosis is often used as a differential marker for microglia. When activated, microglia assume a more amoeboid form upregulate macrophage scavenger receptors and actively phagocytose cell and tissue debris (Thanos et al., 1996; Becher et al., 2000). This occurs in a number of situations ranging from normal development, where they phagocytose apoptotic neurons, to a variety of degenerative, traumatic or inflammatory conditions in the CNS (see reviews Nakajima and Kohsaka, 1993; Thanos et al., 1996). There is, however, some speculation that they may also have features resembling DCs and since myeloid DCs and macrophages are derived from the same lineage this is not surprising (Vandenabeele et al., 1999). In the eye, immunohistochemical and histochemical studies of retinal development suggest that monocytes enter the neural retina from the overlying developing vasculature to scavenge debris produced by neuronal cell death. They then differentiate to form a regularly spaced network of microglia in the plexiform layers (Hume et al., 1983; Sanyal and De Ruiter, 1985; Thanos et al., 1996). The discovery of macrophages in the peripheral subretinal space in the developing eye (McMenamin and Loffler, 1990; McMenamin, 1999a) led to the proposal that another route of entry for microglial precursors in the retina (besides the vessels at the optic nerve head) was the rich vascular bed of the developing ciliary body, which is homologous to

the choroid plexus. This proposal is supported by immunohistochemical studies of the human foetal retina (Diaz-Araya et al., 1995). Microglia secrete proteases and generate free radicals and nitric oxide species. Indeed some have termed them the ‘deadly’ killers (Banati and Graeber, 1994) based on evidence from in vitro cytotoxicity studies of activated microglial interaction with tumour cells, neurons and oligodendrocytes. It is possible that microglia, during ontogeny may play a role beyond that of ‘cellular undertakers’ and may be more actively involved in mediating cell death similar to the speculated role of the macrophages that remove the tunica vasculosis lentis surrounding the foetal lens (Lang and Bishop, 1993). It is clear that strict regulation of this cytotoxic function in vivo is crucial if these cells are to be effective and not cause unwanted damage to the CNS itself. Incertaininflammatoryresponsesintheretina, microglia are likely candidates for the synthesis of pro-inflammatory cytokines and chemokines (Zhang et al., 2000). However, their role in the steady state may be to down-regulate or limit inflammatory responses via their recently demonstrated ability to induce apoptosis of activated T cells (vide infra).

Retinal microglia may act as APCs in vitro, but they are not prominent as APCs in the inflamed retina and they are not DCs Since DCs, currently considered to be the most potent APCs in the immune system, were reportedly absent from CNS (including the retina) and supporting tissues, considerable effort has been made to identify cells which can act as APCs in this environment. Microglia, on the basis of their morphology and lineage, initially appear to be good candidates. There is evidence that human microglia can stimulate resting T cells (Ulvested, 1994; Williams et al., 1994), however, studies with ex vivo rat brain microglia prepared using FACS technology by gating on a population of CD45low, MHC class II cells have shown they are poor APCs in the presence of MBP-specific CD4 T cells, as measured by T cell proliferation and IL-2

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secretion (Ford et al., 1995). Some authors have documented APC activity by microglia in vitro, although these have tended to require stimulation by interferon-gamma (IFNγ) or LPS (Cash et al., 1993; Cash and Rott, 1994; Matsubara et al., 1999). In addition, our recent data (see above) suggest that microglia downregulate expression of MHC class II and co-stimulatory molecules in response to IFNg rather than increase the expression of these molecules. By contrast it has recently been proposed that MHC class II expression on microglia is directed at cutting short T-cell activation and proliferation and inducing apoptosis thereby minimizing the inflammatory response in the CNS (Ford et al., 1996). In the inflamed retina, infiltrating marrow-derived myeloid DCs appear to be the predominant APC (Jiang et al., 1999) (see below). The migratory phase of the DC life cycle is vital to their sentinel function (see reviews, this volume). Therefore, if retinal microglia were the equivalent of DCs in the retina one would predict a short half-life and high turnover. However, a quantitative analysis of the normal rate of turnover of retinal microglia obtained using radiation chimera models indicates very low turnover (months) (Zhang et al., 1997) similar to CNS microglia (Lassman et al., 1993). Thus on the criteria of turnover, microglia do not appear to have a life cycle akin to DCs. In conclusion, on the basis of immunophenotypic characteristics, assays of antigen presenting function, extremely low turnover rates and their phagocytic capacity most authors have concluded that microglia represent specialised resident tissue macrophages of the CNS parenchyma and not DCs. There is, however, some speculation that they may also have features resembling DCs and since myeloid DCs and macrophages are derived from the same lineage this is perhaps not surprising (Vandenabeele et al., 1999).

PERIVASCULAR CELLS Vessels in the CNS are characterised by a perivascular concentration of astrocytic foot

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processes which forms the ‘glia limitans’ external to the endothelium (Figure 29.2). A small subpopulation of perivascular macrophages or ‘perivascular cells’ is associated with these vessels. Their phenotype indicates they are a less down-regulated form of macrophage than microglia (Graeber et al., 1989) with some of the characteristics of interstitial macrophages. In the retina, Dick et al. (1995) using flow cytometry described a population of ED2/class II and ED2/class II cells that could represent perivascular cells. Some authors regard microglia close to vessels as a distinct subpopulation of macrophages from perivascular macrophages (Provis et al., 1995). Perivascular macrophages in the brain are derived, like other macrophages, from precursor cells in the bone marrow, but appear to have a relatively short turnover compared to parenchymal microglia (Lassman et al., 1993). No specific data are available on turnover of retinal perivascular macrophages, but it is likely they are similar to those in the brain. Brain and retinal perivascular macrophages have attracted interest as potential APCs in the brain parenchyma and retina almost by default, since there is little evidence for such a role for retinal microglia (see above). In addition, in clinical and experimental inflammation of the brain and retina, the initial site of lymphocyte accumulation in the parenchyma appears to be the perivascular space suggesting that this may be the initial site of uptake and processing of antigen for transfer to naïve or memory T lymphocytes in the lymph node. A fast-replenishing population of cells is required for such a function which effectively excludes microglia and leaves few other cell types apart from the perivascular macrophage, assuming that in fact they do have a rapid turnover. It is possible that in conditions of established inflammation there might be some degree of antigen presentation by local resident cells, such as microglia, but their role would be relatively minor since under these circumstances there is active and instant recruitment of professional bone marrow-derived DCs (Jiang et al., 1999).

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MACROPHAGE POPULATIONS IN THE EYE OTHER THAN RETINAL MICROGLIA Hyalocytes A population of macrophages (ED2 in rats) is situated in the subhyaloid space (between the inner retinal surface and the vitreous ‘membrane’). They are considered scavengers and probably arise from the population of macrophages that phagocytose the hyaloid vessels and tunica vasculosa lentis during development. It is worth noting that they may act as a source of contamination in ‘retinal preparations’ if all remnants of the vitreous are not carefully removed. They also appear to have anti-proliferative properties for other cell types such as retinal pigment epithelium (Lazarus et al., 1996).

Macrophages in the iris, ciliary body and choroid Conventional histological studies reveal macrophages in the human iris stroma. They phagocytose melanin shed from iris pigment epithelium throughout life especially during iris movements where it comes into contact with the lens. More recently, immunohistochemical studies of wholemounts have revealed that the normal rat and mouse iris contains a rich network (600–800 cells/mm2) of resident tissue macrophages (McMenamin et al., 1992b, 1994; McMenamin, 1999b). These cells are ED1, ED2 and ED3 (rat) or F4/80, SER4 in the mouse. Macrophages of a similar phenotype are also distributed in the connective tissue of the trabecular meshwork and stroma of the ciliary body and ciliary processes in the rat, mouse and human eye (McMenamin et al., 1992b, 1994; McMenamin, 1999b). Like the anterior uveal tract the connective tissue stroma of the normal rodent choroid possesses a rich population of ED1, ED2 and ED3 (rat) or F4/80, SER4 (mouse) resident tissue macrophages (approximately 600 cells/mm2) that display a largely perivascular distribution (Forrester et

al., 1993, 1994; Butler and McMenamin, 1996; McMenamin, 1999b) (Plate 29.5). The distribution pattern of the resident tissue macrophages close to vascular beds suggests a guardian role at the blood–tissue interface. Recent data from our laboratory expand our earlier functional analysis of resident APCs within the iris (Steptoe et al., 2000) and provide insight into the relative roles of these two populations, namely macrophages and DC, in primary and secondary immune responses. Several groups have shown that activated macrophages associated with body cavities or mucosal surfaces secrete a range of soluble mediators that inhibit T-cell proliferation (Pavli et al., 1990; Soesatyo et al., 1991; Holt et al., 1993). In in vitro assays of freshly isolated resident tissue macrophages it was found that these cells exhibited a phenotype which lacked lymphocytostatic properties, but possessed the ability to ingest, process and effectively present soluble antigen to antigen-specific T cells. This is consistent with previous immunomorphological studies which defined the phenotypeofthesecellsas‘resident’tissuemacrophages (McMenamin et al., 1994). These new functional datasuggestthatirismacrophages,unlikemucosal or ‘body cavity’ macrophages do not produce nitric oxide in response to T cell-derived signals, suchasIFN-γ.Thismaybedueto their exposure to high concentrations of TGF-β and/or calcitonin gene-related peptide (CGRP).Whether uveal tract macrophages possess CD14 and thus the potential to bind LPS or LPS-binding protein is unknown. If this were the case, these macrophages could rapidly initiate the inflammatory cascade in the eye via subsequent cytokine release as a consequence of contact with bacteria or bacterial products leaking from fenestrated ocular vessels in the ciliary body and choroid. In the choroid, macrophages are poor APCs, but recent studies suggest that they may modify the primary mixed leukocyte response induced by DC (Forrester et al., unpublished observations). This may be of particular importance in light of their location at the interface between the BOB and the fenestrated vascular beds of the

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choroid and ciliary processes where inflammatory infiltrates are frequently noted in clinical and experimental uveitis (see Forrester and McMenamin, 1999, for full discussion).

DCs IN THE RETINA AND UVEAL TRACT OF THE EYE Distribution and phenotype Following the discovery that MHC class II expression was important to the outcome of transplantation and in the aetiology and pathogenesis of autoimmune and inflammatory mediated diseases, many groups were quick to investigate the pattern of expression of this molecule in normal and diseased ocular tissues. Most of these studies, performed on conventionally sectioned tissue, either failed to reveal any MHC class II cells or revealed only occasional, scattered cells in the normal eye (see McMenamin and Forrester (1999) for review). More recently a contiguous network of MHC class II DCs has been described in the iris and ciliary body (Knisely et al., 1991; McMenamin et al., 1992b, 1994). In all species examined, iris DCs display a variety of forms from pleomorphic to the characteristic highly dendriform morphology with multiple, often branched cytoplasmic processes and indented nucleus (Figure 29.6). The cells are regularly spaced from their neighbours, but do not display as strong a predilection for the perivascular orientation as resident tissue macrophages. The density of the DC network (400–600 cells/mm2) is similar to other well recognised DC populations (e.g. skin 700–800/mm2, tracheal epithelium 670–880/mm2, oral mucosa 160–890/mm2). Double colour immunohistochemical studies (McMenamin et al., 1994) have revealed that DC do not co-localise with antimacrophage monoclonal antibodies and only a small subpopulation of macrophages express MHC class II in the normal rat (McMenamin et al., 1994) and mouse eye (McMenamin, 1999). Immunoelectron microscopic and confocal studies have revealed that DCs in the ciliary processes are

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intraepithelial on the vascular aspect of the tight junctions that form the blood–aqueous barrier. Thus they are ideally situated at this interface to sample antigens either from intraocular compartment or blood-borne antigens arriving via the fenestrated vascular bed. Studies in mice have shown that only a few DCs are CD80, CD86 and β2 integrin and even then only weakly positive (McMenamin, 1999a) supporting the suggestion that ocular DC are in the ‘immature’ stage of their life cycle in which their primary role is antigen capture. Furthermore, in view of the known function of DCs in the induction of tolerance, it is also possible that the primary role of ocular DCs is to promote tolerance rather than immunity as has been described for another immunologically privileged tissue, the liver (Knolle and Gerken, 2000).  Low to moderate densities of MHC class II dendritiform cells have been identified in the rat (McMenamin and Holthouse, 1992) and human (Flügel et al., 1992) trabecular meshwork and around episcleral vessels and collector channels which serve to drain AqH into the venous blood. DCs at these sites would be ideally located to sample antigens exiting the eye and indeed recent experiments using in vivo video fluorescence microscopy following intracameral injection of fluorescent labelled antigens reveals that much of the antigen is deposited or accumulates in the trabecular meshwork and conjuctival/ episcleral tissue (Figure 29.7). The precise nature of these cells is yet to be determined but preliminary investigations suggest conjuctival/limbal Langerhans cells (DCs) and resident tissue macrophages both take up antigens. It has been known for some time that proteins injected into the AC leak from limbal vessels (Sherman et al., 1978), making it feasible that DC and macrophages around collector channels and episcleral vessels have access to intracameral antigens. These cells could then migrate, via conjunctival lymphatics, to draining submandibular lymph nodes, thus bypassing the ‘camero-splenic axis’. Support for this postulated route has recently come from the elegant chimera experiments of Egan et al. (1996) in which antigen-specific T cells in the submandibular lymph nodes

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FIGURE 29.6 Wholemount of mouse iris and ciliary processes (CP) stained with M5/114 (anti-MHC class II). A, low power; B, high power of individual DC in iris; C, high power of individual DC in ciliary processes (CP). DENDRITIC CELLS IN THE PERIPHERY

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FIGURE 29.7 Single captured video frame from in vivo video fluorescent microscopy of a rat eye which had been injected with Cascade blue-Dextran (70 KDa) 5 days previously. Note the staining in the iridocorneal angle and limbal region as well as scattered cells in the iris. Photographed with 20 objective lens.

clonally expand following intracameral injections. This may represent another pathway, besides the spleen, whereby antigens in the eye become accessible to the systemic immune system. Recent studies from our laboratory using a GFP-labelled DNA vaccine incorporating the human C5 protein which is expressed, but not secreted by cells taking up the vaccine, indicate that after instillation into the anterior chamber of the eye the label is detectable in CD11c positive eye-derived DCs in the draining lymph node, thus strongly supporting the above mode of antigen traffic from the eye to the lymph node inside DCs (Forrester et al, experiments in progress). It must also be considered however whether leakage of episcleral and limbal vessels is a consequence of the ocular injury response due to the invasive nature of intracameral injections.

Ontogeny, migration capacity and function of anterior uveal tract DC The development of Ia DCs in the aqueous outflow pathways (McMenamin and Holthouse, 1992) and the iris (Steptoe et al., 1997) of the rat

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eye may occur more slowly than that observed in other tissue sites. It appears that DCs infiltrate  the iris primordium as Ox62/Ia or Ialo precursor cells, which subsequently develop in situ into DCs bearing high levels of Ia antigen (Ox62/Ia) (Steptoe et al, 1997). In the eye the rate of development of Ia iris DCs was greatest between postnatal days 13 and 21 which coincides with eye-opening (day 14) a major event in the development of ocular function which may impinge on the intraocular environment via components of the photodetection cascades, alteration in neuropeptide levels or via exposure of the globe to the external environment/ exogenous antigens. Bone marrow ablation studies have shown that iris DC have a half-life of 2–3 days in comparison to 10 days for epidermal Langerhans cells (LCs) (Steptoe et al., 1996). It appears that iris DCs like those in the respiratory and alimentary tracts (Schon-Hegrad et al., 1991) respond within 12–24 hours to inflammatory signals by altering their phenotype, increasing their density (via immigration) and turnover time (McMenamin and Crewe, 1995). The net effect of this response is presumably to enhance the efficiency of immune surveillance at crucial times of viral or bacterial infection. Functional studies have shown that iris DCs purified by positive immunomagnetic selection have the functional capacity to stimulate naïve T cells in a mixed lymphocyte response (MLR) similar to conventional DCs, such as LCs (Steptoe et al., 1995). Iris DCs, like other ‘immature’ DCs in peripheral tissues which are weak APCs, require appropriate cytokine maturational signals, such as GM-CSF in order to display potent stimulatory capacity in an MLR assay. This evidence taken together with the recent studies of APC activity of freshly isolated iris macrophages compared to fresh iris DCs suggest the two cell populations perform complementary but discrete roles in the eye: macrophages acting as local APCs in secondary immune responses, whilst DCs perform a sentinel function trapping antigen in the eye and subsequently migrating to the spleen or draining lymph nodes thereby regulating

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antigen-specific immune responses. To confirm this hypothesis it is necessary to know whether DCs in the iris, ciliary body and outflow pathways have access to ocular autoantigens such as S-antigen or IRBP. There is evidence that S-Ag is detectable in human AqH (Zaal et al., 1986) and more recent in situ hybridisation evidence has shown that S-Ag cDNA is present in porcine iris and ciliary body (Singh et al., 1996). Thus it seems plausible that local DCs would have access to these autoantigens in addition to other endogenous autoantigens (e.g. lens proteins) or exogenous antigens derived from unicellular and multicellular pathogens which invade the intraocular compartments. Recent experiments performed in our laboratories have aimed to determine the ability of ocular DCs to capture antigens both in vivo and in vitro. These studies (unpublished) have revealed that in freshly isolated iris explants exposed to a variety of fluorescent-labelled antigens of different molecular weights (e.g. Cascade blue-Dextran 70 kDa; Texas red-Dextran 70 kDa; Texas redBSA) DCs can capture antigen within 30 min. Over time, labelled DCs migrate from the iris explants into the culture medium. DCs in iris explants cultured for 24 or 48 hours before antigen exposure are less avid in their uptake of antigen. Iris macrophages also capture antigen and internalise it in large phagosomes, but they migrate less readily from the explants. Labelled

FIGURE 29.8 Double immunofluorescence of iris DC 30 min after intracameral injection with Texas-red labelled BSA. Left panel, Ox6 cells, FITC fluorochrome; right panel, same field photographed at Texas red wavelength. Note many Ox6 cells contain antigen even after only 30 min exposure.

antigens injected into the anterior chamber in vivo are also readily taken up by DCs and macrophages which can be identified directly by in vivo fluorescent video microscopy (Figure 29.7) and by subsequent immunostaining of iris specimens from these eyes (Figure 29.8). These studies have shown that DCs and macrophages in the iris are capable of trapping antigen. What remains to be determined though is whether one or both cell types migrate to either the spleen and/or cervical lymph nodes. Studies to clarify this are currently underway in our laboratories.

Dendritic cells in the normal choroid and their relevance to ocular autoimmune Networks of DCs have been described in the choroid of the rat (Forrester et al., 1993; Butler and McMenamin, 1996) and mouse (McMenamin, 1999a). In the rat, choroidal DC (746 ± 37 cells/mm2), like those in the iris, express MHC class II and in addition there is a small population of OX62 round precursor cells indicating that their turnover is likely to be similar to that in the iris (Steptoe et al., 1996), however specific studies of choroidal DC turnover have not been performed although preliminary observations in our laboratory using rat chimeras suggest that in common with iris DCs they are replenished by bone marrowderived precursors. Choroidal DCs have at least two phenotypes: a very large veiled cell with extensive ruffled membrane and a smaller more typical cell with dendritic processes and high levels of cell surface MHC class II antigen (Forrester et al., 1994). Both cell types are located perivascularly in the choroid and immunoelectron microscopic studies have shown that they possess fine processes which lie directly adjacent to the basal aspect of the retinal pigment epithelial cells and Bruch’s membrane. Therefore although DCs are normally excluded from tissue sites (like the retina) which contain potent autoantigens, their strategic location on the basal aspect of the RPE would place them in an ideal position to sample retinal proteins

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which are continually being phagocytosed by the RPE. The normal presence of low levels of serum antibodies to S-antigen indicates some APC-antigen exposure must occur (Forrester et al., 1989) and may in fact represent a mechanism for tolerance rather than immunity. Functional studies of rat DCs isolated from preparations of the entire posterior segment (retina, choroid and sclera) suggest they are functionally similar to iris DCs as assessed by mitogenesis assays and allostimulatory capacity in MLR assays (Choudhury et al., 1994). Relatively pure populations of DCs can be isolated from the rat and human choroid by culture of choroidal explants in vitro in the presence of foetal calf serum, from which motile cells migrate into the medium to be harvested and purified by immunomagnetic selection. Freshly isolated rat choroidal DCs fail to present antigen in the MLR but after 24 hours culture they develop full MLR reactivity (Forrester et al., in preparation). If choroidal DCs are isolated from cultured explants, rather than by enzyme digestion, they express potent APC activity when ‘freshly’ harvested. This activity is not increased by culture with GM-CSF, TNFα or IL-4 and has been attributed to possible release of cytokines from other cells in the choroidal explant such as macrophages and/or retinal pigment epithelial cells (see below). However, it should be borne in mind that treatment of DCs with certain proteases such as dispase, used in many studies to purify and isolate DCs from whole tissues, removes surface antigen activity, including MHC class II, possibly causing failure of freshly prepared cells to respond in the MLR. Choroidal DCs obtained from cultured explants act as potent APCs in both allogeneic and syngeneic MLR and can present antigen to primed T cells (Forrester, in preparation). In addition, freshly isolated DCs can present antigen to naïve T cells as shown by the polarisation assay (Forrester, in preparation). In this assay a requirement for accessory molecules appears to be lacking and recent studies in a cluster assay suggest that this initial priming of naïve T cells may be dependent on a recently discovered DC surface molecule with TNF-α-like properties, RANKL (Anderson et al., 1997).

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As stated earlier whilst the eye does appear to have a degree of immunological privilege, inflammatory responses are mounted against both autoantigens and infectious agents (see reviews, Nussenblatt, 1991; Forrester and McMenamin, 1999). Experimental autoimmune uveoretinitis (EAU) is a good example of a photoreceptor-specific autoimmune disease which can be induced in several animal models by a variety of photoreceptor autoantigens. The histopathology of EAU resembles some human uveitic conditions such as sympathetic ophthalmia, intermediate uveitis, birdshot chorioretinitis, etc. EAU is mediated by CD4 T cells, and both lymphocytes and macrophages constitute the earliest effector cells which initially infiltrate the retina and cause lysis of the photoreceptors, followed by phagocytosis and eventual destruction of the whole outer retina (see reviews, Forrester, 1992; McMenamin et al., 1993). The photoreceptor outer segments are presumed to be the primary target of the autoimmune attack in these models and in the human disease. Two central questions remain unanswered regarding the aetiology of endogenous posterior uveitis and animal models of these conditions. First, where is the earliest site of T cell entry into the eye and second does antigen presentation occur within the eye and if so which cells act as APCs in the eye? Whilst DCs may be initiators of primary immune responses, macrophages are considered potent APCs in inflammatory autoimmune processes and are known effector cells in models of uveitis causing much of the tissue damage. Several authors have observed anterior segment changes prior to retinal disease in many models of ‘posterior’ uveoretinitis (see review, McMenamin et al., 1993). Recently Prendergast and colleagues (1998) reported a biphasic response in T-cell recruitment to the retina following intravenous infusion of 10  106 labelled SP35 (S-Ag specific) cell lines in the adoptive transfer EAU model. The PKH-26 fluorescent labelled cells peaked at 24 hours and 96/120 hours following injection. The first peak represented nonspecific entry of activated T blasts, as this was also noted with Con-A stimulated T cells in control experiments and was

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confirmed by in vivo video scanning laser ophthalmoscopy, whereas the second peak in T cells was interpreted as evidence of activation following their interaction with Ag-bearing APCs. The commencement of inflammation in the anterior segment, which preceded retinal changes, was postulated by these authors to be due to the interaction of Ag-specific T cells with the rich APC network (DCs and macrophages) in the iris and ciliary body (Prendergast et al., 1998). An alternative explanation for these events might be that the first wave of activated T cells ‘conditions’ the retina via mediators such as chemokines for the second wave of T cells which have been activated by marrow-derived DCs migrating from the eye to activate T cells in the lymph node. The alternative view, i.e. that antigen is presented to T cells in the eye, has little supporting evidence. Most of the various candidates for presentation of autoantigens in uveitis have included vascular endothelium, retinal pigment epithelium and Muller cells, however, evidence for these cells as major activators of activated or naïve T cells compares poorly to the recent evidence for blood-borne DC (Forrester et al., 1995; Jiang et al., 1999). Whilst there have been several previous studies which have reported MHC class II upregulation on retinal pigment epithelium and retinal vascular endothelium in EAU models (Chan et al., 1986a; Liversidge et al., 1988), sympathetic ophthalmia and uveitis (Chan, et al., 1986b) there have been only limited reports of upregulated MHC class II expression on cells of the choroid or retina in diseased states. It has recently been demonstrated that there is an obvious increase in the density of MHC class II (Ia) cells in the iris and choroid in the early stages of EAU (Butler and McMenamin, 1996). While some of the increase may be accounted for by MHC class II activated T cells or macrophages it probably represents enhanced recruitment of DC precursors from the circulation or an upregulation of Ia on previously Ia DCs. There is evidence that a population of DC precursors (OX62 Ia) exists within the choroid and iris (vide supra). These may provide a rapidly inducible source of DCs in the event

of local inflammatory episodes. The concurrent increase in density of Ia DCs and T-cell infiltration in the uveal tract at the time of commencement of uveoretinitis would provide the appropriate environment for local activation of uveitogenic T cells. The crucial importance of the interaction between the T-cell receptor and peptide-bearing MHC class II molecule in induction of autoimmune disease has underpinned the rationale of experimental attempts to disrupt the complex using anti-Ia antibody therapy and therefore ameliorate diseases such as EAU (Rao et al., 1989) and EAE (Jonker et al., 1988; Gautam et al., 1992) The partial success of such studies supports the concept that MHC class II expression is involved in the induction and perpetuation of EAU.

FACTORS REGULATING DC FUNCTION IN THE EYE Despite the location and widespread distribution of DCs in the uveal tract and their proven ability to capture antigen in the anterior chamber and migrate from the eye, it is self-evident that their interactions with T cells in the normal state causes a tolerogenic rather than an immunogenic response. This is illustrated at the experimental level by the ACAID model (see review Streilein, 1999). If this were not the case and ocular DC induced immunogenic responses, they would have the potential to cause local damage to delicate ocular tissues by initiating inflammatory disease in the eye. The failure of DC to activate T cells which may enter the eye may represent one aspect of so-called ‘immune privilege’ in the eye but may more generally represent an illustration of how local microenvironment factors modulate immune responses. A number of such factors operate in the eye including the expression of FasL on the tissues of the eye. FasL is known to induce apoptosis of infiltrating Fas+ lymphocytes thus acting as a potent, although not the only, mechanism of maintaining immune privilege (Griffiths et al., 1995). Recently a pro-apoptotic low molecular weight molecule present in normal aqueous

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humour has been described which effects NK cells, T cells (activated and resting) and macrophage–monocytes, but spares resident ocular cells (D’Orazio et al., 1999). Of the potential immune effector cells, macrophages were found to be most resistant to this molecule. This may represent one mechanism protecting the resident networks of macrophages (and possibly DCs, although these were not investigated) in the uveal tract. Other molecules present in the aqueous humour, such as alpha-melanocyte stimulating protein, TGFβ and glucocorticoids, have also been shown to have immunosuppressive effects and likely contribute to minimising inflammatory-mediated damage of the anterior segment and thus helping to maintain a clear visual axis (see review, Streilein, 1999). In other organs macrophages can downregulate DC function, as is the case in the lung (Thepen et al., 1992), and gut (Pavli et al., 1990) possibly via production of TNFα and nitric oxide. Recent data suggest that ED2 iris macrophages lack the lymphocytostatic activity of mucosal or ‘body cavity’ macrophages (Steptoe et al., 2000). Choroidal macrophages are poor APCs in isolation, but may significantly augment, rather than downregulate the MLR initiated by choroidal DCs (Forrester et al., in preparation). Furthermore, cultured DCs appear to cluster with large macrophages, indicating that there is significant cross-talk between these two cell types. Retinal pigment epithelium (RPE) cells appear to release factors which inhibit APC function. These factors include prostaglandins and nitric oxide (Liversidge et al., 1994). In the eye, RPE cells are also known to release GM-CSF which is a survival factor for DCs (Crane et al., 1999). Production of GM-CSF by RPE cells is induced by other cytokines such as TGFβ, TNFα and IL-1. GM-CSF may not only affect DC function in the choroid, but if it were released into the retina might modulate the function of microglia since it has been shown to inhibit MHC class II expression by these cells (Hayashi et al., 1995). Thus it may have a proinflammatory effect on the CNS by blocking the recently discovered T-cell apoptosis-inducing effect of microglia (see above).

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Other chemokines released by RPE cells include IL-8 and RANTES, but these are produced only after exposure to pro-inflammatory cytokines such as IL-1 and INFγ (Kuppner et al., 1995; Crane et al., 1998). In addition, recent studies by our group have shown that RPE cells release stromal cell derived factor-1 (SDF-1) and also express the cognate receptor CXCR4 (Crane et al., 2000). SDF-1 is a chemokine for T lymphocytes and also acts as a costimulator via the CXCRr 4. T cells activated in this way release IL-4 and IL-10, IFNγ and IL-2 and proliferate in response to IL-2. At high concentrations, SDF-1 may act as a negative chemoattractant for T cells, i.e. actively repel migratory T cells (Poznansky et al., 2000). This has implications for the eye in preventing the migration of T cells into sites of inflammation where SDF-1 concentrations are elevated. RPE cells also preferentially release IL-6 when exposed to a combination of IL-1 and TGFβ (Kuppner et al., 1995). IL-6 has significant effects on DCs and may promote autoimmune disease by permitting DCs to process and present cryptic peptides (Drakesmith et al., 2000). IL-6 is also considered to act as a ‘TH2’ cytokine and this has led to the notion that RPE cells orchestrate the immune response at the blood–retinal barrier, promoting a TH1 type response under certain circumstances and a TH2 or even a null (?TH3) type response when the immune response is being down-regulated (Forrester et al., 1995).

RECRUITMENT OF DCs AND MONONUCLEAR CELLS TO THE EYE DURING INFLAMMATION Although DCs and activated macrophages are absent from the neural retina under normal circumstances, large numbers of the latter type are found in models of intraocular inflammation in which they appear to act as effector cells in tissue damage (McMenamin et al., 1993; Forrester et al., 1998). The recruitment of DCs into the eye likely depends on changes in

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adhesive interactions between leukocytes and the endothelium and on the release of specific chemokines such as MCP-1 and MIP-1a (Crane et al., in preparation) which allow transendothelial migration of mononuclear cells. Recent studies have shown that infiltration of sites of tissue damage by inflammatory cells, presumably containing small numbers of antigen specific T cells, allows release of chemokines by tissue cells such as astrocytes (Barna et al., 1994; Hurwitz et al., 1995). Monocyte chemotactic protein-1 (MCP-1) has been detected in astrocytes in the acute phase of EAE (Hayashi et al., 1995) and can be induced by release of T-cell cytokines, such as TNFα. In contrast MIP-1α appears to be released from microglia rather than astrocytes. T lymphocytes appear to secrete a further chemokine, TCA3, which directly attracts macrophages and even induces chemotaxis in microglia. Other chemokines produced by these cells include RANTES and MIP-1β. In EAE, most of the RANTES produced appeared to be derived from T cells, while some was produced by astrocytes and microglia. However, at the peak of disease all three chemokines appeared to be produced predominantly by perivascular cells (Miyagishi et al., 1997). Our recent data confirm the release of MCP-1 and MIP1α in EAU where they appear preferentially located on the vascular endothelium (Crane et al., in preparation). Release of TNF by infiltrating T cells and macrophages also has a direct effect on the expression of adhesion molecules by the endothelium, thus adding a further dimension to the recruitment of inflammatory cells in CNS inflammation (Korner et al., 1995). Interestingly, adhesion molecule expression commences prior to the onset of clinical disease, suggesting that the endothelium may regulate the traffic of T cells to the site of inflammation (Ma et al, 1996). However, as described above recent studies in EAU using direct observation of retinal vessels in the confocal scanning laser ophthalmoscope reveal that, although nonspecifically activated T cells will adhere to a normal nonactivated endothelium in small numbers, both the endothelium and the T cells

require to be activated before transendothelial migration and infiltration of the tissue occurs (Hossain et al., 1998). Furthermore, the requirement for adhesion molecule upregulation in the development of EAU has further been confirmed using this in vivo model of leukocyte tracking (Hossain et al., in preparation). Recent studies (Navratil et al., 1997) indicate that the repertoire or combinations of adhesion molecules expressed by CNS endothelium does not differ from other endothelium suggesting that the adaptation of the brain (and retinal) endothelium is due to expression of a few highly specific molecules, possibly yet to be discovered.

CONCLUSION AND FUTURE LINES OF RESEARCH From the preceding discussions it is apparent that under normal conditions, DCs are excluded from the neural retina. However, there is now extensive evidence that they are well represented in the tissues that provide its physical and physiological support, i.e. the choroid, ciliary body and iris. Despite the difficulties of isolating pure populations of DCs from small ocular tissue samples it has been shown that after appropriate maturational signals they are potent APCs capable of activating naïve T cells. In vivo however, ocular APCs may have a tolerising role since they are present in an immature state some with low MHC class II expression and generally low levels of costimulatory molecule expression. On the other hand, freshly isolated iris macrophages are potent APCs in models of secondary immune responses by comparison with freshly isolated iris DCs. Thus it appears that macrophage and DC populations in the iris exhibit the classical functions characteristic of these cell types in many organs and likely perform dichotomous, but complimentary tasks. In particular, uveal macrophages may act as a ‘first-line’ APC in secondary immune responses, whereas their adjacent neighbours, the MHC class II DCs exhibit the features characteristic of other nonlymphoid DCs and act primarily as sentinels, capturing antigen

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prior to their migration to draining lymphoid organs where they mature into potent APCs capable of orchestrating either primary immune or immunoregulatory responses. The heterogeneity of DC populations has attracted a lot of interest lately and is an area of potentially fruitful research with regard to uveal tract DCs. For example, are subpopulations of DCs responsible for presenting tolerogenic or immunogenic signals and if so, could this knowledge be used in the prevention and treatment of autoimmune conditions affecting the eye? For example, could some DC/antigen combinations be tailored or modulated (using cytokine signals) to sensitise animals and prevent induction of autoimmune conditions in the eye whilst another DC/antigen mixture be used to induce disease. Whilst such experimental approaches will help unravel the mechanisms controlling ocular immune responses the application of this knowledge to the human condition will continue to encounter the difficulty of treating pre-existing conditions. Perhaps strategies for the limitation of further tissue damage is a more realistic goal.

ACKNOWLEDGEMENTS The authors would like to thank Dr Ray Steptoe for useful discussions. Dr McMenamin acknowledges support from the NH&MRC and ARC. Professor Forrester would like to thank Lynne Lumsden for technical assistance in DC studies and Janet Liversidge, Isobelle Crane and Maria Kuppner for work with RPE cells.

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Poznansky, M.C., Olszak, I.T., Foxall, R. et al. (2000). Nat. Med. 6, 543–548. Prendergast, R.A., Iliff, C.E., Coskuncan, N.M. et al. (1998). Invest. Ophthalmol. Vis. Sci. 39, 754–762. Preston, S., Wright, G.J., Starr, K., Barclay, A.N. and Brown, M.H. (1997). Eur. J. Immunol. 27, 19111918. Provis, J.M., Penfold, P.L., Edwards, A.J. and van Driel, D. (1995). Glia 14, 243-256. Rao, N.A., Atalla, L., Linker-Israeli, M. et al. (1989). Invest. Ophthalmol. Vis. Sci. 30, 2348–2355. Rowland, L.P. (1985). In: Kandel, E.R. and Schwartz, J.H. (eds) Principles of Neuroscience, New York: Elsevier, pp. 837–844. Ruedl, C. and Hubele, S. (1997). Eur. J. Immunol. 27, 1325–1330. Sanyal, S. and DeRuiter, A. (1985). Cell. Tissue Res. 241, 291–297. Schon-Hegrad, M.A., Oliver, J., McMenamin, P.G. and Holt P.G. (1991). J. Exp. Med. 173, 1345-1356. Sedgwick, J.D. (1995). Immune surveillance and autoantigen recognition in the central nervous system. Aust. NZ J. Med. 25, 784–792. Sedgwick, J.D. (1997). Biochem. Soc. Trans. 75, 673–679. Sedgwick, J.D. and Hickey, W.F. (1997). In: Keane, R.W. and Hickey, W.F. (eds) Immunology of the Nervous System, New York: Oxford University Press, pp. 364–418. Sedgwick, J.D., Ford, A.L., Foulcher, E. and Airriess, R. (1998). J. Immunol. 160, 5320–5330. Sherman, S.H., Green, K. and Laties, A.M. (1978). Exp. Eye Res. 27, 159–173. Singh, A.K., Kumar, G., Shinohara, T. and Shichi, H. (1996). Exp. Eye Res. 62, 299–308. Soesatyo, M., Biewenga, J., van Rooijen, N., Kors, N. and Sminia, T. (1991). Res. Immunol. 142, 533–540.

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Steinman, R.M. (1991). Annu. Rev. Immunol. 9, 271–296. Steptoe, R.J., Holt, P.G. and McMenamin, P.G. (1995). Immunology 85, 630–637. Steptoe, R.J., Holt, P.G. and McMenamin, P.G. (1996). J. Neuroimmunol. 68, 67–76. Steptoe, R.J., Holt, P.G. and McMenamin, P.G. (1997). Invest. Ophthalmol. Vis. Sci. 38, 2639–2648. Steptoe, R.J., McMenamin, P.G. and Holt, P.G. (2000). Ocular. Immun. Inflamm. 8, 177–187. Streilein, J.W. (1993). Curr. Opin. Immunol. 5, 428–432. Streilein, J.W. (1999). Chemical Immunology 73, pp. 11–38. Thanos, S., Moore, S. and Hong, Y. (1996). Prog. Ret. Eye Res. 15, 331–361. Thepen, T., McMenamin, C., Oliver, J., Kraal, G. and Holt, P.G. (1992). Eur. J. Immunol. 21, 2845–2850. Ulvestad, E., Williams, K., Bjerkvig, R., Tiekotter, K., Antel, J. and Matre, R. (1994). J. Leuk. Biol. 56, 732–740. Vandenabeele, S. and Wu, L. (1999). Immunol. Cell. Biol. 77, 411–419 Wilbanks, G.A., Mammolenti, M. and Streilein, J.W. (1991). J. Immunol. 146, 3018–3024. Williams, K., Ulvestad, E. and Antel, J.P. (1994). Eur. J. Immunol. 24, 3031–3037. Yang, P., de Vos, A. and Kijlstra, A. (1999). Invest. Ophthalmol. Vis. Sci. 40, 3067 (letter). Zaal, J., Doekes, G., Breebaart, A.C. and Kijlstra, A. (1986). Curr. Eye Res. 5, 763–775. Zhang, J., Wu, G., Ishimoto, S., Pararajasegaram, G. and Rao, N.A. (1997). Invest. Ophthamol. Vis. Sci. 38, 1848–1857. Zhang, G.X., Baker, C.M., Kolson, D.L. and Rostami, A.M. (2000). Mult. Scler. 6, 3–13.

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PLATE 29.3 Frozen section of rat retina in EAU (day 11). Immunohistochemical localisation (red–brown reaction) of MCP in vascular endothelium.

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PLATE 29.5 Rat choroidal wholemount stained with mAb ED2 (resident tissue macrophages, green) and RT97 (anti-neurofilaments, red). Note the largely perivascular distribution of macrophages and the ‘cobblestone’ appearance of the hexagonal autofluorescent retinal pigment epithelial cells. Confocal microscopy, bar 20 µm.

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30 Dendritic cells in the reproductive tract C. Allen Black1 and Michael Murphy-Corb 2 1

Magee Womens Research Institute, and 2 University of Pittsburgh, PA, USA

O sacred receptacle of my joys, Sweet cell of virtue and nobility Titus Andronicus William Shakespeare

INTRODUCTION

While most research has been focused on the female reproductive tract, DCs are also found in the mammary glands, the placenta and the male reproductive tract although the biological role for DCs in these organs has been less well described. In the female reproductive tract, DCs are involved in antigen acquisition, immune surveillance and immune effector mechanisms. DCs play an important role in both detection and response to sexually transmitted diseases (STDs), including human papillomavirus (HPV) and human immunodeficiency virus (HIV ). Indeed, subversion of DC function in the reproductive tract may be an important survival factor for STD pathogens. Additionally, DCs may serve a crucial role in maintaining tolerance to normal flora inhabiting the reproductive tract, since a healthy reproductive tract is heavily colonized with bacteria.

DCs, also termed Langerhans’ cells, are the primary cells involved in immunosurveillance of mucosal barriers including the reproductive tract, and form a contiguous network lining the skin, gut and reproductive tract. DCs in the female reproductive tract were first described histochemically in 1966 by Thierry (Langerhans, 1868; Thiery and Willighagen, 1966). The phenotype of dendritic processes and ATPase staining corresponded to the cellular architecture originally described in skin Langerhans’ cells. These observations were later confirmed using electron microscopy, which showed that cervical DCs contain Birbeck granules, a defining feature of many DCs (Birbeck et al., 1961; Younes et al., 1968). Even though a functional role for the cervical DCs was not ascribed, Younes noted an increase in the density of cervical Langerhans’ cells during neoplasia (Younes, 1969).

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DCs IN THE FEMALE GENITAL TRACT Distribution and function The human female reproductive tract can be categorized into three zones, each with differing functions: the vulvo-vaginal region, the cervix and the uterus. DCs are found contiguously throughout all of the regions although the relative concentrations vary. The vulvo-vaginal epithelium is neither absorptive nor secretory, although antigen can traffic to some degree through the epithelial boundary (Parr and Parr, 1990a, 1990b, 1994). The outer layer of the skin in this region is lined by dead squamous epithelial cells. Underneath the outer layer is the parabasal layer where the epithelial cells are continuously renewed at rates that vary over the menstrual cycle (Sjöberg et al., 1988). DCs have been demonstrated in this region, as well as deeper into the stroma and lining the vessels in the deep mucosa (Edwards and Morris, 1985; Johansson et al., 1999). The primary function of the vaginal tissue of the lower genital tract is to protect against shear force during copulation. There are twice as many vulvar DCs per millimeter of tissue as vaginal canal DCs although the significance of this difference is not known (Edwards and Morris, 1985). Unlike the vagina and vulva which have squamous epithelial linings, the cervical canal, the transformation zone and endocervix are lined by columnar epithelial cells. This columnar layer can vary in thickness but does not reach a thickness comparable to the vaginal epithelium, and is more permeable to antigen flow than the vaginal epithelium (Hiersche and Nagl, 1980; Roelofs et al., 1983; Parr and Parr, 1994). The cervix has been shown to be absorptive and secretory, capable of both absorbing antigen and transporting immunoglobulin, mucus and cytokines into the cervical canal (Gilks et al., 1989; Mattsby-Baltzer et al., 1998). The cervix also acts as a channel into the uterus for sperm and as a consequence, sexually transmitted diseases. Thus, the cervix serves as a sentinel to the upper reproductive tract. In accordance with this role,

the cervix has numerous lymphocytes including DCs found interdigitating between the epithelial cells in both the mid-epithelium and the basal epithelial layers (Roncalli et al., 1988; Di Loreto et al., 1989; Poppe et al., 1998). In some cases the processes can extend into the deeper basal stromal layer and gird the capillaries (BonillaMusoles et al., 1987). The density of cervical basal and parabasal DCs exceeds that of the vagina by almost 10-fold (Edwards and Morris, 1985). Interestingly, S100 DCs can also be found lining the connective tissue of most of the capillaries throughout the body (Edwards and Morris, 1985). It is not known if endothelial DCs are the same as those participating in local mucosal immunity. Phenotypic analysis by numerous groups has demonstrated that most DCs in the reproductive tract are HLA-DR, CD1a, CD1c, Fc gamma (II, III), S100, CD4, and membrane ATPase (Younes et al., 1968; Bjercke et al., 1983; Tay et al., 1987; Cristoforoni et al., 1995). In the mouse the vagino-cervical DC phenotype is similar and expresses Ia, F4/80, NLDC-145 and CD45, but not Mac-1, Moma-1, or Moma-2 (Parr and Parr, 1991). There is no known difference between these cells and DCs found in the skin. It is well established that immunization throughout the mucosal immune system results in distant mucosal responses (Kutteh, 1999). DCs resident in the rodent vagina can home into draining mucosal lymph nodes (Parr and Parr, 1990b). However, the signals inducing homing of the DCs from the blood into the mucosal site are unknown. In human ectocervical and endocervical tissue, the intestinal mucosal addressin of MAdCAM-1 is not expressed, a finding that suggests lymphocyte homing may be different in the genital tract compared to the intestine (Johansson et al., 1999). Notably, DCs in the reproductive tract mucosa are rarely seen associated with T cells. This indicates that at the mucosal level, the DC may not be involved in antigen presentation until it has homed back into the lymph node (Parr et al., 1991a). However, parabasal DCs are capable of phagocytosis of apoptotic vaginal epithelial cells in the rodent, so DCs may have limited effector functions in the mucosa (Parr et al., 1991b).

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DCs are also abundant in the uterus. They have been identified in the endometrium, the myometrium and serosa of the uterus. The endometrium where implantation occurs, varies in thickness over the menstrual cycle as does the total number of interdigitating DCs (Bulmer and Sunderland, 1983). In the rat, the DCs are especially dense around the endometrial glands with the overall density increasing significantly with estradiol injections (Kaushic et al., 1998). Wira has shown that the function of APCs is differentially regulated by sex hormones in the uterus and vagina although the cells that were examined were primarily epitheloid and not purified DCs (Wira and Rossoll, 1995a). The role of the DC in immunosurveillance at the uterine mucosa includes not only foreignantigen uptake but potentially semi-allograft antigen processing during pregnancy. In the pregnant uterus at day 6 post-implantation, fewer lymphocytes including DCs are found and by day 10, the area around the conceptus is devoid of DCs. However, after day 10, DCs begin to reappear at the maternal–fetal interface sometimes near the invading trophoblasts (Head and Billingham, 1986). Presumably, the decreased DC frequency and antigen presenting capability is a result of downregulation to prevent fetal allograft responses. Thus, it is somewhat paradoxical that APCs are stimulated in a progesterone dominant environment, but are depleted during pregnancy (Head and Billingham, 1986; Head and Gaede, 1986). A toleragenic role of a DC in pregnancy is consistent with its ability to regulate graftversus-host disease. In the developing fetus, uterine, cervical (including transition zone), vaginal and vulvar DCs can be detected at birth (Morris et al., 1983). The function of DCs in the oviduct and ovary is largely unexplored. However, immunohistochemical studies have demonstrated DCs in rodent ovarian follicle, the rodent oviduct as well as the anterior pituitary which regulates ovarian function (Brannstrom et al., 1993). DCs have also been identified in ovarian teratomas (Mandel et al., 1994).

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Placental DCs have been identified immunohistochemically in humans (Sutton et al., 1983). DCs can be differentiated from fetal Hofbauer cells and placental macrophages by a higher level of HLA DR expression. They also express HLA-A, -B and -C, but HLA-G expression has not been tested. DCs are present in the chorionic villi, decidua and the subepithelial amnion of third trimester placentas. One study of human placentas describes factor XIIIa-positive cells that have a stellate morphology and HLA-DR expression. However, these cells appeared at 7–9 weeks prior to fetal hematopoiesis (Trimble et al., 1992). A functional role for these cells was not determined.

Sex hormones and DCs In one study, the number of cervical DCs was found to increase during the proliferative phase of the menstrual cycle, but most human studies have not controlled for stage of cycle (Edwards and Morris, 1985). In rodents, however, the endocrinological effects have been closely studied (Young, 1985; Young et al., 1985; Head and Gaede, 1986; Parr and Parr, 1991). An increase in the number of DCs in vaginal tissue after both estradiol and progesterone administration has been observed in ovariectomized rats, with progesterone being the most potent (Parr and Parr, 1990b; Kaushic et al., 1998). Notably, T cell levels remained constant. This observation contrasts with what is seen in the human where no difference in the concentration of DCs between pre- and postmenopausal women is found in the vagina or uterus (Morelli et al., 1992a; White et al., 1997). APC function as assessed by whole stromal cell cultures, shows that diestrus (progesterone dominant) vaginal tissue is more effective at antigen presentation than proestrus (estrogen dominant) tissue (Wira and Rossoll, 1995b). Receptors for progesterone and estrogen have not been demonstrated in DCs; however, myeloid derived cells in the same family such as macrophages and monocytes have been shown to have these receptors (Vegeto et al., 1999).

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Antigen transport and reproductive tract DCs Antigen flow through the vaginal mucosa has also been studied in the rodent. During the estrus cycle, fluorochrome labeled protein-antigen was found to cross the vaginal epithelial cell boundary by uptake from DCs at diestrus (progesterone dominant), but not at estrus (estrogen dominant) (Parr et al., 1991a). The cells were subsequently shown to home into the caudal and iliac lymph nodes draining the region. However, these models are of limited use, as the rodent vaginal wall is disrupted by neutrophils emigrating into the vaginal lumen at diestrus (Black et al., 1998). No direct evidence for vaginal uptake of large proteins through the vaginal wall in humans exists. However, work in primate models has shown that antigen can perhaps pass through the vaginal wall at least to immunize or infect with viruses. Presumably, DCs mediate this type of antigen transfer. One group has demonstrated that SIV can penetrate the vaginal mucosa of rhesus macaques where the cervix has been removed to form a blind pouch, which indicates that the vaginal epithelium alone may be sufficient for antigen uptake (Miller et al., 1992). Antigen uptake from the reproductive tract includes phagocytosis, pinocytosis and receptormediated endocytosis (Parr et al., 1991a, 1991b; Steinman, 2000). However, the molecular signal that induces DC homing back into the lymph node is unknown. It is likely, however, that as in homing with skin DCs, TNFα could be involved as it has been detected in cervical mucus and in reproductive tract tissues (McMaster et al., 1992; Nuovo et al., 1993; Mattsby-Baltzer et al., 1998; Perfettini et al., 2000). DCs are not found in the normal shed epithelium, a finding which suggests that DCs remain at the local site even during epithelial cell turnover.

MALE REPRODUCTIVE TRACT DCs Little is known about DCs in the male reproductive tract. However, Morelli has shown that DCs

form a filamentous network in the squamous epithelium of the penis just as in other epithelial tissues (Morelli et al., 1992b). This work has been extended to describe DCs that reside in the distal tip of the penis, the mucosa of the meatus and fossa navicularis, whereas the urethra proper contains many macrophages within the lamina propria and epithelium, but no DCs (Pudney and Anderson, 1995). This distribution also occurs in the rodent and nonhuman primate penis (Quayle et al., 1994; Hussain and Lehner, 1995). In the testes, MHC class II antigen was identified on dendritic-like cells between the seminiferous tubules and on vessel endothelium, although its expression was limited (Haas et al., 1988; Itoh et al., 1995). Although the function of DCs in the testes is unknown, they may be responsible for maintenance of immunoprivilege at this site (Streilein, 1993). Testosterone has a suppressive effect on DCs in rodents; males have decreased DC density in the skin compared to females (Koyama et al., 1987). Orchiectomy results in increased skin DC density. Whether the hormone effects on DCs in males are due to a testosterone receptor in DCs or a secondary effect is unknown.

CERVICAL DCs IN SEXUALLY TRANSMITTED DISEASE AND OTHER DISORDERS Human papillomavirus Currently, most known dysplasia and cancer of the cervix is associated with HPV infection, primarily of subtypes 16 and 18 while genital warts (condylomata) are caused primarily by subtypes 6 and 11 (Roche and Crum, 1991). The virus is tropic for the differentiating squamous epithelial cells, which are near the same anatomical site as numerous DCs. When DCs were first described immunohistochemically in the cervix, a profound increase in the density of DCs was identified in cervical dysplasia (Thiery and Willighagen, 1966). Contradictory reports showing either increases or decreases in DC

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density have been published, but more recent reports suggest that depletion of DCs in the cervix is most closely associated with HPV infection and neoplasia. In condylomata, either a normal density of DCs or decreased number of DCs is observed relative to adjacent vulval sites. When DCs are found near infections or warts they exhibit an abnormal morphology with shortened processes and a more rounded appearance (Di Girolamo et al., 1985). HPV has not been shown to directly infect DCs, so microenvironmental factors may play an important role. Two hypotheses for this observation have been posited. Potentially, the number of DCs is being reduced by direct viral effects. Alternatively, the DCs may increasingly traffic back into the lymph node or may no longer be recruited into the site due to some negative chemotactic stimulus (Di Girolamo et al., 1985). However, no direct evidence supporting these hypotheses exists. In cervical intraepithelial neoplasia (CIN), conflicting results regarding DC depletion have been reported. The progression from CIN I to CIN III correlates with either an increase or decrease in DCs by immunohistochemistry (Younes et al., 1968; Puts et al., 1986; Tay et al., 1987; Hachisuga et al., 1989; Poppe et al., 1996; al-Saleh et al., 1998). Some evidence indicates that these discrepancies may be due to infection by different subtypes of HPV. At low copy numbers, HPV 18 has been strongly associated with depletion of DCs while depletion in HPV 16 infection only occurs when copy number is high (Hawthorn et al., 1988). This indicates that the viremia alone is not responsible for the loss of DCs and changes are more likely related to neoplastic progression. Additionally, even though HLA-DR is upregulated in keratinocytes, it is not upregulated in DCs but HLA-DP is, indicating that some discrepancies could be explained if class II alone is being used as a marker for DCs in the cervices as opposed to S100 or CD1a (Hughes et al., 1988). When S100 is used as a marker for DCs, all reports show decreases in the DC cell density. Tay has defined two DC populations in the cervix: 65% that are positive for CD1 and the other 35% positive for S100. The S100-

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staining cells were associated with inflammatory foci, but both types were decreased in neoplasia (Tay et al., 1987). Roncalli has posited three separate DC populations (Roncalli et al., 1988). Nevertheless, it is unknown why the DCs are found at lower numbers in neoplasms, but this decrease could reflect the hyperproliferation of the squamous cells, which causes DCs to be outnumbered and compressed. al-Saleh and others have described an aberrant cytokine environment where IL-4 and IL-10, both TH2 type cytokines, are upregulated, perhaps from infiltration of B cells, and these cytokines may inhibit DC function (al-Saleh et al., 1998). Additionally, activational cytokines such as TNFα have been shown to be absent in metaplastic disease, indicating that the site may not have a sufficiently inflammatory environment to induce DC maturation or activation, nor do the DCs in the lesion express co-stimulatory molecules such as CD50 or CD86 (Mota et al., 1998). Theoretical involvement of DCs, as well as T and B cells, has been postulated as a causative factor in neoplastic transformation. Specifically, depletion or aberrant activation of any of these cells leads to reduced immunosurveillance and escape of malignant cells (Bell et al., 1995; Edwards et al., 1996). The actual role of the DC in HPV-associated cervical cancer is unknown. Conflicting reports exist on the prognostic significance of DC density and histology in the lesion and progression to cancer. One group has shown that the greater infiltration of DCs correlates positively with regression of the lesion in both squamous cell cervical carcinoma and cervical adenocarcinoma (Nakano et al., 1989). However, in condylomata of the penis and vulva that are unlikely to become cancerous, DC depletion is also noted, indicating that the loss of DCs alone is not sufficient to cause progression to cancer in HPV infection (Arany et al., 1998). Finally, it has been shown that HPV infection leads to strong humoral and cell-mediated immune responses systemically, thus antigen presentation function as a whole is not impaired in an HPV infection. The density of DCs in the cervix can also be decreased by environmental factors. In

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the cervices of normal women who smoke, decreased numbers of DCs are observed in HPVinfected and -uninfected women (Barton et al., 1989; Poppe et al., 1995). It has been hypothesized that the local concentrations of nicotine, cotinine and other tobacco constituents cause this depletion since cervical mucus of smokers can contain these chemicals. This decrease is theorized to reduce the local immunosurveillance capability of the cervix against HPV infection. Evidence exists for this hypothesis in epidemiological studies. Women with HIV who are immunocompromised have a higher rate of HPV-induced squamous epithelial lesions with the severity correlating to the degree of immunosuppression (Barberis et al., 1998; Six et al., 1998). DC-based immunotherapy has been suggested as a possible treatment of cervical cancer, and it has been demonstrated that peripheral blood DCs can be expanded ex vivo from cervical cancer patients using FLT-3 ligand, but it is unknown where the reintroduced DCs will home after i.v. injection (Hubert et al., 1998). In situ activation of DCs by local administration of Sizofirin, a fungal polysaccharide, into cervical tumors has been attempted (Nakano et al., 1996). It increased the activation state of DCs in the neoplasia while increasing DC infiltration in 10% of patients, but decreasing infiltration in 35%.

Chlamydia and herpes The role of DCs in other STDs besides HPV and HIV infection has not been well studied; however, in chlamydial infections no increase in DC infiltration or abnormal pathology was observed. However, ex vivo expansion of bone marrow DCs, followed by a pulse with killed chlamydia and re-injection i.v. protects animals from genital infection (Su et al., 1998). The response correlated to TH1 type induction of cell-mediated immunity. In genital herpes infection, local increases in DCs at the sites of infection have been noted although the significance of this finding is unknown (Syrjanen et al., 1986; King et al., 1998).

Other disorders Decreased numbers of cervical and dermal DCs have been documented in Sjogren’s syndrome, an autoimmune disorder (Oxholm et al., 1986). Patients with chronic renal failure also have decreased DCs in the cervix but treatment with hemodialysis restores the normal DC density (McKerrow et al., 1989). Depletion of cervical DCs with corticosteroids has not been directly demonstrated. However, it is likely as both topical and systemically administered steroids are known to deplete dermal DCs (Krueger and Emam, 1984).

DCs IN HIV TRANSMISSION IN THE REPRODUCTIVE TRACT HIV is now the leading cause of death from sexually transmitted diseases, with heterosexual transmission accounting for over two-thirds of the more than 50 million infections (Mann et al., 1992; UNAIDS, 1996). In the female, the vagina, cervix and uterus are all possible portals of entry for sexually acquired HIV. Since the vaginal wall is generally impermeable to large antigens, it is suspected that active transport is more likely than passive diffusion through breaks in the epithelial barrier (Wu and Robinson, 1996). Unlike most other STDs, however, HIV replication is not limited to the reproductive tract. Thus, the detection of replicating virus in mucosal tissues in chronically infected individuals does little to resolve how virus is transported across the epithelium (Howell et al., 1997). Bonilla-Musoles first proposed that viruses and bacteria in the genital tract are taken up by the cervical DCs and transported back into the lymph nodes (Bonilla-Musoles et al., 1987). With respect to HIV, this hypothesis is attractive given the substantial evidence regarding HIV infection in DC:T cell conjugates. A pivotal role for DCs in the transport of HIV across mucosal barriers is supported by several lines of evidence, much of which has been derived in experiments whereby macaques were

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atraumatically exposed to SIV. Atraumatic exposure of the vaginal vault with a high dose of SIV has been shown by several groups to induce a systemic infection as evidenced by detectable virus in the periphery, seroconversion and AIDSlike disease. However, 3 to 5 logs more virus is required for infection compared to intravenous inoculation (Miller et al., 1989, 1994; Trichel et al., 1997). Macaques that become chronically infected as a result of vaginal exposure have both replicating virus in the mucosal tissues and CTL responses in the vaginal lamina propria (McChesney et al., 1998; Miller et al., 1998). Interestingly, when more limiting doses are used, a localized infection is produced which is associated with a failure to isolate virus from the periphery and a lack of seroconversion but a detectable virus-specific immune response (McChesney et al., 1998; Wilson et al., 1999). Both MHC class I restricted CTL in the peripheral blood and/or the jejunal lamina propria in one study or SIV-specific secretory IgA responses in vaginal secretions in another have been identified in vaginally exposed uninfected macaques (Van Cott, personal communication; Wilson et al., 1999). Interestingly, in both studies, the immune responses observed were restricted to the viral gp120. The induction of env-specific CTL by exposure of the mucosa of the vaginal vault to limited doses of SIV is similar to the responses observed following atraumatic exposure of the columnar epithelium of the colonic and rectal mucosa (Murphey-Corb, unpublished); (Murphey-Corb et al., 1999). Although not directly determined in these studies, a role for DCs as APCs in the induction of these responses is consistent with their ability to induce CTL, an immune function that could operate effectively in blocking the dissemination of low doses of virus. Mucosal challenge of monkeys that were transiently infected as a result of exposure of either the intestinal or vaginal mucosa has further shown that mucosal CTL, but not antibody, correlate with protection against challenge with a higher dose of heterologous virus (Murphey-Corb et al., 1999; Wilson et al., 1999). These studies are reminiscent of humans who remain uninfected

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despite repeated sexual contact with infected partners, and perhaps illuminate the requirements for an efficacious mucosal vaccine for HIV (Rowland-Jones and McMichael, 1995). Attempts to identify the first cells infected with SIV following vaginal inoculation have been performed by pathological examination of mucosal tissues obtained at sacrifice of monkeys at early times after vaginal SIV exposure. The predominant infected cell in the vaginal submucosa was not the DC, but the T cell (Zhang et al., 1999). T cell infection was detected by day 3, but DC infection was not detected until 1 week after exposure. At later times, infected cells were observed in cervical lymph nodes and other reproductive tract tissues. Nevertheless, in other work using immunohistochemistry and in situ hybridization of cytospin preparations, a high percentage of vaginal DCs has been shown to express SIV RNA as early as 18 hours postexposure (Miller, 2000). Taken together, these studies provide provocative evidence for the role of DCs in the transport of virus across the mucosa of the female reproductive tract. The apparent selection for macrophagetropic (nonsyncytium-inducing) variants of HIV observed in women following sexual exposure further argues for a role of the DC in virus transport across the mucosa given the monocyte/ macrophage lineage of these cells. It has been shown that immature DCs may become infected with HIV and can express CD4 and CCR5, albeit at lower levels than T cells (Granelli-Piperno et al., 1996; Rubbert et al., 1998). DCs also do not express CXCR4 and are not infectable by X4 tropic clones (Weissman et al., 1995; GranelliPiperno et al., 1998). The results of vaginal inoculation of female macaques with culture filtrates of SIV containing a complex genetic mixture of variants however, confounds this issue. Sequence analysis of variants present in the peripheral blood of vaginally exposed females has demonstrated no selection for a particular genotype (Trichel et al., 1997; Sodora et al., 1998). The discrepancy between the outcome of human and simian vaginal exposures may reflect the difference in coreceptor requirements between the two viruses (CXCR4-

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dependent SIV isolates are extremely rare and have not been studied in vaginal infections). Alternatively, the selection observed in human transmissions may not be due to the barrier imposed by the mucosa, but, rather, on selective amplification of a specific genotype in the donor tissues, or once the virus enters the submucosa of the recipient. This hypothesis is supported by studies performed by Miller et al whereby SHIVs containing CXCR4-tropic and CCR5-tropic HIV envelope genes had an equal likelihood of infecting vaginally exposed macaques (Lu et al., 1996; Harouse et al., 1998; Miller et al., 1998). The resolution of these conflicting data may reside in the recent report that HIV may bind to DCs independently of CD4 and be delivered to systemically located CD4 cells (Blauvelt et al., 1997). HIV has been shown to adhere to DCs using a member of the C-type lectin family termed DC-SIGN, which normally binds ICAM-3 (Geijtenbeek et al., 2000a, 2000b). Although this interaction does not facilitate DC infection, it may stabilize the virion long enough for transport into the regional draining lymph nodes where it encounters the CD4 lymphocytes. A role for progesterone in the facilitation of viral transport across the mucosa of the female reproductive tract has been suggested by several studies examining human prostitutes using hormone contraceptives (Mostad et al., 1997; Martin et al., 1998). The suppressive effect of estrogen and/or the requirement for exogenous progesterone therapy for successful SIV vaginal infections has also been documented by several groups (Marx et al., 1996; Wilson et al., 1999). Infection by vaginal inoculation to high doses of the primary isolate SIV/DeltaB670 requires exogenous progesterone treatment (5 mg/kg daily for 10 days) (Wilson et al., 1999). In another study, progesterone treatment of ovariectomized macaques using depo-provera implants significantly enhanced vaginal infection, and estrogen (but not progesterone) treatment completely suppressed infection with a primary stock of SIVmac251 (Marx et al., 1996). The exact mechanism whereby progesterone enhances (and estrogen suppresses) the risk of HIV/SIV transmission is unknown. However, it is possible

that progesterone could either augment the activation and trafficking of DCs back into the lymph node and/or regulate the expression of DC-SIGN. Studies using macrophages have shown that HIV co-receptor expression can be downregulated by progesterone and this may apply to DCs as well (Vassiliadou et al., 1999). Work in the mouse has also demonstrated that antigen flow to DCs is increased through the vagina during the progesterone dominant cycle (Parr and Parr, 1990b). For the most part, both human and macaque studies have focussed on the transport of virus across the cervicovaginal mucosa. In vitro studies, however, have demonstrated that uterine cells are also capable of sustaining HIV replication (Yeaman et al., 1998). In one study, infection of fallopian tube, cervical, and ectocervical tissue, but remarkably not vaginal tissue, with HIV1 was achieved (Howell et al., 1997). In this study, HIV-1-positive cells included CD4 T cells, CD14 macrophages and DCs. Thus various sites in the reproductive tract may function as portals of entry for HIV. Further studies in the macaque system should enhance our understanding of the roles of these sites, and the influence of hormone therapy, on the transport of virus at these sites. Although little is known in the macaque system about infection of the male reproductive tract, SIV infection of the penile foreskin and glands has been demonstrated (Miller, 1998). SIV/HIV may cross the stratified squamous epithelium covering these organs, or through the urethral epithelium. The role for DCs in trans-penile transmission is as yet undetermined, but again, they are likely candidates given their suggested involvement at other mucosal sites.

DCs AT OTHER MUCOSAL SITES Since the network of DCs is contiguous, it is important to mention the role of the DC in the common mucosal immune system. The function and distribution of gut and lung DCs is discussed elsewhere, but it is likely that potentially

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any DC can home into a mucosal site. Immunization strategies using DCs may be able to elicit responses at distant mucosal organs such as the eyes, inner ears, gut, lung, lactating breast and reproductive tract tissues. DCs have been demonstrated in the ductile tissue of the breast (Natali et al., 1984). In breast cancer, DCs can infiltrate into the tumor (Zoltowska, 1997). The role these cells play in preventing cancer or mastitis is unknown, but as in the reproductive tract they most likely act as sentinels against viral and bacterial infection.

CONCLUSIONS DCs provide immunosurveillance for all of the mucosal tissues. Their role in immunosurveillance is tightly regulated in the female genital tract, since the uterus must tolerate a semi-allograft in pregnancy as well as provide protection against STDs. In most cases the immunosurveillance role is an important function of DCs, such as in HPV infection and cervical cancer. However, in HIV infection it appears that the virus is capable of using the DC as a conduit into deeper tissues. In the reproductive tract, unlike in the skin and gut, the DCs are under hormonal regulation although it is unknown if this regulation is direct or indirect. The potential exists for the use of DCs both as vaccine adjuvants against STDs and cancer. Since they are part of the common mucosal immune system it may be possible to use locally derived DCs to immunize distant mucosal sites. The role of DCs in other mucosal sites and the male reproductive tract is just as important as the female reproductive tract and will be an active area of research in the coming years.

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31 The identification of dendritic cells in cancer Michael T. Lotze and Ronald Jaffe GlaxoSmithKline Pharmaceuticals, King of Prussia, PA and University of Pittsburgh Medical Center, Pittsburgh, PA, USA

We have a cancer within, close to the Presidency, that is growing. John Dean, from the [Nixon] Presidential Transcripts, March 21, 1973

INTRODUCTION

et al., 1998) and in patients with pancreatic (Dallal et al., 1998) and colon cancer (Clarke and Sikora, unpublished). It seems also (Tsuge T et al., 2000) that the number of DCs arrested in a more immature phenotype is more common in invasive tumors and more normal appearing DCs are often found outside, in this instance the breast tumor. The initial notion, that DCs are present within tumors to pick up tumor antigen and shuttle it to the lymph node and elicit the adaptive immune response, may be a bit too simplistic to explain this curious set of observations. In fact, many individuals have documented the inability of tumor-derived DCs to express appropriate co-stimulatory molecules as well as class II MHC molecules and the characteristic DC processes associated with their nominal ‘immunosuppressed state’ (Gabrilovich et al., 1996a, 1996b; Chaux et al., 1997; Nestle et al., 1997). An alternative explanation is that they are there to provide co-stimulation and cytokines to prevent apoptotic death for recruited effector T

Following the identification of dendritic cells (DCs) as the Langerhans cell in the skin, as the interstitial cell within tissues and lymph nodes, and as the veiled cell in lymph (Hart, 1997), it was subsequently recognized by clinical pathologists that there was a direct relationship between the presence of DCs within various lung tumors, head and neck tumors, breast cancer, cervical cancer, endometrial cancer (Coppolla et al., 1998), esophageal cancer, gastric cancer (Kakeji et al., 1993), Hodgkin disease, lung cancer, melanoma (Enk et al., 1997), mycosis fungoides, prostate cancer, renal cancer (Thurnher et al., 1996) and other tumors (Lotze, 1997) and prognosis (Tables 31.1 and 31.2). It appears that the presence of greater numbers of DCs is associated with a better prognosis (Ambe et al., 1989; Furihata et al., 1992; Inoue et al., 1993, Tsujitani et al., 1993; Tsuge T et al., 2000, for example). We have recently confirmed these findings in tumors of the oral tongue (Goldman Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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TABLE 31.1

Relationship between DC infiltration of head and neck cancers/oral cavity cancers and prognosis

Tumor

Reference

Larynx Nasopharyngeal Larynx Oral Oral Head and neck Head and neck Head and neck Larynx Oropharynx Oral squamous Oral tongue

Otolaryngology 53, 349 (1991) Laryngoscope 101, 487 (1991) Chin. J. Otorhinol. 27, 297 (1992) J. Oral Pathol. Med. 21, 100 (1992) J. Cutan. Pathol. 19, 398 (1992) Cancer Immunother. 36, 108 (1993) Cancer Immunother. 38, 31 (1994) Ann. Otol. Rhinol. Laryngol. 109, 56–62 (2000) In Vivo 8, 229 (1993) In Vivo 8, 543 (1994) J. Oral Pathol. Med. 24, 61 (1995) Laryngoscope (Goldman et al., 1999)

Salivary gland

Arch. Pathol. Lab. Med. (Soma et al., 2001)

cells. Within the tumor microenvironment, particularly in antigen-specific susceptible T cells, the induction of apoptosis following recognition of the tumor is noted, and thus we have the seemingly paradoxical situation of tumors killing T cells instead of T cells killing tumors (Mailliard et al., 2000; Hahne et al., 1996; Niehans et al., 1997; O’Connell et al., 1996). For that reason, means of preventing premature Tcell apoptotic death and the engagement of socalled ‘T-cell futile cycles’ is another goal of the immunologist. DCs, by virtue of their expression of so-called co-stimulatory molecules such as CD80 (B7.1) and CD86 (137.2) and several ‘dendrikines’ including interferon-α and IL-12, may be uniquely capable of preventing premature Tcell death and thus becoming the mediators of T-cell survival (Shurin et al., 1997a).

THE IDENTIFICATION OF HUMAN DCs IN SITU IN FIXED TISSUES DCs, or cells with a dendritic appearance in extraneural tissues, have long been recognized by their shape and their ability to bind silver metals (Marshall, 1956). It is likely that some of the curious cells described by Marshall are the same cells that we recognize today as the ‘dendritic’ antigen-presenting cells, but using a

Present in tumors Markedly improved prognosis Markedly improved prognosis Less with smokeless tobacco Less in tumors PBMC DCs less functional Present in some tumors Rare DCs in tumors Improved prognosis Less in tumors No relationship with prognosis Peritumoral CD1 infiltrate associated with improved prognosis Decreased CD34 DCs and myofibroblasts in malignant tumors

completely different set of criteria. Dendritic, professional antigen-processing cells are bestcharacterized by a combination of attributes: their shape, motility, phenotype, and their striking ability to induce primary T-cell responses. No single feature or phenotypic marker will identify all DCs in all tissues. The identification of DCs in fixed tissues has been problematical for two main reasons. First, the specificity of the markers used to date has been low. DCs share many of their surface and cytoplasmic proteins and enzymes with macrophages, with which they have a close affiliation, and with other activated cells, lymphoid and endothelial. Second, the sensitivity of putative DC staining has been an issue. Markers in use to date have highlighted only subpopulations of DCs, not all DCs, and only recently have markers come into use that have the potential to be more broadly applicable to a wider spectrum of DCs. There are also various types of DCs that have phenotypic and functional heterogeneity, and no single marker is generally applicable across the board. Most of the information available in humans at present pertains to the so-called myeloid DCs. Little information on the human lymphoid DC exists other than a suggestion that the plasmacytoid T cells can be induced to display DC characteristics when incubated with IL-3 but not GM-CSF (Grouard et al., 1997), and that they express IL-

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TABLE 31.2

427

Relationship between DC tumor infiltration and prognosis in malignancy

Tumor

Reference

Arsenical skin Basal cell Basal cell Basal cell

Proc. Natl Cl. Ch. 16, 127 (1992) Br. J. Dermatol. 130, 273 (1994) Br. J. Dermatol. 127, 575 (1992) Dermatol. Surg. 26, 200–203 (2000)

Breast Breast Bronchoalveolar Castleman’s disease

J. Pathol. 163, 25 (1991) Breast Cancer Res. Treat. 59, 141–152 (2000) Eur. J. Cancer 28A, 1365 (1992) Am. J. Surg. Pathol. 24, 882–888 (2000)

Cervix Cervix Cervix, stage III Cervix/penile Cervix/HIV Endometrial Endometrial

Am. J. Clin. Pathol. 99, 200 (1993) Cancer 70, 2839 (1992) In Vivo 7, 257 (1993) J. Urol. 147, 1268 (1992) Gynecol. Oncol. 48, 210 (1993) Hum. Pathol. 29, 455 (1988) Pathol. Int. 50, 616–619 (2000)

Esophageal Esophageal Gastric Gastric Gastric, stage III Gastric, EBV assoc.

Virchows Arch. 61, 409 (1992) In Vivo, 239 (1993) Int. Surg. 77, 238 (1992) Cancer 75, 1478 (1995) In Vivo 7, 233 (1993) Histopathology 36, 252–261 (2000)

Hepatoma

Cancer Lett. 148, 49–57 (2000)

Hodgkin disease Lung Lung Melanoma Melanoma/nevi

Am. J. Clin. Pathol. 101, 761 (1994) Pathology 25, 338 (1993) J. Clin. Invest. 91, 566 (1993) J. Invest. Dermatol. 100, 269 (1993) Melanoma Res. 9, 521–527 (1999)

Mycosis fungoides/SS Paget’s disease Prostate Skin tumors Thyroid (papillary) Thyroid (papillary)

In Vivo 7, 277 (1993) Br. J. Dermatol. 142, 1190–1194 (2000) Prostate 19, 73 (1991) Arch. Dermatol. 131, 187 (1995) Z f. Chir. 117, 603 (1992) Am. J. Pathol. 156, 831–837 (2000)

Less compared with normal skin Less in tumorsa ?Improved Close proximity of stromal mast cells and dermal dendrocytes surrounding BCC nests ?Improved prognosis Clear stage and grade relationship No effect Localized clonal proliferation of follicular DCs Less in HPV tumors Improveda Marked improved prognosisa Less with HPV infection Less in AIDS Langerhans infiltration favorable S100/silver stained nucleolar organizer Langerhans cells negatively correlated with FIGO grade Marked improved prognosis Direct relationship to grade Marked improved prognosisa More in tumor draining lymph nodes Marked improved prognosisa 7/56 tumors EBV, EBER correlated with prominent lymphoid reaction (P  0.0002) and numerous S100positive dendritic cells CD83 DCs in liver tissues were significantly lower in HCC compared with cirrhosis FDC improved prognosisa Marked improved prognosisa Related GM-CSF production Inverse with tumor thickness Decreased DR and CD1a DCs in sun-exposed skin in the vicinity of naevi Marked improved prognosisa Many DCs Improved prognosis Less in tumors No effect Most DCs had an immature phenotype and were located at the invasion edge of the tumor

a

Indicates statistically significant difference when compared to normal or fewer DCs.

3Rα (Olweus et al., 1997; Cella et al., 1999; Facchetti et al., 1999). Importantly, too, even the best-characterized subtypes of DCs, such as Langerhans cells, undergo a life cycle that

involves differentiation, maturation and functional activation that may be accompanied by changes in enzyme activity and phenotype at each stage.

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When trying to assess the significance of DCs in cancer tissue, the challenge has been to find markers that will be informative in fixed and embedded tissues so that the vast repositories of pathology departments can be tapped. Markers applicable to isolated cells or frozen tissues are dealt with elsewhere (Hartgers et al., 2000). New markers that may be applicable to tissues for the demonstration of DCs or their subsets include CD83 (Zhou et al., 1992; Zhou and Tedder, 1995) fascin (p55) (Mosialos et al., 1996), CMRF-44 (Hock et al., 1994) and CMRF-56 (Hock et al., 1999) and high expression of the IL-3α receptor on the plasmacytoid T cell. Published experience with these markers in fixed tissues (other than fascin) is very limited at this point. The most commonly applied markers for DCs in human tissues identify the HLA II molecules and the associated invariant chain, expressed in high density on the cell surface (Steinman, 1991). Many anti-HLA class II antibodies will react with DCs in frozen tissues, less well in fixed, embedded tissue, but are not specific for DCs, being also expressed on macrophages, B cells, endothelial cells and other assorted activated cells, including T cells and epithelia. The high concentration of class II MHC molecules on DCs, both constitutive and following inflammatory induction (Cella, 1997a), has been used to identify them by resorting to high dilutions of anti-HLA II antibodies, effectively diluting out the staining of competing cells. A number of antibodies have been specifically produced to demonstrate class II MHC in formalin-fixed tissues; most widely used is LN3, which recognizes the DR subregion (Marder et al., 1985). Other antibodies produced specifically to react with human DCs also bind the class II MHC molecule, such as RFD-1, which reacts with an HLADQ-associated antigen, and RFDR1, reacting with HLA-DR, -DQ and -DP (Poulter et al., 1986). The use of anti-class II MHC antibodies as DC markers in fixed tissues suffers from the limitations of low specificity, and it is not possible to know that any stained cell is, in fact, a DC. The S100 family of intracytoplasmic calciumbinding proteins is implicated in a wide range of cellular functions, and has traditionally been

used to demonstrate dendritic populations such as Langerhans cells and the lymph node interdigitating DC (Takahashi et al., 1982; Uccini et al., 1986). Specificity of staining when polyclonal antibodies are used is low since there are many other cell types, including S100-reactive lymphoid cells, nerve, fat, chondrocytes and melanocytes that also express S100 proteins, but in the context of tissue reactions these other cells can be visually discriminated. As other DC markers become available, it is also becoming evident that the use of polyclonal anti-S100 protein antibodies reveals only subpopulations of DCs. DCs contain largely homodimeric S100β with two β chains, and there are monoclonal antibodies to the S100β. Neural tissues contain mostly S100α, but S100α itself is heterodimeric with an α and a β chain, so that crossreaction is still possible. Most available S100 monoclonal antibodies are poorly reactive in fixed tissues. CD68 is a lysosome-associated molecule, the human homologue of mouse macrosialin, a member of the LAMP family with a macrophage-specific mucin-like domain. Antibodies to CD68 stain tissue macrophages in a vacuolar cytoplasmic pattern (Weiss, 1994). Even though there are CD68 antibodies that recognize the presence of a paranuclear dot of CD68 in DCs (Betjes et al., 1991), CD68 cannot be considered a specific marker for DCs because macrophages, developing myeloid cells, and other tumor cells express the antigen (Gloghini et al., 1995). Some of the immune accessory molecules, such as CD40, CD80, CD86 and the adhesion molecules, CD11c, ICAM-1 and ICAM3 (CD54 and CD102), may be present in high concentration on DCs at various times, but the sensitivity and specificity of their detection in tissues are too low to allow these to be used as unique markers of DCs. There are markers that are normally expressed only on certain DC subsets. CD1a outside of the thymus serves as a good marker of the Langerhans cell in tissue (Murphy et al., 1981), although it should be borne in mind that CD1a will be expressed at a low level on the surface of monocytes stimulated with GM-CSF (Kasinrerk et al., 1993) and on some monocytic and

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myelomonocytic leukemias (Misery et al., 1992). The Lag antibody demonstrates an antigen, Langerin (CD 207) associated with the Langerhans cell Birbeck granule (Kashihara et al., 1986), an organelle considered to be the hallmark of the Langerhans cell, but is reactive only in unfixed tissue (Valladeu et al., 2000). Follicular dendritic cells (FDCs) are marked by their expression of CD21, CD35, and other FDCrelated antibodies, such as Ki-M4. S100 and fascin are variably demonstrable on follicular DCs (Said et al., 1997). Dermal and interstitial DCs express a tissue transglutaminase, factor XIIIa (Cerio et al., 1989a). An antibody, Ki-M9, that identifies a putative lymph node sinus DC has been described (Wacker et al., 1997) and these same cells express factor XIIIa and fascin. Newer molecules of interest for the study of DCs in situ include p55 (fascin), CD83, Dec-205 (Witmer-Pack et al., 1995) RelB, CMRF-44 and CMRF-56. CD83 appears to be unique in blood DCs (Zhou et al., 1995). Tissue application in frozen tissue reveals strong staining of lymph node paracortical interdigitating cells (Zhou et al., 1992) and freshly isolated Langerhans cells. A population of smaller lymphoid cells also stains, and Hodgkin cells stain with both frozen and fixed tissues (Sorg et al., 1997). CD83 and CMRF44 are demonstrable on frozen sections, in normal skin, mostly in association with CD83positive lymphocytes (McLellan, 1998). CD83positive DCs have been localized in various liver diseases (Tanimoto et al., 1999). In fixed lymph node, CD83 identifies only a tiny proportion of the cells identified by the other markers mentioned previously. Fascin, a 55 kDa actin-bundling protein has widespread reactivity on tissue DCs, interdigitating, follicular, thymic, splenic and dermal, but is not detected on Langerhans cells in situ (Said et al., 1997). The usefulness of fascin as a tissue marker is, however, limited by the low specificity, since other cell types, most notably capillary endothelial cells, also express the antigen. Interestingly, Hodgkin cells have been shown to express both fascin (Said et al., 1997) and CD83 (Sorg et al., 1997). Its application to the differential diagnosis of histiocytic disorders

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in children reveals that Langerhans cell lesions can be distinguished from so-called indeterminate cell histiocytomas because the first have Birbeck granules, are Lag, CD1a, and are fascin, and the second are CD1a, fascin, but lack Birbeck granules (Jaffe et al., 1998). Juvenile xanthogranuloma cells, presumably dermal or interstitial DCs and those DCs expressing factor XIIIa, also mark with fascin. CMRF-44, a monoclonal antibody to a DC surface membrane, identifies activated bloodderived DCs after a brief period in culture (Fearnley et al., 1997). Detailed tissue exploration with this antibody and the OX40 ligand in humans (CD134) (Ohshima et al., 1997), remains to be described. Mannose receptors are important for internalization of glycosylated ligands, a property of both macrophages and DCs in their phase of antigen uptake. Mannose receptors can be demonstrated on immuno-suppressed macrophages but not on monocytes or activated macrophages, and in tissues, on thymic cortical DCs, lymph node paracortical DCs, and on bone marrow macrophages (Noorman et al., 1997). Consequently, mannose receptor can demonstrate some DCs in tissue but does not distinguish them from macrophages. CMRF-56 is found on isolated Langerhans cells and dermal DCs (Hock, 1999); its relevance to fixed tissues is not yet established. RelB (NFκB) expression is detectable in the mature postmigration DC, and has been detected by immunohistology for the protein and by in situ hybridization for the mRNA (Clark et al., 1999). Both are demonstrable in tonsillar and interdigitating lymph node DCs, but in only a few dermal DCs and not in Langerhans cells. The antibody alpha-p65 revealed the presence of NFκB also in inflamed intestinal epithelial cells and macrophages (Rogler, 1999).

DCs AND THEIR DISORDERS The various subtypes of DCs in humans can be characterized in situ by a combination of topography and phenotype. Table 31.3 lists the

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TABLE 31.3 cells

Markers used to characterize dendritic Target

Clone/ Antigen

Precursor

IL-3R-α

Immature

HLA-DR paranuclear CD68 paranuclear CD1a Fascin lo S100

LN3 KP-1 010 p55 S100β

Mature

HLA-DR surface, hi CD68 granular Fascin hi S100 Rel B, nuclear CD83

LN3 KP-1 p55 S100β Rel-B HB15A

available markers for immunohistochemistry and Table 31.4, the major forms and the lesions that can now be ascribed to the dendritic cell types.

DC-RELATED DISORDERS: PROLIFERATION(S) Langerhans cell histiocytosis Many aspects of Langerhans cell disease and other DC proliferative disorders have been TABLE 31.4

summarized in a publication stemming from the Nikolas symposium, an annual ‘think-tank’ dedicated to the histiocytoses and in recent reviews (Pritchard et al., 1994; Egeler and D’Angio, 1998; Arceci, 1999; Jaffe, 1999). Basically, Langerhans cell histiocytosis (LCH), is a clonal disorder in which abnormal Langerhans cells accumulate at various body sites not usually known to harbor them (Willman et al., 1994; Yu et al., 1994). Diagnosis in all of the varied clinical forms, localized, multifocal and visceral, rests on identifying the Langerhans cell in the tissue lesions, which has until recently required the demonstration within the cells of the characteristic Langerhans cell granule or Birbeck granule. Now, the demonstration of the CD1a molecule at the cell surface, in a lesion that has the morphologic hallmarks of Langerhans cell disease, is sufficient to confirm the diagnosis (Favara et al., 1997). The 010 antibody has the required sensitivity and specificity to demonstrate CD1a in paraffin-embedded tissues, and will mark LCH cells (Emile et al., 1994, 1995). The cells of LCH have the phenotype of immature Langerhans cells because 86 LCH cells also retain their Birbeck granules, while activated Langerhans cells tend to lose theirs. HLA-DR class II is intra-cytoplasmic and not membrane in distribution. In addition to CD1a, LCH cells will express paranuclear

DCs and their disorders

Dendritic cell

Phenotype

Disorder

Langerhans cell

CD1a, S100, LCG, Langerin HLA-DR paranuclear

Langerhans cell histiocytosis

Indeterminate cell

CD1a, S100, fascin

DC histiocytoma indeterminate

Interdigitating DC

S100, fascin HLA-DR, hi

DC histiocytoma, IDC type

Lymphoid DC precursor

IL-3R-α, CD68

Not known

Dermal dendrocytes (and subtypes)

Factor XIIIa, fascin, CD68

Xanthogranuloma family Dermal dendrocytomas

Follicular DC

CD21, CD35, Ki-M4, S100/, fascin

DC histiocytoma, FDC type

Sinus DC

Ki-M9, fascin, CD68, S100

?Rosai–Dorfman disease

Abbreviations: LCG – Langerhans (Birbeck) granules of the Langerhans cell DENDRITIC CELLS IN DISEASE

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CD68, S100 protein and CD4, but not fascin or CD83 (Jaffe et al., 1998). The Lag antigen identifies the intracytoplasmic Birbeck granule but antibody detection is possible only in frozen tissues (Kashihara et al., 1986). There are quantitative differences in staining that reveal that LCH cells are immature DCs. The HLA-DR staining is intracytoplasmic, and CD68 is present as a paranuclear dot. Fascin expression is low, nuclear Rel-B is absent and CD83 is not expressed.

Secondary DC reactions Clusters of Langerhans cells have been described in the tissues in a number of conditions, usually designated as LCH arising in association with the underlying disorder. Such collections are described in malignant lymphomas (Willman et al., 1994), the thymus in myasthenia gravis, and in association with other tumors (Egeler et al., 1993; Burns et al., 1983). There is no information on the clonality of the process, but it is best thought of as a reactive hyperplasia of Langerhans cells, possibly an immune reaction to the tumor, and not an independent disease process because no evidence of LCH exists at other sites (Favara et al., 1997).

Juvenile xanthogranuloma and related disorders The cutaneous lesion juvenile xanthogranuloma (Hernandez-Martin et al., 1997) and its systemic counterparts, which can involve deep soft tissues, brain and meninges as well as other organs (Freyer et al., 1996) appear to be lesions of the dermal and interstitial DCs (De Graaf et al., 1992; Sangueza et al., 1995; Nascimento, 1997). The histology encompasses a mixture of angular histiocytes, lipid-filled xanthomatous forms, and multinucleated Touton-type giant cells. The in situ phenotype is generally fascin, factor XIIIa, CD68 (especially when using PG-M1 antibody), S100 and CD1a. It is likely that their phenotype changes over time as the dendritic histiocytes become lipidized and take on more macrophage-like characteristics

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while losing their dermal dendritic features. Healing lesions are fibrotic and lose the macrophage population, and are generally referred to as dermatofibromas. The same dermal DC type is involved in a number of other dermal and soft tissue masses and some systemic disorders that have been given a variety of names over the years. It is likely that cellular xanthogranulomas, benign cephalic histiocytosis, papular xanthoma, progressive nodular histiocytosis, spindle cell xanthogranuloma, xanthoma disseminatum and Erdheim–Chester disease are the same lesion with varying site or clinical presentation (Zelger et al., 1995). An important feature that they share is their biological behavior, which tends to be self limiting, albeit slowly, with regression over months to years, so that aggressive chemotherapy should be averted by correct diagnosis. Xanthogranuloma can behave much like Langerhans cell disease in its systemic manifestations, but xanthogranulomas may also be noted in patients with LCH and myelomonocytic leukemias, accentuating the close relationship between the disorders. Xanthoma disseminatum is a normolipemic mucocutaneous variant of the juvenile xanthogranuloma family that affects young adults with many dermal lesions and involvement of mucosa, often that of the upper airways (Weiss and Keller, 1993). Brain involvement occurs and diabetes insipidus, a hallmark of Langerhans cell disease, is seen transiently in up to 40% of patients. The histopathology and phenotype is that of juvenile xanthogranuloma (factor XIIIa, CD68, S100, CD1a). Erdheim–Chester disease has histopathologic features very much like the xanthogranulomas and is distinguished by the symmetrical metaphyseal and diaphyseal osteosclerosis demonstrable on X-ray.

Non-Langerhans dendritic cell histiocytomas There are a number of descriptions of dermal or soft tissue lesions that have an appearance and phenotype very similar to the Langerhans cell lesions, but differ in that Birbeck granules are

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not demonstrable, so-called indeterminate cell lesions (Wood et al., 1985; Sidoroff et al., 1996; Manente et al., 1997). Since Birbeck granules can only be demonstrated in 2–69% of Langerhans cells in LCH (Mierau et al., 1986), this distinction alone seems tenuous, but the demonstration of fascin in these lesions, but not in Langerhans cell disease, adds another distinguishing feature (Jaffe et al., 1998). The biological behavior of the dermal and soft tissue lesions appears to be benign, but intracranial lesions have recurred with more aggressive growth. The phenotype is generally CD68, HLA-DR, fascin, variably S100, and FXIIIa, CD1a, and Langerhans granule. Lesions of this kind have been referred to as dermal dendrocytomas (Cerio et al., 1989a, 1989b; Nickoloff et al., 1990), although they are not limited to the skin and may even be intracranial (DC histiocytomas) (Jaffe, 2000). Some of these lesions have a ‘mature’ DC phenotype, when contrasted with LCH. CD1a is absent in the mature lesions, HLA-DR staining is of the high membrane type, CD68 is present as a diffuse granular pattern, fascin staining intensity is high and Rel-B nuclear stain is variable.

Monocytic and myelomonocytic leukemias with dermal lesions of dendritic phenotype Adults are described who have myelomonocytic or monocytic leukemias that have pre-DC phenotype or develop one in culture (Misery et al., 1992; Srivastava et al., 1994). In some of these patients there are dermal (Lauritzen et al., 1994) or disseminated DC lesions that have the features of Langerhans cell disease (Chaux et al., 1989). In one of these (Takahashi et al., 1992), a serum factor was isolated that caused a human monocytic cell line to mature to interdigitatingtype DCs.

Dendritic cell sarcomas Soft tissue and lymph node-based lesions have been described, some of which have the biological potential for recurrence, metastasis and

resistance to chemotherapy, sufficient for them to be deemed malignant. The lesions are generally composed of spindled cells without other distinguishing morphological features, but on phenotyping have been assigned to one or other of the DC forms, so that follicular DC sarcomas Ki-M4, CD21, CD35 (Chan et al., 1997; Masunaga et al., 1997), interdigitating DC sarcomas, HLA-DR, CD68, S100 (Meittinen et al., 1993; Nakamura et al., 1994; Rousselet et al., 1994; Gaertner et al., 2001) and Langerhans cell sarcomas CD1a (Lauritzen et al., 1994) are described. The International Lymphoma Study Group has defined histiocytic and accessory (dendritic) cell lesions by phenotype (Grogan, 2000).

DC-RELATED DISORDERS: DYSFUNCTION Wiskott–Aldrich syndrome The syndrome is X-linked and manifests with thrombocytopenia, eczema and immune deficiency. Wiskott–Aldrich syndrome protein (WASp) regulates the actin cytoskeleton (O’Sullivan, 1999). Binks et al. (1998) have suggested that the clinical syndrome could be accounted for on the basis of DC dysmotility and abnormal cell trafficking.

Identification of DCs within human tumors Many of the studies designed to evaluate DC infiltration into tumors were done 5–10 years ago, before the explosion of interest in DC biology. What had become apparent in many murine experimental studies was that DCs were either decreased in number of dysfunctional, associated with decreased expression of a number of co-stimulatory molecules. It has also been envisioned that DCs could be dysfunctional based on failure to generate them in the bone marrow (Gabrilovich et al., 1996a, 1996b, 1996c), failure to migrate across endothelial barriers that

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did not permit their emigration (Piali et al., 1995), rapid destruction of such cells as demonstrated recently by Michael Shurin in our group (Shurin et al., 1997b), or enhanced migration out of the tumor site related to increased expression of molecules promoting DC maturation and migration such as TNFα. Some investigators (Halliday et al., 1991, 1992) have also suggested that the tumor represents a ‘black hole’ for DCs and that they succumb to apoptotic death at those sites, explaining the paucity of DCs in poor-prognosis tumors. Interestingly, they observed an enhanced number of DCs in nonimmunogenic tumors and attributed this to a cytokine which recruited DCs into a site but prevented their emigration, possibly IL-10 (Din et al., 1997). The notion that hematopoietic malignancies could provide differentiated DCs presenting antigen derived from malignant hematopoietic precursors (Costello et al., 2000) in the setting of human acute myeloid leukemia was recently examined. Interestingly CD34/CD38 leukemia precursors, in contrast with CD38 progenitors, failed to differentiate into DCs when cultured under appropriate conditions seemingly required for antigen recognition. Circulating DCs, derived from hematopoietic precursors, can also be studied following bone marrow transplantation (Galy et al., 2000). Recovery of naïve CD4 and CD8 T lymphocytes was quite delayed for many months following transplant, whereas DC subsets rapidly normalized when measuring circulating levels of the antigen-presenting CD11c DCs. This suggests that the immunosuppression observed following ablative chemotherapy is of T-cell origin and not attributable to the circulating DCs. Malignant B cells have a specific sensitivity to anti-apoptotic signals (Ghia and CaligarisCappio, 2000) that favor their survival (in part provided by CD40 and DC), whereas they have become insensitive to pro-apoptotic signals. The role of DCs in both promoting as well as modifying and inhibiting tumors is thus suggested. Most of the observations in human tumors have been descriptive and allowed pathologists the opportunity to suggest that the increased number of DCs being associated with good

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prognosis might be attributable to direct lysis of tumors by DCs. This seemed implausible initially, but we now have some data from experimental models in the mouse (Shimamura and Baar, unpublished) that suggest that DCs may indeed cause limited apoptotic death in some tumors. This may be related to their ability to rapidly take up both particulate antigen as well as free, unattached cells (to each other and to the stoma and basement membrane). Furthermore, Fas mediated pathways wherein either lymphoid or myeloid DCs expressing FasL could be envisioned to mediate Fas crosslinking and induction of apoptotic death. This is being actively tested at this time. Our group has also demonstrated that several tumors including prostate (Pirtskhalaishvili et al., 2000) induces progressive suppression of DCs. Coincubation of human DCs with prostate cancer cell lines led to high levels of DC apoptosis. Stimulation of DCs with CD40 ligand (CD154), IL-12 or IL-15 prior to coincubation with prostate cancer cells resulted in increases in DC survival suggesting a possible therapeutic approach.

Other findings related to tumors In patients with lung cancer (Nakajima et al., 1985; Almand et al., 2000) S100-positive Langerhans-like cells have been identified and shown to be at highest density in bronchoalveolar (alveolar II) and well and moderately differentiated squamous cell carcinoma. There are fewer cells observed in small-cell lung cancer and poorly differentiated squamous cell carcinoma. Similar findings have been observed in esophageal squamous cell carcinoma, in which a dense infiltration of class II-positive DCs is associated with an improvement in survival rate (P  0.01) (Furihata et al., 1992; Imai and Yamakawa, 1993). In patients with gastric cancer, improvements in survival were limited to patients with stage III disease (Tsujitani et al., 1993), with marked infiltration associated with greater survival time (P  0.001). In colorectal cancer (Ambe et al., 1989), the grade of S100positive DC infiltration was related to the density of lymphocytic infiltration in the tumor (P 

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0.05). Patients survived longer when they had more than 30 cells per 10 high-power fields (HPF) than those with fewer cells (less than 30 cells/HPF) (P  0.001). In bladder carcinoma (Inoue et al., 1993) it appears that the most important factor affecting prognosis was distant organ and/or lymph node metastasis (P  0.01 as well as number of S100 DC, with a hazard ratio of 0.26 (P  0.01). Interestingly, class II expression of the tumor was also a prognostically important factor (P  0.05). This probably relates to the presence of interferon gammaexpressed locally. A recent evaluation of malignant ascites revealed that DCs identified as lineage-negative, class II-positive cells were present in up to 3% of the mononuclear cells (Melichar et al., 1998) but expressed less class II and low levels of CD80, again consistent with a nonstimulatory or immunosuppressed phenotype. The concentration of neopterin, reflecting macrophage activation, in ascitic fluid correlated inversely with the number of lineage-negative HLA-DR cells found in ascites (Spearman correlation coefficient 0.44; P  0.05) and directly with the concentration of IL10 in ascitic fluid (Spearman correlation coefficient 0.40; P  0.05). Thus factors associated with the tumor microenvironment appear to influence both the number of DCs and their expression of costimulatory molecules. Recently, it has become clear that the killer inhibitory receptor family of molecules, which had previously been identified in T cells and NK cells, also exists as closely related family members, so-called LIRS or ILT-3 (Cella et al., 1997b). These molecules may be critically important for class I detection by inflitrating DCs, which may lead to unique DC effector functions and which we are now exploring in the laboratory. Savary and colleagues investigated flow cytometry as a means of evaluating the number and maturation/activation status of DCs in peripheral blood mononuclear cells (PBMC of both normal donors and cancer patients (Savary et al., 1998). DCs were identified as HLA-DR lineage cells (less than 1% mononuclear cells). They had high forward light-scatter characteristics and coexpressed CD4, CD86 and CD54 sur-

face antigens, but lacked the lineage-associated surface markers of T cells, B cells, monocytes, granulocytes or NK cells. Interestingly, the frequency of DC-like cells in PBMC of chemotherapy-treated cancer patients was lower than that of normal individuals (mean  SE 0.36  0.05%, 0.14  0.06% and 0.75  0.04%, respectively). There are other novel markers that could be used to distinguish DCs from other cells. These include CD1, S100 staining, high expression of the MHC class II molecules, and, most recently, expression of the p55 actinbundling protein, fascin (see above), a cytosolic marker that was found within EBV-transformed B cells (Sonderbye et al., 1997), within both follicular and nonfollicular DCs, but more recently, also within the Reed–Sternberg cells in all but the lymphocyte-predominant forms of Hodgkin disease (Pinkus et al., 1997). Immune dysfunction and profound decreases in mature DCs in the peripheral blood were noted in 93 patients with advanced breast, head and neck, and lung cancer (Almand et al., 2000). This is associated with accumulation of cells lacking markers of mature hematopoietic cells. Such immature DCs were noted most frequently in patients with advanced stage and duration of disease. Following tumor removal, partial reversal of these changes is noted. Interestingly, the immature cells noted in the peripheral blood were closely associated with increased plasma level of VEGF but not other cytokines and were decreased in patients who received antivascular endothelial growth factor antibodies. Patients (30) with gastrointestinal malignancies had a clear decrease in T-cell numbers associated with diminished mature DCs but no clear relationship with the number of immature DCs, again pointing to the important interactions between DCs and T cells (Lissoni et al., 2000). Only mature DC deficiency was associated with lymphocyte subset alterations in cancer patients.

CONCLUSION Much has been learned about the various manifestations of DCs in human tissues and in

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REFERENCES

particular in the setting of cancer over the last 30 years since the DC was first defined as an identifiable cell type. Future challenges will be to understand the natural biology of these cells as they traverse tissues, the manifestations of arrested biology revealed by neoplastic DCs, and the errant findings of DCs within tumor and how their recruitment, maturation or emigration is modified. Of particular interest will be the determination of the role of the more recently identified lymphoid DC and how it migrates through various tissues and, in particular, whether a nominal tumor-pathologic variant might be identified. Tissue markers for DCs of varying kind are needed for a number of reasons. Clearly, in order to understand the role of the DC in the immune regulation of cancer it is important to know which are antigen-acquiring and which are the effector DCs. To gain insight into the nature and biology of lesions made up of DCs it requires that there be standardized ways of looking at these cells and categorizing them. This is an area of evolution, and DC phenotyping in situ lags behind the functional understanding of DC capabilities.

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32 Dendritic cells in rejection and acceptance of solid organ allografts Anthony J. Demetris, Noriko Murase, John J. Fung From the Thomas E. Starzl Transplant Institute, University of Pittsburgh Medical Center, Pittsburgh, PA, USA

This grasping of the whole is obviously the aim of science as well, but it is a goal that necessarily lies very far off because science, whenever possible, proceeds experimentally and in all cases statistically. Experiment, however, consists in asking a definite question, which excludes as far as possible, anything disturbing or irrelevant. It makes conditions, imposes them on Nature, and in this way forces her to give an answer to a question devised by man. She is prevented from answering out of the fullness of her possibilities since these possibilities are restricted as far as practicable. . . . The workings of Nature in her unrestricted wholeness are completely excluded. If we want to know what these workings are, we need a method of inquiry which, imposes the fewest possible conditions, or if possible no conditions at all, and then leaves Nature to answer out of her fullness. Synchronicity: An Acausal Connecting Principle C.G. Jung

INTRODUCTION

organ, but for many disorders, this simply represents a local manifestation of a more pervasive problem. An exception to this generalization is organ-specific toxic injury or organ-based metabolic diseases, where replacement of the defective organ corrects a systemic problem and brings about a true cure. A systemic perspective is also of considerable importance in the study of transplantation physiology and immunobiology. For example, insulin secreted by pancreatic islet allografts can regulate recipient blood sugar; a new liver will receive blood and nutrients from the recipient intestines, from which it will synthesize cholesterol that can be used in the construct new ‘recipient’ cells. Fortunately, neither basic

The field of solid organ transplantation is based largely on the concept that replacement of an irreversibly damaged organ in an otherwise ‘healthy’ recipient can significantly prolong survival or even cure some diseases. Although the hope for clinical success is based on this premise, many allograft recipients suffer from recurrence of the original disease. For example, chronic viral hepatitis type B and C almost invariably attack the new liver after hepatic replacement. This unfortunate reality illustrates the importance of a systemic, or nonlocal perspective in transplantation biology: a specific disease may primarily manifest in a single Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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researchers nor clinical physicians worry about compatibility between certain donor and recipient physiologic or biochemical systems. Most of these complex systems have universal or ‘nonlocal’ properties that enable an allograft to spontaneously adjust to its new environment – one size fits all. The most important exception to the above generalization is the immune system. Its ‘local’ or nonuniversal properties prevent spontaneous integration of donor hematolymphoid cells with those of the receipient, and vice versa. Instead, the immune cells from the two different individuals react violently with one another, which leads to significant problems for the transplant surgeon/clinician. Unless this ‘rejection’ reaction is controlled with immunosuppression, it will eventually destroy the allograft. In experimental animals, allograft acceptance occurs ‘naturally’ under some circumstances, or can be engineered predictably, without using immunosuppression. Unfortunately, this success is not yet achievable in humans: most require life-long immunosuppression to prevent rejection of the allograft, except for a fraction (~20%) of liver recipients. After taking immunosuppressive medication for several years after transplantation, these few fortunate patients are able to discontinue all immunosuppressive drugs without rejecting their allografts. Thus, it is biologically possible to transplant a vascularized organ between two unrelated individuals without long-term immunosuppression, but it is rare. Much of transplantation research is devoted to discovering how to make this routinely possible. Transplantation biology could be thought of as an experiment conducted to investigate the ‘local’ or nonuniversal properties of the immune system, including MHC antigens and the consequences of expression of these and other important molecules on different cell types. Since donors and recipients frequently have different MHC antigens and sex chromosomes, these molecules can be used as markers to identify individual cells and trace their location and behavior in functional assays at various times after transplantation. Discoveries made using

these technologies have brought together two previously unlinked research fields in transplantation immunology: the (1) deleterious; and (2) potentially beneficial functions of donor hematolymphoid cells. The sections below overview both lines of investigation and highlight recent discoveries that place dendritic cells (DCs) at a pivotal junction on the road to allograft rejection or acceptance.

OVERVIEW OF ORGAN-BASED IMMUNITY AND DC FUNCTION Before directly proceeding to a discussion of the role of DCs in transplantation, first, it is important to briefly overview organ-based immune physiology. This refers to the interactions between a dynamic network of hematolymphoid cells, especially DCs and their precursors, which are present in the interstitium of all solid organs, with the central lymphoid organs and regional lymph nodes. The organ-based interstitial cells are derived primarily from bone marrow progenitors; locally based hematopoietic stem cells and precursors in various stages of maturation can also contribute to this pool (Steinman et al., 1997, 1999). There is considerable variation in the absolute number, relative proportions and phenotypes of hematolymphoid cells among the commonly transplanted organs (Murase et al., 1995; Holzinger et al., 1996). This variation is primarily related to physiological function. For example, organs in direct contact with the external environment, such as the lungs and intestines, have an exaggerated complement of mature T- and Blymphocytes that expand locally after contact with commensal bacteria in neonatal life. These large local lymphocyte populations intermix with DCs, and arrange spontaneously into organized ‘mucosal associated lymphoid tissue’. Their structure is essentially identical to that seen in lymph nodes, complete with organized B cell follicles with follicular DCs and germinal centers and interfollicular T-cell zones, rich in interstitial DCs, discussed in more detail below.

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Livers have less mature T cells than intestines and lungs, but more macrophages (Kupffer’s cells) and NK cells consistent with the liver’s role as the ‘first line of defense’ against intestinal pathogens, and a filter of various opsinized material, including microorganism, activated platelet aggregates, and coagulated proteins (Seki et al., 2000). Vascularized organs also contain immature monocytic/myeloid DC precursors, scattered throughout the interstitial connective tissue, and rare mature DCs. Even those organs that are not considered to normally harbor a significant population of hematopoietic cells, such as the heart and kidneys, contain such cells in the interstitium (Steptoe et al., 2000). There is a greater absolute number of mature DCs in livers than ‘leukocyte-poor’ organs such as the pancreas, heart and kidney, but they comprise a lower proportion of the total leukocyte pool in the liver (approximately 15–25%) compared to these other organs (approximately 40–60%) (Steptoe et al., 2000). A hierarchical arrangement of DC density is arranged as follows (from most to least): heart, kidney, pancreas and liver (Matsuno and Ezaki, 2000; Steptoe et al., 2000). In tissue sections, immature DCs are scattered throughout the interstitium of organs, and are concentrated near efferent lymphatic vessels in the adventitia of arteries. They are also more plentiful near epithelial-lined conduits that are in contact with the external environment, such as in the mucosal associated lymphoid tissue of the lungs (Gong et al., 1992) and intestines (Inaba et al., 1994); and portal tracts of the liver (Prickett et al., 1988; Demetris et al., 1991). Except for DCs in the T cell zone of lymph nodes, spleen, thymus and mucosal associated lymphoid tissue (Iwasaki and Kelsall, 2000), most hepatic DCs (Lu et al., 1994) and other organ-based DCs have an ‘immature’ phenotype (Austyn et al., 1994), characterized by phagocytic capacity (Austyn et al., 1994; Lu et al., 1994; Steinman et al., 1997), inefficient stimulatory capacity in MLRs (Austyn et al., 1994; Steinman et al., 1997), lower density of MHC class II compared to mature DC (Lu et al., 1994), and low or absent co-stimulatory

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molecule expression (Inaba et al., 1994; Steinman et al., 1997). Immature DCs monitor the local environment and internalize protein antigens into late endosomes and lysosomes rich in MHC class II molecules. MHC class II–peptide antigen complexes are not formed unless the DCs are exposed to various signals precipitated by local injury/stress, such as tumor necrosis factor family members (TNF, CD40 and TRANCE), bacteria, certain viruses and very likely, RP-105 and other Toll family members (Steinman et al., 1997, 1999). The capacity of late endosomes and lysosomes to produce MHC class II–peptide complexes can be strictly controlled during DC differentiation. This helps to coordinate antigen acquisition and inflammatory stimuli with formation of TCR ligands. The increased ability of mature DCs to load MHC class II molecules with antigen contributes to the more than 100-fold enhancement of immune responses observed when immature and mature DCs are compared (Inaba et al., 1997, 1998, 2000; Steinman et al., 1997). During their final stage of maturation, DCs redistribute their MHC class II products from intracellular compartments to the plasma membrane and they regulate the initial intracellular formation of immunogenic MHC class II–peptide complexes (Inaba et al., 1997, 1998; Steinman et al., 1997). These mature DCs then migrate to central lymphoid organs (e.g. spleen) and regional lymph nodes via the circulatory system and lymphatics, respectively, where they stimulate a T-cell response (Steinman et al., 1997). DCs in the T-cell areas of peripheral lymphoid organs such as the spleen, lymph node and Peyer’s patch are called interdigitating cells (IDCs). These cells express MHC II, invariant chain, high levels of self-antigens (Inaba et al., 1997; Steinman et al., 1997), accessory molecules such as CD40 and CD86, a multi-lectin receptor for antigen presentation called DEC205, the integrin CD11c, several antigens within the endocytic system and, in the human system, molecules termed S100b, CD83 and p55 (Steinman et al., 1997, 1999).

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Thus, DCs separate in time and space two important functions: antigen capture in the periphery and antigen presentation in the organized lymphoid tissues (Steinman et al., 1999). In general, DCs capture and present antigen from a variety of endogenous and exogenous sources, including dying cells (Inaba et al., 1998, 2000; Steinman et al., 1999), and are crucial for the initiation of primary immune responses of both CD4 and CD8 T lymphocytes. Although they act as ‘nature’s adjuvant’, DCs can also mediate tolerogenic reactions (Banchereau, 1997). Recent studies suggest that there may be at least three pathways of DC differentiation (and possibly more). The first is epidermal DCs or Langerhan’s cells (Steinman et al., 1997). The second are called myeloid DCs and are classically thought of as immunostimulatory DCs. The third pathway is derived from lymphoid precursors, which may function as regulatory DCs (Austyn, 1998). In some instances, lymphoid DCs can express Fas-ligand (Suss and Shortman, 1996), and appear to be involved in the induction of central, as well as peripheral tolerance (Austyn, 1998; Steinman et al., 1997). Mice deficient in the NFKB transcription factor relB have defects in DC maturation and secondary lymphoid tissues are absent (DiMolfetto et al., 1997). These mice also appear to have impaired negative selection in the thymus, and defective immune responses in vivo (DiMolfetto et al., 1997). Numerous attempts are being made to exploit both the immunogenic and tolerogenic properties of mature and immature DC, depending on the clinical need. In general, investigators are attempting to coordinate in time and space the antigen acquisition and presentation functions of DCs for the purpose of treating diseases. For example, immature DCs can be pulsed with tumor antigens and then allowed to mature with the hope that defective tumor immunity will be enhanced. Precursor cells collected from the peripheral blood after mobilization with exogenous growth factors and cytokines, can be expanded and/or manipulated (e.g. gene transfer, cytokine treatment), ex vivo, then reintroduced

into the same recipient. Theoretically, this approach could be used to treat a variety of conditions, including cancer, autoimmune disorders, infections and allograft rejection.

HEMATOLYMPHOID AND DENDRITIC CELLS AS POTENTIATORS AND FACILITATORS OF REJECTION Acute ‘cellular’ rejection When an organ becomes an allograft, the donor hematolymphoid cells carried with it become known as ‘passenger leukocytes’. Emigration of donor cells out of the allograft marks the beginning of a very interesting series of events, which is still incompletely understood. Snell (1957) was the first to recognize that the hematolymphoid cells are especially immunogenic; Steinmuller (1967) provided experimental evidence in support of this concept. He showed that chimeric donor skin allografts (passenger leukocytes allogeneic and epidermal cells syngeneic to recipient), were permanently accepted, but the recipients became sensitized to the hematopoietic cells. Several other manipulations such as lethal irradiation of the donor and reconstitution with allogeneic bone marrow (Guttmann et al., 1969); culturing organs before transplantation (Lafferty et al., 1975); and ‘parking’ allografts in intermediate immunoincompetent hosts (Lechler and Batchelor, 1982a, 1982b), also reduces the passenger leukocyte load in donor organs. The subsequent prolongation of allograft survival brought about by such treatment further supports the contention that the passenger leukocytes are more immunogenic than parenchymal cells – at least for the short term. Lechler and Batchelor (1982b) provided a key piece of evidence bringing DCs to the forefront as the passenger leukocyte prototype. They showed that the immunogenicity of longsurviving ‘parked’ renal allografts retransplanted into secondary recipients could be restored by the injection of donor strain DC (Lechler and

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Batchelor, 1982b), but not T or B lymphocytes. These findings were consistent with observations about the potent stimulatory capacity of DCs originally made by Steinman et al. (Steinman and Cohn, 1973, 1974; Steinman et al., 1974) and later confirmed by many others. Thus, DCs became known as stimulators of acute rejection, which led to the suggestion that there are at least two ways for the recipient to ‘recognize’ or become sensitized to donor alloantigens: ‘direct’ presentation by intact donor DCs and ‘indirect’ presentation of donor allopeptides processed by recipient DCs (Lechler and Batchelor, 1982a). Both of these pathways have the potential to activate recipient lymphocytes after organ transplantation. Reactions occur where donor and recipient hematolymphoid cells are mixed together: in the allograft (peripheral sensitization) and in recipient lymphoid tissues (central sensitization) (Demetris et al., 1991). In the allograft, recipient lymphocytes–donor DC clusters appearing within allografts several days after transplantation, is given as morphologic evidence for direct presentation (Forbes et al., 1986; Demetris et al., 1991). The distribution of recipient mononuclear cells during early acute rejection reflects the intra-organ distribution of donor DCs. Donor passenger leukocytes including DCs also spontaneously leave the allograft, either hematogenously, or via intact efferent lymphatics that drain to donor regional lymph nodes transplanted en bloc with the allograft (Fung et al., 1989; Murase et al., 1991). Larsen et al. (1990) was the first to show that donor DCs from heart allografts migrate hematogenously to the recipient spleen. The number of donor cells appearing in recipient lymphoid tissues is much greater after liver than heart transplantation (Demetris et al., 1991) and the phenotypes are more varied. After liver transplantation donor class II B cells and DCs localize to the cortex of the lymph nodes and the marginal zone and peri-arterial lymphatic sheath of the spleen, where their appearance is associated with a proliferative response in the recipient lymphoid cells (Demetris et al., 1991, 1993).

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Direct alloantigen presentation is thought to precipitate acute rejection, which is an explosive or vigorous immune response, often associated with enhanced production of TH1-type cytokines (IL-2, γ-IFN, TNFα, GM-CSF, IL-3) and chemokines (Li, X.C. et al., 1998; Zhai et al., 1999). Several lines of evidence support the contention that acute rejection is dominated by direct allorecognition (Hornick et al., 1998; Suciu-Foca et al., 1998; reviewed in Shirwan, 1999). These include: (1) the high precursor frequency of T cells recognizing allogeneic MHC molecules directly; (2) marked amelioration or absence of acute rejection in allografts depleted of donor antigen presenting cells (APC) before transplantation; (3) enhancement of acute rejection using pretreatment of donors with agents that increase the number of mature organ-based donor myeloid DCs (Steptoe et al., 1997); and (4) ability of T-cell lines specific for direct recognition of allogeneic MHC molecules to induce acute rejection in immunocompromised hosts (Shirwan, 1999). Recent studies suggest that the explosive immune reaction early after transplantation might be less dependent on direct presentation than previously thought (Albert et al., 1998; Bedford et al., 1999; Inaba et al., 1998, 2000). These studies show that DCs efficiently acquire MHC antigens from dead or apoptotic allogeneic cells and effectively present them in an MHCrestricted fashion (i.e. indirect presentation) to syngeneic lymphocytes (Albert et al., 1998; Bedford et al., 1999; Inaba et al., 1998, 2000). Although a mixed lymphocyte response is assumed to offer convincing evidence of direct antigen presentation, depletion of responder DCs from the reaction mixture significantly diminishes, but does not completely eliminate its vigor (Bedford et al., 1999). Thus, a significant component of the intense immune reactivity seen in the allograft and recipient lymphoid tissues early after transplantation, and for which potent immunosuppression is required, might be related to the mass migration of donor hematolymphoid cells and their subsequent apoptosis and processing by recipient DCs. More research is needed to determine the relative

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contributions of direct and indirect antigen presentation to acute rejection. Any damage within the allograft, including injury from rejection (Meyer et al., 2000) or ischemia, enhances the rate of passenger leukocyte exchange between the recipient and the allograft. For example, recipient DCs and a variety of accessory cells (Penfield et al., 1999) migrate into the renal interstitium as a result of ischemic and mechanical injury sustained during the transplant operation (Penfield et al., 1999). Consequently, more recipient accessory cells and DCs are available to phagocytize and process donor antigen(s) from apoptotic cells and ‘indirectly’ present donor allopeptides locally (peripheral) or within the draining lymph nodes or spleen (central) (Penfield et al., 1999). Detailed studies of DC precursors have shown that immature liver DCs, in contrast to mature myeloid DCs (Morelli et al., 2000), cluster with IL-10 and IL-4-secreting mononuclear cells and stimulate IL-10, but not γ-IFN production (Khanna et al., 2000). The implication is that donor-derived immature DCs might cause a TH2-type ‘tolerogenic’ cytokine response, and at least partially explain the immune privilege of hepatic allografts (Khanna et al., 2000). Conversely, increasing the number of mature myeloid DCs in the donor organ before transplantation using FLt3 ligand treatment can augment the TH1 response and increase the vigor of acute rejection. The heightened reactivity can overcome spontaneous acceptance of liver allografts in experimental animals (Steptoe et al., 1997). Although a TH1 (rejection) versus TH2 (tolerance) paradigm in transplantation immunology is attractive for several reasons, it is an overly simplistic representation of how immune reactions correlate with conditions within the allografts (Strom et al., 1996). Before leaving a discussion of the allograft, it would be remiss not to mention that liver sinusoidal endothelial cells also function as APC, but these cells do not migrate from the allograft. Nevertheless, they constitutely express all of the molecules necessary for antigen presentation (CD54, CD80, CD86, MHC class I and class II and CD40) and can stimulate both CD4 and CD8 T

cells (Knolle and Gerken, 2000). Similar to immature DCs, resident liver sinusoidal endothelial cells can also activate naïve T cells, but they often do not mature into effector T cells and show a cytokine profile and a functional phenotype that is compatible with the induction of tolerance (Knolle and Gerken, 2000). After treatment with γ-IFN, microvascular endothelial cells from the heart can express CD40 and the CD40 Ligand (CD40L) interaction can induce the expression of the adhesion molecules VCAM-1 and E-selectin and the co-stimulatory molecule CD80, but not CD86 (Jollow et al., 1999). Since the expressed CD80 is functional in both allo-MLR assays and accessory-cell dependent mitogen proliferation assays, microvascular endothelial cells might also play an important role in initiating and maintaining allograft rejection (Jollow et al., 1999). Given the potent allostimulatory properties of DCs discussed above, it was reasonable to consider that removing or masking the allostimulatory antigens before transplantation might ameliorate or prevent acute rejection. This approach was first attempted using irradiation or monoclonal antibodies directed at generic surface antigens, such as leukocyte common antigen (CD45) (Goldberg et al., 1995), or MHC class II (Krzymanski et al., 1991; Pollak and Blanchard, 2000). Although these masking experiments prolonged allograft survival, the long-term benefits were disappointing, which may in part, be related to the possibly false assumption that acute rejection is dependent primarily on direct presentation. In addition, experiments with rat liver allografts unexpectedly showed the opposite effect: disabling the passenger leukocytes enhanced rejection. Studies showed that if a donor liver was irraditated before transplantation, it was now rejected, instead of being accepted (Sun et al., 1995; Bishop et al., 1996; Sharland et al., 1998). The implication was that activation-induced apoptosis might play an important role in allograft acceptance or so-called ‘tolerance’ induction. This experimental finding confirmed clinical observations made by Starzl (Starzl et al., 1963; Starzl, 1964) several decades earlier that allograft

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acceptance or tolerance was a result of activation-induced ‘clonal stripping’. The idea resurfaced and gained popularity when donor hematolymphoid cells (Starzl et al., 1992, 1993a; Starzl and Zinkernagel, 1998), particularly DCs (Demetris et al., 1992), were found to persist in long-term allograft survivors, some of whom were off all immunosuppression. Indeed, recent studies in mice show that active, co-stimulation-mediated or passive (removal of growth factors) apoptosis of alloreactive T cell is required for peripheral ‘tolerance’ to allografts (Li, Y. et al., 1999; Wells et al., 1999; Zheng et al., 1999). The importance of this concept was further underscored by showing that calcineurin-inhibiting drugs, like cyclosporine and possibly tacrolimus, can actually inhibit tolerance induction by blocking activationinduced apoptosis (Wells et al., 1999). This does not occur with the immunosuppressive drug, rapamycin, which blocks the proliferative component of IL-2 signaling, but does not inhibit priming for activation-induced cell death (Wells et al., 1999). Armed with a better understanding of DC surface molecules involved in functional activation of lymphocytes, researchers began to more specifically target co-stimulatory molecule interactions between DC and T lymphocytes, such as B7-1 (CD80)-CD28 and CD40-CD40L interactions (Table 32.1). The hope was to chaperon DC–lymphocyte interactions such that their rejection-provoking activities were minimized and tolerogenic properties maximized. The results of this line of experimentation showed that blocking co-stimulatory molecules with monoclonal antibodies or with gene transfection, could significantly diminish acute rejection and prolong graft survival (Table 32.1). Various attempts at controlling the donor DC–recipient lymphocyte interactions could be crudely thought of as a donor-specific T cell depleting/modulating reagent. Unfortunately, many of the allografts still suffered from chronic rejection, and thus, tolerance was not achieved (Table 32.1). This outcome was predictable because many of the ‘tolerated allografts’ were inflamed (Steurer et al., 1995; Azuma et al.,

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1996), or showed allograft vasculopathy (Steurer et al., 1995; Azuma et al., 1996; Russell, M.E., 1996; Chandraker et al., 1997), which are signs of chronic rejection. An interesting study by Lakkis et al. (2000) further illustrates the importance of DC migration to secondary lymphoid tissues (and perhaps the indirect presentation alluded to above) on allograft survival and tolerance induction. Mice ‘naturally’ lacking peripheral lymphoid tissue because of a gene mutation in NF-KB fail to reject cardiac allografts if they are also subjected to splenectomy before transplantation (Lakkis et al., 2000). Instead the allografts are immunologically ‘ignored’, but tolerance is not achieved (Lakkis et al., 2000). Finally, it is important to remember that passenger leukocyte populations contain hematopoietic stem cells, as evidence by their ability to reconstitute lethally irradiated experimental animal recipients (Murase et al., 1996) and to convert the blood type of human recipients (Collins et al., 1993). Thus, donor progenitor cells will have access to the recipient bone marrow, regional and distant lymph nodes and even to the thymic medulla, where they can engraft in small numbers.

Chronic rejection Chronic rejection can be broadly defined as a largely indolent, but progressive form of allograft injury characterized primarily by persistent, but patchy inflammation of the allograft, fibrointimal hyperplasia of arteries, interstitial fibrosis, and destruction and atrophy of parenchymal elements and organ-associated lymphoid tissue (Demetris et al., 1997). The term chronic implies a temporally prolonged course and in general, chronic rejection develops over a longer period of time compared to acute rejection. However, chronic rejection clearly develops in many cases from inadequately controlled acute rejection and in patients not compliant with immunosuppressive therapy. The discussion here will be limited to the role of DCs in chronic rejection; readers interested in a broader perspective of

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Summary of the recent explosion of interest in chaperoning dendritic cell–lymphocyte interactions in transplantation Method

Findings/Conclusions

Activation of allogeneic T cells involves B7-1 co-stimulatory molecules and can be inhibited using CTLA4-Ig, which blocks the interaction with CD28 on T lymphocytes.

Infusion of CTLA4-Ig with and without donor bone marrow in normal and IL-4-deficient mice.

CTLA4-Ig induces allograft acceptance and skews recipient response toward a TH2 profile (increased IL-10). However, the effects of co-stimulatory moleculeblockadearestillseeninIL-4-deficient mice, so generation of a TH2-type immune response is not obligatory for CTLA4Ig-induced graft acceptance. The protective effect of co-stimulatory molecule blockade is diminished by cyclosporine and augmented with donor antigen (hematopoietic cells). Chronic rejection is diminished, but not eliminated, with co-stimulatory molecule blockade.

Larsen, 1996

Mice

Blockage of DC–T lymphocyte interactions and T cell activation.

Simultaneous CTLA4-Ig and anti-CD40.

Simultaneous, but not independent, blockade of the CD28 and CD40 pathways effectively aborts T cell clonal expansion in vitro and in vivo; promotes long-term survival of fully allogeneic skin grafts; and inhibits the development of chronic vascular rejection of primarily vascularized cardiac allografts.

Pearson, 1997 Zheng, 1997

Mice

Dissect individual components (B7-1, B7-1a, B7-1cyt II, and B7-2 molecules on APC and CD28 and CTLA4 molecules on T cells.

anti-B7-1 and/or anti-B7-2 or CTLA4-Ig.

Anti-B7-1 and anti-B7-2 modestly prolong graft survival; increased survival was obtained with either a combination of anti-B7-1 and anti-B7-2 or CTLA4-Ig. Treatment with CTLA4Ig induces long term cardiac allograft survival in B7-1(/) recipients regardless of donor status; anti-B7-2 mAb leads to indefinite allograft survival only when the recipient and donor both lack B7-1. Thus, both B7-1 and B7-2 have an important role in allograft rejection in the mouse vascularized cardiac allograft model.

Lu, 1999a O’Rourke, 2000 Takayama, 2000

Mice

Myeloid DC transduction with CTLA4Ig, which blocks interaction of CD80 and CD86 on DC with CD28 on T cells.

Transfection of myeloid DC with CTLA4-Ig.

CTLA4Ig-DC show impaired capacity to stimulate naïve allogeneic T cell proliferation and cytotoxic T lymphocyte (CTL) generation (Lu et al. 1999) and is reversed by exogenous IL-2 (Takayama, 2000). Altered DC induce donor-specific inhibition of alloimmune reactivity, inhibition of interferon (IFN)-γ production, augmentation of IL-4 and IL-10 secretion (Takayama et al. 2000) and prolong islet allograft survival (O’Rourke et al. 2000).

Gao, 1999 Lu, 1997a

Mice

CD40 ligation on APC by CD40L induces CD80 (B7-1) and CD86 (B7-2).

TGFβ-cultured DCs  antiCD40L mAb or DC after longterm mixed leukocyte cultures.

Inhibition of CD40–CD40L pathway appears important in regulation of allogeneic DC–T-cell functional interaction in vivo; its blockade increases markedly the potential of co-stimulatory moleculedeficient DCs of donor origin to induce long-lasting allograft survival.

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Intention/Targets

Mice/Rats

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Species

Chandraker, 1997 Lakkis, 1997 Larsen, 1994 Larsen, 1992 Lin, 1997 Pearson, 1996 Pearson, 1994 Russell, M.E., 1996 Steurer, 1995

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TABLE 32.1

Pretreatment facilitates a tolerogenic cascade similar to that in spontaneously tolerant liver recipients that involves: (1) chimerism-driven immune activation, succeeded by deletion of host immune responder cells by apoptosis in the spleen and allograft that is linked to interleukin-2 deficiency; and (2) persistence of comparatively large numbers of donor-derived leukocytes.

Lu, 1997b

Mice

Dissect individual components B7 (CTLA4) and FasL stimulatory pathways on DC.

CTLA4-Ig treatment of normal and FasL-deficient (B6.gld) DC.

CTLA4Ig-treated myeloid DCs fail to stimulate primary MLR and induce increased apoptosis of alloactivated T cells. CTLA4Ig treatment also increased the level of DNA fragmentation induced by FasL-deficient DCs. Co-stimulatory (B7-CD28) and T cell death-inducing pathways may play important counter-regulatory roles in dictating the outcome of (allogeneic) DC–T cell interactions.

Thomas, 1999

Macaque monkeys

NF-κB inhibition with delay in DC maturation.

DSG and anti-CD3 immunotoxin.

Regimen induced ‘tolerance’ and a polarized TH2-type response in plasma cytokines. In DSG recipients, mature DCs were significantly reduced in day 5 lymph node biopsies, with complete repopulation by 30 days.

Min, 2000

Mice

Increasing ability of DC to effect activation-induced apoptosis using FasL.

Transfecting of BM-derived DC with Fas ligand (FasL).

Markedly augmented DC capacity to induce apoptosis of FAS cells, which inhibited allogeneic MLR in vitro, induced hyporesponsiveness to alloantigen in vivo and significantly prolonged the survival of fully MHC-mismatched vascularized cardiac allografts.

Morelli, 2000

Mice

Increasing the number of mature liver DC.

Flt3 ligand (FL) administration.

FL treatment of donor livers increases the numbers of myeloid(CD8α)andlymphoid(CD8α) DCs,which induce strong TH1 and Tc1 responses and lead to rejection of liver allografts that are normally accepted.

Giannoukakis, 2000

Mice

Inhibition NF-κB-dependent transcription of co-stimulatory genes with delay in DC maturation.

Decoy ODN containing binding sites for NF-κB.

Prolongation of heart allograft survival.

Li, X.C., 1999 Li, Y., 1999 Wells, 1999 Zheng, 1999

Mice

Co-stimulation-mediated activation-inducted deletion may be required for tolerance induction.

Combination of co-stimulatory molecule blockade and administration of donor hematopoietic cells and cyclosporine or rapamycin.

Combined pretransplant DST with anti-CD154 mAb treatment may be attractive for clinical deployment, and strategies aimed to selectively block CD28 without interrupting CTLA4/B7 interaction might prove highly effective in the induction of tolerance. Deletion of activated T cells through activation-induced cell death or growth factor withdrawal seems necessary to achieve peripheral tolerance across major histocompatibility complex barriers.

ALS, anti-lymphocyte; DSG, deoxyspergualin; ODN, double-stranded oligodeoxyribonucleotides; DST = donor specific transfusion.

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Immature donor DC  antiCD40L mAb infusion before transplantation.

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Inhibition of CD40/CD80/CD86 upregulation on immature DC.

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chronic rejection are referred to recent reviews (Demetris et al., 2000; Womer et al., 2000). Infiltration of the allograft by recipient accessory cells, including macrophages and DCs, is a hallmark feature of chronic rejection (Demetris et al., 1997, 2000; Murase et al., 1999). Increased transcripts for TH1 cytokines, such as TNF-α, GM-CSF and IFN-γ and chemokines suggest that strong TH1 activation during severe acute rejection, which causes an influx of monocytes/macrophages and DC precursors, might explain the tendency for severe acute rejection to evolve into chronic rejection (reviewed in Demetris et al., 2000). Oguma et al. (1988) first showed recipient S100 DC infiltrated arteries affected by obliterative arteriopathy and the interstitium of chronically rejecting renal allografts. Subsequent studies verified these findings and showed that the number of recipient DCs in chronically rejecting organs directly correlates with the overall severity of inflammation (Demetris et al., 1997; Kuffova et al., 1999; Leonard et al., 2000). Moreover, recipient DCs are concentrated amidst, and apparently organize the lymphoid aggregates typical of chronic rejection (Oguma et al., 1988; Demetris et al., 1997). The implication is that the recipient DCs coordinate indirect alloantigen presentation, both locally in the allograft and after migration to recipient lymphoid tissues (Bradley, 1996). Reasonably strong evidence supports a role for indirect presentation (and recipient DCs) in chronic rejection (Bradley, 1996; Vella et al., 1997; Ciubotariu et al., 1998; reviewed in Shirwan, 1999). Included is: (1) ongoing immunologic injury in the allograft, despite disappearance of donor hematolymphoid cells (Demetris et al., 1997); (2) influx of activated recipient macrophages and DCs into chronically rejecting organs (Demetris et al., 1997); (3) an important role of alloantibodies, mediated by B cells serving as APCs for CD4 T cells generating these antibodies; (4) a high incidence of CD4 Tcell responses to donor MHC allopeptides via indirect recognition in patients with chronic rejection (Vella et al., 1997; Ciubotariu et al., 1998); and (5) evidence that allografts depleted of passenger leukocyte (e.g. donor irradiation

and reconstitution with recipient hematopoietic cells, epidermal and corneal allografts) are still chronically rejected (Sayegh and Carpenter, 1996; Krasinskas et al., 2000). Indeed, allografts that show persistent acute rejection (Vella et al., 1997; Ciubotariu et al., 1998; Suciu-Foca et al., 1998) and those that evolve toward chronic rejection (Vella et al., 1997; Ciubotariu et al., 1998), show evidence of increased indirect and diminished direct alloantigen presentation (Hornick et al., 1998). Interestingly, inflammatory infiltrates in chronic rejection are often arranged into nodular aggregates, some of which contain germinal centers (Demetris et al., 2000). This arrangement is reminiscent of the development of secondary lymphoid tissue outside of organs, discussed above, and in some autoimmune disorders (Zinkernagel et al., 1997), reinforcing the notion that chronic antigenic stimulation occurring outside the lymph nodes can result in the development of intra-organ lymphoid tissue (Zinkernagel et al., 1997). It is tempting to speculate that coalescence in time and space of DC antigen acquisition, maturation and presentation within the organ occurs because the intraorgan immune network is disrupted (see below).

PROBLEMS WITH REESTABLISHMENT OF NORMAL PHYSIOLOGY (ORGAN-BASED IMMUNE NETWORK) IN ALLOGRAFTS Organ-based immune networks are naturally present in all vascularized organs, and exaggerated in organs that are in direct contact with the external environment. Cells comprising these networks are dependent on the bone marrow progenitors and to a lesser extent, from local progenitors or stem cells, for replenishment. Thus, replacement by recipient cells after transplantation is an expected finding (Porter, 1969; Iwaki et al., 1991; Murase et al., 1991). We have just discussed one reason why this does not occur smoothly (see Acute Rejection). In addition, donororganharvestingandreimplantationphys-

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ically disrupts the efferent lymphatic channels. This blocks an important emigration route for either donor or recipient DCs and lymphocytes that traverse the allograft interstitium, until connections with regional lymph nodes are reestablished 2–3 weeks after transplantation (Malek andVrubel, 1968; Kline andThomas, 1976; Cuttino et al., 1978). In fact, efferent lymphatic disruption contributes to a ‘re-implantation’ response, which alone can precipitate activation oftheintra-organimmunenetwork(Cuttinoetal., 1978; Kline and Thomas, 1976). Once recipient T cells are activated within the allograft and rejection effector mechanisms begin to damage the organ, further problems arise. Rejection disrupts lymphatic and capillary microvascular endothelial junctions and increases lymph fluid production, which retards some immune cell traffic and lymph flow from the allograft (Malek and Vrubel, 1968; Kline and Thomas, 1976; Cuttino et al., 1978). Thus, DC precursors that receive maturation signals from the ongoing damage, may not have access to migration routes out of the organ, differentiate in situ, and stimulate a local inflammatory focus (Malek and Vrubel, 1968; Ruggiero et al., 1994). Unless rejection is interrupted by increased immunosuppression, an endless cycle of immune activation and damage develops. The problem becomes even worse in organs with mucosal-associated lymphyoid tissue (Murase et al., 1990, 1991). A large number of donor and recipient lymphocytes and DCs participate in a ‘bidirectional in vivo mixed lymphocyte response’, which occurs both in the allograft and in draining lymph nodes (Fung et al., 1989; Murase et al., 1990, 1991). An orderly transition from donor to recipient cells in the intra-organ immune network is dependent on preventing architectural damage during the changeover (Murase et al., 1991). This usually requires potent immunosuppression, which may not be conducive to tolerance induction (see Acute Rejection). However, inadequate control of immune reactivity during the transitional phase has the potential to disrupt the normal architecture, both within the organ and secondary lymphoid tissues (Kline and Thomas,

1976; Murase et al., 1990, 1991; Ruggiero et al., 1994; Demetris et al., 1997). Once irreversible structural damage has occurred, it may be impossible to re-establish a normally functioning intra-organ immune network and immune architecture of regional lymph nodes or organassociated lymphoid tissues. Consequently, an important aspect of ‘nonlocal’ organ physiology: efficient and effective immunologic response to environmental antigenic challenges, becomes compromised (Steinman et al., 1997; Zinkernagel et al., 1997; Starzl and Zinkernagel, 1998). The separation in time and space of the two important DC functions (antigen acquisition and presentation to lymphocytes) is no longer separated in space. Therefore, microbial and other local antigenic challenges, may be ineffectively cleared and persist in the organ. The local inflammatory milieu damages the allograft and enhances allorecognition and injury. The end result is a downward spiral of declining organ function, continued problems clearing local antigens/ infections and persistent inflammation. Eventually, the allograft fails from chronic rejection. Indeed, it is tempting to speculate that failure to re-establishtheintra-organandorgan-associated lymphoid tissues is related to a lack of robust tolerance and accounts for the frequent association between infection and chronic rejection (Whitehead et al., 1993; Heemann et al., 1996; Siddiqui et al., 1996).

TOLERANCE INDUCTION Solid organ transplantation is a clinically successful short-term solution to end-stage organ disease. The major obstacle to greater long-term success is chronic rejection and consequently, the need for morbidity-producing long-term immunosuppression. Simply stated, the major problem is a lack of robust or true tolerance. Two sources of donor reactive T cells must be eliminated or effectively controlled to achieve robust tolerance: (1) mature T cells in the periphery; and (2) newly developing T cells that arise from bone marrow precursors. From the above

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discussion, we have seen that donor DCs can be manipulated into effective short-term donorspecific T cell-depleting reagents, by enhancing activation-induced T cell apoptosis in the periphery. This might remove many or even most pre-existing donor reactive mature T cells, but alone, mature T-cell depletion does not bring about clinically acceptable drug-free allograft acceptance in most cases. Lakkis et al. (2000) have shown that even when all secondary lymphoid tissue is absent, recipients are not robustly tolerant, the allografts are simply ‘ignored’. Given the ‘antigenically rich’ environment of outbred humans, the relatively short lifespan of DCs in the periphery, and their pivotal role in immune responses, it is unlikely that manipulation of either donor or recipient DCs alone, will prove to be a practical or effective strategy for robust tolerance induction. On a practical basis, very few human allografts are continually ‘ignored’; the majority of recipients still require life-long immunosuppression to prevent chronic rejection (Demetris, 2000). This is also true for human liver allograft recipients, who are relatively resistant to chronic rejection (Demetris, 2000). Even under the best of circumstances, this ‘favorable’ population still requires chronic immunosuppression in 80% of long-term survivors (Mazariegos et al., 1997). Attempts to achieve allogeneic tolerance without engineering an integration of the donor immune system (Coutinho, 1989) or stopping development of new donor reactive T cells from precursors, will likely fail to achieve robust tolerance. Nevertheless, at a practical level, we have learned that passenger leukocyte (especially DC) migration, and activation-induced clonal deletion and graft adaptation (replacement of the intra-organ immune network) have made clinical solid organ transplantation the success that it is today (Starzl, 1964). The discipline is based on the premise that allograft must be protected by potent immunosuppression from the vigor of the early immunologic storm caused by the bi-directional leukocyte migration (and activation). Several months thereafter, it is possible to significantly reduce these potentially lethal levels of immunosuppression because many of

the donor reactive T cells have been eliminated. This enables recipients to survive long term on lower levels of immunosuppression. However, the field of transplantation, and especially patients, are eager for even better results. The ideal outcome is long-term survival with a functioning allograft and no immunosuppression, which can be achieved in only a few human (mostly liver) allograft recipients (Starzl et al., 1993a, b, and c; Mazariegos et al., 1997). Although these few patients might be simply considered ‘lucky’, a surprising finding suggested a more hopeful explanation (Starzl et al., 1992, 1993a). At the request of a journal editor, longsurviving experimental animal (and then human) organ allograft recipients were examined to determine if donor hematolymphoid cells persisted in these survivors, some of whom were chronically free of immunosuppressive therapy (Demetris et al., 1992; Ricordi et al., 1992; Starzl et al., 1992). Very rare donor cells were scattered throughout the lymphoid and nonlymphoid tissues of the recipient, and some had the characteristics of DCs (Demetris et al., 1992; Ricordi et al., 1992). This finding was contrary to common assumptions at the time, that DCs were only powerfully immunogenic and responsible for direct alloantigen presentation, neither of which may be correct. Characteristics of the cells identified included strong expression of MHC class II antigens and a dendritic-shape, as well as localization to the interstitium of organs, paracortex of lymph nodes, thymic medulla (Demetris et al., 1992, 1993; Lu et al., 1995), and the periarterial lymphatic sheath of the spleen – sites where DCs normally reside. More detailed studies by Lu and Thomson (Lu et al., 1994; Thomson et al., 1995) and others, conclusively showed that DCs and their precursors were included among the multilineage passenger leukocytes persisting in successful organ allograft recipients. The suggestion was made that ‘microchimerism’ or the persistence of even trace numbers of donor hematopoietic cells was necessary, but alone, not sufficient for the induction of tolerance (Starzl et al., 1992, 1993a, 1993b, 1993c). Since donor cells persisted for

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decades in some patients, the implication was that donor stem cells carried with the organ had engrafted after transplantation (Rugeles et al., 1997; Nierhoff et al., 2000). The important conceptual point was that given their location and phenotype, the donor hematolymphoid cells appeared to have successfully integrated into the recipient immune system, similar to the mixed chimeras made by irradiation in experimental animals (see below). However, donor cells are rare in microchimeras (immunohistochemistry and PCR are needed for detection), whereas they are easily detectable by flow cytometry in mixed chimeras, and this likely affects the contribution of various tolerance inducing mechanisms (Figure 32.1). For example, it is unlikely that the rare DCs in microchimeras that make their way to the recipient thymic medulla would mediate enough negative selection to remove all donor reactive T cells. Consequently, there would be a greater dependence on peripheral mechanisms, which are less robust. Subsequent studies confirmed that passenger leukocytes persisted in experimental animals and in humans, but other investigations did not

FIGURE 32.1 Diagram illustrating the relative contributions of various tolerance mechanisms according to the amount of hematopoietic chimerism. In neonatal or mixed chimeras produced by preconditioning, donor cells usually comprise 1% or more of all hematolymphoid cells and central tolerance mechanisms predominate. In contrast, donor cells comprise <0.05% in microchimera and peripheral mechanisms predominate.

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necessarily agree with the interpretation that they played a role in tolerance induction (Caillat-Zucman et al., 1994; Schlitt et al., 1994; Suberbielle et al., 1994; Burlingham et al., 1995; Bushell et al., 1995; Sivasai et al., 1997). Individual patients were found that had apparently accepted solid organ allografts, but were not microchimeric, and conversely, other patients were microchimeric, but rejected their allografts when immunosuppressive coverage was inadequate (Sivasai et al., 1997). Both of these situations are easily understandable within the framework of microchimerism. First, detection of microchimeric cells in the peripheral blood is episodic: a single measurement at one point in time is of questionable value (Elwood et al., 1997). For example, CD34 donor cells can be detected in the bone marrow of liver allograft recipients, but the presence of these cells does not correlate with the chimeric status in peripheral blood (Nierhoff et al., 2000). In addition, it is unlikely that all donor reactive T cells are purged from the recipient T cell repertoire after solid organ transplantation, as mentioned above. Conversely, ‘ignorance’ can resemble tolerance, but it is not robust and therefore, is usually only a temporary situation of the allograft. Since the advent of transplantation immunobiology, tolerance achieved by hematopoietic chimerism has been widely recognized as the ideal and most robust form of tolerance (recent comprehensive reviews see Exner et al., 1999; George et al., 1999; Wekerle and Sykes, 1999). Owen (1945) was the first to show that nonidentical twin cattle sharing a placental circulation develop chimeric immune systems, each composed of hematopoietic cells from both individuals (Owen, 1945). Shortly thereafter, it was shown that these chimeric twins were tolerant to skin grafts from their twin siblings (Anderson, D. et al., 1951; Billingham, R.E. et al., 1952). In the fetus or in immunodeficient recipients, engraftment of allogeneic hematopoietic cells can be achieved without immunosuppression (Roncarolo et al., 1988, 1991; Touraine et al., 1993). In mature immunocompetent individuals, some type of recipient conditioning is

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required. Typically, irradiation and/or cytoreductive chemotherapy is combined with T celldepleting antibodies and/or conventional pharmacologic immunosuppression. The intent is to provide a ‘clean slate’ (Wekerle and Sykes, 1999) for the newly developing T cell repertoire; to prevent rejection of the donor cells by mature recipient T cells; and facilitate long-term engraftment of the donor hematopoietic cells (Wekerle and Sykes, 1999). The ‘conditioned’ recipient is then treated with an infusion of donor bone marrow alone, or a combination of donor and recipient bone marrow (Main and Prehn, 1955; Monaco et al., 1966; Ildstad and Sachs, 1984; Ildstad et al., 1985; Sykes, 1996a and b; Sykes et al., 1997). Two types of chimerism are achieved with these protocols (Wekerle and Sykes, 1999), ‘full chimerism’, where the entire recipient hematolymphoid system is replaced by that of the donor; and ‘mixed chimerism’, where a functionally integrated immune system is composed of various proportions of hematolymphoid cells from the donor and recipient. A third type of chimerism called ‘microchimerism’ briefly mentioned above, is similar to mixed chimerism (Starzl et al., 1992, 1993a, 1993b, 1993c). It refers to trace populations of multilineage donor hematopoietic cells that persist in a fraction of stable, unconditioned, long-term vascularized allograft recipients treated only with conventional immunosuppressive regimens (Starzl et al., 1992, 1993a, 1993b). Although some investigators are quick to draw a clear distinction between microchimerism and mixed chimerism (Wood and Sachs, 1996; Wekerle and Sykes, 1999), in our opinion, the only difference is in the proportion of donor cells and in the relative contribution of different ‘tolerance-inducing’ mechanisms. The robust tolerance associated with mixed chimerism is strictly dependent on the persistence of donor cells (chimerism) (Sharabi et al., 1992; Khan et al., 1996). It is mediated primarily by central deletion (Ildstad and Sachs, 1984; Ildstad et al., 1985; Sykes, 1996a, 1996b), although peripheral mechanisms also likely contribute to the process (Thomas et al., 1987; Tomita et al., 1994), similar to mechanisms of

‘self’ tolerance (Anderson, G. et al., 1996; DeKoning et al., 1997; Lo et al., 1997). Specifically, successful mixed chimeras show clonal deletion (negative selection) of ‘selfreactive’ T cells in the thymus, which is mediated primarily by DCs (Tomita et al., 1994; Khan et al., 1996; Sykes, 1996a, 1996b). These chimeras lack donor responsiveness in a mixed lymphocyte reaction; accept donor-specific allograft(s) without immunosuppression, and are resistant to chronic rejection (Colson et al., 1995; Orloff et al., 1995). Instead of delivering manipulated DCs or DC precursors, which have a limited lifespan and a strong tendency to differentiate, engraftment of donor hematopoietic stem cells creates a continuous ‘factory’ of multilineage donor cells in the recipient. Donor cells can participate alongside recipient cells in a fully-functional, integrated immune system. Donor DCs are able to migrate into the recipient thymic medulla and mediate negative selection of T cells potentially reactive against the allograft. Donor precursor cells can also mature into lymphoid or myeloid DCs and migrate into peripheral lymph nodes and into the interstitium of organs where they can mediate immunogenic or tolerogenic reactions, discussed above. Unfortunately, the mixed chimera approach to tolerance induction, which is so successful in small experimental animals, has not gained widespread clinical acceptance. The major drawback is the significant morbidity and mortality associated with the recipient conditioning (Sykes, 1996a, 1996b; Jankowski and Ildstad, 1997) needed to ensure engraftment of the donor hematopoietic cells. The subsequent threat of graft-versus-host disease is another serious consideration, particularly when major MHC barriers are crossed, as is currently the practice in solid organ transplantation (Billingham, R. and Brent, 1956; Russell, P.S. 1962). Efforts are being made therefore, to modify the protocols to reduce morbidity-producing conditioning, without compromising, or even enhancing, donor stem cell engraftment. These modifications include lower doses of irradiation; the use of CD3, class IIdim/inter, CD45, CD45R,

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CD8, but αβ-TCR and γδ-TCR negative hematolymphoid facilitator cells (Kaufman et al., 1994; Gandy et al., 1999) and osteoblast facilitator cells (El-Badri et al., 1998); higher doses or repeated infusions of donor bone marrow (Ricordi et al., 1995; Jankowski and Ildstad, 1997; Sykes et al., 1997); co-stimulatory molecule blockades (Wekerle and Sykes, 1999); depleting anti-CD4 and anti-CD8 monoclonal antibodies, local thymic irradiation, high-dose bone marrow cells (Sykes et al., 1997) and the blocking recipient NK cell activity (Kawai et al., 2000). The crucial test for tolerance-inducing regimens is their ability to lessen or eliminate the need for immunosuppression, while preventing or at least ameliorating chronic rejection in subhuman primates and clinical trials. In monkeys, tolerance induction using a combination of total body irradiation, thymic irradiation, antithymocyte globulin, donor bone marrow transplantation, and a 4-week course of cyclosporine showed that splenectomy may also be required to induce B cell tolerance and prevent the production of alloantibodies (Kawai et al., 1999). A patient with multiple myeloma and renal failure was recently treated with simultaneous kidney and bone marrow transplantation from an HLA-identical sibling (Spitzer, 1999). Using cyclophosphamide, antithymocyte globulin, thymic irradiation and cyclosporin given for 73 days after transplantation, mixed chimerism was easily detectable in this recipient until 105 days after transplantation, but thereafter hematolymphoid chimerism was nondetectable, or fell into the microchimeric range. Nevertheless, the patient was weaned from all immunosuppression with normal renal allograft function, 14 weeks after discontinuance of the cyclosporine (Spitzer et al., 1999). Several centers are currently evaluating the efficacy of clinical protocols in which vascularized organ allograft recipients are given an infusion of unfractionated donor bone marrow at the time of transplantation (Barber et al., 1991; Shapiro et al., 1995; Garcia-Morales et al., 1998). In some studies, additional donor bone marrow infusions are given after transplantation

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(Garcia-Morales et al., 1998). Usually, no specific conditioning therapy is used, and the patients are treated with conventional immunosuppressive agents. The intent is to enhance donor hematopoietic stem cell engraftment. An additional side-effect could be that the DCs in the infusion might act as a ‘donor-specific’ T cell-depleting reagent, via activation-induced apoptosis. If the number of engrafted donor stem cells is increased and the number of donor reactive T cells decreased, perhaps progress will be made towards a more robust form of tolerance needed in clinical transplantation. Results so far show that these treatment protocols are relatively safe and can enhance chimerism, albeit modestly (approximately 1% at 2 years) in most recipients (Barber et al., 1991; Shapiro et al., 1995; Garcia-Morales et al., 1998). A trend has been noted toward donorspecific hypo- or unresponsiveness (Barber et al., 1991; Shapiro et al., 1995; Garcia-Morales et al., 1998) and perhaps a lower incidence of chronic rejection (Barber et al., 1991). However, there are no reports of weaning trials in these patients, or a decreased need for immunosuppression. Perhaps it is still too early to tell, but given the low level of chimerism in most of the treated recipients, it is likely that more modifications to enhance engraftment will be needed.

WHAT HAVE WE LEARNED? What then has been learned about the immune system, MHC antigens and role of DCs in the experiment of transplantation? First, it is abundantly clear that the immune system has ‘local’ properties, which greatly complicate attempts to transfer organs between unrelated people. Transplantation has also taught us that DCs play a pivotal role in causing the immune complications in transplantation immunobiology,preciselybecausetheyhaveanimportantrole in defining the local properties of immune systems. But herein also lies the solution. DCs mediate negative selection of the developing T cell repertoire in the thymic medulla (Brocker et al., 1997; Delaney et al., 1998), probably via direct

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presentation of their strongly expressed MHC antigens and whatever peptides they might carry. Thus, donor lymphoid DCs in the thymic medulla have the potential to ‘redefine the immunologic self’; the same is true in the periphery. However, given the complexities of DC antigen acquisition, maturation, migration and presentation, it is probably easier to strive for donor hematopoietic stem cell engraftment and let ‘nature take its course’, than to try to control or chaperon these processes for tolerance induction. Efficient functioning of chimeric immune system (Roncarolo et al., 1988; Touraine et al., 1993) suggests that the ‘immunologic self’ is not completely ‘hard-wired’ in our genes; although it is likely that certain combinations of MHC antigens may never be compatible in a chimeric immune system. Consequently, the local properties of the immune system and MHC antigens were probably not designed for the purpose of preventing allogeneic transplantation, but to ensure a diversity of immunological responsiveness in individuals and thus adaptability at a species level (Hughes and Yeager, 1998). The key to understanding and thus controlling the immune system for the purpose of transplantation, will come from an appreciation of both its local and nonlocal properties so that the latter can be exploited. Better long-term success of organ transplantation will require either a selective blockade of the generation of new donor reactive T cells, or successful co-transplantation and assimilation of the donor immune system, which induces tolerance ‘naturally’.

ACKNOWLEDGEMENTS This work was supported by NIH 1 R01 DK49615-01, NIH RO1 AI40329-02, NIH RO1 AI 38899, NIH RO1 DK54232.

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Pearson, T.C., Alexander, D.Z., Corbascio, M. et al. (1997). Transplantation 63, 1463–1469. Penfield, J.G., Wang, Y., Li, S. et al. (1999). Kidney Int. 56, 1759–1769. Pollak, R. and Blanchard, J.M. (2000). Transplantation 69, 2432–2439. Porter, K.A. (1969). In: Starzl, T.E. (ed.) Experience in Hepatic Transplantation, Philadelphia: W.B. Saunders, pp. 422–471. Prickett, T.C.R., McKenzie, J.L. and Hart, D.N.J. (1988). Transplantation 46, 754–761. Ricordi, C., Ildstad, S.T., Demetris, A.J., Abou el-Ezz, A.Y., Murase, N. and Starzl, T.E. (1992). Lancet 339, 1610–1611. Ricordi, C., Karatzas, T., Selvaggi, G. et al. (1995). Ann. N.Y. Acad. Sci. 770, 345–350. Roncarolo, M.G., Yssel, H., Touraine, J.L. et al. (1988). J. Exp. Med. 168, 2139–2152. Roncarolo, M.G., Bacchetta, R., Bigler, M., Touraine, J.L., de Vries, J.E. and Spits, H. (1991). Blood Cells 17, 391–402. Rugeles, M.T., Aitouche, A., Zeevi, A. et al. (1997). Transplantation 64, 735–741. Ruggiero, R., Fietsam, R. Jr., Thomas, G.A. et al. (1994). J. Thorac. Cardiovasc. Surg. 108, 253–258. Russell, M.E., Hancock, W.W., Akalin, E. et al. (1996). J. Clin. Invest. 97, 833–838. Russell, P.S. (1962). Transplantation: Ciba Foundation Symposium (eds Wolstenholme, C.E.W., Cameron, M.P., London, J. Churchill, A.), Boston: Little Brown and Co. Sayegh, M.H. and Carpenter, C.B. (1996). Int. Rev. Immunol. 13, 221–229. Schlitt, H.J., Hundrieser, J., Ringe, B. and Pichlmayr, R. (1994). N. Engl. J. Med. 330, 646–647. Seki, S., Habu, Y., Kawamura, T. et al. (2000). Immunol. Rev. 174, 35–46. Shapiro, R., Rao, A.S., Fontes, P. et al. (1995). Transplantation 60, 1421–1425. Sharabi, Y., Abraham, V.S., Sykes, M. and Sachs, D.H. (1992). Bone Marrow Transplant 9, 191–197. Sharland, A., Shastry, S., Wang, C. et al. (1998). Transplantation 65, 1370–1377. Shirwan, H. (1999). Transplantation 68, 715–726. Siddiqui, M.T., Garrity, E.R. and Husain, A.N. (1996). Hum. Pathol. 27, 714–719. Sivasai, K.S., Alevy, Y.G., Duffy, B.F. et al. (1997). Transplantation 64, 427–432. Snell, G.D. (1957). Ann. Rev. Microbiol. 11, 439–458. Spitzer, T.R., Delmonico, F., Tolkoff-Rubin, N. et al. (1999). Transplantation 68, 480–484. Starzl, T.E. (1964). Host–Graft Adaptation. Experience in Renal Transplantation. Philadelphia, PA: W.B. Saunders Company, pp. 164–170. Starzl, T.E. and Zinkernagel, R.M. (1998). N. Engl. J. Med. 339, 1905–1913.

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33 Dendritic cells and autoimmunity Ranjeny Thomas Centre for Immunology and Cancer Research, University of Queensland, Princess Alexandra Hospital,Brisbane, Australia

Neither a borrower nor a lender be: For a loan oft loses both itself and friend; And borrowing dulls the edge of husbandry. This above all, – to thine own self be true . . . Hamlet William Shakespeare

INTRODUCTION

It has been proposed that dendritic cells (DCs) are the critical decision-making cells in the immune system (Fazekas de St Groth, 1998). Through their role in the generation of central and peripheral tolerance, as well as in priming immune responses and stimulation of memory and effector T cells, DCs are likely to play essential roles in the initiation and perpetuation of autoimmunity and autoimmune diseases. This chapter will examine the evidence for the hypothesis that DCs are upregulated by nonspecific and inflammatory stimuli that can lead to presentation of processed and/or endogenous self-antigens in genetically susceptible individuals or strains. After priming by DCs, a number of factors may be involved in establishing autoimmunity and perpetuation of disease pathology. These may vary between organ sites and diseases.

Autoimmunity results from a breakdown in the regulation in the immune system resulting in an inflammatory response directed at self-antigens and tissues. If ongoing, such a response leads to tissue destruction and its resultant detrimental effects. The pathological damage can be in one or several organs, or generalized in various tissues in the body. Organ-specific autoimmune diseases include the endocrinopathies, such as insulin-dependent diabetes mellitus (IDDM), and Hashimoto’s thyroiditis. Generalized autoimmune disease is best exemplified by systemic lupus erythematosus (SLE). Between these extremes is a spectrum, containing a number of diseases with varying components of organ specificity. For example, autoimmune arthritides such as rheumatoid arthritis (RA) have joint-specific and systemic features (Roitt et al., 1992).

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DENDRITIC CELLS AND THE BALANCE BETWEEN TOLERANCE AND AUTOIMMUNITY Loss of tolerance to self-antigens is necessary for generation of autoimmunity. Tolerance is actively maintained centrally in the thymus (Ardavin, 1997). Here, T cells reactive to self-antigen presented by medullary DCs are deleted by negative selection above a threshold of affinity for the antigen (Kappler et al., 1987). In the periphery, tolerance may be active, through deletion or regulation of self-reactive T cells, or passive, due to ignorance of certain self-antigens. Peripheral self-reactive T cells may be controlled in a number of ways. Deletion of self-reactive cells has been shown to occur in lymph nodes draining uninflamed peripheral organs and tissues (Lo et al., 1991; Kurts et al., 1997). In other circumstances, T-cell receptor (TCR) signaling may lead to functional unresponsiveness or anergy (Rocha et al., 1993). However, self-antigens may also be ignored because of lack of presentation in the thymus, or lack of recognition because of presentation below threshold levels of TCR affinity (Hogquist et al., 1994; Heath et al., 1998; Anderson et al., 2000). In the periphery, such antigens may remain sequestered from presentation by DCs, so that potentially autoreactive T cells are not actively regulated (Barker et al., 1977). Recently it has also been proposed that some cases of ignorance may result from presentation of different epitopes of the same MHC class I-restricted self-antigen by DCs in the thymus and non-APCs in the periphery. While a number of mechanisms may overcome or break tolerance, each one is unlikely to lead to overt autoimmune disease on its own. These mechanisms relate to priming of autoreactive T cells, precursor frequency of autoreactive T cells, peripheral regulation of autoreactivity and genetic predisposition. Because of the presumed lack of T-cell regulation, a productive autoimmune response is most readily primed by activated DCs to ignored self-antigens. Priming may lead subsequently to a systemic response. Alternatively, priming may

occur in lymph nodes draining an organ, such as the pancreas, with subsequent perpetuation of organ-specific autoimmunity. The precursor frequency of self-reactive T cells may be very high in certain pre-autoimmune conditions and in self-reactive TCR transgenic models (Waase et al., 1996; Durinovic-Bello, 1998; Anderson et al., 2000). In some cases, genetic mutations may lead to ineffectual mechanisms of self-reactive T-cell deletion or regulation. For example, mutations in Fas or Fas-ligand in both mice and humans lead to systemic autoimmunity as observed in lpr and gld mice, and patients with autoimmune lymphoproliferative syndrome (ALPS) (Jackson and Puck, 1999). In other cases, boosting the precursor frequency of self-reactive T cells, as in TCR transgenic animals, can precipitate overt autoimmune disease (Kouskoff et al., 1996). These data raise the importance of T-cell regulation for prevention of self-reactivity. Genetic background is an important predisposition to autoimmunity. In some animal models, specific mutations lead to a highly likely outcome of auto-immune disease. Mutations in the same human genes translate to a small number of cases restricted to certain families. Examples include the ALPS syndrome due to mutations in Fas, Fas-ligand or caspase 10 (Jackson and Puck, 1999; Wang et al., 1999). However, in many cases, known susceptibility relates to the major histocompatability complex (MHC) genes, as well as other genes that may alter the outcome of immune responses that are normally tolerated (McDevitt, 1998; Ridgway and Fathman, 1999). Such genes are being discovered, in large-scale genome wide screening of multiplex families in a number of auto-immune diseases, and in screening of genetic susceptibility to autoimmunity in a number of animal models (Griffiths et al., 1999). There is evidence that mutations in the coding region or promoter sequences of various cytokine genes and receptors may predispose at least some strains and individuals to autoimmunity (Abraham et al., 1999). MHC genes not only restrict the antigenic epitopes presented, but also shape the repertoire of T cells available through the process of selection. Most autoimmune diseases

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are now recognized to be polygenic, and in most cases this relates to the requirement for a number of features of immune regulation to be abnormal in a susceptible individual or strain before an initiating event triggers the development of overt pathology.

DENDRITIC CELL ACTIVATION AND PRIMING OF AUTOIMMUNE DISEASE Dendritic cells For the priming of immune responses it is clear that DCs must be functionally differentiated. Pro-inflammatory signals stimulate differentiation and migration of peripheral DCs via afferent lymphatics to draining lymph nodes, with associated upregulation of molecules required for efficient antigen presentation. Cytokines, including GM-CSF, TNFα and IL-1, patternrecognition molecules and phagocytosis itself may provide differentiating signals to immature DCs, and in some cases to peripheral monocytes (Medzhitov et al., 1997; Randolph et al., 1998; Thomas and Lipsky, 1996b; Corinti et al., 1999). Upregulated molecules include MHC molecules, co-stimulatory molecules, adhesion molecules and CCR7, which are important for the interaction of DCs with T cells. Furthermore, upregulation of CCR7 directs the appropriate migration of DCs to the T-cell area of the lymph node for the generation of antigen-specific T cells (Cyster, 1999). In addition to their role in T-cell priming, DCs have been shown to regulate B-cell growth, differentiation and immunoglobulin class switching, in a role that is separate to that played by follicular DCs (Dubois et al., 1997). Since this role is as yet ill-defined, this chapter will be confined to consideration of the role of DCs in stimulating auto-reactive T-cell responses.

Antigen and antigen persistence A number of autoimmune animal models are initiated by administration of self-antigen in

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combination with adjuvant (Sobel et al., 1984; Wooley et al., 1989). Administered antigen is taken up by resident DCs or Langerhans cells (LCs) and presented to naïve T cells bearing appropriate T-cell receptors in the draining lymph nodes. Recently, it has been directly demonstrated that transferred DCs bearing endogenous self-antigen, or pulsed ex vivo with self-antigen, can prime T cells that precipitate autoimmune disease (Ludewig et al., 1998; Dittel et al., 1999). The capacity of DCs pulsed with tumor antigen to induce some types of autoimmunity provides further evidence for the principle that self-antigen-loaded DCs can initiate a chain of events leading to autoimmune disease (Nestle et al., 1998). At least one model stresses the requirement for repeated priming of the response by administration of the antigen in the context of DCs (Ludewig et al., 1998). We have shown a similar capacity of bone marrowderived DCs pulsed with methylated BSA to prime antigen-induced arthritis (AIA) in C57Bl/6 mice. Although systemic priming of T cells occurs after DC administration, arthritis occurs only after administration of the antigen to the knee joint effector site, thereby generating a local immune response (Figure 33.1) (S. Zehntner, unpublished data). A similar scenario occurs in the transgenic rat insulin promoter (RIP) lymphocytic choriomeningitis virus glycoprotein (LCMV GP) model of IDDM, in which the LCMV GP transgene is expressed in the beta cells of the pancreas. Diabetes ensues only after activation of GP-specific cytotoxic T lymphocytes (CTL) either by LCMV infection or by repeated transfer of DCs from H8 mice that present the appropriate GP epitope endogenously (Ludewig et al., 1998). In the models of AIA and RIP-GP IDDM, organ specificity is related to localized expression of specific antigens. However, in some cases, organspecific antigen is endogenous, rather than experimentally directed. An example is type II collagen – the autoantigen in collagen-induced arthritis – whose expression is limited to the joint cartilage and vitreous gel of the eye (Wooley et al., 1989). Perpetuation of the immune response against

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Bacterial antigen or pro-inflammatory factors, such as peptidoglycan, may be persistent because of the longevity of macrophages.

Systemic autoimmunity

FIGURE 33.1 Priming of antigen-induced arthritis using DCs pulsed with mBSA. Either bone marrowderived DCs pulsed with or without the antigen mBSA, mBSA and complete Freund’s adjuvant, or mBSA alone were injected subcutaneously to prime C57Bl6 mice. Arthritis was induced in one knee joint by subsequent injection of mBSA. The contralateral control knee joint was injected with PBS. Both knees were evaluated for arthritis after 5 days.

mBSA in AIA is associated with an autoimmune response amplified to self-epitopes. This model closely resembles the human disease Reiter’s syndrome, in which bacterial antigen persists predominantly in the joints. Approximately 70% of patients bear the class I MHC molecule, HLA-B27, which is likely to be an important presenting element. Patients with disease have an immune response not only to bacterial, but also to self-antigens (Gaston, 1998; Keat, 1999). Improvement of some patients with prolonged antibiotic treatment suggests that the persistence of antigen drives the autoimmune reaction in these individuals (Keat, 1999). Evidence from various forms of arthritis, including RA, suggests that the gut and the joints may be particularly susceptible to persistent deposition of bacterial products and to uptake by resident macrophages and dendritic cells, possibly in response to damage (Melief et al., 1995).

However, these models of persistent local antigen reflect only a small proportion of human autoimmune disease. How is spontaneous autoimmune disease initiated in the absence of deliberate self-antigen priming, and in diseases such as RA and SLE, with varying degrees of systemic autoimmunity? We have previously proposed a model in which RA is initiated by an autoreactive response to endogenous selfantigen presented by DCs differentiated in the presence of pro-inflammatory cytokines (Thomas and Lipsky, 1996a). Priming to such self-antigen would occur in the context of inflammation and would be perpetuated in genetically susceptible individuals to full-blown autoimmune disease. This hypothesis is supported in vitro by evidence of upregulation of endogenous self-antigen presentation by jointderived synovial DCs leading to stimulation of auto-reactive T cells in the autologous mixed lymphocyte response (AMLR). Since the AMLR stimulated by PB DCs can also be upregulated by differentiation of DCs in the presence of GMCSF, TNFα and CD40-ligand, it is clear that normal DCs are able to increase their presentation of endogenous self-antigen. Such antigen may access the endogenous processing pathway in a number of ways, including processing and binding of endogenous peptides to MHC molecules during their assembly and export to the DC surface (Thomas et al., 1994), but also through ingestion of apoptotic bodies and immune complexes. The proliferative response of blood T cells after stimulation by DCs isolated from diseased tissue sites, including RA synovial fluid and psoriatic dermis, is higher than the response stimulated by autologous blood DCs (Nestle et al., 1994; Thomas, et al., 1994). Recent evidence further demonstrates that immature DCs can be activated by mechanical disruption and mechanically disrupted tissues as well as by ingestion of necrotic cells, and in some settings,

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apoptotic cells (Gallucci et al., 1999; Rovere et al., 1998a, 1998b, 1999). Therefore, DCs may be activated nonspecifically in response to tissue damage, necrosis, apoptotic cells or bacterial products that may be present at sites of trauma, damage, irradiation and infection. Of importance, experiments using necrotic cells demonstrated not only that the activated DCs could present self-antigen provided by necrotic cells, but also that endogenous self-antigen presentation by the DCs ingesting necrotic material was upregulated (Gallucci, et al., 1999). The efficiency with which apoptotic cells upregulate DC function could be increased either by opsonization of the apoptotic cells with antiphospholipid antibodies or by increasing the load of apoptotic cells for ingestion (Rovere et al., 1998a, 1998b, 1999). DCs may be exposed to a high load of apoptotic cells in vivo in settings of viral infection, in which the extent of cell death overwhelms the capacity of macrophages to ingest apoptotic cells. Other settings might include high-dose irradiation, and the spleen in diseases such as hemolytic anemia (Pacini et al., 1999). A number of models of autoimmune polyarthritis resembling RA demonstrate that jointspecific self-antigen priming is unnecessary for the initiation of polyarthritis. First, the K/BxN TCR transgenic mouse line spontaneously develops autoimmune arthritis resembling RA (Kouskoff et al., 1996). The target of both initiating T cells and pathogenic immunoglobulins has been identified to be a ubiquitous self-antigen, glucose-6-phosphate isomerase, a glycolytic enzyme (Matsumoto et al., 1999). Once intitiated, the pathology is driven almost entirely by autoantibodies directed against this self-antigen (Korganow et al., 1999). This model has provided the clearest example that autoimmune polyarthritis may develop in the absence of a jointspecific T-cell response. Second, spontaneous arthritis in BALB/c mice with a homozygous targeted mutation in the IL-1ra gene, as well as in mice transgenic for a deregulated form of human TNF-α, both develop spontaneous arthritis (Keffer et al., 1991; Horai et al., 2000). While the reason for joint specificity in these models is not yet known, the previously dis-

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cussed evidence regarding tissue damage suggests that the role of day-to-day trauma and perhaps uptake of bacterial products in the joints may be of relevance. Pro-inflammatory effects of trauma or wearand-tear to joint structures should be detectable in synovial membrane from patients with joint degeneration, particularly of bone and cartilage. This occurs progressively with aging in osteoarthritis, and can be exacerbated by several factors, including injury. By immunohistochemical staining, we explored whether DC infiltration was associated with tissue degeneration in osteoarthritis, or with crystal deposition in gout. While DC infiltration of osteoarthritic synovial tissue was significantly less than that of RA tissue, T cells, B cells and activated DCs were present in a perivascular location in the synovial tissue of patients with osteoarthritis (Plate 33.2A and B) and gout (data not shown) (Pettit et al., 2001). There is no primary autoimmune lesion causing degenerative and crystal-induced arthritis – rather the inflammation results from stimulation of the innate immune system by uric acid or calcium pyrophosphate crystals or by degenerative damage to joint tissue leading to the generation of a tissue inflammatory response (Di Giovine et al., 1991). These data provide evidence against the proposition that pathogen-associated molecular patterns (PAMPs) must stimulate the innate immune system (Medzhitov and Janeway, 1997). Moreover, they imply that nonspecific stimuli of synovial inflammation might trigger or prevent resolution of a more intense and prolonged autoimmune response in genetically susceptible individuals.

Viral infection The functional differentiation of DCs may be upregulated by infection with viral or other pathogens through local production of pro-inflammatory mediators such as TNFα, lipopolysaccharide (LPS) and other patternrecognition molecules, and the induction of interferon-α production by the natural interferon-producing cells (NIPCs), or plasmacytoid

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DCs (Di Rosa et al., 1998; Siegal et al., 1999). In this situation, priming and cross-priming of both exogenous and self-antigens may occur. Viral infections have been associated with a number of autoimmune diseases. While there may be a direct pathogenic role in rare cases (di Marzo Veronese et al., 1996), chronic viral infections may have a number of effects that increase susceptibility to autoimmunity nonspecifically. First, there may be high load of apoptotic cells available for capture by immature DCs. Second, these cells will be ingested in the context of inflammation which can promote maturation of DCs, lymph node migration and priming. Third, viral peptides may mimic self-peptides, leading to priming of cross-reactive self-antigens (Wucherpfennig and Strominger, 1995; Oldstone, 1998). Fourth, viral particles and their expressed genes may be processed by the endogenous pathway, and cryptic self-antigens may be generated by upregulation of endogenous protein synthesis. Tolerance may be overcome by priming to cryptic or subdominant self-epitopes (Moudgil and Sercarz, 1994). Recent experiments highlight another mechanism for the production of cryptic selfepitopes, especially in inflammation. IL-6 was found to alter the processing of antigens in the endosomes of DCs by altering their acidification. In this way, previously cryptic or subdominant epitopes could become dominant when processed and presented by DCs differentiated in the presence of IL-6 (Drakesmith et al., 1998). Due to the immunostimulatory effect of unmethylated CPG motifs within prokaryotic DNA, the immune system is able to distinguish unmethylated prokaryotic DNA from methylated self-DNA sequences, by induction of cytokine production in vitro and in vivo (Manders and Thomas, 2000). The immunostimulatory sequences also lead directly to the maturation of DCs (Jakob et al., 1998; Sparwasser et al., 1998). Systemic autoimmunity, and evidence of autoreactivity to multiple self-antigens characterize the autoimmune disease, SLE. Antibodies to double-stranded DNA are frequently found in SLE, and can form immune complexes with DNA (Pisetsky et al., 1997). Therefore, DNA con-

taining unmethylated CPG motifs has been suggested to be important in the disease pathogenesis (Vallin et al., 1999). Increased levels of endogenous IFNα have been demonstrated in SLE, and an SLE-like syndrome can be precipitated by IFNα therapy (Ytterberg and Schnitzer, 1982; Fritzler, 1994). Of interest, a high molecular weight factor found in SLE serum was determined to be sensitive to DNAse-1 and capable of inducing endogenous interferon-α (IFNα) from NIPCs in SLE patients (Vallin et al., 1999). Upregulation of myeloid DC maturation and function by IFNα has also been described (Luft et al., 1998). The data suggest the possibility that immune complexes of DNA containing immunostimulatory CPG motifs and anti-DNA antibodies might induce IFNα production by NIPCs, that can in turn enhance myeloid DC maturation. The overexpression of IFN-α suggests the possibility that a persistent bacterial or viral infection might lead to chronic inflammation and eventual autoimmune disease, particularly in those individuals with genetic mutations in key regulatory pathways. The number of circulating NIPCs, or plasmacytoid DCs, was found to be reduced in the blood of patients with SLE, and it was hypothesized that these cells had been recruited to inflammatory sites (Cederblad et al., 1998).

REGULATION OF DC TO MAINTAIN TOLERANCE Homeostatic cytokines If DCs can indeed readily present self-antigen in various contexts in the course of day-to-day events, it follows that there must be homeostatic regulation of DCs to reduce the risk of spontaneous upregulation of DC function, and susceptibility to autoreactivity. To date, there is indirect evidence that both immunomodulatory cytokines and active suppression by T cells may regulate DC function in health. Candidate downregulatory or homeostatic cytokines include IL-1 receptor antagonist (IL1ra), IL-10 and TGF-β. Spontaneous autoim-

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mune arthritis was recently demonstrated in BALB/c mice with a homozygous IL-1ra deficiency (Horai et al., 2000). There was associated upregulation of IL-1β, TNFα and IL-6 mRNA expression in these animals. The data suggest an important role for IL-1ra in the maintenance of homeostasis and prevention of overt autoimmune disease in the absence of deliberate selfantigen priming. It is important to stress that the autoimmune pathology only occurred in animals of BALB/c, and not of C57Bl/6 strain, highlighting the additional importance of genetic background for the development of overt pathology in the face of dysregulated immune responses. In the skin, TGF-β is the dominant Langerhans cell (LC) differentiation factor, and has a major immunomodulatory effect on LC maturation. It is of relevance that its downregulatory effects on LC maturation can be overcome by CD40 ligation, through cognate interactions with T cells, but not by TNF-α, IL-1β or LPS (Geissmann et al., 1999). Thus, exposure of epidermal LC to pro-inflammatory factors produced in response to infective or traumatic stimuli is unlikely to be sufficient to induce priming of an autoimmune response by LC unless migrated LC make a productive interaction with self-reactive T cells in the draining lymph nodes. The critical importance of CD40-ligand in the generation of autoimmune diseases has been highlighted previously in animal models, and by the demonstration of functional expression at the effector site in various diseases, including RA and SLE (Koshy et al., 1996; MacDonald et al., 1997).

Regulatory T cells IL-10 plays a similar counter-regulatory role, and IL-10/ mice also develop spontaneous autoimmune inflammatory colitis (Kuhn et al., 1993). The development of spontaneous inflammatory colitis in this and a number of other models, has been shown to relate to an altered homeostatic relationship between colonic bacteria, antigen-specific TH1, and CD45RBdim CD4 T regulatory 1 (Tr1) cells (Groux and Powrie, 1999). Tr1 cells can be induced in vitro

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by repetitive presentation of antigen in the presence of IL-10. Tr1 clones demonstrate a differentiated memory T-cell phenotype, proliferate poorly, secrete IL-10, but little IL-2 or IL-4, and, when antigen-activated inhibit development of colitis induced in SCID mice by transfer of CD45RBbright CD4 T cells (Groux et al., 1997). This suppression is dependent on IL-10 and TGFβ (Groux and Powrie, 1999). Similar populations of anergic T cells have been generated in vitro in response to DC in the presence of IL-10 (Caux et al., 1994). We have previously demonstrated the enrichment of a population of CD4 CD45RBdim T cells in RA synovium, with similar characteristics to the described Tr1 T cells (Thomas et al., 1992). Furthermore, the CD27 subset of CD45RBdim T cells in PB produces high levels of IL-10 upon activation, and this subset is enriched in the RA synovium (Tortorella et al., 1995; Kohem et al., 1996). Peripheral enrichment of anergic T cells has also been described in other autoimmune diseases and autoimmune models (Kouskoff et al., 1996). The data suggest that a population of CD45RBdim CD4 cells with some characteristics of Tr1 cells differentiates in autoimmune disease, presumably as a result of regulatory mechanisms stimulated by the chronic inflammatory process. In this regard, it is of relevance that IL10 is abundant at the effector site of several autoimmune diseases (Cush et al., 1995). IL-10 inhibits generation of DCs from monocytic precursors, and inhibits the function of peripheral precursor DCs (Buelens et al., 1997; De Smedt et al., 1997; MacDonald et al., 1999). In contrast, mature DCs found at the effector site in RA SF are resistant to IL-10-induced immunosuppression because of lack of cell surface IL-10 receptor expression (MacDonald et al., 1999). Therefore, in an IL-10-abundant environment (Cush et al., 1995), peripheral DC immaturity and effector site DC resistance to suppression would be expected. In this regard, evidence for peripheral DC suppression in autoimmune disease has been generated in a number of models, including spontaneous rodent and human IDDM. It has been noted that peripheral DC function is

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reduced in rats and mice with diabetes, and in patients and prediabetic relatives of patients with IDDM (Jansen et al., 1995; Takahashi et al., 1998; Delemarre et al., 1999). This reduced function is related to immaturity of DCs in the periphery. Blood DCs from these patients, as well as those with other autoimmune diseases, including RA, SLE and Sjogren’s syndrome, have been shown to have a reduced ability to stimulate the autologous MLR and to present nominal antigen in vitro, but to stimulate normal allogeneic responses (Thomas et al., 1994). The DC immaturity and functional suppression for autologous, but not allogeneic, T cells suggest active suppression of DC by autologous peripheral T cells. The findings in pre-autoimmune- diseased relatives further suggest that specific genetic factors might affect the balance of T-cell subsets, costimulatory molecules and/or critical regulatory cytokines. While it is not clear whether the same APC is responsible for expansion of autoreactive CD45RBbright and differentiation of regulatory CD45RBdim T-cell subsets, preliminary evidence suggests that DCs can stimulate T cells capable of preventing autoimmune disease. Thus, lymphoid organ DCs bearing appropriate self-

FIGURE 33.3

antigen could prevent the onset of clinical disease after transfer into prediabetic animals, suggesting that the transferred DCs are capable either of stimulating regulatory T cells or deleting potentially autoreactive T cells in a prediabetic host (Clare-Salzler et al., 1992; Shinomiya et al., 1999). As opposed to lymphopenic animals such as SCID, in which the T-cell subsets clearly have opposing effects on autoimmunity (Powrie et al., 1994), in T cell-replete animals, the capacity of DC or T cells to suppress established autoimmune disease has not yet been demonstrated. DC constitutively drain peripheral tissues into afferent lymphatics in large numbers, and these DC may contain apoptotic inclusions derived from somatic cells, to which T cells may be tolerized in the LN (Pugh et al., 1983; Huang et al., 2000; Kurts et al., 1997). The nature of the APC capable of stimulating regulatory T cells in vitro or in vivo is poorly understood. However, two in vitro models suggest that regulatory T cells can be generated by undifferentiated monocytes or DC (Groux et al., 1997; Jonuleit et al., 2000). Regulatory T cells confer antigenspecific regulation, although bystander effects also occur. It is likely that they regulate not only

Model of peripheral tolerance to endogenously-processed self-antigen. DENDRITIC CELLS IN DISEASE

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activated T cells but also APC, potentially through a number of mechanisms (Roncarolo et al., 2000). Considering this emerging concept of peripheral tolerance maintained actively through populations of regulatory cells, it might by hypothesized that for perpetuation of immunity against self-antigens in autoimmune disease, an induced immune response must be sufficient to overcome constitutive regulation against somatic cell damage by regulatory T cells, primed by ‘regulatory’ DC that present selfantigen via an endogenous antigen processing pathway (Figure 33.3). Conversely, preventing or stopping such an induced autoaggressive effector response will require reinstitution of the control on self-antigen presentation, derived from regulatory T cells.

PERPETUATION OF AN INITIAL AUTOIMMUNE RESPONSE Based on the likelihood of nonspecific DC activation, the relative ease with which T cells respond to ubiquitous or cross-reactive selfantigen, and the ease with which autoimmune disease can be initiated by vaccination with DCs, it seems probable that self-limiting autoimmune events are generated frequently in any individual. Furthermore, such events may be more frequent in some individuals than others. Thus, perhaps more important to the pathological outcome than the initiation of autoimmunity are the factors perpetuating autoreactive responses and leading to disease.

Persistent or recurrent priming to self-antigen Persistent priming of self-antigen is associated with persistent antigen at the site of inflammation. This may be related to poor clearance of an antigen as in large immune complexes, ubiquitous self-antigen, persistent chronic nonimmunogenic infections and persistent drainage of DCs from the site to lymph nodes (Edwards and Cambridge, 1998; Ludewig et al.,

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1998; Matsumoto et al., 1999). Persistent antigen deposition ensures that effector T cells are persistently restimulated. Persistent stimulation of autoreactive T cells may also result from a recently discovered caspase 10 mutation in some patients with the autoimmune lymphoproliferative syndrome (ALPS). Due to resistance of the DCs bearing this mutation to TRAILmediated apoptosis, these patients accumulate DCs in the lymph nodes (Wang et al., 1999). In the RIP-GP model of IDDM, repetitive DC immunization with the relevant self-antigen was required for the eventual progression to IDDM. In this model, disease outcome was determined by the number of autoreactive CTL present during antigenic stimulation by DCs. Repeated DC priming also led to de novo formation of inflammation-associated lymphoid structures in the pancreatic islets (Ludewig et al., 1999). Such lymphoid structures are seen in a number of autoimmune diseases, including Hashimoto’s thyroiditis, Sjogren’s syndrome and RA (Thomas et al., 1999). In all cases, the lymphoid tissue contains not only bone marrow-derived DCs, but also follicular DCs within B cell follicles (Randen et al., 1995; Stott et al., 1998; Ludewig et al., 1999). This mouse model provides evidence that priming of a self-specific immune response by repetitive immunization with antigen-loaded DCs alone can lead to the formation of such organized lymphoid tissue. The data suggest that DCs may drive this response. In support of the role of DCs in disease perpetuation, it has been further demonstrated in chimeric models in the NOD mouse that bone marrow-derived antigen presenting cells are required for perpetuation of IDDM. In contrast, antigen presentation by islet β cells is not required (Lo et al., 1993). The data also suggest that after DC infiltration of the lymphoid tissue of autoimmune effector sites, their progress is not arrested, as in lymph nodes, and that the DCs should migrate and prime self-reactive T cells in draining lymph nodes. Preliminary data from our laboratory demonstrate a rapid turnover of DC migration into autoimmune synovial effector sites, and thence to draining lymph nodes, where selfantigen priming takes place (S. Zehntner,

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unpublished data). Thus, the generation of organized lymphoid tissue and its associated chemokines may itself be very relevant to provision of ongoing priming of the self-antigenspecific response. While not all autoimmune effector sites contain organized lymphoid tissue, the infiltration of effector sites by activated DCs surrounded by T cell and B cell aggregates, sometimes associated with FDCs is almost invariable. Furthermore, as was seen in Plate 33.2A and B, such rudimentary organization is common to chronic inflammatory sites even in the absence of an autoimmune pathogenesis. Clear evidence of the infiltration of effector sites in autoimmune diseases by differentiated DCs has been obtained in several animal models (Ziegler et al., 1992; Suzuki, 1995). Moreover, there is accumulating evidence that DCs and macrophages are the first cells to enter autoimmune sites and that these DCs are able to produce cytokines, including TNF-α and IL-12 (Dahlen et al., 1998). The combination of prostaglandin E2 (PGE2) and TNF-α can dramatically upregulate IL-12 production by DCs, thereby stimulating the generation of interferon-γ-producing TH1 effector T cells in the lymph nodes (Rieser et al., 1997). PGE2 may be particularly relevant in this regard in particular locations, including the synovial fluid, pleural and pericardial cavities, that are frequently subject to autoimmune pathology. Paradoxically, in the absence of TNF-α, PGE2 downregulates the ability of DCs and macrophages to produce IL-12, and may be important for homeostatic regulation of DCs at these sites (Panina-Bordignon et al., 1997). The evidence that DCs at effector sites are consistently clustered with T cells but that they enter autoimmune sites even before T cells suggests the capacity of DCs to chemoattract T cells to autoimmune effector sites. The stimulus for entry of DCs into damaged organs is unknown but raises the possibility that damage, or tissue necrosis is chemoattractive for DCs. Early chemoattraction of DCs into infective sites has been previously shown (McWilliam et al., 1994). To examine specificity of DC infiltration into a

wide variety of autoimmune effector sites, work from our laboratory has determined that the DC differentiation marker RelB was 88% specific for detection of mature DC in formalin-fixed human tissues (Pettit et al., 2000). Using this marker, we have shown that the inflamed joint synovial tissue in RA patients is infiltrated by differentiated DCs which are invariably found in association with lymphocytes, predominantly T cells. However, B cell infiltration and aggregates are usually found adjacent to areas of DC infiltration. DCs are typically found in a perivascular location (Pettit et al., 2000). Recent studies demonstrate that the number of infiltrating DCs is related to the patient’s clinical disease activity (Plate 33.2C and D) (Pettit et al, 2001). These data provide clear evidence that infiltration of the effector site by DCs is related to clinical expression of this auto-immune disease. The data suggest that infiltration of the effector site by DCs is important for the perpetuation of RA. As discussed, this may be due to continuous priming to self-antigens by migration of effector site DCs to draining lymph nodes, or to production of pro-inflammatory cytokines and chemokines, leading to tissue damage. Such DC infiltration associated with infiltration of T and B cells was observed at effector sites in a large number of human autoimmune disease tissues including spondyloarthritis, Hashimoto’s thyroiditis, SLE nephritis, Sjogren’s syndrome and ulcerative colitis (Plate 33.2E and F). Similar infiltration by DCs also occurs in the dermis in delayed type hypersensitivity reactions, however these sites were devoid of B cell infiltration, suggesting an important role for B cells in the persistence of inflammation (data not shown).

B cell amplification of autoimmunity As in the RIP-GP model, human autoimmune lymphoid aggregates are characterized by the presence of FDCs. The demonstration of fibroblast-like synoviocytes from patients with RA with intrinsic properties of follicular DCs suggests that the synovial microenvironment may itself allow the development of FDCs from nonhemopoietic resident precursor cells (Shimaoka

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et al., 1998; Lindhout et al., 1999). This may also be the case in the salivary glands in Sjogren’s syndrome, and perhaps other tissues. The presence of FDCs in inflammatory sites indicates that the B cells have developed a high capacity for affinity maturation and the ability to generate high affinity immunoglobulin. Furthermore, bone marrow-derived DCs also play a role in immunoglobulin class switching and B cell differentiation (Dubois et al., 1997). The production of specific auto-antibody may be exquisitely important in some autoimmune situations. For example, in the K/BxN transgenic model, as little as 100 µl of serum is sufficient to transfer disease to susceptible animals. In this model, the auto-antibody specificity is not related to rheumatoid factor (Korganow et al., 1999). In contrast, rheumatoid factor clearly plays an important pathogenic role in RA (van Zeben et al., 1993). Generation of auto-antibody is particularly important for formation of immune complexes, which can deposit in blood vessels and other tissues leading to macrophage and polymorphonuclear leukocyte stimulation and therefore tissue damage. In addition, the presentation of endogenously processed self-antigen by myeloid DCs may be enhanced if the antigen is taken up as immune complexes (Sallusto et al., 1994). At least in some cases, auto-antibody can also function as an opsonin to apoptotic cells (Rovere et al., 1998a). This may be important in the pathogenesis of the antiphospholipid syndrome.

CONCLUDING REMARKS DCs are likely to play a dominant role in initiation and perpetuation of autoimmunity. The specific mechanisms of autoimmune disease generation are likely to be different according to the pathogenesis of individual diseases. Many possibilities for immunotherapy are now available for clinical experimentation. Use of T-cell depletion, oral tolerization, bone marrow transplantation and cytokine inhibition have provided insight into the pathogenesis of some autoimmune diseases. The prospect of

immunotherapy using modified DCs is exciting. Complex issues with regard to the T-cell repertoire, antigen-non-specific mechanisms of tissue damage, secondary regulatory effects and the role of the innate immune system raise questions about how best to use dendritic cell immunotherapy (Wagner et al., 1998). The next 5 years will see its use in animal models and even clinical trials for prevention and treatment of allograft reactions and autoimmunity. For efficient clinical translation, careful design and analysis of the effect of such interventions on interacting T cells and B cells will be essential.

ACKNOWLEDGEMENTS I thank Allison Pettit and Simone Zehntner for inclusion of their unpublished data, and Allison Pettit and Ian Frazer for critical reading of the manuscript.

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PLATE 33.2 Dendritic cells infiltrate autoimmune and non-autoimmune effector sites. Formalin-fixed osteoarthritis synovial tissue (A, B) rheumatoid arthritis synovial tissue before treatment (C), or after successful treatment (D), thyroid tissue from a patient with Hashimoto’s thyroiditis (E) or colon biopsy from a patient with ulcerative colitis were stained with either RelB (brown) and Ulex (red, A, B, C, D), or RelB (brown) and HLA-DR (red, E, F). Approximately 80% of cells staining for RelB in the nucleus are DCs. (B) is representative of approximately 15% of OA perivascular aggregates. Black arrows denote blood vessels, white arrow denotes thyroid follicle. OA, osteoarthritis; RA, rheumatoid arthritis; HT, Hashimoto’s thyroiditis; UC ulcerative colitis.

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C

H

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34 Interaction of dendritic cells with bacteria M. Rescigno1, M. Urbano1, M. Rittig 2, S. Citterio 1, B. Valzasina 1, F. Granucci 1 and Paola Ricciardi-Castagnoli 1 1

University of Milano-Bicocca, Italy; and 2 University of Erlangen, Germany

It is no measure of health to be well-adjusted to a profoundly sick society Krishnamurti

DCs AS SENTINELS TO ALERT THE ADAPTIVE IMMUNE RESPONSE

different types of effector adaptive responses are directed by the innate immune system after recognition of particular groups of pathogens (Medzhitov and Janeway, 1997). The central role of dendritic cells (DCs) in the induction of adaptive immune responses towards infectious agents has been extensively described (Steinman, 1991; Ibrahim et al., 1995; Banchereau and Steinman, 1998), but, recently, a new role of DC as a link between the nonantigen- and the antigen-specific responses has been proposed (Figure 34.1). DCs are indeed widely distributed especially in tissues that interface the external environment, such as the skin, the gut and the lungs (Sertl et al., 1986; Nestle et al., 1993; Nelson et al., 1994), where they can perform a sentinel function for incoming pathogens, and have the capacity to recruit and activate cells of the innate immune system upon inflammation (Foti et al., 1998; Sallusto et al., 1998; Fernandez et al., 1999; Rescigno et al., 1999). Uptake of pathogens induces a state of activation which eventually leads to the

Many of the bacteria that enter the body via the oral or the airway routes, do not survive to the inhospitable microenvironments that they encounter. Still, some microorganisms have evolved mechanisms to subvert this first line of defence and to infect the host. Thus, mammals have developed second lines of defence, consisting in innate and adaptive immune responses, to protect themselves from infectious microorganisms. The innate response is phylogenetically more ancient and, for a long time, it has been considered to be broadly directed to microorganisms. However, the discovery of a new class of receptors, involved in recognition of patterns characteristic of groups of microorganisms (the toll-like receptor family) has re-evaluated the role of the innate immune system as a discriminating system. Indeed, there is increasing evidence that the induction of Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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THE COMPLEXITY OF MICROORGANISMS AND THEIR STRATEGIES TO EVADE IMMUNE RESPONSE

FIGURE 34.1 DCs as a bridge between adaptive and innate immune responses.

migration of the antigen-loaded DCs to the lymphoid organs where the cells of the adaptive immune response can be alerted (Moll et al., 1993). Thus, understanding the interaction of bacteria with DCs, and the early molecular events resulting from this interaction may shed some light on the mechanisms of initiation of the immune response to infectious agents and on aspects of invasiveness, pathogenicity and the persistence of certain bacteria.

FIGURE 34.2

The interactions of DCs with microorganisms are extremely complex. The reasons for this complexity are to be discovered in two characteristics of bacteria: (1) their antigenic complexity; and (2) the strategy developed by bacteria to evade the immune response of the host. This chapter aims to bring together some of the unifying principles governing the interaction of bacteria and DCs. It will do so by reference to a limited number of microorganisms which have been chosen as representatives of three major bacterial groups according to their cell envelope, namely gram-positive, gramnegative and mycobacteria. A common feature of the three groups is the inner protoplasmic membrane (PM) and a peptidoglycan (PG) cell wall. In addition, gram-negative bacteria, exemplified here by Salmonella thypimurium, have an outer membrane containing proteins and lipopolysaccharides (LPS) which are antigenic. The outer membrane often contains pili, or fimbriae formed of type-specific protein antigens which can mediate attachment to the host cell surface or be anti-phagocytic (Figure 34.2). Mycobacteria have an outer membrane of mycolic acid (MA) residues and phenolic

Model of gram-positive, gram-negative and mycobacteria cell wall. DENDRITIC CELLS IN DISEASE

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glycolipids (PGL) linked through arabinogalactan (AG) to the inner peptidoglycan layer; the bacterial antigens are carbohydrate-based, lipoarabinomannans which extend from the protoplasmic membrane to the bacterial surface. Finally, gram-positive bacteria, exemplified here by Streptococcus gordonii, have a cell wall composed predominantly of peptidoglycan (PG), and lipoteichoic acids (LTA) attached directly to phospholipids of the protoplasmic membrane. LTA protrude to the cell surface where they form surface antigens for cell adhesion (Figure 34.2).

Antigenic complexity The antigenic complexity of bacteria is the result of a number of several distinct epitopes, often repeated (as in the case of carbohydrates); however, bacterial protective structures, such as the capsule, may limit epitope recognition. In addition, antigen accessibility can vary because internal antigens may become accessible only after the bacterium has been killed.

Bacterial strategies to evade immune response Bacteria have evolved strategies to overcome the host defence response by either avoiding the action of complement and phagocytosis or by evading antigen recognition. Meningococci and Pneumococci mask surface components that activate the complement by secreting a capsule that covers these activators. Some pathogenic bacteria, such as Pseudomonas or Staphylococci, produce exotoxins that kill neutrophils and macrophages. Pneumococci, instead, avoid phagocytosis because they are covered by antiphagocytic capsules. As we shall see, other pathogenic bacteria do not escape phagocytosis, but have developed strategies to survive inside phagocytes either by inhibiting phagosome– lysosome fusion, or by a rapid escape from the phagocytic vacuole into the cytoplasm, or by the resistance to lysosomal enzymes. Finally, some infectious agents divert lymphocyte function, as in the case of superantigens, or avoid antigen

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recognition by changing their surface antigens, as in the case of Gonococci.

IMMATURE DCs ARE READILY PHAGOCYTIC Since DCs are essential for priming the immune system to antigens, it is important that they are present in tissues where they may encounter bacteria soon after invasion. This is indeed the case. DCs have a privileged distribution in tissues which interface with the external environment and therefore they can serve as efficient sentinels for the uptake of invading microorganisms. Since both the bacterial cell and DC have a net negative charge, specific molecular interactions including bridging molecules are involved in the uptake process. Despite several early reports on the uptake of particulate material and cells by DCs (reviewed in Austyn, 1996), the phagocytic capacity of DCs has long been denied. One reason for this was the technical difficulty until recently, of growing DCs in their immature phenotype. Earlier DC purification, and in vitro cell growth procedures were such that the enriched DCs were in most cases terminally differentiated and had a mature phenotype and function. Only recently have maturation stages of DCs been recognized, and it became possible for immature ‘processing DCs’ to be grown and maintained in vitro (Winzler et al., 1997). At this functional stage, DCs have phagocytic activity (Figure 34.3) which decreases with DC differentiation in vitro. Indeed, several studies have shown that DCs can not only internalise latex and zymosan beads (Inaba et al., 1993; Reis e Sousa et al., 1993; Austyn et al., 1994; Matsuno et al., 1996), but also apoptotic bodies (Parr et al., 1991), as well as microbes such as Bacillus Calmette-Guerin (BCG) (Inaba et al., 1993; Henderson et al., 1997), Saccaromyces cerevisiae, Corynebacterium parvum, Staphylococcus aureus (Reis e Sousa et al., 1993), Leishmania spp. (Blank et al., 1993), and Borrelia burgdorferi (Filgueira et al., 1996). Both gram-positive and gram-negative bacteria, as well as their respective bacterial cell wall

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FIGURE 34.3 Phagocytosis of GFP (green fluorescent protein)-expressing S. typhimurium (B) as detected by confocal microscopy (nuclear section). DC cell shape is shown by phase contrast (A). Phagocytosis of S. gordonii by D1 cells (C) as detected by electron microscopy (EM).

FIGURE 34.4 High motility of TNFα activated D1 cells as shown by time-lapse video microscopy. Frames represent 6 s apart for a total of 2 min of video recording.

components (LPS, LTA) function as DC activators (Riva et al., 1996; Thurnher et al., 1997; Rescigno et al., 1998a). The ability of DCs to phagocytose particulates or bacteria is greatest

in immature, processing DCs, whereas this capacity is reduced, but not abolished, in mature presenting DCs (Henderson et al., 1997). Upon attachment, DCs engulf the bacterium

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FIGURE 34.5 EM micrographs showing conventional phagocytosis of bacteria. Partial (A) and complete (B) degradation of undegraded bacteria in phago-lysosomes.

by actively surrounding it with pseudopodia. This process is started intrinsically when phagocytosis-promoting receptors of the Fc-type are involved, but in the case of complement-type receptors it is dependent on complementary signals. The movement of the pseudopodia in activated DCs involves actin-binding proteins, and it can be blocked by the drug cytochalasin D which stops the polymerisation of actin and inhibits phagocytosis. The rearrangement of the cytoskeleton is associated with DC motility (Winzler et al., 1997). The movement of activated DCs can be observed by time-lapse video microscopy as shown in Figure 34.4. Here it can be seen that in less than 1 min (frames represent 6 s apart) a cell completed the movement of one of its ‘veils’ thus indicating very intense motility. Once a bacterium has been fully internalised in the phagosome, fusion of the phagosome with other intracellular vacuoles or granules takes place. Processing of bacterial molecules for antigen presentation occurs in lysosomes following their fusion with phagosomes (see Figure 34.5). This process may take several hours, as

antigen presentation of bacterial antigens is not observed earlier than 6 h following infection. In addition to the conventional zipper-type phagocytosis, there are a number of other mechanisms by which phagocytes may engulf bacteria. From the various unconventional uptake mechanisms known (Swanson and Baer, 1995; Figure 34.6), DCs have been found to use macropinocytosis and coiling phagocytosis (as shown in Figure 34.7), simultaneously with conventional phagocytosis for the uptake of bacteria (Rittig et al., 1995, 1996). These alternative routes of internalisation will almost certainly result in different routes of antigen processing and presentation. This is because macropinosomes are built differently from phagosomes (Swanson and Watts, 1995) and coiling phagocytosis of Borrelia Burgdorferi delivers these spirochetes into the cytosol (Rittig et al., 1992) from where spirochetal antigens can join the endogenous route of MHC class I presentation (Rittig et al., 1994). Indeed, Lyme borreliosis is one of the bacterial infections which also elicits a cytotoxic CD8 response against the pathogen (Busch et al., 1996).

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FIGURE 34.6 Model of conventional and unconventional mechanisms of bacterial uptake dependent on the microbial model used.

DC SURFACE MOLECULES WHICH CAN MEDIATE THE UPTAKE OF BACTERIA To be internalised, bacteria have first to be attached to phagocytosis-promoting receptors (Figure 34.8). This may take place either via direct interaction between microbial adhesins and phagocytic receptors (non-opsonic uptake); or indirectly via opsonins, for example antibody or complement. These act as bridging molecules between the microbial surface and opsonin receptors of the phagocytes (opsonic uptake). The fusion (zipper model) of the phagosome to form a discrete vacuole is probably under the control of specific fusogenic proteins which have still not been identified. DCs express a moderate level of Fc receptors (FcR) which is not modulated during maturation. DCs also express the Mac-1 molecule (CD11b/CD18; αMβ2 integrin) which is the CR3 complement receptor used for the phagocytosis

of complement-coated bacteria. As with FcR, the surface expression of Mac-1 molecules is not changed during activation of DCs. This is in contrast to monocytes and neutrophils which strongly upregulate Mac-1 expression during differentiation, and in the presence of inflammatory stimuli. As well as having a role in receptor-mediated internalisation, the Mac-1 molecule also mediates adhesion and chemotaxis (Anderson, 1986). Recent studies have shown that Mac-1 is stored in intracellular vesicles which are rapidly mobilised to the cell surface in response to chemoattractants (Miller et al., 1987). Although the presence of the mannose receptor on DCs is still questionable, internalisation and presentation of mannosylated proteins is very efficient in DCs (Tan et al., 1997). The high mannan content in the bacterial oligosaccharide cell envelope in both gram-positive and gram-negative bacteria assures efficient mole-

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FIGURE 34.7 EM micrograph showing morphological appearance of DC macropinocytosis (A) and coiling phagocytosis (B) of bacteria. Bars represent 0.3 µm (A) and 0.6 µg (B).

cular recognition. Therefore it is likely that bacteria use the mannose receptor pathway for internalisation. Bacterial cell products, such as LPS, are inducing activation of DCs both in vivo (Roake et al., 1995) and in vitro (Verhasselt et al., 1997). After the discovery that hyporesponsiveness to bacterial lipopolysaccharide (LPS) was associated to mutations in the Tlr4 gene (Poltorak et al., 1998), several groups have

identified members of the expanding family of Toll-like receptors (TLR) (Medzhitov and Janeway, 1997) as the transducing receptors for precise constituents of the bacterial cell wall. Although many of these studies have been carried out on fibroblasts engineered to express members of the TLR family, it is clear that some receptors are specialised in recognising some structures rather than others and there is great

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FIGURE 34.8 Model of different types of conventional phagocytosis (Fc-type, Opsonic, complement-type and non-opsonic).

variability between mouse and human systems (Table 34.1). In particular, human TLR-2 has been shown to mediate LPS-induced cellular signaling (Yang et al., 1998), whereas mouse TLR-4 mediates both LPS and lypotheicoic acid (LTA) cellular signaling (Takeuchi et al., 1999). By contrast, mouse TLR-2 is mainly involved in recognising either peptidoglycans (PGN) which are typical components of the cell wall of gram-positive bacteria (Takeuchi et al., 1999; Yoshimura et al., 1999), or lipoarabidomannan (LAM) isolated from rapidly growing mycobacteria (Means et al., 1999). Thus it seems that at least in macrophages, distinct members of the TLR family enable the cells to discriminate between different classes of microorganisms. DCs express the TLR4 which plays a major role

in LPS-induced DC maturation, but the absence of this receptor does not compromise the activation of DC induced by whole gramnegative bacteria (Rescigno et al., manuscript in preparation). It will be interesting to dissect out the exact contribution of these receptors in the outcome of the immune response.

INTRACELLULAR GROWTH AND SURVIVAL OF BACTERIA In the normal course of events, the phagocytic vacuole containing the bacteria fuses with a second type of vacuole, the lysosome, giving rise to a hybrid vacuole termed the phago-lysosome

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TABLE 34.1

Microorganisms recognised by members of the Toll-like receptor family

Receptor

Species

Microorganisms or their components

Reference

TLR2 TLR2 TLR2 TLR2 TLR4 and TLR2 TLR2 TLR2 TLR4 TLR2 TLR4 TLR2 TLR4

Human Human Human Human Human Human Human Mouse Mouse Mouse Mouse Mouse

Bacterial lipoproteins Bacterial lipoproteins LPS LPS binding protein Gram and Mycobacterium avium Mycobacterium tubercolosis Lipoproteins, lipopeptides, sonicated B. burgdorferi Peptidoglycan and gram bacteria LPS Yeast and gram bacteria LPS and LTA Peptidoglycan Heat shock protein60?

Aliprantis et al., 1999 Brightbill et al., 1999 Yang et al., 1998 Means et al., 1999 Means et al., 1999 Hirschfeld et al., 1999 Yoshimura et al., 1999 Poltorak et al., 1998 Underhill et al., 1999 Takeuchi et al., 1999 Takeuchi et al., 1999 Ohashi et al., 2000

(Figure 34.5). A number of bacteria have learned to escape lysosomal killing. Three different strategies for enhancing intracellular bacterial survival are known: (1) resistance to the killing action of the lysosomal enzymes; (2) rapid escape from the phagocytic vacuole into the cytoplasm; and (3) prevention of phagolysosomal fusion. The precise molecular basis by which the bacterium interferes with the host cell metabolism is largely unknown. Salmonella typhimurium survives within the intracellular vacuole even after fusion with potentially lethal lysosomes, and a number of genes involved in this resistance have been identified. Following internalisation by macrophages, S. typhimurium increases the synthesis of over 30 different bacterial proteins, including the heat shock proteins GroEL and DnaK, which are also the immunodominant antigens. Interestingly, the bacterial heat shock proteins are not synthesised by S. typhimurium when the bacteria infect epithelial cells, probably because the epithelia cells are not equipped for the intracellular degradation of phagocytosed microbes. Virulence is regulated by a two component regulator/signal system consisting of two genes, PhoP/PhoQ. Mutations of these genes leads to attenuation of virulence due to decreased microbial survival and increased sensitivity to acidic pH. Rickettsia can be found in the cytoplasm within 30 min of being phagocytosed. This is because bacterial phopholipase A activity facilitates the dissolution of the phagosomal membrane and

the escape of the bacterium into the cytoplasm. Shigella flexneri and S. dysenteriae have plasmids encoding haemolysin activity on contact with the cell membrane, whereas Listeria monocytogenes – which is also able to escape from the vacuole – has haemolytic activity involving haemolysin, listeriolysin O (LLO), a member of the thiol-activated family of cytolysins. Cytolysins similar to listeriolysin are produced by a number of other gram-positive bacteria, including Clostridium and Streptococcus. It appears that a general strategy within this group of cytoplasmic parasites is a rapid exit from the phagosome which is mediated by a controlled contact lysis of the membrane. The bacteria which prevent fusion of the endocytic vacuole with the lysosome include M. tuberculosis, which preferentially infects macrophages. If however, the phagocytosed M. tuberculosis are coated with antibody, phagolysosomal fusion does take place, and the bacteria continue to multiply. This resistance is related to the structure of the mycobacterial envelope, which contains peptidoglycolipids not found in gram-positive bacteria. The mycobacteria produce polyanionic glycolipids and sulphatides, which seem to interact specifically with the lysosomal membrane. This causes a reduction in vacuole mobility within the cytoplasm, and this may reduce the chances of lysosome and phagosome fusion. Model antigens expressed in recombinant gram-positive and gram-negative bacteria can be presented on both MHC I and II molecules

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(Svensson et al., 1997; Rescigno et al., 1998; Corinti et al., 1999), and the efficiency of presentation on MHC I molecules is one million times higher than presentation of the soluble protein (Rescigno et al., 1998). Unlike macrophages, this exogenous pathway of MHC I presentation is transporter associated with antigen presentation (TAP)-dependent (Rescigno et al., 1998). This implies that exogenous bacterial antigens introduced by phagocytosis are directed into the classical pathway of MHC I presentation. Indeed, transport of whole bacterial proteins from phagolysosome to the cytosol takes place after phagocytosis of bacteria (Plate 34.9) as happens after immune-complex internalisation (Rodriguez et al., 1999). The capacity of DCs to present bacterial antigens with very high efficiency on both MHC I and II molecules can be exploited to induce strong and long-lasting immunity towards bacteria, as well as nominal antigens of interest. Several examples of partial

protective immune responses achieved by injecting in vivo DCs loaded with microbes have been described against Chlamydiae trachomatis (Su et al., 1998), against Borrelia burgdorferi (Mbow et al., 1997) or against Mycobacterium tubercolosis (Demangel et al., 1999).

BACTERIAL INFECTION OF DCs RESULTS IN CYTOKINE PRODUCTION AND FUNCTIONAL MATURATION A few hours after bacterial infection, DCs synthesise a number of cytokines and chemokines (Rescigno et al., 1997, 1998a). As shown in Figure 34.10, the production of both TNFα and IL-6 was readily detected in DCs infected with either gram-positive or gram-negative bacteria. Interestingly, activation of DCs is achieved particularly when bacteria are alive, indeed, heat-

FIGURE 34.10 Time-course of the cytokines released by DC infected with S. typhimurium. IL-12, IL-10, TNFα and IL-6 production by D1 cells following bacteria stimulation were detected by standard ELISA assays. DENDRITIC CELLS IN DISEASE

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inactivated bacteria although phenotypically induce the maturation of DCs, fail to induce a high production of inflammatory cytokines (Rescigno et al., manuscript in preparation). IL-12 Is produced by myeloid mouse DCs only in a very small number after bacterial encounters as compared to human monocyte-derived DCs (Corinti et al., 1999), but this could be attributed to the different population of DCs analyzed rather than to interspecies diversities. Indeed, IL-12 in the mouse is produced in large amounts by lymphoid DCs rather than by myeloid DCs in response to microbial challenge in vivo (Reis e Sousa and Germain, 1999). Moreover, it has been shown recently that after an initial production of IL-12 which is crucial to induce IFN-γ-mediated responses for the clearance of intracellular pathogens, DCs if restimulated, are refractory to further produce IL-12 in vivo. This confers to DCs an important role of regulators of the inflammatory response besides the one of inducers of immune responses towards pathogens (Reis e Sousa et al., 1999). TNFα production is rapidly induced following infection. It is likely that the phenotypical and functional maturation which occurs in DCs within 24 h of bacteria uptake is the result of cytokine amplification during this response. Indeed, DC activation by TNF-α alone mimics the phenotypical maturation observed after bacterial infection, although the addition of anti-TNF-α antibodies only partly inhibits DC phenotypical and functional maturation. This is consistent with the finding that the pattern of genes induced after activation of DCs by TNF-α and LPS is very different and that the number of genes regulated during either LPS or bacterial activation is highly diverse (Granucci et al., 2001). Indeed, maturation obtained by whole bacteria is quantitatively and qualitatively more pronounced indicating the induction of several transducing pathways, likely via receptors which recognise distinct bacterial components. Furthermore, incubation of DCs with several strains of bacteria resulted in a clear modification of cell surface activation markers. Consistent with acquisition of costimulatory activity during maturation is the upregulation of

483

B7.2 (CD86) and CD40 molecules. This phenomenon is dose-dependent, and it is already maximal at a bacteria : DC ratio of 10 : 1 (Table 34.2). The upregulation of B7.2 and CD40 molecules has also been observed with BCG (Thurnher et al., 1997) and Mycobacterium tuberculosis (Henderson et al., 1997), but it was not observed following the use of inert latex beads of various sizes (Figure 34.11). This is relevant when considering the rational design of new vaccines intended to target and activate DCs. In fact, the upregulation of the co-stimulatory molecule B7.2 and the coordinated translocation of MHC molecules at the cell surface are essential molecular events for the subsequent antigen presentation and activation of both CD4+ and CD8+ T cells. The internalisation of bacteria is associated with maturation of DCs and with increased TABLE 34.2 Bacteria-induced upregulation of costimulatory molecules on DCs

Uninfected D1 cells D1 cells infected with: Salmonella typhi E. coli Lactobacillus Staphylococcus S. aureus S. pyogenes Pneumococcus R6 (not capsulated) Streptococcus Lactococcus Stretococcus gordonii Mycobacterium smegmatis Listeria monocytogenes

CD40

B7.2

/

/

      

      

    

    

FIGURE 34.11 Bacterial products induce DC maturation. Upregulation of the co-stimulatory molecules (CD40 and B7.2) on D1 cells is detected following LPS activation, but not after latex beads phagocytosis (1 or 2 µm diameter).

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stability of MHC class I- and class II-peptide complexes. Thus the half-lives of MHC class IIand class I-peptide complexes change from 10 to 20 h and from 3 to 9 h, respectively. This has important consequences for T cell induction because it increases the chances of DCs to encounter antigen-specific T cells in the draining lymph nodes (Rescigno et al., 1998a).

INTRACELLULAR PATHWAYS LEADING TO DC ACTIVATION TLRs signal through the NF-κB pathway and induce the expression of NF-κB-controlled genes for inflammatory cytokines such as IL-1, IL-6 and IL-8 (Medzhitov et al., 1997). TLRs have been shown to mediate the intracellular signaling of LPS (Poltorak et al., 1998) or of the complex LPSCD14- and LPS-binding protein (LBP) (Yang et al., 1999). Indeed, DC maturation induced by LPS or bacteria is mediated by NF-κB and inhibition of NF-κB blocks phenotypical maturation of DC (Rescigno et al., 1998b; Hofer et al., 2001). Three out of five members of the NF-κB family are rapidly recruited after DC activation (30 min) whereas RelB is translocated into the nucleus only 4 h after bacterial encounter suggesting a role for this member in late events of DC maturation (i.e. antigen presentation and/or DC migration). Interestingly, activation of DCs by LPS promotes survival of the cells in a growth arrested state after deprivation of growth factors (Rescigno et al., 1998b). This response is dependent on mitogen-activated protein (MAP)-kinases of the ERK family but the anti-apoptotic mediators still need to be identified.

DENDRITIC CELLS AS IMMUNISATION TARGETS FOR THE RATIONAL DESIGN OF MUCOSAL VACCINES In recent years it has become clear that antigen delivery is a crucial field of research for vaccine development. Appropriate targeting of an antigen to the immune system can affect the type

of response: local/systemic, humoral/cellular, class I-/class II-restricted, TH1/TH2. Although much still has to be done to fully understand the interaction between DCs and different bacteria, some common features have already been identified. All tested bacterial strains (pathogenic and non) lead to functional and phenotypical maturation of DCs. Thus, as long as bacteria have crossed the epithelial barrier regardless of their pathogenicity become dangerous for DCs. Furthermore, according to how bacteria are internalized by DCs, i.e. conventional versus unconventional phagocytosis, there are different efficiencies of MHC class I and II presentation. Likewise, the capacity of microorganisms to escape the phagolysosome can result in reduced MHC II, but increased MHC I presentation. Also, the capacity of live bacteria, but not of heat-killed bacteria to induce full maturation of DCs has to be considered. Concerning the route of administration of live attenuated bacteria, subcutaneous vaccination of mice with S. typhimurium carrying a tumor antigen is not effective when compared to DCs loaded with the same recombinant Salmonella (Rescigno et al., 2001). Thus, the subcutaneous route might not be the best route for recombinant bacteriabased vaccine development. On the other hand, recombinant bacteria as antigen delivery systems for in vitro loading of DCs is very powerful and should be exploited to induce immunity. The delivery of vaccine antigens to mucosal surfaces is of particular interest, because it can generate local immunity at the major portals of pathogen entry. Vaccination strategies based on mucosal immunisation can also be safer, minimise adverse effects, and make administration easier. Delivery systems based on live microorganisms, such as recombinant bacterial vectors, are especially suitable for mucosal immunisation and they might represent the Trojan horse to fight and overcome cancer.

ACKNOWLEDGEMENTS We wish to thank Donata Medaglini and Gianni Pozzi for providing the bacteria used in

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this study, Jürgen Wehland and Antonio Sechi for time lapse video-microscopy, Patrizia Rovere for confocal microscopy. This work was supported by Biopolo, EC grant No. ERB FMRX CT 960053 and CNR target project in Biotechnology.

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Rodriguez, A., Regnault, A., Kleijmeer, M., RicciardiCastagnoli, P. and Amigorena, S. (1999). Nat. Cell. Biol. 6, 362–368. Sallusto, F., Schaerli, P., Loetscher, P. et al. (1998). Eur. J. Immunol. 28, 2760–2769. Sertl, K., Takemura, T., Tschachler, E., Ferrans, V.J., Kaliner M.A. and Shevach, E.M. (1986). J. Exp. Med. 163, 436–443. Steinman, R.M. (1991). Annu. Rev. Immunol. 9, 271–296. Su, H., Messer, R., Whitmire, W., Fischer, E., Portis, J.C. and Caldwell, H.D. (1998). J. Exp. Med. 188, 809–818. Svensson, M., Stockinger, B. and Wick, M.J. (1997). J. Immunol. 158, 4229–4236. Swanson, J.A. and Baer, S.C. (1995). Trends Cell Biol. 5, 89–93. Swanson, J. and Watts, C. (1995). Trends Cell Biol. 5, 424–428. Takeuchi, O., Hoshino, K., Kawai, T. et al. (1999). Immunity 11, 443–451.

Tan, M.C., Mommaas, A.M., Drijfhout, J.W. et al. (1997). Eur. J. Immunol. 27, 2426–2435. Thurnher, M., Ramoner, R., Gastl, G. et al. (1997). Int. J. Cancer 70, 128–134. Underhill, D.M., Ozinsky, A., Hajjar, A.M. et al. (1999). Nature 401, 811–815. Verhasselt, V., Buelens, C., Willems, F., De Groote, D., Haeffner-Cavaillon, N. and Goldman, M. (1997). J. Immunol. 158, 2919–2925. Winzler, C., Rovere, P., Rescigno, M. et al. (1997). J. Exp. Med. 185, 317–328. Wright, S.D. (1995). J. Immunol. 155, 6–8. Yang, R.B., Mark, M.R., Gray, A. et al. (1998). Nature 395, 284–288. Yang, R.B., Mark, M.R., Gurney, A.L. and Godowski, P.J. (1999). J. Immunol. 163, 639–643. Yoshimura, A., Lien, E., Ingalls, R.R., Tuomanen, E., Dziarski, R. and Golenbock, D. (1999). J. Immunol. 163, 1–5.

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PLATE 34.9 Transport of GFP from the phagosome to the cytosol after uptake of GFP-expressing S. typimurium in a time course. The graph shows the quantification of cytosolic GFP in DCs that have internalised the bacteria.

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35 Dendritic cells during infection with HIV-1 and SIV R. Ignatius1, Ralph M. Steinman1, Angela Granelli-Piperno 1, D. Messmer1, C. Stahl-Hennig 2, K. Tenner-Racz 3, Paul Racz 3, I. Frank1, L. Zhong 1, S. Schlesinger Frankel 4,5 and Melissa Pope 1 1

The Rockefeller University, New York, NY, USA; German Primate Center, Göttingen, The Netherlands; 3 Bernhard Nocht Institute for Tropical Medicine, Germany; 4 Division of Retrovirology, Walter Reed Army Institute of Research and the Henry M. Jackson Foundation, Rockville, MD; and 5 Armed Forces Institute of Pathology, Washington, DC, USA 2

The enemy is anybody who’s going to get you killed, no matter which side he is on. Yossarian in Catch 22 Joseph Heller, 1961

INTRODUCTION

mission and immunodeficiency development (Desrosiers, 1990). Both HIV-1 and SIV infect cells that express CD4 and chemokine receptors (Sattentau et al., 1988; Edinger et al., 1997). Such receptors are found on different leukocytes, including T cells, macrophages and DCs. DCs can express additional newly described receptors that may also be important in SIV/HIV infection (Samson et al., 1998; Ignatius et al., 2000). DCs obtained from the blood and tissues of macaques exhibit comparable phenotypes and functions to their human counterparts (O’Doherty et al., 1997; Pope et al., 1997a; Ignatius et al., 1998) (Table 35.1). Therefore, as a model for HIV-1, the macaque system is being closely examined to gain further insight into the involvement of DCs in SIV transmission, pathogenesis, and vaccine development.

Dendritic cells (DCs) can drive virus replication in collaboration with CD4 T cells as evidenced from in vitro and in vivo studies. DCs may acquire viruses and/or viral antigens (e.g. infection with pathogenic versus attenuated virus, uptake of Ab-coated virus or apoptotic-infected cells) for the subsequent activation of virusspecific immunity and this may impact disease development. Virus-carrying DCs, while driving immune activation, could directly destroy an essential group of cells required to combat the ensuing infection, by actively promoting the transmission of virus to CD4 T cells (Plate 35.1). The SIV-macaque model is one of the best animal models available to study HIV-1 transDendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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TABLE 35.1

Phenotype of macaque dendritic cells

Immature 

HLA-DR CD86 CD80 CD83/ CD40 CD25 p55 CD68 diffuse DC-LAMP DEC-205/ CD4 CD14

Mature HLA-DR CD86 CD80 CD83 CD40 CD25 p55 CD68 spot DC-LAMP spot DEC-205 CD4 CD14/

In review, we will provide an overview of the in vitro studies investigating the interactions between HIV-1 and SIV with different human and macaque DC populations and relate this to in vivo studies documenting where virus is located in situ. There are three main areas in which DCs most significantly foster virus transmission and spread (at acute and chronic stages of infection): epithelial surfaces, peripheral lymph nodes (LNs) and mucosal-associated lymphoid tissues (MALT). Summarized herein is the evidence that DCs can amplify immunodeficiency virus transmission and subsequent replication within these tissue locations, and how this may contribute to the immunity elicited in response to infection.

DENDRITIC CELLS AT THE EPITHELIAL SURFACES AND SEXUAL TRANSMISSION OF INFECTION Maturation and migratory characteristics of dendritic cells at body surfaces Skin-derived DCs have been used as a model to study the interactions between immunodeficiency viruses and DCs within the mucosal epithelia that are exposed to virus during sexual transmission. DCs located within the stratified squamous epithelium (SSE) of the epidermis, Langerhans cells (LCs), are similar to those DCs

positioned in the epithelial coverings of the vagina, ectocervix and anus (Plate 35.2). The dermis and subepithelial regions contain additional DCs (Lenz et al., 1993), that lack markers typically expressed by LCs, i.e. Birbeck granules, CD1a antigens, and a newly described C-type lectin Langerin (Valladeau et al., 2000). Detailed studies of skin DCs have lead to the concept that there are at least two stages in their function (Plate 35.3) (Schuler and Steinman, 1985; Romani et al., 1989; Streilein and Grammer, 1989). Skin and other tissues (e.g. blood, lung, bone marrow, mucosal epithelia) contain immature antigen-capturing DCs that, with the appropriate inflammatory stimuli, become mature T cell-stimulatory DCs. Mature DCs produce large amounts of IL-12, have diminished endocytic capacity, and upregulated expression of adhesion and costimulatory molecules (Steinman, 1999). Mature DCs also express the more recently defined lysosome-associated molecule, DC-LAMP, which is homologous to CD68 (Table 35.1 and Plate 35.4) (de Saint-Vis et al., 1998). In contrast, immature DCs express lower amounts of these molecules (except CD68, Table 35.1), have a greater capacity to endocytose antigens, including viruses, by a variety of routes and process them within intracellular compartments for the subsequent presentation to CD4 or CD8 T cells by matured DCs. DCs undergo another important change during maturation, which is that the cells begin to migrate (Plate 35.2) (Larsen et al., 1990; Richters et al., 1994; Pope et al., 1995a; Lukas et al., 1996). Migration of DCs from the skin is modulated by both TNF-α and IL-1β (Stoitzner et al., 1999). Epidermal and dermal DCs leave the skin via the dermal lymphatic vessels (Lukas et al., 1996) to traffic to the draining LNs. This phenomenon has been exploited to isolate DCs from skin explants in vitro, where epidermal LCs and dermal DCs migrate into the surrounding medium, making them accessible for experimentation (Richters et al., 1994; Pope et al., 1995a). Considerable work has also been performed using immature and mature DC populations derived from blood monocytes (below). Support for the use of monocyte-derived DCs as a model

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system came from in vitro studies by Randolph et al. (1998). This provided the first direct evidence that blood monocytes can be induced to become mature DCs in the absence of exogenous cytokines, after having migrated in an ablumenal to lumenal direction across an endothelial barrier (much like that being crossed during migration via the lymphatics in vivo). This occurs within 2 days and is most dramatic after the cells have phagocytosed particles within the subendothelial collagen layer. Furthermore, recent data from in vivo murine studies documented that a percentage of inflammatory monocytes migrating to an injected skin site take up the injected particles, carry them to the LNs, and differentiate into DCs (Randolph et al., 1999). A report from Robert et al. (1999) shows that in mice, DCs can rapidly move from blood into the peripheral tissues. These cells may be identical to the peripheral blood CD1a, CD11c DCs that were shown to be LC precursors (Ito et al., 1999). Steady-state DC migration and migration activated in inflamed tissues, are controlled by specific chemokines and their respective receptors. During DC maturation, chemokine receptor expression is modulated, thereby influencing their responsiveness to various chemokines (Sallusto and Lanzavecchia, 1999; Sozzani et al., 1999). As a result, specific chemokines have been shown to facilitate (1) DC migration to the T cell areas of lymphoid tissues; (2) efficient DC–T cell interactions; and (3) rapid activation of T cell immune responses (Sallusto and Lanzavecchia, 1999; Sozzani et al., 1999). As mentioned earlier, however, this also expedites the spread of virus by virus-carrying DCs directly to the LNs. Furthermore, chemokine receptor expression by DCs significantly influences their susceptibility to infection (Plate 35.3, below).

Replication of CCR5-using HIV-1 isolates in immature dendritic cell populations To determine the capacity of skin-derived immature DCs to capture HIV-1 in situ, Reece et al. (1998) used sheets of skin (containing

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epidermis and dermis), abraded the epidermal surface, applied CCR5-using macrophage-tropic (R5) or CXCR4-using T cell-tropic (X4) HIV-1 isolates, and cultured the skin explants for 2 days. The emigrated DCs were collected and tested for the capacity to transmit infection to activated T blasts. When R5 HIV-1 was used, DCs transmitted an infection to T cells. In contrast, DCs did not capture X4 HIV-1 applied to the abraded epidermis. Somewhat analogous findings were reported by Zaitseva et al. (1997), when they documented that CCR5 but not CXCR4 expression was detected on the surface of immature LCs isolated fresh from healthy skin. When R5 and X4 HIV-1 were added to the isolated LCs, R5 virus fused with the immature LCs, whereas X4 virus only fused after the LCs had been cultured for a short period, when CXCR4 also began to be expressed at the cell surface. As a more accessible system to compare immature and mature DCs, blood monocytederived immature and mature DCs have been examined (Delgado et al., 1998; Granelli-Piperno et al., 1998). Both blood monocytes and mature DCs are difficult to infect with R5 HIV-1, but immature DCs can be productively infected (Granelli-Piperno et al., 1998). At the level of viral DNA, blood monocytes and mature DCs only form early RU5 transcripts, while immature DCs complete reverse transcription, showing strong LTR-gag signals (Granelli-Piperno et al., 1997, 1998). Even though functional CCR5 and CXCR4 are expressed (Sozzani et al., 1997; Delgado et al., 1998), R5 HIV-1 predominantly replicates in the immature blood-derived DCs. Placing the V3 loop from the R5 HIV-1 BaL into the X4 virus also facilitates replication. However, in the presence of high doses of X4 HIV-1, replication in immature DCs has been reported (Zaitseva et al., 1997). Studies from Frank et al. (1999) revealed that virus isolated from infected immature DC cultures carries molecules expressed by the DCs, whereas virus obtained from DC-T cell cocultures displays a pattern of T cell-associated molecules on the virus surface. Therefore, while R5 virus can replicate in immature DCs,

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replication appears to occur preferentially in T cells in mixed cultures. These findings are relevant to transmission of virus in vivo, since (1) immature DCs are located in the blood and within the epithelia lining the organs exposed to HIV-1 during sexual and perinatal transmission, and (2) R5 strains are primarily transmitted during clinical HIV-1 infection (Zhu et al., 1993). Therefore, immature DCs within the blood and at the body surfaces may be one of the principal cell types contributing to this phenomenon.

Immunodeficiency virus growth in mature dendritic cell-T cell mixtures isolated from the body surfaces The inherent migratory capacity of DCs has been exploited to obtain clean mixtures of epidermal and dermal DCs and T cells (Richters et al., 1994; Pope et al., 1995a). The T cells are almost entirely of a memory phenotype (CD45RO, CD45RA/weak, CD62L), and the emigrated mature DCs express HLA-DR, CD86, p55, CD83, and DC-LAMP (Plate 35.4). Langerin expression is detected in the LC subset of the emigrated cells (Plate 35.4). Purified mature skin-derived DCs do not produce detectable HIV-1 in vitro, but the DC-CD4 T cell mixtures are permissive to both R5 and X4 HIV-1 isolates (Pope et al., 1994, 1995b, 1997b; Zaitseva et al., 1997). Even SIV-HIV chimeric viruses, SHIVs, can replicate in the skin DC-T cell milieu (Figure 35.5). Low-level infection of mature skin DCs (100 HIV-1 DNA copies per 5  104 DCs) is notable since: (1) upon interacting with memory CD4 T cells rapid virus replication ensues; (2) the infectivity of the DCs remains for up to 2 days; and (3) the ability of the HIV-1-bearing DCs to amplify infection in the presence of T cells is sensitive to AZT. Therefore, even small amounts of virus associated with mature DCs could contribute to HIV-1 replication in vivo. The CD4 T cells that interact with the DCs to support viral replication in vitro can be syngeneic and do not need to proliferate (GranelliPiperno et al., 1995; Pope et al., 1994, 1995b). No

FIGURE 35.5 Replication of SHIV 162 in cutaneous dendritic cell-T cell mixtures. Emigrated skin DC-T cell mixtures were pulsed with SHIV 162 with the equivalent of 5, 1 or 0.2 ng/ml of SIV p27 per 105 cells for 1.5 h. The cells were washed and the cultures monitored for virus replication over the next 9 days. Every other day, supernatants were collected, frozen at 20C, before reverse transcriptase (RTase) activity was measured (Cameron et al., 1992). Results are expressed as the mean counts per minute (c.p.m.), of RTase activity per µl of culture supernatant. SHIV 162 was kindly provided by Drs Leonidas Stamatatos and Cecilia Cheng-Mayer of the Aaron Diamond AIDS Research Center, Rockefeller University, New York.

mitogens or exogenous cytokines need to be added to the cultures, and cell staining with the Ki-67 cell cycle antigen is infrequent. However, skin T cells are memory cells that express the activation antigen CD69. As expected, HIV-1 infection of the skin DC-T cell mixtures is also dependent on CD4, since the blocking anti-CD4 mAbs 5A8 and 6H10, but not the non-blocking mAb L120, prevent HIV-1 infection (Figure 35.6A). Specific chemokines have also been shown to interrupt infection with HIV-1 isolates (Granelli-Piperno et al., 1996; Ayehunie et al., 1997; Rubbert et al., 1998). In addition, HIV-1 infection can be blocked by nonspecific agents like sulfated polysaccharides such as dextran sulfate (Figure 35.6B), which likely act by preventing HIV-1 adsorption to the cell surface or by interfering with the positively

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FIGURE 35.6 Anti-CD4 monoclonal antibodies and sulfated polysaccharides inhibit HIV infection of dendritic cell-T cell mixtures. Skin DC-T cell mixtures were pre-incubated (30 min at 37C) with (A) 10 µg/ml of the antiCD4 mAbs 5A8, L120, or 6H10 (Moore et al., 1992; and M. Pope, A. Trkola, and J.P. Moore, unpublished observations) or (B) 10 µg/ml dextran (D) or dextran sulfate (DS) (M. Pope, R. Pearce-Pratt, and D. Phillips, unpublished observations). Untreated cells (Med.) and cells pretreated with 10 µM AZT (AZT) were included as internal controls. The cells were then pulsed with 100 TCID50 of the subtype E isolate of HIV-1, 92TH009, for 90 min in the presence or absence of the specific reagents. Washed cells were cultured for 36 h in the presence of the indicated reagent before the DNA was extracted for PCR analysis (Pope et al., 1994). PCR for the HIV gag sequence was performed to document infection and the HLA-DQ sequence amplified to verify the integrity of the samples. A standard curve of controls was included to estimate the virus copy number;   0; 0  100, 1  101, 2  102, 3  103, 4  104 HIV copies.

charged V3 domain of the virus envelope gp120 region (Lederman et al., 1989; Callahan et al., 1991). In the monkey, it has been possible to obtain DCs and T cells from different sites (O’Doherty et al., 1997; Pope et al., 1997a; Ignatius et al., 1998) and thus appreciate the extent to which the findings in the skin system can be extended to other tissues. SIV-permissive mixtures of mature DCs and T cells have been isolated from the skin and vaginal and nasopharyngeal epithelium of healthy macaques (Pope et al., 1997a). Both dual-tropic SIVmac251 and T cell-tropic SIVmac239 replicate in these leukocyte mixtures. Comparable observations were also made using DC-T cell mixtures isolated from the mucosae of the ectocervix or vagina of women undergoing hysterectomy or vaginal reconstructive surgery (Hladik et al., 1999), where significant in vitro replication of several HIV-1 isolates has been detected. Mature blood-derived macaque DCs also collaborate with CD4 T cells to drive SIV replication (Ignatius et al., 1998). Furthermore, the ability of DCs pulsed with SIV to transmit infection to CD4 T cells is preserved for a day

after exposure to the virus (Figure 35.7). The propensity for DCs to promote virus replication is not dependent on the origin of the cells, because T cells from either the skin, blood, spleen or LN support SIV replication in the presence of mature DCs derived from the skin or blood (Ignatius et al., 1998). Interestingly, the CD45RA memory cells are much more permissive when cultured with immature DCs than are the CD45RA naïve populations (MP and RI, unpublished observations). Similar to human skin DC-T cell cultures, macaque blood-derived DC-T cell mixtures sustain replication of SHIV chimeric viruses, SHIV89.6 and SHIV89.6P (Figure 35.8). SHIVs comprise an SIV backbone with an HIV-1 envelope, are infectious in vivo (Li et al., 1995), and are, therefore, useful to study HIV-1 envelopespecific reagents. For example (Figure 35.8), the ability of SHIV-pulsed macaque DCs to transmit infection to autologous T cells was blocked in the presence of the neutralizing anti-HIV envelope mAbs 2F5 and 2G12. Identical observations have been reported for blocking the transmission of HIV infection from human blood and skin-derived DCs to T cells (Frankel et al., 1998).

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FIGURE 35.7 Infectious SIV is retained on macaque dendritic cells for up to a day after exposure to the virus. Purified mature DCs (obtained from the blood of three healthy macaques; AR 82, K080 and 6H8) were pulsed with SIVmac251 (800 TCID50 per 105 cells), washed, and cultured immediately or 17–20 h later with the permissive cell line, CEM174 (1 DC per 10 CEM174 cells). After the 17–20 h culture, the DCs were washed again prior to coculture. Productive infection was evaluated by measuring the release of SIV p27 (ng/ml) into the culture supernatants by ELISA.

In vivo evidence for dendritic cell infection within the mucosae

FIGURE 35.8 Anti-HIV-1 envelope monoclonal antibodies inhibit transmission of SHIV from dendritic cells to T cells. As described for the human system (Frankel et al., 1998), mature macaque DCs were pulsed with SHIV 89.6 (1000 TCID50 per 105 cells) or SHIV 89.6P (100 TCID50 per 105 cells), washed and cultured overnight. After culture, the cells were washed again and pre-incubated with medium (No Ig), human Ig (HIg), or the neutralizing anti-HIV-1 envelope Abs 2F5 and 2G12 (Trkola et al., 1995) at 25 µg/ml for 30 min. Autologous CD4 T cells were added at a ratio of 1 DC per 10 T cells with additional Abs (or not) to maintain the 25 µg/ml concentration. Supernatants were collected for the next 11 days and monitored for the presence of SIV p27.

Therefore, the DC-T cell systems in the human and macaque represent biologically relevant models to dissect the requirements for virus replication and how this could influence virus transmission in vivo.

Because DCs are located at the surface epithelia they are in the prime position to encounter virus that breaches the mucosa during the earliest stages of infection. Furthermore, the powerful capacity of DCs to promote virus growth upon interaction with CD4 T cells, implicate them as a critical factor in the onset and spread of infection. Due to the difficulties in accessing sufficient mucosal tissue samples from HIV-1infected individuals, considerable work has been carried out using the SIV-macaque model. Virus can be applied to the mucosa in question and samples collected at controlled times after exposure. Earlier studies provided indirect evidence that DCs within the vaginal mucosa are one of the first cells infected by SIV (Spira et al., 1996). Rare SIV-positive cells with dendritic morphology can be seen within and underlying the mucosal epithelia just days after vaginal application of the virus. However, definitive documentation that these cells are DCs or LCs, has not been provided. One of the limiting factors for DC identification has been the availability of DC-specific reagents. However, new anti-DC reagents that recognise monkey cells are now available (e.g. DC-LAMP, Langerin, DEC-205) that will advance the identification of DCs in situ.

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More recent studies examining acute infection across the mucosal epithelia suggest that CD4 T cells are in fact the major source of infectious virus during the acute stages of infection. Specifically, Zhang et al. (1999) documented SIV RNA-positive CD4 T cells in the underlying mucosal tissue within days of vaginal infection with SIV. CD4 T cells were also the site of most virus production in the first days after infection across the tonsillar mucosa (Stahl-Hennig et al., 1999). These data suggest that once the infection is established, the predominant source of new virus production is the CD4 T cells. However, in the first days after inoculation the virus may be trapped by cells within the mucosa (e.g. DCs, monocytes, epithelial cells) which do not produce significant (easily detectable) amounts of virus. These cells could subsequently transmit infection to the more permissive CD4 T cells in which the virus is amplified (and hence detected). This scenario is supported by a recent report from Hu et al., that SIV RNA-positive LCs can be detected in the mucosal epithelia of macaques within 18 hours of vaginal application of virus (Hu et al., 2000). Similar studies have been performed on animals chronically infected with SIV. Previously, SIV-positive cells had been identified within the uterus, cervix and vagina of chronically infected animals, being most numerous in the submucosal regions of the ectocervix and vagina (Miller et al., 1992; Miller, 1998) and more infrequent in the SSE (Miller et al., 1992). Conclusive identification of SIV-positive DCs isolated from the mucosal epithelia of the reproductive tract was recently documented (Hu et al., 1998; Miller and Hu, 1999). Relatively few SIV-infected DCs are detected, compared to the numbers of infected CD4 T cells, especially within lymphoid compartments. SIV protein-positive DCs are not seen, which may reflect a general paucity of infected DCs or that actively infected DCs have died. In chronically infected male macaques, numerous SIV-positive (T cells and macrophages) are found throughout the reproductive tract, while infected cells (possibly LCs) are rarely detected within the SSE of the foreskin (Miller et al., 1994). Therefore, DCs may have a

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greater role in retaining virus and then spreading it to the more permissive populations of CD4 T cells, rather than producing large amounts of virus themselves (below).

Summary From extensive in vitro studies examining skinand blood-derived immature and mature DCs as models for the DCs within the mucosae, it is apparent that DCs efficiently advance HIV-1 and SIV replication in collaboration with CD4 T cells. Immature DCs may capture virus more efficiently than mature DCs and can replicate the virus in vitro, however rapid virus expansion occurs when the DCs transmit it to T cells. In vivo observations support the notion that DCs TABLE 35.2 surfaces

Immunodeficiency virus at the mucosal

Acute SIV infection Possible SIV-positive DCs in submucosa of genital tract mucosae Active replication of SIV in CD4 T cells Chronic SIV infection Rare SIV-positive DCs, numerous SIV-infected CD4 T cells genital tract epithelia nasopharyngeal mucosal tissue

could capture virus crossing the mucosae and subsequently carry it to serve as a source of virus to be amplified most effectively by surrounding CD4 T cells within the local tissues or in the draining lymphoid compartments (Table 35.2).

DENDRITIC CELLS AND VIRUS LOAD IN PERIPHERAL LYMPH NODES Dendritic cell subsets in the T and B cell areas LNs are a major depot for viral RNA and gag protein (Biberfeld et al., 1985; Tenner-Racz et al., 1986, 1988; Schuurman et al., 1988). To

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Dendritic cell subsets in the lymph

B cell areas FDCs CD45 Retain immune-complexed antigens Present native antigens to B cells GCDCs CD45 Bind immune-complexed antigens Stimulate T and B cells T cell areas DCs (DC1s) CD45 Retain antigens as MHC-peptide complexes Present processed antigens to T cells (TH1) pDC2s/DC2s CD45 Localize around HEVs of inflamed lymph nodes Secrete type I IFNs in response to enveloped viruses Activate T cells

understand these findings, one must appreciate that there are distinct classes of DCs in lymphoid organs (Table 35.3), which are critical for immune activation and, therefore, impact virus replication. Follicular dendritic cells (FDCs) located in the B cell areas are a mesenchymal type of cells that retain antigens in a native form, as extracellular immune complexes bound to Fc and C3 receptors. In contrast, bone marrowderived DCs, predominantly in the T areas retain processed antigens as MHC–peptide complexes on their cell surfaces. The dramatic distinction between the types of DCs in the B and T cell areas changed with the report from Grouard et al. (1996) of a leukocytetype DC population within the germinal centers (GCs) of stimulated or secondary B cell follicles. These GCDCs are CD11c, CD3, CD4, large, irregularly shaped cells. Isolated GCDCs are not fully mature, but develop high levels of costimulatory molecules within a day of culture. Interestingly, GCDCs also express surface κ and λ light chains, indicating a capacity to carry immune complexes. Most notably, GCDCs are powerful stimulators of allogeneic T cells, unlike GC B cells, and provide critical signals for the GC reaction of B cell expansion, plasma cell differ-

entiation, and isotype switching (Dubois et al., 1999). Another level of complexity to the DC system has been added with the identification of a new DC precursor, originally termed the plasmacytoid cell (Grouard et al., 1997). These precursors, also called pDC2, are CD11c, CD123 (IL-3Rα), CD4, lineage marker negative, express the immunoglobulin-like transcript receptor (ILT3), but lack ILT1 and are present in blood and inflamed lymphoid tissues (Cella et al., 1999; Siegal et al., 1999). Distinct from myeloid DCs (DC1s which are CD11c, CD123/, ILT1, ILT3), pDC2s are responsive to IL-3 and secrete large amounts of IFN-α following exposure to enveloped viruses (Siegal et al., 1999). Expressing CXCR3 and CD62L, pDC2s can respond to inflamed tissue signals and migrate across the high endothelial venules (HEVs) within the LNs (Cella et al., 1999). In response to IL-3 and CD40L stimulation, pDC2s develop into mature DC2s and activate T cells in vitro, preferentially eliciting TH2-like responses, compared to the TH1 responses induced by the myeloid DC1s (Rissoan et al., 1999). Ongoing studies will reveal what this DC subpopulation contributes to HIV-1 pathogenesis.

Acute SIV infection of the T cell areas of the lymph node During acute infection in the SIV-macaque model, wild-type virus replicates chiefly within the T cell areas of the lymphoid tissue (Plate 35.9) (Reimann et al., 1994). The cells that show strong in situ hybridization (ISH) signals for SIV RNA are primarily T cells, since most do not double label for macrophage and DC markers. Similar observations were recently made in the draining LNs of animals infected with virus: (1) via tonsillar application of small amounts of pathogenic SIV (Stahl-Hennig et al., 1999) or attenuated SIV (in preparation); and (2) following i.v. infection with attenuated SIV (in preparation). Therefore, it appears that once an infection is founded, irrespective of the route of exposure and the virulence of the SIV strain

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used, acute virus growth occurs within the T cell areas of the LNs. Possibleexplanationsfortheprominentgrowth of virus within the CD4 T cell compartment include: (1) infection of activated T cells by cellfree virus within the LN; or (2) transmission of virus to the T cell areas by virus-carrying DCs. The former seems improbable, since cell-free virus is more likely to encounter resting, nonpermissive T cells than activated ones. More efficient virus spread would occur between virus-carrying DCs and CD4 T cells. Transmission would be further amplified when the CD4 T cells are being activated by the virus-bearing DCs (below).

Infection of the B cell areas of lymph nodes during chronic infection with HIV-1 and SIV A very different picture is seen in the LNs of those chronically infected with HIV-1 or SIV (Plate 35.9). The cells with the most abundant viral RNA are found in the B cell areas, especially the GCs. Viral RNA is found in two predominant sites when detected by ISH (Schuurman et al., 1988; Tenner-Racz et al., 1988). One consists of a diffuse labeling over the FDC network, which likely represents Ab–virion complexes bound to the FDC surface (Tenner-Racz et al., 1988). The other location of viral RNA comprises mainly CD4 T cells that label strongly for viral RNA. The bulk of the virus identified within the GCs, is believed to represent virus that is trapped via immune complexes on the surface of the FDCs and possibly also the more recently described GCDCs, which can also bind Abcomplexed substances. Interestingly, recent work revealed that in patients taking antiretroviral therapies whose plasma viremia was dramatically reduced, the high levels of virus localized to the B cell areas of their LNs also drastically decreased (Cavert et al., 1997; Wong et al., 1997). This indicates that the virions trapped within the LN FDC network and possibly on the GCDCs are rapidly and continuously replaced during infection. Even at end-stage disease, CD4 T cells are a major source of HIV-1 production (van der Endeet

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al., 1999). Transmission of infection to the GC T cells may occur via two mechanisms: (1) Infection of GC T cells by the virus trapped on FDCs, since there is evidence that FDC-associated virus can be infectious (Heath et al., 1995); (2) GCDCcaptured virus could be transmitted to CD4 T cells that are being activated by the antigen-presenting GCDCs. The latter seems a more effective way of transmitting virus to the GC T cells considering these cells interact with activated T cells, whereas FDCs are not known to do so.

Augmented virus growth when virus is transmitted from dendritic cells to antigen-responding T cells Early studies by Cameron et al. (1992, 1994) provided a model for what might happen when DCs capture virus at the periphery, and subsequently migrate as maturing DCs to the LN T cell areas to present viral antigens and elicit antiviral immunity. Specifically, when mature DCs were pulsed with HIV-1 and then used to present superantigen or alloantigen, the virus only replicated when T cells were added. This was also reported by Weissman et al. (1996), when DCs presented tetanus toxoid to reactive T cells. In such stimulated cultures, the DC:T cell

FIGURE 35.10 Active SIV replication in allogeneic dendritic cell-T cell co-cultures. Purified mature DCs were generated from the blood monocytes of a healthy macaque and pulsed with SIVmac251 (800 TCID50 per 105 cells) for 1.5 h at 37C. The cells were then washed and cultured alone (DC-SIV) or mixed with syngeneic or allogeneic CD4 T cells (DC-SIV  T) at various DC:T cell ratios. Culture supernatants were assessed for the release of SIV p27 over the ensuing 2 weeks.

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ratio was small (1:30–1:100), relative to that used in syngeneic, antigen-independent mixtures (1:3–1:10). Related observations have been made in the SIV-macaque system, where strong SIV replication ensues in allogeneic mixtures of DCs and T cells even at lower DC numbers (Figure 35.10). When HIV-1-pulsed DCs stimulated T cell activation, the subsequent productive infection occurred primarily in the responding T cells, i.e. most of the viral gag protein was localized in MHC class II-weak T cells rather than in MHC class II-strong DCs (Cameron et al., 1992, 1994). Therefore, it is likely that during acute infection with HIV-1 and SIV, virus-carrying DCs that reach the LN would transmit virus most effectively to responding virus-specific CD4 T cells in the T cell areas.

Virus transmission from dendritic cells to T cells: dendritic cell entrapment versus infection In the preceding sections we have summarized the capacity of DCs to transfer HIV-1 and SIV to T cells, and described that this may happen in both the acute and chronic phases of virus replication in the LNs. Such transmission is enhanced during antigen presentation, but readily occurs in the ostensible absence of foreign antigens. Does the DC simply trap virus, or must the virus enter the DC prior to transmission and active replication in the T cells? Initial studies indicated that the ability of virus-pulsed skin-derived DCs to transmit infection to autologous T cells is sensitive to treatment with AZT (Pope et al., 1995b). This suggested that a low-level infection of the mature DCs, not detected as a productive infection, is required for the subsequent transmission and amplification of virus in the T cells. This observation was verified in more recent studies. DCs obtained from individuals carrying the ∆32 mutation in CCR5 are unable to be infected with R5 HIV-1 isolates and do not transmit infection to T cells (Granelli-Piperno et al., 1998). Similarly, normal DCs pulsed with R5 HIV-1 cannot transmit infection to ∆32 mutant T cells.

Therefore, these studies suggest that both the DCs and T cells must express functional CCR5 for infection with R5 HIV-1 isolates to occur in the DC-T cell co-cultures. As expected, X4 HIV-1 replication occurs in the presence of DCs and T cells carrying the ∆32 CCR5 mutation. Using VSV-G pseudotyped HIV-1 that undergoes only one round of replication, mature DCs were found to replicate virus following signaling via CD40L (Granelli-Piperno et al., 1999). Together these data confirm that DCs can capture virus, replicate at least low levels, and upon interaction with CD40L expressed by T cells amplify infection by transmitting virus to CD4 T cells (Plate 35.3). The HIV-1 life cycle is blocked in mature DCs (Granelli-Piperno et al., 1997). R5 and X4 viruses can enter the DCs via CD4 and chemokine receptors, yet after reverse transcription is initiated, the majority is not completed (Pope et al., 1994, 1995b; Granelli-Piperno et al., 1996, 1998). It is possible that, after entering via the chemokine receptors, a small proportion of the virus may reverse transcribe while most remains in an uncoated state close to the DC surface. Upon T cell contact, the virus-infected DC then directs the virus to the T cell (Plate 35.3). The identification of a novel molecule, DCSIGN (Geijtenbeek et al., 2000a), that is selectively expressed by DCs and can bind HIV-1, reveals a unique infection-independent mechanism through which DCs can amplify virus infection (Plate 35.11). DC-SIGN, which is identical to the previously described high affinity HIV-1 gp120 binding C-type lectin (Curtis et al., 1992), is strongly expressed by blood-derived immature DCs and by DCs within the mucosal tissues, skin and LN (Geijtenbeek et al., 2000b). Geijtenbeek et al. demonstrated that DCs bind HIV-1 via DCSIGN (independent of CD4 and CCR5) and efficiently facilitate infection in surrounding CD4 CCR5 Tcells.DC-SIGN-boundvirussignificantly enhances infection of permissive T cells in the presence of extremely small amounts of virus, at a dose that would otherwise not infect the T cells. Furthermore, DC-SIGN-expressing transfectants (unable to be infected themselves) pulsed with HIV-1 preserve their ability to transmit infection

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to activated T cells for up to 4 days after exposure to the virus, indicating the longevity of DC-SIGNcaptured HIV. The localization of DC-SIGNexpressing DCs at the body surfaces, implies that virus crossing the mucosae could be efficiently trapped by DC-SIGN, not necessarily resulting in DC infection. However, it is important to note that it is the submucosal DCs that express DCSIGN and LCs in the outer epithelium lack DCSIGN. Therefore, if LCs in the epithelia do indeed capture the first virus breaching the mucosa (Hu et al., 2000), receptors other than DC-SIGN must be involved here. Once the virus reaches the submucosal region, either via LC-borne transmission or directly through a damaged mucosal surface, then the DC-SIGN-bearing DCs may play an important role. Entrapped virus could then be rapidly transmitted to nearby CD4 T cells within the mucosae or the draining LNs (after migration) to augment virus replication in the CD4 T cell compartments. Using a mouse model, Masurier et al. (1998) have documented that HIV-pulsed DCs are rapidly routed to the LNs following vaginal or intravenous administration. This suggests that even in the absence of infection DCs can readily direct infectious virus to the lymphoid tissues.

Summary HIV-1 and SIV continuously replicate in the LNs (Plate 35.9). During acute infection, replication is predominantly in the T cell areas, while in chronic infection, actively infected cells are most prominent in the B cell areas. In each case, it is possible that virus is efficiently transmitted to the CD4 T cells via DCs, and that the interactions between DCs and T cells drive replication in the latter, particularly when the T cells are responding to antigenic stimulation. It is fascinating to note that, although mature DCs capture virus they do not readily replicate virus, but effectively transmit infection to T cells (Plate 35.3). The intriguing studies documenting the expression of DC-SIGN, a molecule that efficiently entraps HIV-1 on the DC in the absence of viral infection, and then transmits infection to surrounding CD4 T cells provides a

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mechanism through which virus-carrying, but noninfected DCs can dramatically intensify virus spread (Plate 35.11).

IMMUNODEFICIENCY VIRUSES IN THE MUCOSALASSOCIATED LYMPHOID TISSUES Dendritic cells and the mucosal-associated lymphoid tissue MALT is abundant in the gut, including the oral and rectal mucosa, and is, therefore, potentially targeted by HIV-1 following sexual and perinatal exposure. MALT exhibits many principles of a LN with B and T cell areas and the major subtypes of DCs: FDCs and GCDCs within the GCs and DCs within the T cell areas (Plate 35.12). One of the major distinctions of the MALT is the presence of a specialized pathway for antigen uptake and delivery to the underlying lymphoid tissues that utilizes M cells within the apical or dome epithelium (Neutra, 1999). Light and electron microscopy also show lymphocytes forming a ‘lymphoepithelium’ (LE). The specialized antigen-transporting LE of the MALT is magnified in the tonsils at the upper end of the GI tract (Plate 35.12). These include the adenoids or nasopharyngeal tonsils at the back of the nose, the palatine tonsils at the back of the throat, and the sublingual tonsils in the rear of the tongue. The tonsil surfaces invaginate to form deep crypts, along which the epithelium is very different from that seen at the remaining tonsil or pharyngeal surfaces. Most of the pharyngeal surface is lined by a SSE, in which there are DCs, but few lymphocytes. In the crypts, however, the epithelium becomes much thinner and more ramified, there are antigentransporting M cells, DCs and many more lymphocytes. DCs have been noted lying beneath the M cells of the MALT epithelia (Iwasaki and Kelsall, 1999). In the human (Frankel et al., 1996, 1997) and monkey (Hu et al., 1999; Stahl-Hennig et al., 1999), DCs can be identified within and just

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below the LE. Because DCs within the LE are constantly exposed to activation stimuli, and because lymphocytes seem to traffic continuously through this region, a cellular environment is apparent that resembles the leukocytes that emigrate from skin explants and are so permissive to HIV-1 and SIV.

Chronic virus production at the surfaces of the mucosal-associated lymphoid tissue of the tonsils Studies by Frankel et al. (1996, 1997) documented the presence of virus-producing syncytia within and just beneath the tonsillar LE of HIV1-infected individuals. The only infectious agent detected within these tissues was HIV-1, which was evident within large stellate cells at the MALT surface when the tissue was stained for HIV-1 gag protein. Specifically, HIV-1-infected cells were only found associated with the LE and not the SSE (Frankel et al., 1997). Further examination revealed that the HIV-1infected cells were multinucleated and could be labeled for antigens expressed by DCs, S100 and p55, suggesting that DCs were involved in the formation of these virus-producing syncytia. In a separate study, HIV-producing syncytia staining for macrophage (CD68 and HAM56) and DC (p55 and S100) markers were identified in pediatric and adult tonsil specimens (Orenstein and Wahl, 1999). Furthermore, infected giant cells containing DCs are quickly produced and abundant when HIV-1 (Pope et al., 1994; Frankel et al., 1997) or SIV (Pope et al., 1997a; Ignatius et al., 1998) is added to DC-T cell co-cultures. SIV-positive cells within the tonsillar MALT of chronically infected monkeys have been identified, however multinucleated syncytia were infrequent (Hu et al., 1999). The majority of cells producing virus were CD4 T cells surrounded by numerous mature DCs. Rare SIV-positive DCs were identified. It is possible that a lack of syncytia reflects that the tonsils were not enlarged (unlike the human studies), suggesting that an underlying stimulation may be critical for the amplified replication of virus within syncytia in these tissues.

At mucosal surfaces, the DCs are probably being activated continuously by stimuli entering from the environment via the M cells, and these DCs have ready access to memory T cells. Therefore, the cellular milieu resembles that described for the permissive DC-T cell mixtures that are isolated from the body surfaces, creating a locale in which virus can propagate on a chronic basis.

Rapid spread of acute SIV infection within the mucosal-associated lymphoid tissue Infection across the oral mucosa is a potential mechanism of transmission when exposed to the virus perinatally or via oral sex (Ruprecht et al., 1999). Previous studies have reported the infection of neonate and adult macaques via the oral route (Baba et al., 1999). However, the exact site of virus entry is difficult to pinpoint. Depositing approximately 1 ml of virus into their mouths, infected animals and therefore, virus could have accessed the mucosa of the oral cavity but also mucosae along the gut once the volume was swallowed. A recent report documented the ability of virus transmission to occur across the tonsillar epithelium (Stahl-Hennig et al., 1999) (Plate 35.13). Stahl-Hennig et al. demonstrated that atraumatic application of SIV directly on to the palatine tonsils of macaques resulted in their infection. During this acute stage virus-infected cells were not detected until 2–3 days post infection at which point numerous virus-producing cells were identified within the T cell areas of the MALT. Virus continued to replicate and spread to the draining and ultimately distal lymphoid compartments and the blood. CD4 T cells appeared to be the major origin of virus production within the tissues. Therefore, virus readily crossed the tonsillar epithelia, but remained undetected in the first days after transmission, where it might have been trapped by leukocytes (DCs, macrophages), until it was amplified by the CD4 T cells and spread to the neighboring tissues (Plate 35.13). Similarly, one of two newborn macaques exposed to only 128 TCID50 of SIVmac251 applied directly to the palatine tonsils in 8 

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TABLE 35.4 Virus isolation from the lymph nodes of a monkey infected via the tonsils Days of co-culture

3

7

12

Lymph node Posterior mesenteric Anterior mesenteric Iliac Inguinal Axillary

0.31 0.24 0.15 0.63 0.24

109.6 82.4 76.4 77.5 74.7

98.2 108.8 97.3 102.6 90.4

LNs were obtained at necropsy approximately 6 months after tonsillar exposure to 128 TCID50 of SIVmac251. Isolated LN cells were co-cultured with the permissive T cell line, CEMx174, and the supernatants measured for the presence of SIV p27 protein by ELISA. SIV p27 levels (ng/ml) are shown after 3, 7 and 12 days of co-culture.

2.5 µl volumes rapidly became infected (M.P. and R.I., unpublished observations). Disease progressed quickly and the animal had to be euthanized within 6 months of tonsillar infection. At necropsy, infectious virus was readily detected in the LNs (Table 35.4). Hence, transmission of small amounts of virus across the tonsillar mucosa results in a rapid dissemination of virus to promptly establish systemic infection in adult and newborn monkeys. Extensive replication of SIV within the T cells of the gut MALT has also been reported (Veazey et al., 1998, 2000). As a result of the vast growth a large percentage of the memory CD4 T cells within the MALT were destroyed. Furthermore, selective loss of the CCR5-expressing memory T cells of the intestinal mucosa was also reported in animals infected with the R5 SHIVSF162P, but not the X4 SHIVSF33A (Harouse et al., 1999). Thus, the milieu of DCs and memory T cells within the MALT facilitates the expeditious replication of virus in vivo, leading to the destruction of the memory CD4 T cell pool.

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from stimuli crossing the LE that contains both DCs and T cells. This DC-T cell environment is supportive of virus replication during all stages of infection. Viruses transitting these mucosal tissues may get trapped by DC-SIGN-bearing DCs (Plate 35.11) and efficiently transmitted to surrounding memory CD4 T cells.

IMMUNITY TO IMMUNODEFICIENCY VIRUSES AND THE POTENTIAL ROLE OF DENDRITIC CELLS In the previous sections, consideration was given to the role of DCs in the transmission and replication of immunodeficiency viruses. However, DCs most likely also have important roles in immune activation against viral infection, particularly in the stimulation of virus-specific CD4 and CD8 T cells. Immunodeficiency virus infections represent a battleground in which the virus replicates continuously, yet the levels of virus and CD4 T cells can remain steady for long periods of time. This ‘stalemate’ may depend on the fact that DCs are driving virus replication within the CD4 T cell pool, while eliciting antiviral resistance via activation of CD8 T cells (Plate 35.1). In fact, virus-specific CD8 T cells could ultimately eliminate virus-carrying DCs, thereby diminishing the availability of potent antigen-presenting DCs to maintain and expand immunity. Therefore, understanding the requirements for virus replication versus immune activation in the DC environment is critical to advance our knowledge of the regulation of this disease, to allow us to develop antiviral strategies to prevent or clear infection.

Summary Significant virus replication within the MALT has been described at the acute and chronic stages of infection with HIV-1 and SIV. In addition to the peripheral LNs, the MALT may be particularly important because of the content of typical B and T cell areas, as well as the chronic activation

Dendritic cells mediate resistance to HIV-1, especially at the level of CD8 T cells There are potentially numerous ways in which DCs can capture viral antigens and induce the strong CD8 T cell responses that can be

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observed in HIV-1 infection. The first is by direct infection, particularly at the mucosal surfaces (above). However, there could be other pathways that do not require infection of the DCs. In influenza, DCs can present nonreplicating virus (Bender et al., 1995), and there is an abundance of noninfectious virus in the plasma of HIV-1-infected individuals (Piatak et al., 1993). DCs can also present antigens derived from apoptotic cells (Albert et al., 1998). In HIV1 infection, many infected T cells die and possibly these are taken up and presented by DCs. Recent work also documented the efficient uptake and presentation of immune-complexed antigens by DCs, but not by macrophages or B cells (Regnault et al., 1999; Rodriguez et al., 1999). Immune-complexed HIV-1 is abundant in vivo and if picked up by DCs, this should result in the efficient induction of T-cell responses. Such a mechanism may be most efficient if noninfectious, immune-complexed virions are encountered, but also may be effective in directing infectious particles to antigen-processing pathways. These presenting pathways, especially those involving noninfectious virus and dying infected cells, would allow DCs to present HIV-1 for immune activation, but one would not necessarily detect active infection of DCs. The identification of vaccine strategies for HIV-1 is one of the highest priorities. One approach is genetic immunization with DNA vaccines (Leitner et al., 1999). It has been shown that DCs present the antigens encoded by DNA vaccines and can even carry small amounts of vaccine DNA (Casares et al., 1997; Akbari et al., 1999). This suggests that if DNA targeting to the DC could be amplified during vaccination, stronger antiviral responses might be targeted. The inclusion of CpG motifs into the constructs may further enhance the effectiveness of DNA vaccines by activating DCs (Hartmann et al., 1999; Jakob et al., 1999). Other encouraging strategies under investigation include the use of viral vectors (Rolls et al., 1994; Zhou et al., 1994; Caley et al., 1997; Tubulekas et al., 1997; Fang et al., 1999) such as poxviruses, adenovirus, alphaviruses, and VSV-pseudotyped virus to

directly target the DC system for the induction of potent immunity. Work is also being initiated in the macaque system where one can readily test potential vaccine strategies for their ability to induce virus-specific immunity, the ultimate test being the outcome of virus challenge. Therapeutic applications of DCs in infected individuals can also be examined. Recent work has revealed that DCs transfected with SIV gag-expressing adenovirus can activate SIV-specific T cells in vitro (Zhong et al., 2000). Approaches using DNA vaccines, various recombinant viral vectors, inactivated and attenuated viruses are being examined. How such strategies interact with the DC system will be critical to understand.

Dendritic cell involvement in attenuated vaccines One potential vaccine strategy is the use of live attenuated virus. A live attenuated form of SIV, containing a deletion in the nef gene (SIV delta nef ) was initially proposed as a potential vaccine, protecting against subsequent challenge with wild-type SIV (Daniel et al., 1992). Even though the nef-defective SIV is not completely protective and is associated with the risk of disease development, this attenuated virus does induce immunity (Gauduin et al., 1999) that initially controls infection and significantly retards the development of disease. SIVspecific T cells are readily detected in the peripheral blood and LNs of these animals (in preparation) (Gauduin et al., 1999; Zhong et al., 2000). However, one of the major obstacles facing HIV vaccine researchers is knowing what immune responses are required to prevent the onset of infection and to clear an established infection. Understanding the pathology of an attenuated virus may aid in these investigations. It is possible that, unlike the wild-type SIV (above), the nef-defective SIV would not replicate well in the DC-T cell milieu, but replicate sufficiently to provide viral antigens for the DCs to present to activate the T cells in the absence of overwhelm-

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in vivo, in which we can begin to dissect the requirements for virus growth versus immune stimulation. Such information will help us understand the factors that influence virus transmission, disease development, and the subsequent immunity induced.

FIGURE 35.14 Defective replication of SIV delta nef in immature macaque dendritic cell-T cell cocultures. Mixtures of immature DCs and autologous CD4 T cells (1 DC:10 T cells) were exposed to either SIVmac239 wild-type (wt) or SIVmac239 delta nef (delta) (5  103 TCID50 per 105 cells). Infection was monitored over the following 2 weeks. The peak SIV p27 production values (ng/ml) for 14 separate experiments are shown.

ing virus replication. We recently completed a study that revealed that SIV delta nef exhibits significantly impaired replicative capacity in the immature DC-CD4 T cell milieu (Figure 35.14) (Messmer et al., 2000). In contrast, SIV delta nef grows normally in the presence of mature DCs and T cells, comparable to the replication of the wild-type virus in the presence of either immature or mature DCs. Although, nef-deficient virus replicates if the cultures are activated with a superantigen, the ability of the wild-type virus to grow in the immature DC-T cell environment cannot be attributed to activation of either the DCs or the T cells as assessed at the phenotypic level. Therefore, the low level replication of attenuated SIV in the presence of immature DCs may reflect a situation that occurs in vivo, where the immune system is stimulated and not immediately destroyed by vigorous virus replication, as seen with wild-type virus (Plate 35.15). The signals provided by mature DCs or by nef in the immature DC environment, which facilitate virus replication, are under investigation. There is clearly a fine balance between virus replication and immune destruction versus immune activation. This immature DC-CD4 T cell coculture represents an in vitro primary cell system, much like what the virus will encounter

A SYNOPSIS OF HOW DENDRITIC CELLS CONTRIBUTE TO HIV-1 DISEASE This chapter has concentrated on some of the more recent literature on the potential roles of DCs in driving the replication of HIV-1 and SIV. Herein, we divided the discoveries into anatomical compartments, and tried to relate the in vitro observations to the morphological studies of the DC–virus interactions in situ and how these contribute to virus replication and antiviral immunity. The mechanisms employed by DCs to promote virus replication may vary at different tissue sites (Table 35.5). At the body surfaces, TABLE 35.5 genesis

Dendritic cells in HIV-1/SIV patho-

Infection Body surfaces Immature migrating DCs capture and replicate R5 HIV-1 Possible transmission to local CD4 T cells Lymph nodes In acute infection, virus-carrying DCs circulating from the blood and body surfaces transmit virus to CD4 T cells, especially HIV-specific T cells In chronic infection, GCDCs (and FDCs) entrap immune-complexed virus and transmit infection to CD4 T cells Mucosal surfaces of the MALT Activated DCs within the LE can drive virus replication and transmit infection to memory CD4 T cells Immunity DCs process viral antigens captured via infectious or noninfectious mechanisms Antigen-bearing DCs activate CD4 and CD8T cells and B cell responses

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immature DCs may effectively capture and replicate virus. During acute infection, DCs could transmit virus to T cells, especially those T cells that are responding to viral antigens, locally and in the T cell areas of the lymphoid tissues. In chronic infection, while DCs can carry virus, the bulk of the virus is probably produced as a result of DCs interacting with T cells in the GCs of the lymphoid organs and in the LE of the MALT. The report that DCs can capture HIV via a newly identified molecule, DC-SIGN, and efficiently transmit infection to CD4 T cells, provides a mechanism whereby noninfected, viruscarrying DCs can also amplify virus infection within the CD4 T cell pool. Therefore, even with different types of immunodeficiency virus, DCs could be a driving force to catalyze virus replication in several pertinent anatomical compartments. In addition, CD4 T cells die during their infection mediated by DCs, thereby resulting in the critical loss of helper T cells during infection with immunodeficiency viruses. Accompanying their role in supporting virus transmission and spread, DCs are likely a crucial determinant in the induction of virus-specific immune responses to control virus replication (Figure 35.16). Identifying the factors facilitating virus replica-

FIGURE 35.16 Dendritic cell-associated virus induces virus-specific immune activation while fostering virus replication. DCs can capture virus via several routes: (1) infection; (2) entrapment via molecules like DC-SIGN; (3) uptake of infected apoptotic cell debris, or (4) after FcR-mediated ingestion of immune-complexed viruses. Presentation of virus by DCs to the immune system results in an incessant battle between immune stimulation and virus replication.

tion and development of immunity in the context of the DC will advance the design of blocking agents and immune-based strategies to prevent HIV-1 infection and/or disease development.

ACKNOWLEDGEMENTS M.P. was supported by grants from the NIH (AI40877 and AI44335), amfAR, The Campbell Foundation, The Dorothy Schiff Foundation, The Rockefeller Foundation, and The Irma T. Hirschl Trust. The work does not represent the opinions of the U.S. Government or the respective Foundations. Special thanks go to Judy Adams for computer expertise and Heidi Cleven, Loreley Villamide and Erin Mehlhop for technical assistance.

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PLATE 35.1 Immunity versus infection in the dendritic cell-T cell milieu. DCs presenting viral antigens will activate CD4 and CD8 T cell responses. However, DCs can transmit infection to CD4 T cells, especially those cells responding to antigen, to augment virus replication.

PLATE 35.2 Dendritic cell distribution within the stratified squamous epithelia and migration to the draining lymph nodes. Immature DCs positioned within and underlying the SSE survey the periphery capturing foreign antigens that they traffic via the dermal lymphatics to the T cell areas of the draining LNs. Matured DCs then present the antigens to CD4 and CD8 T cells to induce antigen-specific immunity.

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PLATE 35.3 Characteristics of immature and mature dendritic cells relevant to antigen presentation and virus replication. Immature DCs have the capacity to efficiently capture and process antigens. Upon maturation, the capacity to endocytose particles is diminished, but the T cell stimulatory activity is enhanced. The improved ability of mature DCs to activate T cells is facilitated by the increased expression of MHC, adhesion, and costimulatory molecules. Modulation of the expression of certain chemokine receptors during DC maturation (1) encourages DC movement to the T cell areas of the LNs; (2) favors DC-T cell interactions; and (3) influences the susceptibility of DCs to infection with HIV-1.

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PLATE 35.4 Langerin and DC-LAMP expression by cutaneous dendritic cells. Fresh LC suspensions were prepared from trypsinized epidermal sheets (fresh LCs). Fresh LCs were also cultured for 3 days at 37C, before being enriched by floatation over a Metrizamide density gradient (cultured LCs). Pieces of split thickness skin were cultured for 3 days and the emigrated leukocytes (containing epidermal and dermal DCs and T cells) were collected (emigrated skin DCs). Cytospins of each population were immunoperoxidase stained for Langerin, DC-LAMP, or CD1a. Langerin was evident in the fresh and cultured LC preparations and in LCs (arrows), but not the dermal DCs (arrowhead) of the emigrated skin DCs. DC-LAMP expression was detected upon maturation (culture) of the cells, present in both epidermal and dermal DCs. Original magnifications 10 and 40.

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PLATE 35.11 Entrapment of HIV-1 by DC-SIGN expressed by dendritic cells. DC-SIGN is a molecule expressed by DCs that can capture HIV independent of CD4 and CCR5 and in the absence of virus entering the DC. DC-SIGN-trapped virus can be efficiently transmitted to CD4 T cells, in which the virus then replicates.

PLATE 35.9 Focal localization of virus production within the lymph nodes at acute and chronic stages of infection. During acute infection most virus is produced by CD4 T cells within the T cell areas. This occurs in the absence of an established anti-viral immune response, i.e. lacking antiviral Abs and CD8 T cells. At chronic stages of infection, the majority of the virus is located in the B cell areas, trapped as immune complexes on FDCs and GCDCs and amplified by remaining CD4 T cells.

PLATE 35.12 Cellular distribution within the mucosal-associated lymphoid tissue of the tonsil. The lumenal surface of the tonsil is covered by a SSE that changes to a more ramified, lymphocyte-rich LE lining the tonsillar crypts. DCs are located within both the SSE and LE, and in the underlying T cell areas of the lymphoid tissue. The LE also contains T and B cells and the antigen-transporting M cells. FDCs and GCDCs are positioned within the B cell follicles.

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PLATE 35.13 Transmission of immunodeficiency virus infection at the mucosal-associated lymphoid tissues. HIV-1/SIV likely crosses the antigen-sampling LE. During the first days of exposure, virus cannot be readily detected and may remain cell-free or cell-associated within the LE. In subsequent days, however, rapid virus replication can be detected, especially within the CD4 T cells.

Plate 35.15 Potential immune implications of attenuated virus replication in the dendritic cell context. Immature DC-T cell mixtures support rapid replication of wild-type SIV, potentially resulting in the extensive immune destruction observed in vivo (particularly in the lymphoid compartments). In contrast, immature DCs cannot drive replication of the attenuated SIV delta nef even in the presence of CD4 T cells. Small amounts of virus produced in this setting could then be presented by the DCs to activate the immune system in the absence of consuming virus infection.

PLATE 38.1 Macrophages and dendritic cells coordinately regulate angiogenesis and antiangiogenesis. The fine balance regulating the microvasculature includes survival and inhibitory factors which are abundantly produced by macrophages and DCs. During normal homeostatic periods within a tissue, macrophages produce requisite factors facilitating small vessel integrity. During inflammation with progressive maturation and activation, DCs produce a variety of antiangiogenic factors including interferonα, IL-12 and IL-18. During the healing phase DCs are less abundant and are not activated by resident T cells or other cells, reverting to cells which promote neoangiogenesis and vessel survival. (Modified from Lotze, M.T. et al. (2001).

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36 Interactions of viruses with dendritic cells Marie Larsson, Jean-Francois Fonteneau, Andrew Lee and Nina Bhardwaj The Rockefeller University, New York, NY, USA

Specialized antigen-presenting cells that phagocytose large cellular debris and shuttle the resulting peptide degradation products to their endogenous class I presenting system. Michael Bevan, 1976

INTRODUCTION

Geijtenbeek et al., 2000), mobilization of high levels of antiviral products and expression of specialized antigen processing pathways that induce T-cell mediated immunity (Figure 36.1). It has become evident, however, that the consequences of DC–virus interactions can be a double-edged sword. Viruses can suppress DC function through multiple mechanisms including induction of apoptosis, inhibition of maturation, abrogation of cytokine production, loss of migratory capacity and inhibition of T-cell activation. These effects may account for some of the immunosuppression seen in certain viral infections. Viruses such as HIV-1 and measles virus can also use DCs to facilitate disease pathogenesis. This chapter will explore these dual and contrasting roles of DCs, emphasizing interactions with human cells. For discussion we have selected viruses on the basis of their known interactions with DCs. For a review of DC–HIV and –SIV interactions, the reader is referred to other chapters.

Dendritic cells (DCs) are bone marrow-derived professional antigen presenting cells (APCs) that are poised by virtue of their locations to encounter viruses upon their entry into the human body. Langerhans cells in the epidermis and mucosa, dermal dendritic cells in dermal regions, resident interstitial DCs in tissues and circulating blood DCs can capture viruses and viral antigens and migrate to T-cell rich areas in secondary lymphoid tissues. Upon encounter with inflammatory agents, stimuli or activated T cells, DCs undergo maturation whereupon costimulatory antigens, MHC molecules and MHC–peptide complexes are upregulated, and antigen processing is affected through the immunoproteosome (reviewed in Banchereau and Steinman, 1998). Because of these properties DCs are considered to be key initiators of Tcell responses to viruses. DCs are specialized to interact with and respond to viruses through unique virus binding molecules (e.g. DC-SIGN; Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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FIGURE 36.1 Crosspresentation of dying cells to T cells is dependent upon dendritic cells. In vivo, crosspresentation requires a bone marrow-derived APC, unidentified to date. In vitro, we have shown that immature DCs phagocytose virus-infected dying cells and, following a maturation signal, crosspresent antigen therefrom to T cells.

INNATE IMMUNE RESPONSES Innate immune responses are the first line of defense encountered by invading infectious agents. These include release of antimicrobial factors, phagocytosis by macrophages and neutrophils, lysis of virally infected cells by NK cells and complement-dependent clearance of viruses. The most striking antiviral innate immune response at the level of the DC is the release of interferon alpha by a specific subset of DCs. In blood one finds three subsets of DC precursors: CD14 monocytes, CD11c and CD11c precursor DCs. The CD11c subset is derived from a lymphoid lineage, lacks myeloid markers and can be differentiated from the CD11c subset in its reliance on IL-3 for growth and differentiation. In its precursor form, the CD11c subset is the principal interferon alpha producing cell in the blood. When exposed to influenza virus, Sendai virus, HIV, inactivated HSV or CD40-L, these DCs rapidly produce large

amounts of interferon alpha (4000 U/ml/104 cells, 100-fold more than their monocyte-derived counterparts) (Siegal et al., 1999; Cella et al., 1999a). This cytokine has pleiotropic effects: virus inhibition, NK cell and macrophage activation, enhanced expression of MHC I molecules on infected targets, and heightened CD8 T cell activation. Complementary dual functions have been identified for the CD11c DCs. Following maturation with a monocyte-conditioned medium or CD40-L, these cells activate T cells while skewing responses towards the TH2 phenotype – which may serve to downregulate the cellular immune response. CD11c cells have been located within the lumen and in close proximity to high endothelial venules (HEVs) in inflamed lymph nodes. They express CD62L or L selectin, which is required for adhesion and rolling on HEV endothelium and CXCR3, a receptor for chemokines induced by interferon gamma. These observations suggest that CD11c DCs enter lymphoid tissue via HEVs from blood

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during inflammatory reactions. This response may not be limited to viral infection, as CD11c cells have also been identified within nodes from individuals with Hodgkin’s lymphoma and myeloid leukemias (Salio et al., 1999). CD11c DCs, therefore, appear to provide a critical link between the innate and adaptive immune responses. It will be important to ascertain the full spectrum of their function. DCs also have the capacity to interact with and lyse cellular targets via expression of NKRs and TRAIL (tumor necrosis factor related apoptosis inducing ligand) (Fanger et al., 1999). While the targets thus far have been primarily tumor cells, it is possible that DCs will recognize virusinfected cells either directly or indirectly through activation of NK cells. DCs are known to directly trigger NK cell functions. Cell–cell contact leads to an increase of NK cell cytolytic activity and IFN gamma production (Fernandez et al., 1999). Once activated, NK cells or LAK cells can downmodulate ongoing immune responses by directly lysing DCs. Additional pathways in which DCs participate in the innate immune response are being identified. Beta-defensins, antimicrobial peptides expressed by epithelial cells of the skin and trachea-bronchial lining, are chemotactic for DCs and memory T cells through expression of the chemokine receptor CCR6 (Yang et al., 1999). Although beta-defensins have no homology to LARC, the chemokine ligand for CCR6, they may function to recruit immature DCs (which express CCR6 and lose it upon maturation) and memory T cells to cutaneous or mucosal sites of microbial invasion. Many virally encoded proteins can interact with chemokine receptors. Thus it is likely that more products that affect DC recruitment to inflamed sites will be identified. DCs, formerly only considered in the context of adaptive immunity, are now appreciated to be key participants in innate immune responses to microbes. Further studies will elucidate the full extent of the DC’s role in this arm of the immune response.

ADAPTIVE IMMUNE RESPONSES Influenza Virus DCs play a pivotal role in the generation of T-cell immunity to human viral pathogens (Figure 36.1). Influenza virus is probably the best studied virus with respect to the induction of CD8 T cell responses and its interaction with DCs. Influenza A virus is the prototype of the orthomyxoviridae which are characterized by a segmented negative strand RNA genome that is replicated in the nucleus of the infected cell. This subtype is the most clinically important pathogen since it has been responsible for severe epidemics in humans and domestic animals. Influenza virus–DC interactions DCs are located in the airway epithelia and can be rapidly recruited to enter lung tissue following inhalation of microbes. Thus they are poised to interact with influenza virions that enter the respiratory tract. In vitro, influenza has distinctive effects on DCs depending upon their stage of development and their origin.

Immature DCs Immature DCs, generated by culturing peripheral blood monocytes with GM-CSF and IL-4, are highly susceptible to infection by influenza virus. Exposure to either influenza or doublestranded RNA induces maturation (Cella et al., 1999a). This is characterized by: expression of the maturation associated markers DC-LAMP and CD83 (Cella et al., 1999b; Larsson et al., 2000) increased protein synthesis; upregulation of MHC, adhesion and costimulatory molecules; a two–three-fold increase in the half-life of class I molecules; production of interferon alpha and IL-12 and expression of MxA. The latter, an active GTPase which inhibits the primary transcription of influenza, is dependent upon the autocrine production of interferon alpha. Infection efficiency improves with higher MOIs, but viability decreases secondary to apoptosis.

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The mechanism by which influenza matures DCs remains undetermined. Interferon alpha activates PKR, an activator of gene transcription including NFκB. Expression of the matrix protein and nucleoprotein (NP), may directly activate NFκB and thereby induce maturation, as well as affect activation of genes that sustain DC viability, e.g. Bcl-xL (Wong et al., 1998). Mature DCs LPS and poly I:poly C matured DCs require 10fold more virus to produce comparable levels of influenza proteins. This may be due to the induction of MxA, which presumably functions to limit the cytopathic effect of the virus (Cella et al., 1999a). Interestingly, DCs matured with monocyte-conditioned medium (MCM), or TNF alpha are poor producers of MxA but are more resistant to the apoptotic effects of the virus, suggesting that MxA independent pathways exist for resistance to viral cytopathic effects. A protein with 2–5 oligoadenylate synthesizing (OSA) activity was recently described in maturing murine DCs (Tiefenthaler et al., 1998). OSA inhibits viral replication through the production of short 2 5-digoadenylate that activate 2-5-Adependent RnaseL and activation of this pathway leads to extensive cleavage of single stranded RNA. Whether human immature DCs express OSA in forms that are readily activated upon exposure to maturation stimuli like MCM, LPS or influenza virus remains to be determined. Following infection with MOIs of 2–4, up to 90% of MCM matured DCs express the influenza hemagglutinin, nucleoprotein and nonstructural protein (NS1) by 10–12 h (Bender et al., 1998). In contrast to immature DCs, the infection is nontoxic and minimally productive. Viability is retained for 2 days, at which time DCs still express viral proteins. The sustained synthesis of viral antigens, enhanced half-lives of MHC class I and resistance to reinfection, provides mature DCs with the capacity to activate T cells in lymph nodes and polarize T helper cell responses.

DC subsets Blood derived CD11c and CD11c DCs, as well as monocyte-derived DCs, synthesize IL-8, and TNF alpha in response to influenza (0.05–2 ng/ml/104 cells, (Cella et al., 1999a)). As described above, the CD11c subset is specialized to produce significantly greater quantities of IFN alpha than their monocyte-derived counterparts. Monocyte-derived DCs, but not the lymphoid derived CD11c DCs also produce IL-15, an important factor for the activation and differentiation of CD8 T cells (Ma et al., 2000). While the CD11c subset skews T cell responses towards the TH2 phenotype, monocyte-derived DCs skew responses towards the TH1 phenotype by virtue of their ability to produce IL-12 (Cella et al., 1999a, 1999b). Theoretically, the outcome of influenza infection could be substantially altered by the type of DCs that are infected. Monocytes Influenza virus also readily infects monocytes, although the virus replicates inefficiently in these cells. Infection induces a range of cytokines (GM-CSF, TNF alpha, type 1 interferons, IL-1, IL-6 and CC chemokines (Bender et al., 1998)). However, the majority of cells that are infected undergo apoptosis within 10 h of infection. Transcription factors such as NF-κB and AP-1 are activated but then rapidly decline, a feature that may contribute to the eventual outcome of programmed cell death. Hofmann et al. 1997, have postulated that influenza-induced apoptosis in monocytes serves to limit viral replication and the release of potentially degrading enzymes. Prior to death, monocytes synthesize inflammatory cytokines including chemokines that attract T cells and DCs into the environment (Hofman et al., 1997). As we will discuss below, these features of influenzainfected monocytes may facilitate the phenomenon of crosspriming. Induction of anti-influenza immune responses by DCs Dependence upon stage of maturation Influenza-infected DCs, but not monocytes, can

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induce substantial recall responses from purified blood CD8 T cells. Efficient presentation of antigens to memory CTLp is dictated by the stage of DC maturation and the form of antigen (see below). Monocyte-derived mature DCs are up to 20-fold more efficient than immature DCs in eliciting influenza-specific CD8 responses in ELISPOT assays measuring interferon gamma production (Larsson et al., 2000). Significantly, only mature DCs, but not immature DCs, induce CD8 expansion as assessed by MHC-tetramer binding assays and differentiation of CTLp into cytolytic effectors over 7 days. T cell activation by APCs is determined by several factors, amongst them duration of antigen stimulation, dose of antigen, and presence of co-stimulation (Lanzavecchia et al., 1999). Even at high doses of antigen, however, immature DCs failed to induce sustained proliferative responses and cytolytic T cells indicating an inability to commit memory T cells to enter effector pathways (Larsson et al., 2000). These data emphasize the use of mature DCs for immunotherapy purposes. In fact we have recently shown that mature DCs pulsed with matrix protein peptide rapidly boost CD8 T cell responses in healthy volunteers without the requirement for foreign helper antigens (Dhodapkar et al., 2000). Pathways for presentation of influenza antigens to CD8 T cells Three pathways for presentation of influenza antigens to CD8 CTLs by DCs have been identified. Infectious influenza virus: DCs infected with influenza induce strong antiviral CD8 CTL responses from freshly isolated blood T cells in short-term culture (Bhardwaj, 1998). Few DCs (DC:T cell ratio of 1:50–1:100) are required, the response is accompanied by extensive proliferation of CD8 T cells, and develops in the absence of CD4 helper cells or exogenous lymphokines probably because infection itself matures DCs (Cella et al., 1999b). IL-12, a cytokine that is produced by mature DCs (Bhardwaj, 1998; Subklewe et al., 1999a), enhances proliferative and CTL responses when added exogenously (Heiser et al., 1999). Only low levels of virus

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infection are required to elicit these recall responses from donors (MOIs of 0.01–1). Monocytes and B cells are weak or inactive in inducing these responses (Bhardwaj et al., 1994; Bender et al., 1995; Bhardwaj et al., 1996). However, influenza-infected monocytes can serve as excellent targets in short-term, 5-h CTL assays. Thus DCs serve as effective APCs for the induction of CD8 T cells, while monocytes act as targets for the CTLs that are induced. Non-replicating virus: DCs pulsed with inactivated, nonreplicating forms of influenza virus or with small amounts of influenza matrix protein peptide (nM range), induce CTLs as effectively as DCs infected with live virus. Similar data have recently been obtained for nonreplicating forms of vaccinia that encode influenza or EBV genes (Subklewe et al., 1999a, 1999b) and for avipoxes (e.g. canarypox) that encode HIV genes (Englemayer, et al., 2000). The inactivated forms of influenza virus make little detectable protein within the DCs. Less than 1% of DCs express viral protein, including nonstructural protein 1 (NS1, which is only synthesized in the infectious cycle), indicating that DCs require only small amounts of viral antigens to stimulate T cells. The binding and fusogenic functions of inactivated influenza virus are intact as assessed by standard hemagglutinating (binding) and hemolytic (fusion) assays. To be optimally effective, the inactivated virus must retain its fusogenic activity to presumably access the cytoplasm of DCs (Bender et al., 1995). These findings with nonreplicating viruses are important developments in the design of vaccines where induction of CD8 CTLs from quiescent human T cells is the goal. Virus-infected apoptotic cells: Classically, CD8 CTLs recognize antigens that are synthesized endogenously in the cytoplasm of the target cell, processed and presented as peptideMHC class I complexes (Heemels and Ploegh, 1995). However, there is evidence for an exogenous pathway whereby antigens that are not expected to gain access to the cytoplasm are presented on MHC class I (Song et al., 1997; Specht et al., 1997). The most dramatic example of this is the phenomenon of ‘crosspriming’, first

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described by Michael Bevan in vivo (Bevan, 1976), where antigens from donor cells are acquired by the host bone marrow-derived APCs and presented on MHC class I molecules to CD8 T cells (Figure 36.2). This has been now demonstrated for multiple antigens including histocompatibility, tumor and viral antigens (Bevan, 1976; Pfeifer et al., 1993; Huang et al., 1994; Kovacsovics-Bankowski and Rock, 1995; Reis e Sousa and Germain, 1995; Suto and Srivastava, 1995; Norbury et al., 1997). Crosspriming may therefore provide the immune system with a mechanism to detect and respond to viruses that do not target professional APCs and to antigens expressed by tumors that are inherently poor APCs. The identity of the bone marrow- derived APC-mediating crosspriming and the mechanism by which viral or tumor antigens are acquired has been studied in several laboratories, including ours. Using influenza virus as a model antigen system, we have demonstrated that apoptosis is one trigger for uptake of dying cells by APCs, and that DCs are the APCs that acquire and crosspresent antigens from these sources to T cells

FIGURE 36.2

(Figure 36.2). The DC is critical here as macrophages and B cells do not have this capacity. Other laboratories have recently identified immune complexes and exosomes as additional forms of antigens that can enter this exogenous pathway for crosspresentation (Zitvogel et al., 1998; Regnault et al., 1999). The special features of this exogenous pathway are summarized below ( Albert and Bhardwaj, 1998; Inaba et al., 1998; Albert et al., 1998a, 1998b, 1998c; Sauter et al., 2000). (1) DCs and monocytes both phagocytose influenza-infected apoptotic cells, however, only DCs induce class I-restricted CTLs from resting autologous T cells. Influenza antigen is acquired and presented from many types of infected apoptotic cells including allogeneic and xenogeneic cells, and at low apoptotic cell:DC ratios of 1:10–100 (Albert et al., 1998b). The CTL responses are in many cases equivalent to those generated by influenza virus infected DCs. By both immunolabeling and EM, we have shown that DCs phagocytose the apoptotic cells, which are visualized within compartments that coexpress MHC class I and II molecules, as well as

Possible outcomes of dendritic cell–virus interactions.

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lysosomal markers such as LAMP (Albert et al., 2000). In contrast to DCs, monocytes instead degrade apoptotic cell-derived antigens, release immunosuppressive cytokines such as IL-10, TGF beta (Fadok et al., 1998) and fail to stimulate T-cell responses. (2) Optimal CTL activation following phagocytosis of apoptotic cells by DCs requires two steps: phagocytosis by immature DCs, followed by a maturation signal to enhance costimulatory activity. Phagocytosis is mediated in part by a unique set of molecules: the αvβ5 integrin (a receptor for vitronectin and VEGF), and CD36, the thrombospondin receptor (as antibodies to these receptors block 50% of phagocytosis (Albert et al., 1998c). With maturation, receptor expression is downregulated co-ordinately with apoptotic cell engulfment. Uptake via this receptor complex may influence the trafficking of apoptotic cells to compartments or pathways where class I restricted epitopes are generated and loaded. (3) It was originally thought that only apoptosis charged DCs for crosspresentation of antigens to CD8 T cells. We have since found that DCs fed necrotic cells (cell lysates) also activate CD8 T cells. Optimal activation requires feeding DCs at their immature stage when they have efficient antigen acquisition properties, and then delivery of a maturation signal (Larsson et al., submitted). (4) Uptake of apoptotic and necrotic virusinfected cells by DCs induces CD4 T cell activation, presumably because both forms of antigen readily access the class II restricted antigen processing pathway by phagocytosis. Significantly, this pathway is 3000 times more efficient at activating CD4 T cells than soluble peptide. (5) DCs can distinguish between the two forms of cell death. Whereas immature DCs efficiently phagocytose both apoptotic and necrotic cells, only the latter induce maturation (Sauter et al., 2000). The factors within necrotic lysates with maturation-inducing qualities remain to be identified, but HSPs are a strong candidate (Srivastava, personal communication).

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(6) CD4 help is essential for crosspresentation of virus-infected dying cells in vitro, as demonstrated for protein, self and tumor antigens in animal models in vivo (Bennett et al., 1998; Ossendorp et al., 1998). The helper component can be substituted by CD40-L which presumably enhances DC function through upregulation of cytokines, MHC and costimulatory molecules and adaptation of immunoproteosome function. (7) In addition to tumor and viral antigenexpressing cells, DCs also phagocytose cells expressing self-antigens (e.g. intestinal epithelial cells (Huang et al., 2000)) and display selfpeptide–MHC complex molecules both in vitro and in vivo. This pathway may lead to peripheral tolerance of the T cell repertoire to self in the steady state, particularly if there is an absence of stimuli for maturation (Steinman et al., 1999). By eliminating or tolerizing self-antigen-reactive cells, the immune system can prime T cells to viral antigens contained within dying cells without also priming to self-antigens. In summary, these studies indicate a pivotal role for DCs in inducing immunity to influenza virus. In addition to direct infection, DCs also have the capacity to acquire viral antigens through phagocytosis of dying cells. As influenza is a cytopathic virus, immature DCs may be readily exposed to infected dying cells in the lung (such as epithelial cells, monocytes and DCs themselves). Thus phagocytosis of dying cells may be a physiologic route for the processing and presentation of influenza antigens to T cells in draining nodes. The novel aspect is the requirement for death in the generation of immunogenic material for delivery to the DC. As a consequence of cell death, antigens within cells which by themselves are poor APCs or viruses that fail to infect professional APCs, can gain access to the potent DC system. Upon maturation, these DCs can then elicit the requisite stimulatory responses. We believe this pathway accounts for the in vivo phenomenon of ‘crosspriming,’ whereby antigens derived from virus-infected cells, tumor cells or transplants are presented by host APCs to CD4 and CD8 T cells.

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Measles virus Despite the advent of effective vaccines, measles remains a major cause of morbidity and mortality in the third world. Measles virus (MV), a negative single strand RNA virus, induces both a humoral and cellular immune response that is characterized by the development of both CD4 helper and CD8 cytolytic immunity (Arneborn and Biberfeld, 1983). These responses, are considered to confer lifelong immunity following infection. Acute infection, and even vaccination can be associated with a transient immunosuppression that can be profound and which is characterized by loss of DTH to recall antigens, and heightened susceptibility to bacterial and other secondary infections (Lichtenfels et al., 1994). Strikingly, it has been shown that PBMCs isolated following infection, do not proliferate to recall antigens or mitogens in vitro for several weeks (Schlender et al., 1996). Recent studies of MV infection of APCs have revealed a multitude of effects on these cells, as well as T cells (see Figure 36.1) which provide explanations for these phenomena.

MV–DC interactions MV is acquired through the respiratory tract. Multinucleated giant cells (the Warthin– Finkeldey cells) identified within the nasopharyngeal epithelium and the subepithelial layer of the tonsils during the prodromal stage of measles are a major source of virus replication and spread to other nodes and tissues. In vitro, the interaction of MV has been studied with four sources of DCs. These include Langerhans cells (Grosjean et al., 1997; Steineur et al., 1998), freshly isolated blood dendritic cells (Schnorret et al., 1997), immature DCs derived from blood monocytes with GM-CSF and IL-4 (Fugier-Vivier et al., 1997) and CD34 cord blood progenitors in the presence of GM-CSF and TNF alpha (Grosjean et al., 1997). All sources of DCs are productively infected, albeit at low levels, with various strains of MV, presumably via the CD46 receptor. MOIs of 0.05–0.1 infect 40–100% DCs as determined by expression of the viral proteins

HA and F. However, the infection was characterized by syncytia formation and significant compromise of DC integrity and viability secondary to apoptosis (approaching 45–70%). Infection also causes the downregulation of CD46 and abrogates the production of IL-12 after ligation of CD40 (Fugier-Vivier et al., 1997). Since DCs are normally induced to synthesize IL-12 through CD40 signaling, this suggests that MV infection intersects the IL-12 synthesis pathway through alternative signaling mechanisms. In fact, IL-12 production by LPS- or SACS stimulated monocytes is inhibited after engagement of CD46 by MV, complement components C3b/C4b (natural ligands of CD46), or after ligation of Fc gamma or scavenger receptors. In contrast to these observations, MV has been shown to stimulate IL-12 in DCs after exposure to LPS or SACS (Fugier-Vivier et al., 1997). It is not yet clear whether these differing effects relate to DC maturation state or source, MV strain, or ligation of CD46, as CD46-independent pathways may be used for virus entry. Infection of immature DCs reportedly induces their maturation with a concomitant upregulation of DR, CD40, CD83 and the costimulatory molecules CD80 and CD86 (Fugier-Vivier et al., 1997). This effect of maturation, seemingly at odds with the induction of apoptosis, and inhibition of IL-12 production and T cell function (see below) will require further study. For instance, it has yet to be determined whether the maturation is due to direct infection, or occurs in uninfected cells secondary to maturation factors released by dying infected cells.

Consequences of MV infection on DC and T cell interactions The addition of activated or memory T cells, or ligation of CD40 dramatically increases MV replication (18-fold or so). Replication is associated with extensive apoptosis of both infected and uninfected T cells (leading to 90% cell death after 7 days) and the formation of giant multinucleated cells reminiscent of the Warthin– Finkeldey cells. This is a good example of how a virus controls the apoptotic cascade in cells it

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infects. Apoptosis is prolonged sufficiently to allow extensive viral replication. The mechanisms involved in this apoptosis cascade are being delineated. Recently, Vidalain et al., 2000, have shown that MV infection induces functional TRAIL on monocyte-derived DCs. TRAIL mRNA but not Fas-L mRNA was rapidly upregulated in immature DCs, monocytes and CD3-activated T cells within 8 h of infection. Curiously surface TRAIL could only be detected on monocytes, but not DCs. DCs apparently express TRAIL protein intracellularly, possibly the full length monomeric TRAIL of 33 kDa. MV- infected DCs lyse transformed cell lines, an effect that is blocked by TRAIL-R2:Fc chimera. As alpha interferon treatment upregulates TRAIL in human DCs (Vidalain et al., 2000), it is possible that TRAIL upregulation relies on release of this soluble factor after MV infection. The effects on T-cell function are striking as MV-infected DCs fail to adequately stimulate either mitogenic or allogeneic T-cell responses. Furthermore, relatively few infected DCs and only minimal contact with T cells are required for this effect to be observed (Fugier-Vivier et al., 1997). T cell function is blocked regardless of whether immature or mature DCs are infected (Steineur et al., 1998) but apoptosis is more striking when immature DCs are infected. Besides the induction of apoptosis and inhibition of IL-12, it is likely that MV-mediated inhibition of T cells involves other factors.

MV interferes with the function of other cells MV also adversely affects the function of other cells. Expression of MV proteins F and H by transfection, or via infection of monocytes with UV irradiated particles, were sufficient to inhibit the proliferation of mitogen-stimulated T cells. Direct cell–cell contact was required rather than inhibition by direct virus infection or release of an inhibitory factor (Schlender et al., 1996; Schnorr et al., 1997). This phenomenon is associated with a drastic deregulation of the expression of cell cycle genes essential for the G1/S phase transition (Engelking et al., 1999).

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B cells also succumb to the effects of MV. Ravenel et al., have shown that the nucleocapsid protein of MV binds B cells, through the FcR gamma II receptor and inhibits polyclonal Ig synthesis by as much as 50%. Infection of transformed B cells leads to the release of soluble factors that inhibit the proliferation of antigenspecific T cells and of uninfected B cells (Fujinami et al., 1998). Hoffman et al. (Hoffman et al., 1992) have shown that MV fails to activate NFκB in neuronal cells, due to its inability to induce phosphorylation and degradation of IΚB, the inhibitor of NFκB. This effect may explain the repression of interferon beta and MHC class I gene expression in virus-infected cells, and account for the persistence of MV in neurons and lack of recognition by the immune system. Therefore, MV has a multitude of effects on several cells. Collectively these effects provide potential mechanisms for the virus to confound, evade and damage the immune system.

Role of DCs in vivo It is important to try to reconcile the clinical descriptions of immunosuppression after MV infection and the primary role of DCs as stimulators of immune responses. One possibility is that the Warthin–Finkeldy cells in the submucosal areas of the tonsils and pharynx are syncytia consisting of DCs and T cells. DCs in the lining of mucosal surfaces, as suggested for HIV-1 infection, are most likely the targets for the virus and responsible for viral transmission. Thus DCs may be a reservoir for MV infection and a vehicle to transmit the virus to lymphoid cells in draining lymph nodes. While no studies have directly assessed in vitro or in vivo whether DCs bearing MV antigens directly stimulate antiviral cellular responses, it is tempting to speculate that DCs are also involved in activating these adaptive immune responses, thereby serving a dual function. The CD8 T cell response in particular is important in the clearance of established MV infection. Acute infection and vaccination have been associated with the detection of circulating CD8 effector T cells

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(Jaye et al., 1998). Marked distortions of the T cell repertoire have also been described in the majority of infected patients and in vaccinated individuals where CD8 TCR V-beta expansions were noted to be clonal or oligoclonal with cytolytic activity against MV antigen expressing lines (Mongkolsapaya et al., 1999). These findings are reminiscent of large clonal expansions of CD8 T cells following EBV infection. We speculate that DCs will be essential to activate MV-specific immune responses, although it remains unclear whether infected DCs themselves are the antigen-presenting cells or if crosspresentation of apoptotic infected cells (DCs or epithelia) are the essential elements requisite for T cell activation.

Depending upon the tumors either a full array or limited numbers of latent genes are expressed. For example, in Burkitt’s lymphoma only EBNA 1 is expressed (latency state I) while in NPC EBNA 1, LMP 1 and LMP 2 are expressed (latency state II). EBNA1 is essential for the persistence of the viral genome in replicating EBV-transformed human B cells, indicating why all EBV-transformed tumors express this foreign antigen. Because of the clear oncogenic role of EBV in several tumors, considerable effort is being directed towards understanding how the immune system responds to infection and how to manipulate these responses to eliminate virus-transformed tumors.

Host immune responses to EBV HUMAN HERPES VIRUS Of 100 or so herpesviruses, only a few have been isolated from humans. We will focus primarily on Epstein–Barr virus (EBV) and herpes simplex virus 1 (HSV-1) as their interactions with T cells and DCs have been well studied.

Host immune responses are considered to be critical in controlling the infection both during primary infection and in the carrier state. Most individuals have been infected with EBV by adulthood, and have established humoral and cellular immune responses to EBV antigens. Antibodies to the major virus envelope (gp 340) are considered to neutralize viral infectivity (Rickinson and Moss, 1997).

EPSTEIN–BARR VIRUS In primary infection, EBV replicates in the oropharyngeal epithelium, with expression of the full array of replicative or lytic genes. The virus colonizes the lymphoid system, in particular the B cell system by binding to CR2 (CD21) via gp340 and then through the growth transformation of infected B cell clones selectively expressing latent genes: EBNAs 1, 2, 3A, 3B, 3C, -LP, LMP-1 and 2 (latency state III). A virus carrier state ensues where periodic reactivation of latently infected B cells leads to activation of lytic cycle genes and reinitiation of replicative foci. However, the primary long-lived pool of virus carriers is probably the resting B cell which expresses only EBNA 1. A number of human tumors are associated with EBV infection including immunoblastic lymphoma, Burkitt’s lymphoma, nasopharyngeal carcinoma (NPC), and Hodgkin’s disease.

Cellular immune responses: role of CD8 effector cells The cellular immune response is of major importance in controlling the initial expansion of transformed B cells, viral load and established infection as shown by the emergence of lymphoproliferative disease in T cell immunocompromised patients and the regression of B cell transformed cells following treatment with virus-specific CTLs (Riddell and Greenberg, 1995). The CTLs have the following properties: CD8 phenotype, HLA class I restriction, preferential skewing of CTL responses to EBNAs 3A, 3B and 3C, and highly conserved TCR alpha beta usage (Burrows et al., 1994; Silins et al., 1996; Steven et al., 1996). Interestingly, only subdominant or rare CTL responses have been described to the remaining latent proteins: EBNA1, EBNA2, EBNA-LP, LMP-1 and LMP2

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(Rickinson and Moss, 1997). The hierarchy of EBV antigens towards which CTLs are directed can be explained by several factors. First, certain antigens are poorly processed and presented, e.g. EBNA 1 has a glycine–alanine repeat domain in the protein which inhibits proteasome processing and hence access of antigenic pepides to class I molecules (Levitskaya et al., 1997). LMP1 and LMP 2 are membrane proteins that are also less accessible to cytoplasmic processing. Second, the pattern of immunodominance determined thus far may reflect the fact that most CTLs have been detected by repetitive stimulation of bulk mononuclear cells with EBVtransformed B cell lines (LCLs which express the full spectrum of latent cycle antigens) (Rickinson and Moss, 1997). Paradoxically, the EBV-specific CTLs fail to lyse LCLs expressing EBV antigens, against which they were raised unless antigens are delivered exogenously. One possibility is that the cells have short half-lives of peptide–MHC complexes (Levitsky et al., 1996). LCLs, therefore, may only elicit CTL profiles of antigens which are expressed on their surface (even then not efficiently). LCLs or their equivalents, are not likely to be the APCs that induce CTLs in vivo.

Cellular immune responses: role of CD4 effector cells CD4 T cells are required to initiate and maintain CD8 T cell immunity (Kalams and Walker, 1998). The spectrum and antigenic specificities of class II restricted T-cell responses to EBV antigens which resist or are less accessible to cytoplasmic processing (e.g. EBNA 1, LMP 1 and LMP 2), are under active investigation. A majority of healthy infected donors appear to have CD4 T cells of a TH1 phenotype that recognize EBNA1 and fewer that recognize LMP-1 (Münz et al., 2000). Upon stimulation with antigenpulsed DCs, these CD4 T cells proliferate, secrete cytokines (interferon gamma), and even lyse EBNA1 presenting target cells, including BLCLs. It remains to be seen if EBNA 1-specific immunity in healthy donors is solely maintained by CD4 T cells, or whether

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CD8 T cells are also mediators of protection. Rare CD8 T cell responses to EBNA 1 have been described but their activation appears to require exogenous antigen processing (Blake et al., 1998). LCLs pulsed with soluble EBNA 1 could be readily lysed by specific CTLs. However, the physiological role of these cells is obscure as they would be inactive against EBV-infected cells which do not express endogenously processed EBNA 1 on class I molecules. EBNA 1specific CD4 T cells may contribute to protective immunity by direct recognition of EBNA 1 epitopes on class II molecules, or through sustaining CD8 T-cell responses to other EBV associated antigens.

EBV–APC interactions The presence of EBV antigens in many malignancies offers a potential target for immunotherapy. However, there are several unknowns with respect to CTL generation and conference of immunity in EBV-induced tumors. One relates to immune surveillance in the carrier state. Lytic antigen-specific CTLs likely control the initial infection in the oropharynx and any recrudescences of viral replication. CTLs directed towards latent cycle antigens would also contribute to controlling B cell proliferation. However, as emphasized above the full range of memory CTLs has not been defined as LCLs have been commonly used to determine the spectrum of CTLs expressed in vivo. In the carrier state, most of the EBV-infected B cell pool (about one in 1 million) expresses EBNA1 and therefore would escape CTL detection. Only B cells (in the lytic state or latency III state) would be recognized by CTLs. What therefore are the primary APCs for inducing CTLs in vivo? B cells? Or must EBV antigen expressing cells be acquired by professional APCs which lack infectibility, e.g. DCs to elicit CTLs. Some answers are beginning to emerge from recent studies by Subklewe et al. (1999a, 1999b), comparing human DCs and monocytes as APCs, they found that DCs infected with recombinant vaccinia elicited CD8 T-cell responses specific for the immunodominant EBNA 3A, 3B and 3C

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antigens from PBMCs of most healthy individuals. In a smaller number of donors, recognition of the subdominant LMP-1, LMP2 and EBNA 1 antigens was documented. DCs pulsed with the HLA-B8-restricted EBNA-3A peptide, FLRGRAYGL, elicited CTLs that recognized antigen-pulsed BLCLs. Furthermore, mature DCs pulsed with UV-inactivated virus were also efficient stimulators of EBV-specific CTLs (Subklewe et al., 1999b). In comparison, BLCL were less efficient and required IL-2 to elicit CTLs, perhaps because they present insufficient amounts of endogenously processed peptides. These data suggest that infected B cells by themselves are poor activators of T-cell effector responses. When EBNA 1 specific CD4 T-cell responses were examined, it was found that DCs could process EBNA 1 either from purified protein or from EBNA 1 expressing cells including BLCL via crosspresentation (Münz et al., 2000). BLCL failed to present exogenous sources of EBNA 1, but could present EBNA 1 if infected with vaccinia constructs expressing EBNA 1 with a GA deletion. Altogether these data suggest that DCs induce protective immune responses against EBV by crosspresenting B cells expressing EBV-antigens, such as BLCL and infected blasting B cells. Once activated, these CD4 and CD8 cells lyse infected targets. This new knowledge regarding the ability of DCs to induce CTLs via an exogenous pathway and with unconventional sources of antigens identifies new approaches to target EBV-associated tumors that needs to be explored in the future.

HERPES SIMPLEX VIRUS -1 HSV-1 is a member of alpha-herpes viridae, a neurotropic virus with a wide host range. Infection usually involves mucosal surfaces or squamous epithelial sites, followed by latency in neurons. From there it can periodically emerge to reinfect skin or other tissues, causing recurrent mucocutaneous lesions in about one-third of human populations. HSV-1 efficiently destroys infected cells, inducing apoptosis in

PBMC, CD4 T cells and hepatocytes. Rounds of lytic infection occur at peripheral sites and continue until terminated by host immunity (Daheshia et al., 1998).

Evasion of immune responses by HSV T cells, in particular, CD8 CTLs are necessary to clear HSV1 infections (Cose et al., 1997; Raftery et al., 1999). However, herpes viruses such as HSV and CMV, in particular, have evolved multiple strategies to evade the attack of T cells, enabling them to persist and reactivate within the host (Raftery et al., 1999). As described for EBV, lack of expression of viral antigens during latency is one such strategy. HSV interferes with antigen presentation by preventing the transport of MHC class I molecules to the surface, blocking peptide transporters via the ICP-47 protein, producing cytokine and chemokine homologues that may subvert the inflammatory immune system (Ploegh, 1998; Salio et al., 1999) and inducing apoptosis in the cells it infects, thereby facilitating virus replication. HSV-1 glycoprotein gC (a virulence factor involved in initial attachment by binding heparan sulfate) binds to complement components C3, C3b, iC3b and C3c, accelerates the decay of the alternative pathway C3 convertase, and blocks interaction of C5 and properdin with C3 (Lubinski et al., 1999). Many of these evasion tactics are beginning to emerge at the level of DCs (see below).

HSV–APC interactions HSV-1 initially attaches to cell surface heparan sulfate proteoglycans which is followed by glycoprotein D binding to three specific surface receptors: HVEM, a member of the TNF/NGF receptor family, and Prr1 and Prr2, members of the Ig superfamily (Montgomery et al., 1996). Immature and mature DCs express HVEM, Prr2 is found in moderate levels on immature DCs and low levels on mature DCs and both DCs have traces of Prr1 (Salio et al., 1999). Infection of immature DCs with disabled single cycle HSV-1 led to some increase of MHC class II. Interestingly, bystander maturation of

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uninfected cells occurred, possibly due to the trans effects of virally induced proteins. Infection of immature DCs that were simultaneously exposed to LPS, inhibited DC maturation. Infected DCs failed to upregulate markers associated with maturation such as CD83, costimulatory molecules, and CCR7 (a chemokine receptor that ensures migration of maturing DCs to lymph nodes (Kellermann et al., 1998)). The latter was characterized by lack of responsiveness to its ligand EBV-induced molecule 1-ligand (ELC) (Salio et al., 1999). Furthermore, infected cells fail to synthesize IL12, TNF-alpha, IL-6 and IL-10, but can synthesize low levels of IFN alpha. Finally, infected immature DCs were severely compromised in their capacity to stimulate allogeneic T cells. As nonreplicating HSV-1 was used in these studies it is unclear whether HSV-1 can replicate in DCs, and whether it induces apoptosis as it does in macrophages (Salio et al., 1999). Mature DCs were resistant to HSV-1, requiring 10-fold higher MOIs to obtain the same percentage of infected cells. While MHC class I expression and ELC responsiveness were unchanged, mature infected DCs were also inhibited in their T-cell stimulatory capacity. Unlike MV-infected DCs, HSV-1infected DCs were not inhibitory when mixed with uninfected mature DCs ruling out an active suppressive mechanism. HSV infection leads to the loss of host cell protein synthesis due to disruption of polysomes and degradation of mRNA. The viral protein vhs, the product of UL41 gene is responsible for the shutoff process. This mechanism may account for the block in several of the steps that are associated with the maturation process. However, elucidation of the molecular pathways within DCs that are targeted by HSV-1 will require further study.

Role of DCs in vivo LCs or dermal DCs are likely to come in contact with HSV by direct infection. Primary response may not occur via direct presentation of viral antigens by infected DCs as they fail to undergo maturation, migrate to lymph nodes,

or stimulate T cells. Therefore, phagocytosis of virus-infected DCs by immature DCs which simultaneously undergo bystander maturation may be the predominant mechanism by which antiviral immunity to HSV-1 is generated.

POXVIRUSES Vaccina virus Vaccinia, the prototype poxvirus, is a large enveloped DNA virus that is a member of the orthopox virus family. Vaccinia virus consists of three different forms of infectious virions: the most abundant intracellular mature virions (IMV), intracellular enveloped virus (IEV) and extracellular enveloped virus (EEV). These different forms of virions are responsible for various routes of infection. Vaccinia envelope proteins A27L and D8L bind to the cell surface. The A27L is required for fusion of virus by mediating binding to cell surface glycosaminoglycans (GAGs), mainly heparan sulfate. Another envelope protein D8L, which binds to chondroitin sulfate has also been implicated in vaccinia entry (Hsiao et al., 1999). A wide variety of cell types can be infected with vaccinia, the most permissive being epithelial cells. Despite the obscure origin of vaccinia, its successful use as a vaccine to eradicate smallpox has made it a valuable tool to study virus–host interactions, with particular emphasis on its application as a vaccine vector.

Vaccinia virus– APC interactions Although vaccinia induces strong humoral and cellular immune responses, it remains unclear how these actually develop in vivo. The APCs targeted by scarification (the usual method of immunization), have yet to be defined. Furthermore, poxviruses employ many mechanisms to evade the immune system (Moss, 1991; Smith et al., 1997). For example, vaccinia virus encodes apoptosis inhibitors (SP-2), receptor homologs of IL-1beta, TNF-alpha, IFN-alpha, IFN-beta and IFN-gamma (Smith et al., 1997),

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chemokine receptors to capture chemokines, and inhibitors of the IFN signaling pathway (Lalani et al., 1999). The latter include E3L and K3L which block the serine threonine protein kinase PKR and 2–5 oligoadenylate synthetase which inhibit viral replication. Because the natural APC for vaccinia in vivo is unclear, several studies have evaluated the effects of vaccinia–APC interactions in vitro. Vaccinia infects immature and mature DCs, monocytes and BLCLs (Engelmayer et al., 1999). In all but the lattercase,theinfectionisabortive,withonlyearly genes being expressed. The block in viral replication in both immature and mature DCs remains unexplained at this time. High frequencies of immature DCs are infected with vaccinia (70%) compared to mature DCs, infection of immature DCs leads to a block in their maturation within 1 dayinhibitingtheexpressionofproteinsknownto be induced during DC maturation such as CD83, CD86, DR and CD25. This is accompanied by a marked inhibition (upto 90%) in the allogeneic MLR of their T cell stimulatory capacity. Besides the block in maturation, immature DCs also undergo a delayed apoptosis 48 h following infection, possibly due to the production of caspase inhibitors. In productive cells, this delay in apoptosis would allow the virus sufficient time to replicate and facilitate spread of progeny. As with influenza virus, mature DCs are more resistant to these inhibitory effects of vaccinia. How does vaccinia exert these multitude of effects in immature DCs? Production of cytokine receptor homologues may inhibit maturation effects mediated by TNF-alpha or IFN-alpha. Vaccinia may also block intracellular signalling pathways that lead to maturation. The resistance to apoptosis conferred by maturation could be due to higher levels of PKR and 2-5 oligoadenylate synthetase that inhibit the synthesis of proapoptotic products. Furthermore, maturation stimuli such as TNF-alpha or CD40 ligand promote the viability of DCs via upregulation of antiapoptotic proteins, e.g. Bcl-xL (Wong et al., 1998).

Role of DCs in vivo Access to DCs in vivo may be critical for potent immune responses against vaccinia and other pox viruses. Traditionally, vaccinia is delivered by scarification, and local epidermal DCs (Langerhans cells) in the skin may provide a vehicle to traffic vaccinia to draining lymph nodes. Since vaccinia abortively infects DCs, blocks their maturation, and induces delayed apoptosis, it is unclear how vaccinia immunity is generated in vivo. One possibility is that sufficient amounts of immature DCs survive vaccinia infection or may mature before infection. Poxvirus infection induces the release of inflammatory cytokines from infected tissues, which could affect the maturation of DCs from precursors. Another mechanism for generating immunity could be crosspriming. Bystander DCs may ingest vaccinia-infected apoptotic cells and present viral antigenic epitopes to T cells. In vitro data indicating that immature DCs can phagocytose dying vaccinia-infected cells and present the viral antigenic epitopes to T cells supports this route of antigen presentation (Larsson et al., submitted).

CANARYPOX Canarypox is a member of the avipoxvirus family with a natural host range restricted to avian cells. Canarypox is capable of infecting mammalian cells, resulting in a nonproductive infection with early and in some cases late gene expression before the block in replication (Smith et al., 1997). For this reason, this virus has been applied as a vaccine vector against a variety of agents.

Canarypox–DC interactions Engelmayer et al. (2000) have investigated the interactions of canarypox vectors encoding HIV genes with different stages of DCs and the ability of these cells to elicit recall responses from HIV individuals. Like vaccinia, canarypox infected immature DCs to a higher extent than

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mature DCs. Unlike vaccinia, the number of infected cells was low (5–15% as assessed by protein expression) and canarypox did not inhibit DC maturation as assessed phenotypically by markers such as CD83, CD25, CD80, CD86 and HLA DR. Canarypox infection affected DC viability especially of the immature cells rapidly inducing apoptosis within 1 day. The mature DCs resisted the cytopathic effect to a great extent, as in the case of vaccinia virus (Engelmeyer et al., 1999). These effects of avipoxes upon DCs may vary according to the virus strain and origin as recombinant fowlpox virus reportedly infected more than 80% of the DCs (Brown et al., 1999).

DC–T cell interactions Despite a low frequency of DCs infected with canarypox, strong effector CD8 T-cell responses could be elicited from chronically infected HIV individuals, e.g. cytolysis, secretion of IFN-γ and β-chemokines. Furthermore, canarypoxinfected DCs induced responses that were at least as potent as the ones induced by vaccinia vectors or peptides. Addition of exogenous cytokines was not necessary to elicit these CTLs, although the presence of CD4 T cells was required to obtain and maintain effective CTL responses. Strikingly, canarypox-infected DCs were able to directly stimulate HIV-specific IFNγ secreting CD4 helper responses from purified CD4 T cells. Therefore, targeting canarypox vectors to mature DCs may enhance anti-HIV CTL and CD4 helper responses in vivo.

ADENOVIRUS Adenoviruses are encapsulated double-stranded linear DNA viruses that normally infect terminally differentiated epithelial cells and which cause acute upper respiratory infections. In permissive cells, the projecting adenovirus fiber-knob protein mediates binding to coxsackie-adenovirus receptor (CAR) on the cell surface, and internalization is facilitated by interaction of the adenovirus base with either αvβ3 or αvβ5

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integrins (Pina and Green, 1999). Adenovirus has drastic effects on the metabolism of host cells as the virus usurps almost the entire synthetic machinery of its host. Inhibition of cellular DNA synthesis begins 6–8 h after infection, and there is a concomitant rise in viral protein RNA synthesis (Nelson and Krensky, 1998). Because wild-type adenoviruses have been shown to downregulate MHC molecules and induce lytic infections, adenovirus has been substantially modified to produce vectors that can also accommodate foreign DNA (Pina and Green, 1999). Together with adeno-associated viruses they have been extensively studied as vectors for immunizations. Some preliminary work has explored the interaction of recombinant adenovirus vectors with DCs, and stimulation of CD8 T cell responses

Adenovirus –DC interactions Adenovirus infects a wide variety of cells but efficient transfection requires the CAR, which the DC lacks. In general it is necessary to use a high multiplicity of infection to get acceptable infection in cells lacking the CAR (100 MOI) (Diao et al., 1999). DCs can be readily infected with replication-deficient recombinant adenoviruses derived from adenovirus type 5 with MOIs of 100. As with other viruses, a difference in transfection efficiency and gene expression between immature and mature DCs, has been reported. Greater than 90% of immature DCs were transfected compared to only half of the mature DCs, in part because the mature DCs downregulate the integrins that are utilized for virus entry (Zhong et al., 1999). Transfection was stable as virally encoded GFP was detectable even 6 days after transfection and transgene expression could be prolonged after exposure to TRANCE, a DC survival factor. Notably, recombinant viruses did not alter the DC’s capacity to undergo maturation, produce cytokines (e.g. IL-12), or stimulate T cells. Similarly, Diao et al. (1999) reported no significant effect on DC morphology, immunophenotype or potency in allogenic lymphocyte proliferation assays, with the exception of an increase in CD86. The

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upregulation of CD86 increased with rising MOIs of adenovirus. Another report suggests that infection by adenovirus (AdTG6401/ Ad5CMVβ-gal) enhances the immunostimu-latory capacity of immature DCs through the upregulation of costimulatory molecules such as CD80, CD86, CD40 and MHC class I and II molecules (Rea et al., 1999). The infection failed to induce IL-12 secretion, however, and only marginally induced expression of CD83 (Rea et al., 1999). Finally it has been reported that the induction of allogeneic proliferation by adenovirus-infected DCs is unaffected unless high doses of DCs are used (Tuting et al., 2000). Recombinant adenoviruses have been widely used for delivering genes into primary cells and have been successfully applied for antitumor therapy. The advantage in targeting genes to DCs via these vectors is high transfection efficiency in immature DCs compared to retroviral vectors, the ability to mature transduced DCs without loss of function and stable gene expression, which can be further enhanced with the addition of survival factors. An additional theoretical advantage is that targeting to DCs, by reducing exposure to envelope proteins, will reduce the problem of neutralizing antibodies that are an inherent feature of repetitive adenovirus injections and which compromise immunogenicity to encoded antigens.

SUMMARY DCs, ‘nature’s adjuvants’, are key APCs for the induction of protective immune responses to viruses. DCs in peripheral tissues can be directly infected by viruses thereby allowing viruses that do not gain entry to the central lymph nodes to be transported to the T cell areas of draining nodes. In some instances (e.g. influenza), infection directly leads to DC maturation providing a mechanism for the coupling of antigen presentation and T cell activation. For those viruses that fail to infect DCs directly, such as EBV, crosspresentation of virus-infected dying cells provides a mechanism by which potent antiviral CTLs can be generated. This

feature appears to be unique to the DC. Virus infection may also induce bystander maturation of DCs. In this way, uninfected DCs could phagocytose virus-infected dying cells, while simultaneously undergoing maturation. DCs are now recognized to be important bridges between innate and adaptive immune responses via their ability to rapidly produce significant quantities of antiviral proteins, lyse cellular targets and simultaneously skew the T cell helper response. Because of the DC’s strategic role in the induction of antiviral immune responses, it is not surprising that viruses have evolved mechanisms to exploit DCs by interfering with their function. The challenge in the future will be to develop strategies for the development and maintenance of protective immunity against viruses that have endured in human hosts.

ACKNOWLEDGEMENTS Some of the studies cited here were supported by NIH grants (AI39516 and AI44628), the SLE Foundation, and the Burroughs Wellcome Trust.

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Steineur, M., Grosjean, I., Bella, C., Kaiserlian, D. (1998). Eur. J. Derm. 8, 413–420. Steinman, R.M., Turley, S., Mellman, I., Inaba, K. (1999). J. Exp. Med. 191, 411–416. Steven, N.M., Leese, A.M., Annels, N.E., Lee, S.P. and Rickinson, A.B. (1996). J. Exp. Med. 184, 1801–1813. Subklewe, M., Chahroudi, A., Bickham, K. et al. (1999a). Eur. J. Immunol. 29, 3995–4001. Subklewe, M., Chahroudi, A., Schmaljohn, A., Kurilla, M.G., Bhardwaj, N. and Steinman, R.M. (1999b). Blood 94, 1372–1381. Suto, R., Srivastava, P.K. (1995). Science 269, 1585–1588. Tiefenthaler, M., Marksteiner, R., Hofer, S. et al. (1998). J. Leuk. Biol. 2, 83. Tuting, J., Steitz, J., Bruck, J. et al. (2000). Gene Ther. 7, 249–254. Vidalain, P., Azocar, O., Lamouille, B., Astier, A., Rabourdin-combe, C. and Servet-Delprat, C. (2000). J. Virol. 74, 556–559. Wong, B. R., Josien, R., Lee, S.Y., Vologodskaia, M., Steinman, R.M. and Choi, Y. (1998). J. Biol. Chem. 273, 28355–28359. Yang, D., Chertov, O., Bykovskaia, S.N. et al. (1999). Science 286, 525–528. Zhong, L., Granelli-Piperno, A., Choi, Y. and Steinman, R.M. (1999). Eur. J. Immunol. 29, 964–972. Zitvogel, L., Regnault, A., Lozier, A. et al. (1998). Nat. Med. 4, 594–600.

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37 Dendritic cells in allergy A.E. Semper, A.M. Gudin, J.A. Holloway and S.T. Holgate School of Medicine, Southampton University, Southampton, UK

We all know people who are affected by a condition such as hay fever, asthma or eczema. The costs to public health and the economy are massive and growing, and a major research effort has been underway for many years to understand these diseases. Despite much progress, allergic diseases still present significant puzzles, which hinder the development of effective therapies. Allergy and asthma P. Campbell and U. Weiss

INTRODUCTION

sonal rhinitis (hayfever) in the upper airways and allergic conjunctivitis in the eye. At its most acute, the allergic response manifests itself as the lifethreatening systemic condition of anaphylaxis. It is the role of dendritic cells in these atopic, allergic diseases that is the subject of this chapter. Of the allergic diseases, asthma has the greatest socio-economic impact, and has consequently received the most intensive research effort. For this reason, many of the examples quoted in this chapter are drawn from the study of respiratory allergy.

In current usage, the term allergy is restricted to describing the inappropriate response of susceptible individuals to common innocuous environmental antigens, or ‘allergens’. These responses are characteristically triggered by immunoglobulin E (IgE) and susceptible individuals typically show a genetic tendency to develop elevated levels of (allergen-specific) IgE, a trait termed ‘atopy’. This has led to the terms allergy and atopy being used interchangeably. Allergic diseases are increasing in prevalence and are now a major source of disability throughout the developed world (Holgate, 1999). Atopic asthma in the lungs and allergic eczema (atopic dermatitis) in the skin are the most serious and prevalent of the allergic diseases: in Britain, asthma now affects one in four children under the age of 14 years and one in five have eczema. Other common allergic diseases are perennial and sea-

Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

EARLY- AND LATE-PHASE ALLERGIC RESPONSES AND THE DEVELOPMENT OF CHRONIC INFLAMMATION Atopic allergic diseases share a common triggering mechanism, namely immunoglobulin E. In an

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individual previously sensitised to an allergen, allergen-specific IgE antibodies are bound to high affinity IgE receptors (FcεRI) on the surface of mast cells in target tissues, such as the skin or lung. Upon re-exposure to allergen, receptorbound IgE molecules on the sensitised mast cells are cross-linked, causing mast cell degranulation and release of mediators such as histamine, tryptase, prostaglandins and leukotrienes. These mediators cause increased vascular permeability, local oedema and itching resulting, for example, in the wheel and flare reaction that occurs within 10–20 min of introducing allergen into the dermis of the skin. This acute, immediate, type 1 hypersensitivity response is frequently followed by late phase reactions (LPR) that typically peak at 6–12 hours following allergen exposure, then gradually decline over the next 12–24 hours. During the late phase response, the site of allergen exposure shows marked infiltration by eosinophils and CD4 T helper (Th) cells (Azzawi et al., 1992). Using short, linear peptides of the cat allergen Fel d 1, which did not cross-link FcεRI, it has recently been shown that MHC/peptide engagement of allergen-specific T cells was sufficient to elicit late phase responses in the absence of a prior IgE-mediated immediate reaction (Haselden et al., 1999), thus highlighting the key role played by the T cell in the delayed, ‘type IV’, cell-mediated arm of the atopic hypersensitivity response. Over the past two decades, it has become apparent that even in the absence of recent allergen provocation, there is an underlying persistent chronic inflammation in the target organs of allergic diseases. This chronic allergic inflammation is characterised by the same cell types as the LPR, namely eosinophils (Bruijnzeel-Koomen et al., 1988a; Djukanovic et al., 1990) and CD4 T cells (Frew and O’Hehir, 1992; Poston et al., 1992; Metz et al., 1996). Both cell types show signs of activation (Azzawi et al., 1990; Bradley et al., 1991; Wilson et al., 1992; Robinson et al., 1993). Whereas the activated eosinophils probably contribute to disease chronicity by damaging epithelial and epidermal barriers, thus rendering them more vulner-

able to environmental antigens, much attention has focused on the activated CD4 T cell as the possible orchestrator of the chronic allergic immune response.

TYPE 2 T HELPER CELLS ORCHESTRATE THE CHRONIC ALLERGIC INFLAMMATORY RESPONSE The CD4 cells present at sites of allergic inflammation (Del Prete, 1992; Robinson et al., 1992; Thepen et al., 1995) and allergen-specific CD4 T cells cloned from these sites (Wierenga et al., 1990; van Reijsen et al., 1992), are skewed towards the TH2 phenotype. In man, TH2 cells are characterised by production of interleukins (IL) -4, -5, 6, -9, and IL-13 and low to negligible amounts of IFN-γ. These are indeed the cytokines expressed at the sites of allergic disease (Kapsenberg et al., 1991; Kay et al., 1991; Bentley et al., 1993) and their production has been attributed to allergenactivated T cells (Krishnaswamy et al., 1993; Ying et al., 1993; Huang et al., 1995). These TH2 cytokines are thought to be responsible for much of the pathophysiology of allergic diseases, with particular attention being focused on IL-4, IL-5 and IL-13 (Wills-Karp, 1999). IL-4 and IL-13 can both bind to the α chain of the IL-4 receptor (IL-4Rα) and hence share many biological functions. In vitro, IL-4 can (1) promote TH2 differentiation; (2) switch human B cell immunoglobulin isotype production to immunoglobulin E (IgE); (3) act with IL-3 as a growth factor for mast cells; and (4) upregulate the expression of vascular cell adhesion molecule (VCAM-1) on endothelial cells which leads to preferential migration of eosinophils into tissues. Progress has been made recently in differentiating between the relative importance of IL-4 and IL-13 in allergic inflammation using mouse models of allergic lung inflammation. Comparing the ability to induce allergic inflammation in the lungs of mice that were deficient in IL-4, IL-13 or IL-4Rα, it was found that mice in which IL-4Rα was knocked out were more resistant to developing the asthmatic phenotype than

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mice deficient in IL-4 or IL-13 alone (Grunig et al., 1998). Whereas the contribution of IL-4 may be in the early induction of TH2 differentiation and IgE isotype switching, IL-13 probably contributes more to the effector stage of the allergic immune response (Corry, 1999). Thus, eosinophilia, mucus overproduction and airways hyperresponsiveness (AHR) still occur in the absence of IL-4, but not in the absence of IL-13 (or IL-4Rα). IL-5 stimulates eosinophil development in the bone marrow, promotes the migration of eosinophils from the blood into tissues and prolongs eosinophil survival in the tissues. It can also activate eosinophils to release their cytotoxic products. Mice in which IL-5 has been knocked-out, fail to mount lung eosinophilia upon antigen challenge (Foster et al., 1996) and mice in which IL-5 is constitutively expressed show peribronchial eosinophilia, goblet cell hyperplasia, epithelial hypertrophy and collagen deposition, i.e. many of the pathological changes characteristic of asthma (Lee et al., 1997).

T-CELL ACTIVATION IN ATOPIC ALLERGIC DISEASE: THE ROLE OF DENDRITIC CELLS The primary response: T-cell sensitisation to allergen The sites affected by allergic disease typically lie at the interface between the body and its external environment, and are exposed to the daily bombardment of allergens, pollutants and pathogens. For an allergen to induce a primary response it must gain access to naïve T cells within the lymphoid tissues. According to current dogma, the dendritic cell (DC) is uniquely adapted to induce the primary response of a naïve T cell to an antigen encountered in the periphery (Steinmann, 2000). The proposed pathway for allergic sensitisation, illustrated in Figure 37.1, is best characterised in the skin where Langerhans cells (LCs)

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are responsible for the uptake, processing and transport of allergens to the local draining lymph nodes where they facilitate their presentation to naïve T cells. Epithelial surfaces elsewhere in the body also contain DC networks, notably in the conducting airways (Holt et al., 1989) and in the eye (McMenamin, 1997). Recently, advances have been made towards demonstrating in animal models whether DCs can really induce local pulmonary immunity. DCs instilled into the airways are able to traffic to local draining lymph nodes (Havenith et al., 1993), where they can drive T-cell priming (Havenith et al., 1993; Lambrecht et al., 2000). Furthermore, Lambrecht et al., (2000) have been able to show that the initial activation and proliferation of T cells is restricted to the draining lymph nodes, but that subsequently the progeny of these cells circulate systemically, displaying the phenotype of effector T helper cells.

The secondary response: establishment and maintenance of tissue inflammation Following priming, the progeny of naïve T cells enter the circulation. From here, they can enter peripheral tissues under the control of sets of adhesion molecules and in response to chemotactic gradients. Once in the tissue, what governs whether these cells immediately execute effector functions or become local memory cells is poorly understood. Nonetheless, it is these cells that will respond to secondary and subsequent challenges with the sensitising antigen. From in vitro studies, it seems likely that the clonal progeny of a stimulated naïve T helper cell will, at best, be only weakly polarised towards a TH2 phenotype, and that if appropriate signals are received in the tissues, this polarised phenotype will be reinforced during subsequent rounds of cell division (Sornasse et al., 1996). The outcome of local antigen presentation is the activation and proliferation of resident memory CD4 T helper cells to yield TH2 effector cells with concomitant release of TH2-like cytokines which lead to B cell IgE production and recruitment of eosinophils and more memory T cells from the blood, thus

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environment epithelium / epidermis

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FIGURE 37.1 Events leading to allergic sensitisation to an environmental antigen (allergen). Intraepithelial dendritic cells take up allergen arriving from the environment. In response to both constitutive and inflammatory chemokines, dendritic cells (DC) migrate through the efferent lymphatics to lymphoid organs. As they migrate they mature so that they are able to efficiently present allergen to naïve T cells (nTh). Activated naïve T helper cells proliferate and release cytokines for B cell help. Their allergen-specific progeny enter the circulation for distribution to peripheral tissue sites where they may act as effector or memory cells. Depending on signals received during antigen presentation by the dendritic cell, these progeny may be weakly polarised towards a TH2 phenotype (Th(2)). Providing sufficient IL-4 (or IL-13) is released on this first round of T-cell activation, B cells will switch to IgE production. Otherwise, Ig isotype switching to IgE may occur during a subsequent round of allergen-dependent activation. Dendritic cells may also participate in T cell–B cell interactions to facilitate B-cell proliferation and antibody production.

perpetuating the allergic inflammation (Figure 37.2). Using an in vitro allergen challenge model it has recently been shown that the T cells resident within asthmatic endobronchial biopsies release IL-5 during the first 24 hours of allergen stimulation (Jaffar et al., 1999). It is questionable how dependent memory T helper cells are on receiving CD28-mediated accessory signals for antigen-dependent activation (Gause et al., 1997) and therefore, it may not be critical to the secondary response that allergen should be presented specifically by DCs. Other candidate antigen presenting cells (APCs) for the stimulation of memory T helper cells in allergic inflammatory reactions are B cells, monocytes/macrophages, and even epithelial cells. However, B cells are present at only very low frequency in the sites most commonly affected by allergic disease (Graeme-Cook et al., 1993) and although epithelial cells can express

MHC class II and molecules involved in T-cell costimulation such as CD40, it seems unlikely that they posses the necessary cellular machinery to pick up, process and present antigenic peptides to T helper cells. Turning to cells of the monocyte/macrophage lineage that are abundant in allergic inflammatory infiltrates, it seems they are unable to induce full and sustained T-cell activation. For example, in the respiratory mucosa, alveolar macrophages, and to a lesser extent, interstitial macrophages, fail to stimulate antigen-dependent Tcell proliferation, although they do support cytokine secretion (Strickland et al., 1996). This has been attributed to nitric oxide (NO) released by the macrophages that inhibit kinases such as Jak3/STAT, thus attenuating full T-cell signalling (Bingisser et al., 1997) although additional mechanisms may exist by which pulmonary macrophages suppress T-cell function (Lee et al.,

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environment

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FIGURE 37.2 The mechanisms underlying cell-mediated allergic inflammation that contribute to the maintenance of chronic inflammation. Memory TH2 (mTh2) cells reside in the tissue. They may be preferentially attracted through the action of specific chemokines. mTh2 cells are activated locally by allergen presented by dendritic cells, although some macrophage populations can inhibit this process, these mechanisms may be less efficient in atopic subjects. Activated TH2 cells become efficient effector cells (eTh2) releasing type 2 cytokines which bring about local eosinophilia, B cell IgE production and may mediate epithelial cell damage. Note the central role played by FcεRI receptors. Allergen and specific IgE are bound to FcεRI on the surface of the DC. This leads to allergen focusing and increased activation of allergen-specific mTh2 cells.

1999). Macrophages also inhibit T-cell activation indirectly by downregulating DC function. Again, this action is mediated by macrophagederived NO which impairs local DC maturation (Holt et al., 1993). Thus, following allergic sensitisation, it is likely that the DC plays a continued role in the local stimulation of memory T cells at the sites of allergic inflammation. In vivo evidence for the importance of lung DCs in the secondary immune response has come from a murine model of TH2mediated airway inflammation. Depletion of DCs from antigen-sensitised mice led to failure to develop eosinophilic infiltration upon reexposure to inhaled antigen (Lambrecht et al., 1998). In man it is more difficult to prove the importance of DCs in the secondary immune response. CD1a cells isolated from the bronchoalveolar lavage fluid of atopic asthmatics, have been shown to activate allergen-specific memory TH2 cells in vitro (Bellini et al., 1993). In vivo, circumstantial evidence suggests a role for DCs in the maintenance of allergic inflamma-

tion. First, DCs within the epithelia lining the interfaces between the body anditsexternalenvironment, are ideally situated to encounter allergens. Second, the intraepithelial DC populations are dynamic and readily respond to inflammatory stimuli. While constitutive chemokines such as macro-phage inflammatory protein (MIP-3α (CCL20)) probably control the steadystate turnover of intraepithelial DCs (Charbonnier et al., 1999), allergens or other environmental agents up-regulate local expression of inflammatory chemo-kines, resulting in recruitment of a pulse of DCs to the site of challenge (McWilliam et al., 1996; Allavena et al., 1999). Indirect evidence that a similar process may operate at sites of allergic inflammation comes from reports of increased levels of known DC chemoattractants (MCP-4 (CCL13), RANTES (CCL5), MIP-1α (CCL3)) in the lung and skin following allergen challenge (Homey and Zlotnik, 1999) and increased numbers of DCs in the lungs of asthmatics (Bellini et al., 1993; Tunonde-Lara et al., 1996) and hayfever sufferers

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(Fokkens et al., 1989). Furthermore, if these DCs produce the CCR4 ligands thymus and activation-regulated chemokine (TARC (CCL17)) or macrophage-derived chemokine (MDC (CCL22)), they might preferentially attract and interact with TH2 cells expressing this receptor (Zlotnik and Yoshie, 2000). A final feature of DCs in human allergic inflammation is their expression of the high affinity IgE receptor (FcεRI). Particularly in atopic dermatitis, but probably also in other allergic diseases, FcεRI on DCs is thought to play a major role in the perpetuation and escalating chronicity of allergic inflammation

FcεRI ON DENDRITIC CELLS: A MOLECULE MEDIATING DISEASE CHRONICITY Structure of FcεRI on dendritic cells FceRI, a member of the antigen receptor superfamily, is a tetrameric complex comprising an α, a β- and two disulfide-linked γ-chains (Turner and Kinet, 1999). The α-chain mediates the high-affinity binding to IgE (Kd10 9 10 10 M). The β- and γ-chains each carry a cytoplasmic signalling immunoreceptor tyrosine-based activation motif (ITAM), but while the γ-chains are clearly obligate for FcεRI-mediated signal transduction, this is not the case for the β-chain, which has been ascribed a role in the amplification of the γ-chain signal (Lin et al., 1996). In rodents, FcεRI is confined to the surface of mast cells and basophils and is always present as an αβγ2 tetramer. However, in humans, while messenger RNA (mRNA) for FcεRI α- and γ-subunits was detected by RTPCR analysis of LCs (Jürgens et al., 1995) and circulating DCs (Maurer et al., 1996), transcripts for the β-chain were not detected (Maurer et al., 1996), or were present in DCs from only a minority of donors (Jürgens et al., 1995). This suggests that DCs might express FcεRI as an αγ2 complex. Indeed, Maurer and co-workers (1996) have shown that FcεRI immunoprecipitated from the surface of human periph-

eral blood DCs lacks the β-subunit. These findings are in keeping with the current hypothesis that both αβγ2 and αγ2 complexes of the human FcεRI exist, with expression of the αγ2 form not being restricted to mast cells and basophils (Turner and Kinet, 1999).

FcεRI expression by dendritic cells and its regulation Early studies demonstrated that LCs from subjects with atopic dermatitis carried surfaceassociated IgE (Bruijnzeel-Koomen et al., 1986), which was subsequently shown to be bound to FcεRI (Bruijnzeel-Koomen et al., 1988b; Bieber and Ring, 1992; Klubal et al., 1997). FcεRIbearing DCs have now been described in various tissue sites including normal skin (Wang et al., 1992), the lung (Semper et al., 1995; Tunon-deLara et al., 1996) and the nasal mucosa (Rajakulasingam et al., 1997) and also on circulating dendritic cells (Maurer et al., 1996; Holloway et al., 2000). In allergic disease, the numbers of FcεRI-positive dendritic cells are increased as shown in the asthmatic lung (Semper et al., 1995, Tunon-de-Lara et al., 1996), as are the numbers of cell surface FcεRI molecules, clearly demonstrated on LCs isolated from atopic dermatitis lesions (Wollenberg et al., 1996). In atopic dermatitis, levels of FcεRI on LCs are significantly higher at lesional sites compared to uninvolved areas, although LCs at nonlesional sites still bear higher receptor numbers than in nondiseased skin (Wollenberg et al., 1996). Indeed, it has been suggested that, amongst skin diseases, high expression of FcεRI on epidermal LCs may be a diagnostic marker of atopic dermatitis (Wollenberg et al., 1995). Several mechanisms can be envisaged to explain the increase in both numbers of FcεRIDCs and levels of DC FcεRI expression in atopic disease. If FcεRI expression is intrinsically high in the circulation in atopy, the enhanced recruitment of DCs in response to chemokines generated at sites of allergic inflammation would bring FcεRI-positive DCs into the tissue. However, levels of FcεRI expression on HLA-DRhi lineage DCs in the peripheral blood of atopic

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asthmatics and nonatopic control subjects, were found to be equivalent (Holloway et al., 2000). Alternatively, it is possible that in atopy, FcεRI is preferentially expressed on a subset of peripheral blood DCs that are selectively recruited to the sites of allergic disease. However, it seems most probable that the inflammatory environment at sites of allergic disease acts to further increase FcεRI on DCs. Two factors characteristic of allergic inflammation, namely IgE and IL-4, have been identified as having a modulatory role on FcεRI expression. On mast cells (Yamaguchi et al., 1997) interaction of FcεRI with its ligand can upregulate receptor expression. A similar relationship is seen for monocytes (Reischl et al., 1996) and data suggest that FcεRI expression by monocytederived DCs may be similarly regulated by receptor ligation (Semper et al., 1998). However, investigation of ex vivo receptor expression on LCs and other DCs is complicated by the fact that these cells are prone to rapidly change their phenotype upon isolation, with FcεRI expression being particularly affected by some cell isolation protocols (Reischl et al., 1996). Nonetheless, it is well established that the amount of total IgE in the serum correlates with receptor expression and IgE loading on monocytes (Maurer et al., 1994) and LCs (BruijnzeelKoomen et al., 1986; Wollenberg et al., 1996). A major advance in understanding the regulation of FcεRI expression was the discovery of the receptor-inducing effect of IL-4. Both the transcription of FcεRI-α and surface expression of the receptor complex is markedly increased by IL-4 in human mast cells (Xia et al., 1997). IL-4 upregulates FcεR1-α mRNA expression in monocyte-derived DCs (Semper et al., 1998) and increases intracellular expression of FcεRI-α in DCs derived from CD34 progenitors (Geiger et al., 2000), yet in both types of in vitro generated DC, this cytokine fails to increase surface FcεRI expression. This may be a consequence of the inability of these cells to produce FcεRI-β, which has a role in stabilising the FcεRI complex for transport through the Golgi apparatus to the cell surface (Turner and Kinet, 1999). However, recent results challenge this idea, as transcripts

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for FcεRI-β are consistently detected in monocyte-derived dendritic cells (Semper et al., 1998) and in circulating dendritic cells (Hartley et al., 1996) and weak immunoreactivity for the β subunit has been seen in monocyte-derived dendritic cells (Semper et al., 1998). Thus, further studies are needed to examine the effect of IL-4 and IgE on DCs, focusing in particular on the modulation of FcεRI subunits, the composition of the surface receptor and the possibility of counter-regulatory mechanisms.

THE FUNCTION OF FcεRI ON DENDRITIC CELLS On LCs, FcεRI carrying specific IgE can bind allergen that is then internalised by receptormediated endocytosis (Jürgens et al., 1995). Uptake of allergen by this route is of superior efficiency and specificity to the alternative modes of antigen capture, namely nonspecific adsorption, fluid phase pinocytosis and macropinocytosis. Thus, uptake of allergen via FcεRI, focuses antigen uptake towards those allergens to which the individual is sensitised. FcεRI-mediated allergen uptake targets the allergen into the MHC class II processing pathway, sensitive to inhibitors of cathepsin S (Maurer et al., 1998). Here the allergens are proteolytically processed into peptides and loaded into the binding groove of MHC class II mol-ecules. The outcome of FcεRI-mediated allergen focusing is that allergen-derived peptides will occupy a greater proportion of the total peptide repertoire displayed by MHC class II molecules at the DC cell surface. This is thought to enhance the activation of allergen-specific T cells as has been shown in vitro, when IgEdependent antigen presentation is mediated by FcεRI on either monocytes or peripheral blood DCs (Maurer et al., 1995, 1996). On a per cell basis, Maurer and co-workers (1996) have suggested that DCs are approximately 10 times more potent than monocytes at FcεRI-mediated IgE dependent allergen presentation. Compared to allergen taken up by bulk phase pathways alone,

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FcεRI-mediated uptake enhanced the activation of allergen specific T cells 100- to 1000-fold (Maurer et al., 1995). The cascade of signalling events caused by ligation of FcεRI by IgE has been wellcharacterised for mast cells and basophils (Turner and Kinet, 1999). Aggregation of surface FcεRI by crosslinking activates Lyn, which is associated with the β-chain. This then phosphorylates the β- and γ-chain ITAMs leading to recruitment of Syk to the γ-chains and its activation. Activated Syk can then interact with several alternative intracellular signalling pathways. In contrast, little is known about FcεRI signalling in human DCs. It has been suggested that due to the lack of a β subunit, the competence of the αγ2 complex on DCs to signal will be compromised in two ways (Turner and Kinet, 1999). First, without the β subunit to assist receptor trafficking to the cell surface, surface receptor densities are lower than on mast cells. Second, without the signal amplifying function of the β subunit, interaction of Lyn with the FcεRI-γ subunits will occur in a less directed way, reducing the overall signalling efficiency of the αγ2 complex. Thus, in DCs, only moderate induction of intracellular signalling pathways by FcεRI crosslinking might be predicted. In one study, this prediction is borne out. Cross-linking FcεRI on LCs induced tyrosine phosphorylation of several proteins in a manner directly proportional to the density of surface receptor expression ( Jürgens et al., 1995). However, this tyrosine phosphorylation did not necessarily lead to an increase in free intracellular calcium that would be indicative of cell activation. Again, cells showing full activation upon FcεRI cross-linking, tended to display higher FcεRI expression and cells from atopic dermatitis lesional skin were consistently activated by FcεRI cross-linking (Jürgens et al., 1995). Preliminary data suggest that FcεRI crosslinking on DCs may induce sufficient intracellular signalling to upregulate molecules that might enhance DC–T cell interaction (Katoh and Bieber, 1998). Factors that might skew T helper cell differentiation towards a TH2 phenotype may also be produced.

In conclusion, it seems reasonable to suppose that, in an atopic environment with elevated levels of IgE and increased expression of FcεRI on DCs, this receptor will, through allergen focusing, exacerbate allergic inflammation by increasing the activation of allergen specific TH2 cells. Through the action of type 2 cytokines, the T cell may in turn further enhance IgE and FcεRI expression on the dendritic cell, thus completing a vicious circle of events that results in chronic allergic inflammation (Figure 37.3). The central importance of IgE in this process is illustrated by clinical trials of a humanised anti-IgE monoclonal antibody directed against the FcεRI-binding domain of human IgE (Kinet, 1999). This agent reduces IgE synthesis leading to downregulation of FcεRI expression (on basophils) and reduces both early and late phase responses in asthmatic subjects. Could this therapeutic agent be reducing FcεRI-mediated antigen presentation by DCs, thus ameliorating the symptoms of the late-phase (and chronic) response?

DENDRITIC CELLS IN TH2 POLARISATION: THEORETICAL ASPECTS Several theories have been advanced recently to explain how dendritic cells might drive TH2 polarisation. In particular, it has been suggested that whereas myeloid DCs promote TH1 differentiation in vitro, CD4 T cells will differentiate along a TH2 pathway under the influence of CD4, CD11c, CD3 blood-derived lymphoid DCs (Rissoan et al., 1999). However, when it is considered that allergens are concentrated at the body’s epithelial surfaces, the most relevant DC to handle these antigens is the intraepithelial DC. These are of myeloid lineage, irrespective of whether they derive from CD34 progenitors via a CD14 CD1a intermediate or whether they are derived from tissue-infiltrating monocytes which differentiate into DCs in situ. For this reason, the section that follows deals solely with the ways in which myeloid DCs might drive TH2 polarisation (see Figure 37.4).

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531

IL-4

Eo

Th2 antigen presentation & Th2 skewing

?IgE IL-4

IL-4 ?

Dc

?IgE

?

Mo\ Mac

Th2

IgE production

B

FceRI mediated allergen uptake

Th2

IL-4

IL-4

IL-4

IgE-mediated FceRI increase

Mc

IgE

IgE

degranulation of effector cells; mediator release

IL-4

FIGURE 37.3 The interconnecting pathways regulating FcεRI expression in allergic inflammation. The known function of FcεRI on dendritic cells is to enhance allergen uptake and activation of allergen specific T helper cells. It may also contribute to the skewing of the TH2 response, thus exacerbating the production of IL-4 and IgE. These factors may further upregulate FcεRI expression on the dendritic cell, as well as on other cell types. Whether FcεRImediated allergen focusing on dendritic cells has consequences for B cell function remains to be demonstrated.

PGE2

IL-10 histamine chemokines

1. Strength of TCR Signal Low Ag dose Low affinity peptide

3. Reducing IL-12 production by autocrine factors IL-10 by microenvironment e.g. PGE2

IL-10 IL-12

nTh

DC

nTh

2. Costimulation Amount of costimulation (immature dendritic cell) Particular receptor/ligand usage e.g. OX40/OX40L e.g. CD30/CD30L e.g. 4-1BB/4-1BBL

IL-4?

IL-4 from e.g. mast cell e.g. NK T cell IL-4

IL-6 IL-10

4. Promoting IL-4 production IL-4 direct from dendritic cell (?) Factors promoting early T cell autocrine IL-4 production e.g. IL-6 IL-4 from other cell types

Key A

allergen

CD4

CD80/86

MHC II

peptide

TCR

IgE

CD28

FceRI

FIGURE 37.4 Mechanisms by which dendritic cells may initiate TH2 skewing during T cell priming. Processes mediated by both cell-contact (1 and 2) and soluble factors (3 and 4) may be involved in the determination of TH2 skewing by dendritic cells. DENDRITIC CELLS IN DISEASE

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During the interaction between an APC and a T cell, several factors have been shown to influence the subsequent pathway of differentiation taken by the T helper cell. These include: (1) antigen dose and the availability of antigen peptide-loaded MHC class II molecules at the APC cell surface; (2) peptide affinity; (3) the nature and level of surface-expressed costimulatory molecules; and (4) the production of soluble mediators by the APC and their regulation by microenvironmental stimuli.

Antigen dose and peptide affinity In some experimental systems, high doses of antigenic peptide cause naïve T cells to differentiate into TH1-like cells, while low doses of the same peptide induce TH2 differentiation (Constant et al., 1995). With the exception of food allergens, environmental allergens are typically delivered to the body at low dose and may thus favour a TH2 response. DCs will efficiently handle even these low doses of antigen, due to highly efficient mechanisms for both nonspecific and receptor-mediated antigen uptake, high levels of MHC class II surface expression and the capacity for serial triggering of T-cell receptors (TCR) by individual MHC–peptide complexes (Valitutti et al., 1995). The peptide affinity for MHC class II or the TCR may also affect T-cell differentiation (Pearson et al., 1997). High affinity peptides have been shown to drive TH1 differentiation in vitro, whereas low-affinity peptides drive TH2 differentiation (Constant and Bottomly, 1997). The route by which the antigen is captured and processed by the DC may affect the affinity of the resulting peptides. For example, using a model native antigen, differences were found in the peptides produced by untreated and IL6-treated dendritic cells (Drakesmith et al., 1998). The duration of TCR stimulation has also been shown to influence the phenotype of polarised T cells. T cells displaying a TH1 phenotype were found following a relatively short period (24 h) of TCR engagement whereas development of TH2 cells required prolonged TCR engagement (96 h)

(Iezzi et al., 1999). It has been suggested that low-affinity peptides favour TH2 development as a result of maintained serial triggering that in effect prolongs TCR engagement (Iezzi et al., 1999). It remains to be determined how this translates into terms of the formation and stability of the immunological synapse between a dendritic cell and a T cell.

Costimulation Much work has focused on the role of the B7 family of costimulatory molecules in the determination of T cell phenotype. Interestingly B7-1 (CD80) and B7-2 (CD86) are differentially expressed and regulated on DCs (Kawamura and Furue, 1995). Early studies suggested that engagement of CD28 can direct naïve T cells to differentiate along a TH2 pathway (Webb and Feldmann, 1995) and even that the preferential differentiation of TH1 or TH2 cells from uncommitted T helper precursors is induced by ligation of CD28 with B7-1 or B7-2, respectively (Kuchroo et al., 1995). However, in terms of TH2 development in allergy, this relationship does not hold true (Green, 2000), although human studies suggest that CD28 ligation is required for the local activation of TH2 cells and the release of type 2 cytokines in the asthmatic airway (Jaffar et al., 1999). Other pairings of costimulatory receptors on T cells and their ligands on APCs have been implicated in the development of TH2 polarisation, notably, CD30 and CD30-ligand (Del Prete et al., 1995), OX40 (CD134) and OX40 ligand (Delespesse et al., 1999; Akiba et al., 2000) and 41BB and 4-1BB ligand (Chu et al., 1997). However, a recent study suggests that the importance of individual costimulatory pairings on TH1/TH2 differentiation is strongly influenced by the number and affinity of peptide/MHC complexes (Rogers and Croft, 2000). Thus, it is the total strength of the integrated signal the T cell receives that determines its differentiated fate, rather than the signal delivered by ligation of a particular costimulatory receptor.

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Soluble factors produced by dendritic cells Under appropriate stimulation conditions, for example the crosslinking of CD40 by CD40ligand that occurs during the interaction between DCs and T cells, DCs may be induced to produce cytokines. The most potent mediator driving TH2 polarisation in vitro is IL-4 (Sornasse et al., 1996; Ohshima and Delespesse, 1997). Although murine DCs recently were shown to produce low levels of IL-4 in response to Rauscher leukemia virus (Kelleher et al., 1999), human DCs are not known to produce this cytokine. The most important cellular source of IL-4 is a subject of debate in the field of allergy research. In established allergic inflammation, mast cells, eosinophils and TH2 cells themselves, could all be sources of IL-4. However, during primary allergic sensitisation, the source of IL-4 is unknown although one candidate cell type is the NK T cell (Porcelli and Modlin, 1999). First identified in mice, but with an analogous CD4 CD8 TCR-α/β human T cell population (Davodeau et al., 1997), these cells are potent and rapid producers of IL-4 upon TCR ligation and promote IgE production in vivo (Bendelac et al., 1996). They recognise novel lipid antigens presented by CD1d, and presumably critically depend on DCs for antigen presentation. Other T cells that recognise lipid and glycolipid components of, for example, mycobacteria, in a CD1-a, -b or -crestricted manner exist. It remains to be determined whether these T cells could also be potent sources of IL-4 when stimulated by DCs. An alternative source of IL-4 in the primary immune response may be the responding T cell itself. IL-6, produced by human DCs in response to inflammatory stimuli (Verhasselt et al., 1997), may induce TH2 differentiation since IL-6 has been shown to initiate IL-4 production by naïve T cells (Rincon et al., 1997). Similarly, in vitro derived DCs have been shown to produce IL-10 (de Saint-Vis et al., 1998) that may directly promote TH2 differentiation by inducing T cell IL-4 production (Umetsu and DeKruyff, 1997). Both inflammatory and constitutive chemo-

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kines of the CC and CXC families are produced by DCs and are able to assist in establishing contact between DCs and T cells. Chemokines may also directly influence the development of TH1/TH2 cells. For example, monocyte chemotactic protein-1 (MCP-1 (CCL2)), produced by DCs following stimulation with LPS, has been shown to enhance IL-4 secretion from T cells (Karpus et al., 1998). MCP-1-deficient mice show a correspondingly impaired TH2 immune response, although mice deficient for the MCP-1 receptor, CCR2, demonstrate defective TH1 responses (Boring et al., 1997).

Suppression of dendritic cell IL-12 production by the microenvironment DCs are potent producers of bioactive IL-12 p70 in response to CD40 ligation (Cella et al., 1996) or following exposure to pathogens including Staphylococcus aureus Cowan strain I and LPS (Hilkens et al., 1997). IL-12 is classically thought of as the most efficient promoter of TH1 differentiation (Heufler et al., 1996). However, it is the amount of IL-12, rather than its absolute presence or absence that determines the resulting effector T helper cell phenotype. Using human monocyte-derived DCs as a model system, it has been demonstrated that pathogen-derived factors, or pathogen-induced tissue responses, influence the immature DC to adjust the level of IL-12 it will subsequently produce upon stimulation (Kalinski et al., 1999). In effect, information on the nature of the antigen encountered by the immature DC in the periphery is provided to the naïve T cells in lymphoid organs not only via the antigen-derived peptide (signal 1) in conjunction with adequate costimulatory molecules (signal 2), but also via a T cell polarising signal, ‘signal 3’ (Kalinski et al., 1999). Mediators known to reduce IL-12 production by DCs that are of particular interest in atopic disease, include prostaglandin E2 (PGE2) and IL10. Human monocyte-derived DCs generated in the presence of PGE2 show reduced IL-12 production and consequently stimulate naïve T cells to differentiate into a TH2 phenotype producing high levels of IL-4 and IL-5 and low levels

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of IFN-γ (Kalinski et al., 1997). In atopic disease, monocytes show increased PGE2 production as do epithelial cells and recently, even monocytederived DCs were reported to produce PGE2 (Kiertscher et al., 2000) that may work in an autocrine fashion to affect IL-12 production. Similar processes may be acting in vivo since IL12 production by monocytes is reduced in patients with allergic asthma (van der Pouw Kraan et al., 1997). IL-10 has been suggested to promote TH2 differentiation as DCs exposed to IL-10 in combination with IL-1β and TNF-α in vitro have reduced IL-12 p70 production (Kalinski et al., 1998) and murine DCs treated with IL-10 have been demonstrated to induce TH2 differentiation (De Smedt et al., 1997). However, IL-10-treated DCs are more commonly thought to be involved in tolerance induction, as immature DCs exposed to IL-10 show a reduced capacity to stimulate CD4+ T cells in an allogeneic mixed lymphocyte reaction (De Smedt et al., 1997; Steinbrink et al., 1997). Several other agents have been described that can suppress IL-12 production, including histamine (van der Pouw Kraan et al., 1998) adrenergic agonists (Panina-Bordignon et al., 1997), MCP-1 to -4 (Braun et al., 2000) and type 1 interferons (McRae et al., 1998). IL-13, which is produced by in vitro derived mature DCs following PMA-ionomycin activation (de Saint-Vis et al., 1998), has been shown to indirectly promote TH2 differentiation by inhibiting the production of IL-1β, IL-1α, and IL-12 (de Waal et al., 1993). If IL-13 is produced by DCs following physiological stimulation, this may provide an additional autocrine route by which DCs can influence TH2 differentiation.

DENDRITIC CELLS AND THE DEVELOPMENTAL PROGRESSION OF ALLERGIC DISEASE Sensitisation to allergens and the establishment of weak TH2 polarisation probably occurs very early in life (Holt, 1999). Allergen-specific T-cell reactivity to aero-allergens such as house dust

mite and birch and timothy grass pollen, and to common food allergens such as cow’s milk and hen’s egg (Warner et al., 1994; van DurenSchmidt et al., 1997; Szepfalusi et al., 1997; Prescott et al., 1998) has been demonstrated at birth. This implies that T-cell priming occurs in the fetus in response to trace amounts of inhaled and food allergens carried in the maternal vasculature. In support of this, the conceptus has been shown to be able to mount an allergenspecific immune response from 23 weeks of gestation (Jones et al., 1996). Critical to understanding how in utero sensitisation leads to the development of allergic disease, is the finding that these proliferative responses of cord blood T cells to innocuous environmental antigens can be demonstrated for most neonates, irrespective of whether they go on to develop atopy. These allergen-specific T-cell responses at birth are also universally skewed towards preferential production of type 2 cytokines (Prescott et al., 1998), perhaps as a result of the low antigen dose that is likely to occur in the fetus (Secrist et al., 1995; Carballido et al., 1997). This has led to the concept that individuals going on to develop allergic disease consolidate the type 2 response, whereas normal individuals lose this bias toward TH2 immunity by a process of immune deviation within the first few months or years of life (Holt, 1999). To understand fully what processes are different in the infant that goes on to develop allergic disease, knowledge of the processes operating in the skin and mucosae of normal subjects is required. Soluble antigens entering the body through the mucosal surfaces normally elicit only transient local secretory immunity that is replaced by long-term peripheral unresponsiveness. Using a murine model of inhalation tolerance, in which aerosolised ovalbumin failed to trigger a productive pulmonary immune response, the ability of different cytokines to break this tolerance in the airways has been tested. Transient expression of GM-CSF in the airway, led to development of lung eosinophilia, goblet cell hyperplasia, increased TH2 cytokine production and led to establishment of antigenspecific T-cell memory (Stämpfli et al., 1998).

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Numbers of DCs and macrophages in the lung were increased and these cells were implicated as being central to the sensitisation process, since GM-CSF could not break tolerance in MHC class II-deficient mice. Additional coexpression of an IL-10 transgene in this model restored tolerance (Stämpfli et al., 1999), although T-cell activation and proliferation still occurred, suggesting that a subset of regulatory T cells resembling the Tr1 regulatory cell (Groux et al., 1997) may have been activated. To what extent the DC is a target of GM-CSF and IL-10 is not fully addressed by Stämpfli and co-workers. However, Stumbles et al., (1998) found that although freshly isolated rat respiratory tract DCs were able to prime low level TH2 responses, following GM-CSF exposure, they became much more potent T cell activators and stimulated a mixed TH1 / TH2 response. One notable phenotypic difference between the fresh- and GM-CSF-treated DCs was reduced IL-10 production following GM-CSF exposure. Thus, DCs exposed to GM-CSF in the local environment may escape the suppressive actions of IL-10 and thus drive a much more productive effector and memory T helper cell response. There is increasing evidence that GM-CSF and IL-10 are important in the aetiology of chronic allergic inflammation. GM-CSF production by epithelial cells is upregulated at the sites affected by allergic disease and significantly less IL-10 is found in the lungs of subjects with asthma (John et al., 1998). This IL-10 defect has also been identified in early life in those children that are starting to show an atopic phenotype (Macaubas et al., 1999).

findings presented in the preceding section, have led to the suggestion that an altered microenvironment at sites such as the bronchial epithelium or the epidermis could predispose an individual to develop allergic disease at that anatomical site (Holgate, 1999). The tissue DC could act as the intermediary between an altered microenvironment and the immune system, being able to respond to factors such as GM-CSF by changing its antigen presentation function. It is not clear in man whether more potent DC function would produce both TH1 and TH2 responses as seen by Stumbles et al. (1998) in the rat. Factors such as the low dose of allergens and the action of gene mutations that are known to predispose towards atopy, may serve to sustain an aggressive TH2-skewed response in man. Thus, by influencing the DC, local abnormalities in epithelial cell function could drive the local development of chronic TH2-cell mediated inflammation, finally leading to the development of the clinical symptoms of allergic disease. In this scenario of escalating local inflammation leading to allergic disease, IL-10 would appear to be a potential therapeutic target for the treatment of allergy. Approaches that aim to upregulate IL-10 production in individuals with, or at risk of developing allergic disease, may reduce TH2 skewing with the ultimate goal of achieving immunological nonresponsiveness (Umetsu and DeKruyff, 1999). This might provide a useful alternative to some of the current therapeutic strategies being considered for the treatment of allergic disease.

REFERENCES FUTURE DIRECTIONS As described above, a challenge in understanding the aetiology of allergic diseases is that, although an individual may be atopic, this condition does not necessarily lead to the development of allergic disease. Furthermore, it is unclear what factors determine the anatomical site that will be affected by allergic disease. The

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38 DCs in wound healing Kristine Kikly and Michael T. Lotze GlaxoSmithKline Pharmaceuticals, King of Prussia, PA, USA

‘. . . the merit of the practical introduction of a method which has revolutionized the treatment of wounds, and changed the whole aspect of modern surgery. I am old enough to have been a dresser in a large general hospital in the pre-Listerian days, when it was the rule for wounds to suppurate, and when cases of severe pyaemia and septicaemia were so common that surgeons dreaded to make even a simple amputation. . . . Lord Lister’s experimental work on the healing of wounds led to results of the deepest moment to every individual subject to an accident, or who has to submit to an operation . . . every wound which suppurated had to be dressed, and there was the daily distress and pain, felt particularly by young children. Now, even after operations of the first magnitude, the wound may have but a single dressing, and the after-pain is reduced to a minimum.’ Man’s Redemption of Man Sir William Osler, Edinburgh, 1910 Presentation to the Natural Association for the Prevention of TB

INTRODUCTION

recruit other inflammatory cells; (2) modulate angiogenesis by production of pro- and antiangiogenic factors (Plate 38.1); and (3) promote or suppress local T cell effector function may be critically important and underappreciated roles for DCs in the setting of wound healing.

Dendritic cells emigrate into tissues and reside there with half-lives from 3 days (pulmonary and GI mucosal sites) up to 30 days in the skin. In addition to their critical role as antigen presenting cells, they have the opportunity to interact with resident stromal cells (fibroblasts, endothelial cells and macrophages), as well as epithelial cells. Additional functions of DCs in regulating local tissue homeostasis during wound healing, cancer, chronic inflammatory diseases and chronic microbial infections have not been carefully studied or, until recently, considered or identified. The ability of DCs to (1) Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

AN OVERVIEW OF WOUND HEALING The process of wound repair is initiated following receipt of an injury culminating in healing of the wound, with the ultimate aim of restoring normal tissue architecture and function. The

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sequence of events following injury, whether from surgical procedures, trauma, burns or other inducers of an inflammatory response, is very similar (Table 38.1). First there is the initiation of the coagulation cascade to attain hemostasis and form a provisional matrix for cell adhesion (Clark, 1995). Byproducts of the coagulation cascade and the complement system drive development of chemotactic factors that recruit neutrophils, initially, and then monocytes (fibrinopeptides, C5a). These cells are responsible for elimination of bacteria, debris and effete cells within the wound. The incoming immune cells, as well as the damaged tissue, secrete a variety of factors leading to new blood vessel formation regulating angiogenesis (bFGF, VEGF, endothelin), some of which are also important in regulating stromal cell proliferation, motility and viability as well as stem cell proliferation (Takakura et al., 2000). Along with the resupply of oxygen and nutrients to the area by the new blood vessels there is mesenchymal cell proliferation and connective tissue matrix TABLE 38.1

remodeling (Riches, 1996). There are many conditions resulting from poor or nonhealing wounds, such as diabetic ulcers, bed sores, chronic venous insufficiency ulcers and lesions of inflammatory bowel disease. The wound healing process occurring in the skin will be described in more detail since more substantial information is known about this organ with respect to normal and abnormal wound healing.

THE SKIN AND THE NORMAL WOUND HEALING PROCESS The skin is the largest organ in the body and provides a protective barrier against the external environment. There are three predominant layers within the skin, the epidermis, the dermis and the subcutaneous fat or hypodermis (Eckert, 1991). The epidermis is the surface layer and is comprised of epidermal keratinocytes which differentiate from basal stem cells. Cells within this basal layer fully mature to become

Summary of skin wound healing biological processes involving macrophages and DCs

Stage

Process

Role of Mφ/DC

References

Inflammatory phase

Inflammation activation

Migration in response to coagulation products and platelet factors Increase of proinflammatory cytokines/chemokines (TNF-α, IL-1β, IL-8) Influx of immune cells Differentiation of monocytes to DCs Antiangiogenesis (IFNα, IL-12, IL-18, IP10)

Sozzani, 1999 Clark, 1995; Abe, 2000; Sallusto, 1999 Clark, 1995 Randolph, 1998; Randolph, 1999; de Saint-Vis, 1998

Inflammation resolution

Secretion of protease inhibitors Downregulation of cytokine/chemokine production

Clark, 1995; Riches, 1996 Sallusto, 1999; Sato, 2000

Cell proliferation and migration

Secretion of stromal factors (IL-1β and TGFβ) Secretion of mesenchymal cell growth factors

de Saint-Vis, 1998; Nissen, 2000 Riches, 1996

Angiogenesis

Secretion of IL-8, PDGF, TGFα, TGFβ, VEGF, bFGF

Nissen, 2000; Riches, 1996

Re-epithelialization

Keratinocyte functions

Secretion of IP10, MIG

Clark, 1995

Remodeling

Scarring

Secretion of TGFβ

Riches, 1996

Granulation phase

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corneocytes in the cornified or surface layer. Interspersed within the keratinocytes are melanocytes, or the pigment producing cells of the skin, as well as Langerhans cells, professional antigen presenting cells. Below and abutting the epidermis is the dermis, which is comprised of fibroblasts producing connective tissue proteins and a variety of different cytokines. This layer provides structural integrity to the skin and support for a network of nerves and vasculature. Macrophages, mast cells and dermal dendritic cells are also found scattered throughout the dermis (WeberMatthiesen and Sterry, 1990). The innermost layer of skin is the hypodermis, providing a cushion of adipose tissue interposed between the skin and bone and muscle. Immediately following tissue injury that results in blood loss, blood clotting occurs, triggered by the release of tissue procoagulant factors from damaged cells and the adhesion of platelets to the damaged area. Clotting continues until the signals for clot initiation, including tissue factors, exposed viable epithelium or open blood vessels, are dissipated. The clot not only prevents further blood loss but provides a matrix for immune cells recruited to the area. Mediators released from platelets promote vasoconstriction, further platelet aggregation and growth factor release. Byproducts of the coagulation (fibrinopeptides, α-thrombin) and complement (C5a, C5a des Arg) cascades, bacteria as well as platelet factors (PF4, PDGF) act as chemoattractants to neutrophils and monocytes (Riches, 1996). DCs express receptors for platelet activating factor (PAF) and C5a and can chemotax in response to these mediators (Sozzani et al., 1999). In an excisional model in rats, elevated levels of mRNA and protein for macrophage migration inhibitory factor (MIF) were detected at 1 hour following wounding and stayed elevated for 24 hours (Abe et al., 2000). MIF appears to be produced by all epidermal cells at the wound margin as compared to just cells of the basal layer in the control. MIF acts as an initiator of inflammation through the regulation of other pro-inflammatory cytokines such as TNF-α, IL-1 and IL-6. These

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cytokines not only act on cells of the immune system, but also on keratinocytes and fibroblasts at the wound site, prompting cell migration, growth and extracellular matrix production. MIF appears to be an important component of the initial cytokine milieu since treatment of mice with anti-MIF antibodies doubles the time to wound healing (Abe et al., 2000). Whether MIF has a direct effect on LCs or dermal DCs is unknown, but clearly the increase in pro-inflammatory cytokines would promote maturation. In addition to the plethora of cytokines and chemokines released following tissue damage, the presence of necrosis will also promote the maturation of DCs at the wound site. Necrotic fibroblasts and blood vessels are themselves able to induce upregulation of class II and costimulatory markers on resting DCs and allows for priming of naïve T cells following migration to secondary nodal sites (Gallucci et al., 1999). Neutrophils are responsible for phagocytosis and killing of bacteria in the wound. If little or no bacterial contamination occurs, neutrophil infiltration only lasts a few days. Monocyte infiltration to the site persists due to the local sustained release of specific monocyte chemoattractants (MCP1, 2, 3). The majority of monocytes mature to a macrophage phenotype within the tissue and are important in debriding the wound through phagocytosis of tissue debris, apoptotic neutrophils and contaminating bacteria. However, using an adoptive transfer model, Randolph et al. demonstrated that approximately 25% of monocytes migrating into a tissue site will differentiate into DCs which then traffic to the lymph node (Randolph et al., 1999). It has also been demonstrated that the reverse transmigration of monocytes through tissue causes differentiation into DCs (Randolph et al., 1998). While the role of Langerhans cells (LC) or dermal DCs at the wound site is unclear there is certainly ample opportunity for resident cells or recruited cells to play a role in wound healing. In an experimental model of wound healing of human skin grafted on to a SCID mouse, LCs were found within the newly formed epithelium 2 days following wounding (Juhasz et al., 1996). There was a slight

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increase in the number of dermal DCs under the woundbed.Theobservation of LCs in the wound early in the process of re-epithelialization supports the accepted hypothesis that LCs take up antigen at an inflammatory site, but also allows the possibility that cytokines secreted by LCs and the underlying dermal DCs would participate in the recruitment of other cells necessary for proper healing. Chemokine and chemokine receptors have key roles in the trafficking of immature and mature DCs. Inflammatory chemokines are responsible for attracting cells to a site of inflammation and immature DCs express CCR1, CCR2, CCR5, CCR6 and CXCR1. Once at the site, maturation signals in the form of necrotic cells, LPS, TNF, IL-1β, etc. rapidly induce (6 hours) the production of MIP-1α and MIP-1β while RANTES slowly accumulates over a 24-hour period (Sallusto et al., 1999). Maturing DCs are able to produce large quantities of MIP-1β, up to 1 pg/cell/h (Sallusto et al., 1999). These chemokines would recruit more monocytes and lymphocytes into the area. Maturing DCs can also produce large amounts of IL-12 and IL-18, perhaps regulating the T-cell response in the area (de Saint-Vis et al., 1998; Heufler et al., 1996). Epithelial cells also respond to the chemoattractant signals generated by the wound, proliferate and migrate across the open wound. An influx of immune cells begins to move to the wound site within several hours. Transforming growth factor β (TGFβ) isoforms are increased by 1 day post wounding. Both TGFβ1 and TGFβ3 are produced by human epithelial keratinocytes. The ratio of TGFβ1 to TGFβ3 appears to contribute to the rate and quality of scar formation (O’Kane and Ferguson, 1997). TGFβs are also produced by platelets, activated macrophages, lymphocytes and DCs (de Saint-Vis et al., 1998). The mechanisms responsible for altering the ratio of TGFβ1 to TGFβ3 are not clearly defined. Although several cell types can produce various TGFβs, the influx of macrophages, many of which produce this cytokine, appears to be crucial for normal healing. In a full-thickness linear skin incision rat model of glucocorticoidinduced wound healing deficit, exogenous TGFβ

was able to reverse inhibition of healing (Pierce et al., 1989). Glucocorticoids also nearly eliminate the influx of neutrophils and macrophages and cause a delayed and reduced density of fibroblasts. Local application of TGFβ increases the breaking strength values by almost 200% after 1 week. This appears to be due to the increased fibroblast influx and activation induced production of procollagen type I noted in the TGFβ-treated mice (Pierce et al., 1989). TGFβ is also an important cytokine for the development of epidermal LCs (Strobl et al., 1997). Increases in TGF-β at the wound site may promote the maturation of LCs in the area. TGFβ1 has also been shown to influence immature DCs by causing the upregulation of expression of CCR1, CCR3, CCR5, CCR6 and CXCR4 and blocking the upregulation of CCR7 in response to TNFα (Sato et al., 2000). TGFβ1 did not change the DCs ability to express MHC or costimulatory markers or the ability to stimulate T cells, however, the change in chemokine receptor expression could affect the normal trafficking of the DCs. Angiogenesis is also initiated, most likely by tissue hypoxia and release of growth factors from damaged tissue and wound macrophages (Nissen and DiPietro, 2000). Fibroblast growth factor-2 (FGF-2 or basic FGF) levels increase immediately after skin injury and are most likely derived from preformed FGF-2 released following platelet degranulation, from damaged cells or subsequent to degradation of the extracellular matrix. The heparin-binding protein FGF-2 stimulates endothelial cell activation and induces angiogenesis in vivo. FGF-2 null mice have delayed wound healing, impaired reepithelialization and collagen formation implying that the early increase in FGF-2 sets off a cascade of events critical for wound healing (Ortega et al., 1998). The result of this initial burst of FGF-2 is the initiation of angiogenesis and the production of other angiogenic factors, such as vascular endothelial cell growth factor (VEGF). VEGF levels in wounds increase over days, probably due to the production by keratinocytes, endothelial cells, fibroblasts and macrophages. The local increase in VEGF

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concentration acts as a chemoattractant and mitogen for endothelial cells and promotes angiogenic activity (Nissen and DiPietro, 2000). FGF-2 and VEGF are important angiogenic mediators observed to be present during wound healing. Wound macrophages have also been shown to produce TNF-α, PDGF, TGFα, TGFβ, IGF-1 as well as VEGF (Nissen and DiPietro, 2000). The production of angiogenic factors produced by LCs or dermal DCs has not been carefully studied however, they are known to produce both TGFβ and IL-8. Successful wound healing depends on the interaction of macrophages, fibroblasts and blood vessels and DCs. Macrophages or DCs provide growth factors promoting angiogenesis and fibroplasia, fibroblasts make extracellular matrix allowing for the organized regrowth of cells and blood vessels carry oxygen and nutrients required to sustain the process. As new cells grow into the wound space there needs to be extracellular matrix remodeling to allow for the formation of a collagenous scar. The enzymes responsible for the remodeling come from macrophages and endothelial cells.

ABNORMAL WOUND HEALING Abnormal or prolonged wound healing can occur due to underlying disease pathogenesis, such as diabetes or chronic venous insufficiency, or as a result of wound size or depth, i.e. for thermal burns. Infection can also contribute to prolongation or deviation from the normal wound healing process. It has long been assumed that the elderly have problems with nonhealing wounds, because healing is defective in this population. Careful studies recently performed have cast doubt on this notion (Ashcroft et al., 1995). Clearly, there are changes in the skin of elderly individuals, particularly in sun-exposed skin. The number of LCs decrease with age by 50% in sun-exposed skin and 14% in non-sun-exposed skin from the ages of 24–65 years (Ashcroft et al., 1995). The number of melanocytes also decreases and the collagen

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organization changes with age. In both humans and mice, elderly subjects had a delayed influx of monocyte/macrophages and lymphocytes and slower re-epithelalization (Ashcroft et al., 1997, 1998). While the collagen deposition was slower in the old mice, the scar quality was improved. Young animals formed dense scar tissue quickly and the older animals formed a scar with an organization more like normal tissue (Ashcroft et al., 1997). The changes in the inflammatory response and scar quality are similar to those observed in fetal healing (Cowin et al., 1998). Since otherwise healthy elderly individuals are able to heal normally, albeit slightly slower, there must be other contributing factors for nonhealing wounds. Poor nutrition, certain medications and stress could impact on wound healing directly or indirectly through suppression of the immune system and increased rate of infection. Diabetes and chronic venous insufficiency (CVI) can result in nonhealing ulcers, particularly in the leg and foot area. Venous reflux or venous hypertension (i.e. raised venous pressure) are the main hemodynamic abnormalities associated with the skin changes of CVI. Approximately 1% of adults will be affected by venous ulcerations at some time in their lives (Cheatle et al., 1991). The exact mechanism responsible for initiation and maintenance of the ulcer is not clear, but the ulcers persist for many months and are prone to recur in susceptible individuals. The current theory is that venous hypertension leads to stasis of the blood in the small capillaries of the leg causing white blood cells (WBCs) to become trapped and activated. They in turn activate the endothelial cells in the area, initiate thrombosis and increase permeability. The capillaries become occluded, more WBCs are activated, a fibrin cuff forms around the capillary and there is edema in the area. This results in local ischemia, cytokine and chemokine production and inflammation (Cheatle et al., 1991). The mechanisms for allowing resolution of the local inflammation versus exacerbation and transformation to an ulcer are not understood. A recent study has examined the differences in expression of IL-10 and

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GM-CSF and the presence of APCs in CVI ulcers versus other chronic and acute wounds. HLADR+, CD68+ APCs (macrophages and dermal DCs) were increased in CVI ulcers compared to other wounds. There was a significant correlation between the number of CD68+ cells and CVI ulcer time duration (Li et al., 1998). CVI ulcers also had IL-10 present in the wound tissue in the hypertrophic epidermis in contrast to other wounds where little or no IL-10 was observed (Li et al., 1998). IL-10 is predominantly a TH2 cytokine with anti-inflammatory properties and keratinocytes can produce IL-10 when given an appropriate stimulus. Punch biopsies of human skin demonstrated IL-10 expression in keratinocytes in tape-stripped skin, poison ivy dermatitis and Sezary syndrome, but not in skin from psoriatic plaques (Nickoloff et al., 1994). Interestingly, CD80 expression on LCs is downregulated by IL-10 and upregulated by GM-CSF (Chang et al., 1995). In a small clinical study, CVI ulcer patients were injected with GM-CSF at the wound margins resulting in complete healing of the ulcers (Marques da Costa et al., 1994). One could speculate that the increased levels of IL-10 produced by the keratinocytes in CVI ulcers inhibits the function of the macrophages and LCs (i.e. downregulation of CD80 and perhaps other effects) in the wound leading to abrogation of the normal healing process. Interestingly, in psoriasis, a disease that is characterized by benign hyperproliferation of keratinocytes (too much wound healing) there is no IL-10 found in the lesions (Nickoloff et al., 1994). In normal wound healing, the number of LCs increases at the site and the activated LCs express inducible nitric oxide synthase (iNOS) and subsequent nitric oxide (NO) production (Morhenn, 1997). NO can recruit T cells, activate keratinocytes and promote dilation of blood vessels, all characteristics of psoriatic plaques. IL-10 can inhibit the production of NO perhaps restoring homeostasis between LCs and keratinocytes as normal wound healing is completed. One could hypothesize that psoriatic plaques and CVI ulcers are two sides of the same coin, both result from the imbalance of LCs– keratinocyte interactions.

REFERENCES Abe, R., Shimizu, T., Ohkawara, A. and Nishihira, J. (2000). Biochem. Biophys. Acta 1500, 1–9. Ashcroft, G.S., Horan, M.A. and Ferguson, M.W.J. (1995). J. Anat. 187, 1–26. Ashcroft, G.S., Horan, M.A. and Ferguson, M.W.J. (1997). J. Invest. Dermatol. 108, 430–437. Ashcroft, G.S., Horan, M.A. and Ferguson, M.W.J. (1998). Lab. Invest. 78, 47–58. Chang, C., Furue, M. and Tamaki, K. (1995). Eur. J. Immunol. 25, 394–398. Cheatle, T.R., Sarin, S., Coleridge Smith, P.D. and Scurr, J.H. (1991). Eur. J. Vasc. Surg. 5, 115–123. Clark, R.A.F. (1995). The Molecular and Cellular Biology of Wound Repair, New York: Plenum Press, pp. 3–50. Cowin, A.J., Brosnan, M.P., Holmes, T.M. and Ferguson, M.W.J. (1998). Dev. Dyn. 212, 385–393. de Saint-Vis, B., Fugier-Vivier, I., Massacrier, C. et al. (1998). J. Immunol. 160, 1666–1676. Eckert, R.L. (1991). In: Mukhtar, H. (ed.) Pharmacology of the Skin, Boca Raton: CRC Press, pp. 4–12. Gallucci, S., Lolkema, M. and Matzinger, P. (1999). Nat. Med. 5, 1249–1255. Heufler, C., Koch, F., Stanzl, U. et al. (1996). Eur. J. Immunol. 26, 659–668. Juhasz, I., Simon M. Jr, Herlyn, M. and Hunyadi, J. (1996). Immunol. Lett. 52, 125–128. Li, Q., Doyle, J.W., Roth, T.P., Dunn, R.M. and Lawrence, W.T. (1998). J. Surg. Res. 79, 128–135. Lotz, M.T., Odoux, C. and Luketich, J. (2001). Lung Cancer (in press). Marques da Costa, R., Aniceto, C., Jesus, F.M. and Mendes, M. (1994). Lancet 344, 481–482. Morhenn, V.B. (1997). Immunol. Today 18, 433–436. Nickoloff, B.J., Fivenson, D.P., Kunkel, S.L., Strieter, R.M. and Turka, L.A. (1994). Clin. Immunol. Immunopath. 73, 63–68. Nissen, N.N. and DiPietro, L.A. (2000). In: Rubanyi, G.M. (ed.) Angiogenesis in Health and Disease, New York: Marcel Dekker, pp. 417–427. O’Kane, S. and Ferguson, M.W.J. (1997). Int. J. Biochem. Cell Biol. 29, 63–78. Ortega, S., Ittmann, M., Tsang, S.H., Eherlich, M. and Basilico, C. (1998). Proc. Natl Acad. Sci. USA 95, 5672–5677. Pierce, G.F., Mustoe, T.A., Lingelbach, J., Masakowski, V.R., Gramates, P. and Deuel, T.F. (1989). Proc. Natl Acad. Sci. USA 86, 2229–2233. Randolph, G.J., Inaba, K., Robbiani, D.F., Steinman, R.M. and Muller, W.A. (1999). Immunity 11, 753–761. Randolph, G.W., Beaulieu, S., Lebecque, S., Steinman, R.M. and Muller, W.A. (1998). Science 282, 480–483.

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Riches, D.W.H. (1996). In: Clark, R.A.F. (ed.) The Molecular and Cellular Biology of Wound Repair, New York: Plenum Press, pp. 95–141. Sallusto, F., Palermo, B., Lenig, D. et al. (1999). Eur. J. Immunol. 29, 1617–1625. Sato, K., Kawasaki, H., Nagayama, H. et al. (2000). J. Immunol. 164, 2285–2295. Sozzani, S., Allavena, P., Vecchi, A. and Mantovani, A. (1999). J. Leuk. Biol. 66, 1–9.

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Strobl, H., Bello-Fernandez, C., Riedl, E. et al. (1997). Blood 90, 1425–1434. Takakura, N., Wantanabe, T., Suenobu, S. et al. (2000). Cell 102, 199–209. Weber-Matthiesen, K. and Sterry, W. (1990). J. Invest. Dermatol. 95, 83–89.

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PLATE 35.13 Transmission of immunodeficiency virus infection at the mucosal-associated lymphoid tissues. HIV-1/SIV likely crosses the antigen-sampling LE. During the first days of exposure, virus cannot be readily detected and may remain cell-free or cell-associated within the LE. In subsequent days, however, rapid virus replication can be detected, especially within the CD4 T cells.

Plate 35.15 Potential immune implications of attenuated virus replication in the dendritic cell context. Immature DC-T cell mixtures support rapid replication of wild-type SIV, potentially resulting in the extensive immune destruction observed in vivo (particularly in the lymphoid compartments). In contrast, immature DCs cannot drive replication of the attenuated SIV delta nef even in the presence of CD4 T cells. Small amounts of virus produced in this setting could then be presented by the DCs to activate the immune system in the absence of consuming virus infection.

PLATE 38.1 Macrophages and dendritic cells coordinately regulate angiogenesis and antiangiogenesis. The fine balance regulating the microvasculature includes survival and inhibitory factors which are abundantly produced by macrophages and DCs. During normal homeostatic periods within a tissue, macrophages produce requisite factors facilitating small vessel integrity. During inflammation with progressive maturation and activation, DCs produce a variety of antiangiogenic factors including interferonα, IL-12 and IL-18. During the healing phase DCs are less abundant and are not activated by resident T cells or other cells, reverting to cells which promote neoangiogenesis and vessel survival. (Modified from Lotze, M.T. et al. (2001).

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Nam et ipsa scientia potestas est [Knowledge is power] Francis Bacon

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and Lord, 1995a). In atherosclerotic lesions, the numbers of vascular DCs increase (Bobryshev and Lord, 1995b, 1996a, 1998; Bobryshev et al., 1997a). Advanced atherosclerotic lesions may also be infiltrated by blood DCs through microvessels formed during neovascularization (Bobryshev and Lord, 1998; Bobryshev et al., 1999c).

Dendritic cells (DCs) accumulate in human primary atherosclerotic lesions of the aorta, coronary and carotid arteries (Bobryshev et al., 1996b,1996c; Bobryshev and Lord, 1998) as well as in the aortic wall affected by Takayasu’s disease (Inder et al., 2000) and in abdominal aortic aneurysms (Bobryshev et al., 1998a). DCs have also been detected in veins affected by varicosity and thrombophlebitis (Cherian et al., 1999a) as well as in aortocoronary saphenous vein bypass grafts (Cherian et al., 1999b). Normal veins are free of DCs (Cherian et al., 1999a), and this implies that DCs in diseased veins are recruited from the blood in response to vessel injury. In contrast, in large arteries, DCs are present in the normal nondiseased arterial wall (Bobryshev and Lord, 1995a, 1996a; Waltner-Romen et al., 1998). As these DCs are a basic element of the normal nondiseased arterial wall, they have been designated as vascular DCs (Bobryshev Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

ULTRASTRUCTURAL CRITERIA FOR IDENTIFICATION OF DCs IN THE ARTERIAL WALL Vascular DCs exhibit distinct morphologic characteristics similar to other well-studied DCs (Bani and Giannotti, 1989; Okato et al., 1989; King and Katz, 1990; Naito, 1993; Sprecher and Becker, 1993; Takahashi et al., 1996). Their low electron density cytoplasm contains hypertrophied cisterns of smooth endoplasmic reticulum (Figure 39.1A) which, together with vesicles

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FIGURE 39.1 Ultrastructural criteria for identifying DCs in the arterial wall (A–C). (A) Typical appearance of cisterns of the tubulovesicular system (large arrows); dense granules are shown by small arrows. (B) Atypical dense granules transforming into Birbeck granule-like structures (arrows). (C) Birbeck granule-like structures (large arrows) and atypical dense granules transforming into Birbeck granule-like structures (small arrows). In (A–C), caveoles are marked by stars. Reproduced with permission from Bobryshev, Y.V. and Lord R.S.A. 55-kD actinbundling protein (p55) is a specific marker for identifying vascular DCs. Journal of Histochemistry and Cytochemistry 47, 1481–1486, 1999.

and vacuoles, form the characteristic tubulovesicular system uniquely found in welldifferentiated DCs, in particular in interdigitating DCs (Anderson and Anderson, 1981; Takahashi et al., 1996). The cisterns, tubular and vesicular structures form continuous nets in the cytoplasm and this tubulovesicular system is functionally associated with the extracellular

space (Bobryshev et al., 1997b). The connection with the extracellular space through vesicles, vacuoles and cisterns can allow the entry of different substances into the cistern nets (Bobryshev et al., 1997b). Cisterns of the tubulovesicular system are usually most developed in the cell processes. The cell processes pass in different directions from the cell body

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and in ultrastructural studies, serial sectioning and consequent reconstructive analysis is required to demonstrate the association of cell processes with the cell body (Bobryshev and Lord, 1995a, 1996b; Bobryshev et al., 1997a). Atypical dense granules, which are a characteristic feature of Langerhans cells (Kobayashi et al., 1987), are also present in vascular DCs (Figure 39.1B,C). Atypical dense granules are different from lysosomes (Shamoto, 1970; Koboyashi et al., 1987; Hosokawa et al., 1993; Bobryshev et al., 1997a, 1997b). In primary lysosomes, the lysosomal content is evenly distributed throughout the organelle while in atypical dense granules, an electron-dense core is encircled by a band of material of low electron density (Shamoto, 1970; Koboyashi et al., 1987; Hosokawa et al., 1993; Bobryshev et al., 1997a, 1997b). Some of these atypical dense granules transform in vascular dendritic cells to structures resembling Birbeck granules present in Langerhans cells (Figure 39.1B,C) (Bobryshev et al., 1997a). Both Birbeck granules present in Langerhans cells and Birbeck granule-like structures present in vascular dendritic cells are rodand disc-shaped structures with a lamina through their center (Birbeck et al., 1961; Wolff, 1967; Sagebiel and Reed, 1968; Bobryshev et al., 1997a). However, in Birbeck granules present in Langerhans cells, the central lamina is intermittent (Birbeck et al., 1961; Wolff, 1967; Sagebiel and Reed, 1968) while in Birbeck granule-like structures present in vascular DCs, the central lamina is continuous (Figure 39.1C) (Bobryshev et al., 1997a). In Langerhans cells, Birbeck granules are cytoplasmic organelles involved in antigen processing and presentation (Hanau et al., 1987a, 1987b; Bucana et al., 1992). The presence of atypical dense granules and the tubulovesicular system allow unambiguous identification of vascular DCs in the arterial wall (Bobryshev and Lord 1995a, 1996a, 1996b, 1999; Bobryshev et al., 1996b, 1997a, 1997b; Bobryshev and Watanabe 1997a, 1997b). Similar to DCs in other tissues, DCs in the arterial wall contain only a few lysosomes, exhibit relatively low endocytotic activity, usually show a low level accumulation of lipids in their cytoplasm and

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usually do not transform into foam cells even when they are located among foam cells and lipid-rich debris in atherosclerotic lesions (Figure 39.2) (Bobryshev and Lord, 1995a, 1996a, 1996b; Bobryshev et al., 1997b; Bobryshev and Watanabe, 1997a, 1997b).

IMMUNOHISTOCHEMICAL IDENTIFICATION OF DCs IN THE ARTERIAL WALL In the arterial wall, DCs express 55 kDa actinbundling protein (p55) (Plate 39.3; Plate 39.4A,B) (Bobryshev and Lord, 1999), CD1a (Plate 39.4C,D) (Bobryshev and Lord, 1995a, 1996a; Bobryshev et al., 1996a, 1996b, 1996c; Bobryshev and Lord, 1998), S-100 protein (Bobryshev and Lord, 1995a, 1995b, 1995c, 1998; Bobryshev et al., 1996b; Bobryshev and Watanabe, 1997b) and Lag-antigen (Bobryshev et al., 1997a) which are markers for their immunohistochemical identification in the arterial wall. At present, CD1a and p55 are the most reliable markers for identifying DCs in the arterial wall as CD1a expression is restricted to thymocytes and to DCs while p55 expression is restricted to DCs and to B lymphocytes affected by Epstein–Barr virus (Mosialos et al., 1996). S-100 is expressed by a variety of cell types including glial cells, Schwann cells and ependyma, but none of these are present in the intima, making S-100 a convenient marker for DCs in this location (Bobryshev and Lord 1995a, 1995b, 1998; Bobryshev et al., 1996b). In Langerhans cells, Lag-antigen is located in Birbeck granules and in Birbeck granuleassociated structures (Kashihara et al., 1986). Lag-antigen specifically identifies dendritic cells in the arterial wall but its expression occurs only in a relatively small proportion of vascular DCs (Bobryshev et al., 1997a). In the arterial wall, DCs also express HLA-DR (Bobryshev et al., 1996b; Bobryshev and Lord, 1998), E-cadherin (Bobryshev et al., 1998b), intercellular cell adhesion molecule-1 (ICAM-1) (Bobryshev and Lord, 1997, 1998), vascular cell adhesion molecule-1 (VCAM-1) (Bobryshev et al., 1996c; Bobryshev and Lord, 1998) and CD40

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FIGURE 39.2 A DC (A) and its process contacting a macrophage foam cell (B) in an atherosclerotic lesion (an association between the DC body and its process was revealed by the reconstructive analysis of serial ultrathin sections). Note that the DC contains a large number of cisterns, some of which are hypertrophied (arrows). Note also that the DC contains a few membrane-bound vacuoles and its cytoplasm is free of ‘lipid droplets’. Reprinted from Cardiovascular Pathology, volume 6: Bobryshev Y.V. and Watanabe T. Subset of vascular dendritic cells transforming into foam cells in human atherosclerotic lesions, pp. 321–331, 1997, with permission from Elsevier Science.

(Bobryshev and Lord, 1999) but these molecules are widely expressed by different cell types in atherosclerotic lesions. These plasmalemmal markers would probably be useful for the isolation of vascular DCs from the arterial wall but do not allow specific immunohistochemical identification.

DCs IN THE NORMAL ARTERIAL WALL. ORIGIN OF VASCULAR DCs In the intima and adventitia of apparently normal, nondiseased arteries, vascular DCs are

present in their immature forms (the terms ‘immature’ and ‘mature’ vascular DCs are used to indicate the degree of structural differentiation of the cells and these terms are not meant to define the functional status of the cells) (Bobryshev and Lord, 1995a, 1996a; Bobryshev et al., 1997a; Waltner-Romen et al., 1998). In the intima, immature DCs are distributed along the subendothelial layer and are often in close contact with endothelial cells (Bobryshev and Lord, 1995a, 1996b). These immature vascular DCs can be unambiguously identified by the presence of one or two cisterns of the tubulovesicular apparatus appearing as semilunar crescentic structures concentric with the

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nuclear membrane (Bobryshev and Lord, 1995a, 1996b). The number of vascular DCs expressing immunohistochemical markers such as S-100, Lag-antigen and CD1a, is markedly lower than the total number of DCs identified in the intima using electron-microscopic analysis (Bobryshev and Lord, 1995a, 1996a, 1998). This suggests that the expression of DC immunohistochemical markers is associated with the activation and maturation of vascular DCs. The origin of vascular DCs is unknown. It is generally accepted that DCs derive from bone marrow and migrate into different tissues where they mature, depending on the microenvironmental influences, and develop regionspecific characteristics (Steinman, 1981, 1991; King and Katz, 1990). The hematopoietic development of DCs may allow for several precursor pathways, some closely linked to monocytes (Banchereau and Steinman, 1998; Ferrero et al., 1998; Palucka and Banchereau, 1999). The differentiation of monocytes into DCs has been demonstrated in vitro (Grassi et al., 1998; Randolph et al., 1998), which suggests a possibility that monocytes in the arteria intima may differentiate into both intimal macrophages and DCs. Alternatively, vascular DCs might locally differentiate from unique vascular DC precursors disseminated throughout the arterial intima and the adventitia in embryogenesis. In any case, the ultrastructural peculiarities of vascular DCs (Bobryshev et al., 1997a, 1997b) suggest that in the arterial intima, DC precursors differentiate into a distinct cell type within the DC family. In the normal intima, DCs have been proposed to be an element of vascular-associated lymphoid tissue (VALT), which is probably analogous to the mucosa-associated lymphoid tissue (MALT) of the respiratory and gastrointestinal tracts (Wick et al., 1997). VALT consists of small numbers of immunocompetent and antigenpresenting cells disseminated throughout the subendothelial layer of the arterial intima (Wick et al., 1997; Waltner-Romen et al., 1998). VALT probably screens ‘vascular tissue’ for potentially harmful antigens (Wick et al., 1997).

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CLUSTERING OF DCS IN ATHERO-PRONE AREAS OF THE AORTA The distribution of atherosclerotic lesions in the human arterial tree has been extensively surveyed and athero-prone (AP) as well as athero-resistant (AR) areas of the aorta have been identified (Bhagwat and Robertson, 1973; Sveden and Eide, 1980; Cornhill et al., 1990). In an immunohistochemical study (Lord and Bobryshev, 1999), the distribution of DCs in athero-prone and athero-resistant areas of the nondiseased aorta was compared and it was found that in the intima of athero-prone areas, there were more CD1a DCs than in atheroresistant areas. In the intima of athero-prone areas, CD1a DCs were found to form cell clusters which were mainly located in the subendothelial layer. Electron microscopy of athero-prone areas demonstrated that these cell clusters were formed by vascular DCs through their cellular processes which were filled with cisterns of a well-developed tubulovesicular system (Lord and Bobryshev, 1999). In contrast, no clusters of vascular DCs were observed in the intima of AR areas (Lord and Bobryshev, 1999). In other pathological conditions, the clustering of DCs is thought to be an indicator of autoimmune processes (King and Katz, 1991; Sprecher and Becker, 1993). According to the autoimmune hypothesis of atherosclerosis proposed by Wick et al. (1997), VALT activation by autoantigens is responsible for initiating immune responses in the arterial wall which lead to atherosclerotic alteration. Finding vascular DCs accumulating and clustering in atheroprone areas of the normal aorta adds to other evidence that immune mechanisms may be involved in lesion formation from the very early stages of atherogenesis (Lord and Bobryshev, 1999).

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DISTRIBUTION AND INTERCELLULAR CONTACTS OF DCs IN ATHEROSCLEROTIC LESIONS Immunohistochemical studies have shown that in atherosclerotic lesions, the numbers of DCs increase (Bobryshev and Lord, 1995b, 1996a, 1998; Bobryshev et al., 1997a). The functional significance of DCs in atherosclerosis is unknown, although it is most likely that these cells are involved in antigen processing and presentation as are DCs in other tissues (Bobryshev and Lord, 1995a, 1996a, 1996b, 1998). The migratory routes of vascular DCs might be similar to those known for other well-studied DCs such as Langerhans cells (Steinman, 1991; Banchereau and Steinman, 1998; Palucka and Banchereau, 1999). DCs from atherosclerotic lesions can migrate into lymph nodes where they can activate T cells (Bobryshev and Lord, 1998). The number of interdigitating DCs in para-aortic and jugulodigastric lymph nodes attached to atherosclerotic arterial wall segments have been found to markedly exceed those in the lymph nodes attached to nonatherosclerotic arterial segments (Bobryshev et al., unpublished). In addition to the migratory route into lymph nodes via the lymphatics as veiled cells, DCs from the atherosclerotic intima can migrate to the spleen via the blood. Only some DCs have been suggested to migrate to lymphoid organs while others contact T cells directly within the intima (Bobryshev and Lord, 1998). Atherosclerotic plaques contain inflammatory infiltrates consisting of macrophages and activated T cells (Hansson et al., 1989; Hansson, 1993; Wick et al., 1995). Antigenspecific T-cell activation depends on the interactions of T-cell receptors (TCR) with antigens presented by MHC molecules but how the antigen presentation occurs in atherogenesis is unclear (Hansson et al., 1989; Libby and Hansson, 1991; Hansson, 1993; Wick et al., 1995). The observations that DCs display ICAM-1 and VCAM-1 in atherosclerotic lesions (Bobryshev et al., 1996c; Bobryshev and Lord, 1997, 1998) implies that DCs might interact with T cells lead-

ing to the activation of T cells, since ICAM1/LFA-1 and VCAM-1/VLA-4 interactions are critical in T-cell activation (Ibelda and Buck, 1990; Springer, 1990). Double immunostaining has demonstrated a frequent association of DCs with T cells in the atherosclerotic intima (Plate 39.5) (Bobryshev and Lord, 1998). Serial sectioning of lymphocyte-rich areas of the atherosclerotic intima has shown that DCs, through their processes, form multiple contacts with T cells (Bobryshev and Watanabe, 1997a). In these contacts, DC processes exhibit hypertrophied cisterns of the tubulovesicular system (Figure 39.6), which suggests that the DCs are activated (Bobryshev and Watanabe, 1997a). The mapping of DCs in atherosclerotic plaques has demonstrated that DCs are frequently co-localized with T cells and that more than 90% of DCs colocalizing with T cells are located in neovascularization areas associated with inflammatory infiltrates (Figure 39.7) (Bobryshev and Lord, 1998). In atherosclerotic lesions, DCs express not only high levels of MHC class II but also display on their surfaces CD1a, which has been recognized as an antigen-presenting molecule in the same sense as the classical MHC class I and II molecules (Beckman and Brenner, 1995; Porcelli, 1995; Porcelli and Modlin, 1995). It has been suggested therefore, that an involvement of CD1-restricted T-cell responses in atherosclerosis cannot be excluded (Bobryshev and Lord, 1996b, 1998; Bobryshev and Watanabe, 1997a, 1997b; Melian et al., 1999). In the atherosclerotic arterial wall, two regions where DCs most frequently co-localize with T cells have been identified (Bobryshev and Lord, 1998). The first region is the area of neovascularization associated with inflammatory infiltrates within atherosclerotic lesions. The second region is the area around the inflamed vasa vasorum in the adventitia. In the DC processes contacting T cells, the tubulovesicular system is consistently hypertrophied. In these T-cell-rich areas, DCs, through their numerous processes, also form contacts with other cell types including B lymphocytes (plasma cells), small numbers of which are consistently present in diseased arterial wall (Bobryshev and Watanabe,

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FIGURE 39.6 Contacts of T cells (large asterisks) with DC processes (small asterisks). (A) and (C) are serial parallel sections. (B) is a detail of (A). (D) is a detail of (C). Note well-developed cisterns of the tubulovesicular system in DC processes contacting T cells. In (B) and (D), arrows indicate the zones of close contact between the T cell and a DC process. Reproduced, with permission, from Bobryshev, Y.V. and Watanabe T. Ultrastructural evidence for association of vascular dendritic cells with T-lymphocytes and with B-cells in human atherosclerosis. Journal of Submicroscopic Cytology and Pathology, 29, 209-221, 1997.

1997a). That DCs contact both T and B cells suggests that these DCs differ from other DCs which are associated exclusively with either T cells or B cells, but not both (Bobryshev and Watanabe, 1997a). While in the area of neovascularization and the areas around inflamed vasa vasorum, most DCs are associated with T cells, outside of these areas in atherosclerotic plaques, co-localizing

DCs and T cells have been detected only occasionally (Bobryshev and Lord, 1998). In contrast, in these areas, contacts between DCs and macrophages are common (Bobryshev and Lord, 1998). The DC/macrophage and DC/ macrophage foam cell co-localizations have been shown to be especially prominent around the necrotic core of atheroma. Possibly, this is where antigen-presenting cells acquire most of

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immune information. DCs lack lysosomal activity and DC/macrophage cooperation might compensate for this DC deficiency (Bobryshev and Lord, 1998).

HETEROGENEITY OF VASCULAR DCs

FIGURE 39.7 Schematic representation of the distribution of T cells and DCs in an atherosclerotic plaque. Neovascularization associated with vasa vasorum is represented by branched, twisted double lines.

FIGURE 39.8 Schema illustrating the structural features and a possible relation between the two phenotypes of vascular DCs.

the antigens they will display, as a result of arterial tissue breakdown. In DC processes contacting macrophages and macrophage foam cells, cisterns of the tubulovesicular system are typically hypertrophied (Figure 39.2). The presence of a large number of DC/macrophage contacts around the necrotic core suggests that in atherogenesis, interactions between these two cell types might be essential for processing

Immunohistochemical studies indicate that DCs in the arterial wall display different combinations of cell surface markers which reflect the different stages of cell maturation and, possibly, the differentiation into different phenotypes (Bobryshev et al., 1996a, 1996c; Bobryshev and Lord, 1998). Electron microscopic observations show that in the nondiseased arterial wall and in early atherosclerotic lesions, vascular DCs exhibit different levels of differentiation in their transition from immature to mature forms (Bobryshev and Lord, 1995a, 1995b, 1996a; Bobryshev et al., 1997b). Mature vascular DCs differ by the degree of development of their tubulovesicular system and by the number of atypical dense granules in their cytoplasm (Bobryshev and Lord, 1996b; Bobryshev et al., 1997a, 1997b). According to the ultrastructural characteristics, two phenotypes of vascular DCs have been recognized (Figure 39.8) (Bobryshev and Lord, 1996b). Type I vascular DCs contain only a few atypical dense granules but the tubulovesicular system is well developed. Through their long processes, type I vascular DCs have been shown to form multiple contacts with lymphocytes (Bobryshev and Watanabe, 1997a). In the cytoplasm of type II vascular DCs, cisterns of the tubulovesicular system are present but they are much less developed than in type I vascular DCs. In contrast to type I vascular DCs, type II vascular DCs are rich in atypical dense granules, some of which undergo transformation into Birbeck granule-like structures (Bobryshev et al., 1997a, 1997b). Large numbers of coated and uncoated vesicles, vacuoles as well as multivesicular structures are typical in the cytoplasm of type II vascular DCs (Bobryshev et al., 1997a, 1997b). The ability for antigen capture, antigen

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processing and antigen presentation in the arterial wall might differ between the members of the heterogeneous population of DCs, especially in relation to the two main phenotypes of vascular DCs. How the two phenotypes of vascular DCs relate is uncertain. The comparison of the ultrastructure of the two vascular DC phenotypes with that of other well-studied DCs suggests that type I vascular DCs are similar to interdigitating DCs while type II vascular DCs resemble Langerhans cells.

FOAM CELLS OF DC ORIGIN The vast majority of DCs in atherosclerotic lesions do not transform into foam cells. However, the defense mechanisms against extensive lipid accumulation may be broken in some vascular DCs, causing them to transform into foam cells (Bobryshev and Watanabe, 1997b). A subpopulation of CD1a/S-100 foam cells in human atherosclerotic lesions might, therefore, relate to DCs. These foam cells are distinct from other foam cells as they contain cisterns of the tubulovesicular system and Birbeck granule-like structures (Bobryshev and Watanabe, 1997b). Melian et al. (1999) have found that CD1apositive foam cells in atherosclerotic lesions coexpress CD1b, CD1c and CD1d. Using a co-cultivation of T cells with CD1 monocytederived foam cells, Melian et al. (1999) demonstrated T-cell activation in vitro which supports the hypothesis that CD1-restricted T-cell activation may occur in atherosclerotic lesions (Bobryshev and Lord, 1996b, 1998; Bobryshev and Watanabe, 1997a, 1997b; Melian et al, 1999). In atherosclerotic plaques, CD1 foam cells have been found to strongly express CD68 (Melian et al., 1999) while most nonfoam DCs are negative for CD68 (Bobryshev and Lord, 1996a, 1998; Bobryshev et al., 1996b, 1996c). The vast majority of foam cells in atherosclerotic lesions are negative for CD1a (Bobryshev et al, 1996a, 1996c; Bobryshev and Lord, 1998).

DCs IN ANIMAL MODELS OF ATHEROSCLEROSIS A few observations suggest that DCs might accumulate in aortic lesions in experimental animal models of atherosclerosis (Bobryshev et al., 1999a, 1999b, 1999d; Ozmen et al., 1998). Cells with a DC appearance have been identified in atherosclerotic lesions in apo-E-deficient mice (Bobryshev et al., 1999b). Although mouse atherosclerosis models are very useful for investigating both the contribution of macrophages in atherogenesis and the mechanisms of foam cell formation, mouse atherosclerosis seems not to completely mimic the immune-inflammatory aspects of human atherosclerosis, as the mouse atherosclerotic lesion lacks an accumulation of T lymphocytes (Dansky et al., 1997). In contrast to mouse atherosclerosis, the accumulation of large numbers of both macrophages and T cells in the aortic intima is a feature of experimental atherosclerosis in hypercholesterolemic rats (Haraoka et al., 1995) which may make this model useful for investigating the DC contribution in atherogenesis (Ozmen et al., 1998).

CONCLUDING REMARKS The study of the involvement of DCs in vascular pathology is in its infancy and so far it has been mostly descriptive. Morphological observations suggest that DCs are present in their immature forms in the normal nondiseased arteries and become activated during atherogenesis (Bobryshev and Lord, 1995a, 1995b; Lord and Bobryshev, 1999). Ultrastructural peculiarities of their tubulovesicular system and atypical dense granules (Bobryshev et al., 1997a, 1997b) suggest that vascular DCs represent a separate cell type within the DC family. Isolation of vascular DCs might, therefore, allow the identification of a unique antigen(s) on their surface and thus ultimately lead to new strategies in which vascular DCs will be targeted to deliver biologically active substances to atherosclerotic lesions.

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ACKNOWLEDGEMENTS I thank Prof. Mason W. Freeman for reading the manuscript and for his valuable comments. This work was supported by the NIH grant, 5PO1 DK 50305-10.

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Bobryshev, Y.V., Lord, R.S.A. and Parsson, H. (1998a). Cardiovasc. Surg. 6, 240–249. Bobryshev, Y.V., Lord, R.S.A., Watanabe, T. and Ikezawa, T. (1998b). Cardiovasc. Res. 40, 191–205. Bobryshev, Y.V., Babaev, V.R., Iwasa, S., Lord, R.S.A. and Watanabe, T. (1999a). Atherosclerosis 143, 451–454. Bobryshev, Y.V., Babaev, V.R., Lord, R.S.A. and Watanabe, T. (1999b). J. Submicrosc. Cytol. Pathol. 31, 527–531. Bobryshev, Y.V., Cherian, S.M., Inder, S.J. and Lord, R.S.A. (1999c). Cardiovasc. Res. 43, 1003–1017. Bobryshev, Y.V., Konovalov, H.V. and Lord, R.S.A. (1999d). J. Submicrosc. Cytol. Pathol. 31, 179–185. Bucana, C.D., Munn, C.G., Song, M.J., Dunner, K. Jr. and Kripke, M.L. (1992). J. Invest. Dermatol. 99, 365–373. Cherian, S.M., Bobryshev, Y.V., Inder, S.J., Lord R.S.A. and Ashwell, K.W.S. (1999a). Angiology 50, 393–402. Cherian, S.M., Bobryshev, Y.V., Inder, S.J. et al. (1999b). Cardiovasc. Surg. 7, 508–518. Cornhill, J.F., Herderick, E.E. and Stary, H.C. (1990). Monograph Atheroscler. 15, 13–19. Dansky, H.M., Charlton, S.A., Harper, M.M. and Smith, J.D. (1997). Proc. Natl Acad. Sci. USA 94, 4647–4652. Ferrero, E., Bondanza, A., Leone, B.E., Manici, S., Poggi, A. and Zocchi, M.A. (1998). J. Immunol. 160, 2675–2683. Grassi, F., Dezutter-Dambuyant, C., McIlroy, D. et al. (1998). J. Leuk. Biol. 64, 484–493. Hanau, D., Fabre, M., Schmitt, D.A. et al. (1987a). Proc. Natl Acad. Sci. USA 84, 2901–2905. Hanau, D., Fabre, M., Schmitt, D.A. et al. (1987b). J. Invest. Dermatol. 89, 172–177. Hansson, G.K. (1993). Br. Heart J. 69 (Suppl.), S38–S41. Hansson, G.K., Jonasson, L., Seifert, P.S. and Stemme, S. (1989). Arteriosclerosis 9, 567–578. Haraoka, S., Shimakama, T. and Watanabe, T. (1995). Virchows Arch. 426, 307–315. Hosokawa, S., Shinzato, M., Kaneko, C. and Shamato, M. (1993). Virchows Arch. [B]. 63, 159–166. Ibelda, S.M. and Buck, C.A. (1990). FASEB J. 4, 2868–2880. Inder, S.J., Bobryshev, Y.V., Cherian, S.M. et al. (2000). Cardiovasc. Surg. 8, 141–148. Kashihara, M., Ueda, M., Horiguchi, Y., Furukawa, F., Hanaoka, M. and Imamura, S. (1986). J. Invest. Dermatol. 87, 602–607. King, P.D. and Katz, D.R. (1990). Immunol. Today 11, 206–211. Kobayashi, M., Asano, H., Fujita, Y. and Hoshino, T. (1987). Cell Tissue Res. 248, 315–322. Libby, P. and Hansson, G.K. (1991). Lab. Invest. 64, 5–15. Lord, R.S.A. and Bobryshev, Y.V. (1999). Atherosclerosis 146, 197–198.

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Melian, A., Geng, Y.J., Sukhova, G.K., Libby, P. and Porcelli, S.A. (1999). Am. J. Pathol. 15, 775–786. Mosialos, G., Birkenbach, M., Ayehunie, S. et al. (1996). Am. J. Pathol. 148, 593–600. Naito, M. (1993). Arch. Histol. Cytol. 56, 331–351. Okato, S., Magary, S., Yamamoto, Y., Sakanaka, M. and Takahashi, H. (1989). Arch. Histol. Cytol. 52, 231–240. Ozmen, J., Lord, R.S.A., Bobryshev, Y.V., Ashwell, K.W.S. and Munro, V.F. (1998). Biomed. Res. 19, 279–287. Palucka, K. and Banchereau, J. (1999). J. Clin. Immunol. 19, 12–25. Porcelli, S.A. (1995). Adv. Immunol. 59, 1–98. Porcelli, S.A. and Modlin, R.L. (1995). J. Immunol. 155, 3709–3710. Randolph, G.J., Beaulieu, S., Lebecque, S., Steinman, R.M. and Muller, W.A. (1998). Science 282, 480–483. Sagebiel, R.W. and Reed, T.H. (1968). J. Cell. Biol. 36, 595–602.

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Shamoto, M. (1970). Cancer 26, 1102–1108. Sprecher, E. and Becker, Y. (1993). In Vivo 7, 217–227. Springer, T.A. (1990). Nature 346, 425–434. Steinman, R.M. (1981). Transplantation 31, 151–155. Steinman, R.M. (1991). Annu. Rev. Immunol. 9, 271–296. Sveden, T. and Eide, T.J. (1980). Acta Pathol. Microbiol. Scand. (Sect. A). 88, 97–103. Takahashi, K., Naito, M. and Takeya, M. (1996). Pathol. Int. 46, 473–485. Waltner-Romen, M., Falkensammer, G., Rabl, W. and Wick, G. (1998). J. Histochem. Cytochem. 46, 1347–1350. Wick, G., Schett, G., Amberger, A., Kleindienst, R. and Xu, Q. (1995). Immunol. Today 16, 27–33. Wick, G., Romen, M., Amberger, A. et al. (1997). FASEB J. 11, 1199–1207. Wolff, K. (1967). J. Cell Biol. 35, 468–473.

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PLATE 39.3 Patterns of distribution of p55 cells in the arterial wall (A–D). (A) p55 cells (arrows) in nondiseased aortic intima. p55 cells were visualized using the peroxidase-antiperoxidase technique. E, endothelium; L, lumen of the aorta. (B) p55 cells (arrows) in a neovascularization area in an atherosclerotic plaque in a carotid artery; microvessels are marked by asterisks. (C) p55 cells in the deep portion of aortic atherosclerotic intima on the border with the media. (D) is a detail of (C). In (D), a cluster of p55 cells is shown by open arrows while separately located p55 cells are shown by solid arrows. (B–D) Double immunostaining resulting in brown staining of p55 cells (peroxidase–antiperoxidase technique) and rose staining of α-smooth muscle actin cells (alkaline phosphatase–antialkaline phosphatase technique). Counterstaining with Mayer’s haematoxylin. Reproduced, with permission, from Bobryshev, Y.V. and Lord R.S.A. 55-kD actin-bundling protein (p55) is a specific marker for identifying vascular dendritic cells. Journal of Histochemistry and Cytochemistry, 47, 1481–1486, 1999.

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PLATE 39.4 A group of p55 (A)/CD1a cells (C) in an area of neovascularization in an aortic atherosclerotic plaque seen in consecutive sections. L, lumen of the aorta. (B) is a detail of (A). (D) is a detail of (C). In (B, D), p55/CD1a cells are shown by arrows while a microvessel is marked by stars. Peroxidase–antiperoxidase technique. Counterstaining with Mayer’s haematoxylin. Reproduced with permission from Bobryshev Y.V. and Lord R.S.A. ‘55-kD actin-bundling protein (p55) is a specific marker for identifying vascular dendritic cells’. Journal of Histochemistry and Cytochemistry, 47, 1481–1486, 1999.

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PLATE 39.5 S-100 DCs (brown) located between CD4 cells (rose) in large (A–C) and small (D) inflammatory infiltrates (A, B) in aortic atherosclerotic plaques. (B) is a detail of (A). Double immunostaining; combination of peroxidase–antiperoxidase and alkaline phosphatase–antialkaline phosphatase techniques. Counterstaining with Mayer’s haematoxylin. Reprinted from Cardiovascular Research, volume 37: Bobryshev Y.V. and Lord R.S.A. ‘Mapping of vascular dendritic cells in atherosclerotic arteries suggests their involvement in local immuneinflammatory reactions’, pp. 799–810, 1998, with permission from Elsevier Science.

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40 Clinical trials of dendritic cells for cancer Jeffrey Weber 1 and Lawrence Fong 2 1

USC/Norris Comprehensive Cancer Center, Los Angeles, California, USA; 2 Stanford University School of Medicine, Palo Alto, California, USA

There are many ways of going forward, but only one way of standing still. Franklin D. Roosevelt

INTRODUCTION

An extensive series of preclinical murine experiments demonstrated that DCs function as potent antigen-presenting cells (APCs) and could mediate regression of advanced tumors after antigen-pulsing and adoptive transfer. This preclinical experience, which has been summarized in detail in earlier chapters, has been rapidly translated into the performance of a variety of early-stage clinical trials to provide proof of concept for the notion that large numbers of purified human DCs could be reliably produced for adoptive transfer, that administration of DCs was safe and well tolerated, and that preliminary indication of clinical efficacy could be observed. In the first part of this chapter we will describe early trials of DCs purified directly from fresh blood without exposure to cytokines by the use of affinity purification procedures. The next section will describe a variety of clinical trials from phase I up to phase II in which peripheral blood-derived monocytes and CD34 stem cells were exposed ex vivo to cytokines including GM-CSF, IL-4 and TNFα to produce

The discovery that human dendritic cells (DCs) can be propagated from peripheral blood mononuclear cells, CD34 progenitors and cord blood by prolonged growth ex vivo in serum-less media using GM-CSF, TNFα and IL-4 or affinity purified from fresh blood without cytokines and briefly incubated ex vivo, has facilitated the performance of a number of phase I and II clinical trials in melanoma, prostate and other cancers using DCs pulsed with peptide, protein, RNA and tumor lysate antigens. Preliminary data from trials using other approaches such as DC–tumor cell fusions have recently become available. As our understanding of how the proliferation, antigen-processing and presentation, migration and trafficking of DCs improves, clinical trials will progress from empirically based early phase trials to more rationally based larger scale phase II and III trials. We present here a detailed review of the current clinical trials using DCs for cancer. Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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DCs. Finally, we will list the important unanswered questions in this rapidly growing clinical field and describe how those questions are being addressed in ongoing clinical trials. Since the selection of and availability of antigen for DC pulsing is likely to be critically important for a successful clinical outcome, many of the early trials have included patients with metastatic melanoma and lymphoma. The variety of cloned T-cell antigens present on melanoma cells is the broadest for any histology; patients with metastatic disease often have multiple subcutaneous lesions that are easily and safely resected for the production of tumor lysates; there are no established curative treatments for metastatic melanoma, so that patients with good prognosis M1 and M2 (subcutaneous, lymph nodal and lung) disease can legitimately be accrued to DC trials with low-volume disease without having received prior chemotherapy. Non-Hodgkin’s lymphoma is one of the few tumors which expresses idiotypes, wellestablished tumor antigens that are restricted to malignant cells and have been shown to elicit immune and clinical responses. The follicular and well-differentiated lymphomas also have prolonged survival and an indolent clinical course which facilitates identification of antigens and prolonged immunization regimens in the absence of rapidly progressive disease.

GRADIENT ENRICHMENT OF DCs FROM PERIPHERAL BLOOD Human DCs can be enriched as circulating precursors from the blood by density-based purification techniques. After a period of in vitro culture and maturation, DC precursors become larger and less dense. Gradient solutions lacking potentially immunogenic proteins such as BSA have been employed including Percoll (Pharmacia Biotech, Parsippany, NJ, USA), Nycodenz (Nycomed Pharma, Oslo, Norway), and metrizamide. These different fluids are osmotically active to varying degrees, and have additional stimulatory properties as well.

Commercial devices are being developed that may simplify this process. One such method involves loading patient blood into a closed device that contains solution with a density of 1.0650 g/mL and osmolarity of 320 mOsm. This device is then spun in a centrifuge and the buoyant cells are then harvested (F. Valone, personal communication). The use of density-based isolation is limited by the low frequency of DC precursors in blood, representing around 1% of peripheral blood mononuclear cells (PBMCs), so that a leukapheresis must be performed to generate sufficient numbers of DCs (on average 5  106) for therapeutic vaccination in humans. The use of cytokines such as Flt-3 ligand to expand circulating DCs in vivo allows significantly higher numbers of DCs to be obtained from a leukapheresis. Below we describe clinical trials employing the technique of direct DC isolation without cytokine incubation.

B cell non-Hodgkin’s lymphoma In the first reported DC trial, patients with malignant B-cell lymphoma who had failed conventional chemotherapy were treated with autologous DCs pulsed ex vivo with tumorspecific idiotype antigen (Hsu et al., 1993, 1996). The idiotype immunoglobulin expressed by monoclonal B-cell lymphomas is potentially immunogenic by virtue of its unique idiotypic determinants, which are formed by the combination of the variable regions of immunoglobulin heavy and light chains (Kwak et al., 1992). To prepare idiotype proteins for this clinical study, patients underwent tumor biopsies and the immunoglobulin (idiotype) produced by each tumor was ‘rescued’ by somatic cell fusion techniques and purified from hybridoma supernatants. Peripheral blood leukocyte preparations containing DC precursors were obtained from patients by leukapheresis, and monocytedepleted mononuclear cells were incubated for 36 hours in the presence of either tumor-specific idiotype or a control protein, keyhole limpet hemocyanin (KLH) (Timmerman and Levy, 2000). This incubation technique not only enabled the DC precursors to take up and

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process the exogenous proteins, but also induced their maturation into potent APCs. The idiotype (Id) and KLH-pulsed DCs were then enriched from contaminating lymphocytes on the basis of their differences in buoyant density. The DCs were washed to remove free protein and administered intravenously. This procedure was repeated three times at monthly intervals with a booster immunization given 4 to 6 months later. Two weeks after each DC infusion the patients received subcutaneous injections of Id, which in the absence of DCs or a chemical adjuvant would not be expected to induce an immune response. Ten patients were treated with relapsed, measurable lymphoma. These patients, as well as six not described in the published report, tolerated their infusions well and none experienced clinically significant toxicity during the study. Eight of 10 patients developed Id-specific cellular proliferative responses. T cells from one patient were expanded for several weeks in vitro in the presence of Id protein and shown to lyse autologous tumor hybridoma cells but not an Id-matched, unrelated hybridoma. In contrast to earlier Id vaccination studies, which utilized chemical adjuvants, a humoral response to Id was not detected in our patients, consistent with the interpretation that the Id-pulsed DCs induced a TH1 rather than a TH2 response. Most importantly, two of the patients experienced complete tumor regression, lasting 44 months and 40 months. A third patient developed a ‘molecular’ response with Bcl-2 translocationpositive bone marrow converting to negative following the vaccination and has been clinically tumor free for 62 months ( J. Timmerman, personal communication). A fourth patient experienced a partial response lasting 12 months. Another cohort of 22 patients has been vaccinated while in first remission. The initial 11 patients in this series received DCs loaded with native Id, while the remaining 11 patients were immunized with DCs loaded with Id conjugated to KLH. This latter approach was found to accentuate the generation of Id-specific antibodies in preclinical studies. Ten of 22

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developed Id cellular responses following vaccination. However, five of 11 patients immunized with Id–KLH-pulsed DCs developed antibodies specific for autologous tumor. The clinical assessment of these patients is currently underway with 82% of the patients progression free at a median follow-up of 25 months (J. Timmerman, personal communication). It is worth noting that B-cell lymphoma is unusual among malignancies in that the neoplastic cells express both MHC class I and II antigens and would, therefore, be susceptible to recognition and attack by either CD4 or CD8 T cells, and also by tumor-specific antibodies.

Multiple myeloma At Stanford 26 myeloma patients were vaccinated with Id-pulsed DCs following high-dose chemotherapy and autologous peripheral blood progenitor cell transplantation (PBPCT) (R. Levy, personal communication, Reichardt, et al., 1999). All patients had minimal residual disease after chemotherapy. Three to seven months later, patients received a series of monthly immunizations consisting of two intravenous infusions of Id-pulsed autologous DCs. Each patient’s idiotype was purified from autologous serum, and DCs were enriched either with the conventional gradient approach as described above or a modified closed gradient approach. Twenty-four of 26 patients developed an Idspecific proliferative T-cell response. Four patients developed an Id-specific, cytolytic immune response. Three of these were detected in patients who achieved a complete response with high-dose chemotherapy and had no detectable myeloma protein. PBPCT produced a complete response in three patients and a partial response in 21 patients. Eight of the 21 partial responders developed a decrease in the M spike during or following vaccination. Another trial was performed at the Mayo Clinic in 15 patients with residual low-tumor burden following high-dose chemotherapy and autologous stem cell transplant (Valone et al., 2000). The median M spike in this trial was 0.845 g/dL with a range of trace to 1.2 g/dL.

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Patients were immunized with DCs pulsed with autologous serum as a source of Id for three monthly vaccinations followed by a final vaccination 16–24 weeks later. DCs were enriched from peripheral blood with the closed gradient device. Among 13 evaluable patients there were three pathologic CRs and three PRs. Only one patient had disease progression with a median follow-up of 28 weeks. A multicenter trial utilizing the same approach was also performed in 46 patients with refractory disease, who were not stem cell transplant candidates or had progressive disease following transplant (Valone et al., 2000). Median M component was 2.7 g/dL (range 0.9–5.4), and median number of prior chemotherapy regimens was three (range 1–8). T-cell immune responses to the patients’ M component were observed in one-third of patients and immune responses appear to correlate with clinical response. Among 38 evaluable high tumor burden patients there were no tumor regressions. However, 17 of 38 evaluable patients had minor responses or stable disease lasting 24 weeks and the overall median time to progression was 37.3 weeks. A patient with advanced-stage refractory multiple myeloma who received DCs derived from adherent peripheral blood mononuclear cells and pulsed with Id protein has been described (Wen et al., 1998). Id-specific immune responses were generated, characterized by MHC-dependent T-cell proliferative responses with cytokine release and the production of antiId antibodies. A T-cell line generated after vaccination was also able to lyse autologous Idpulsed targets and recognize fresh autologous myeloma cells. The immune response was associated with a transient minor fall in the serum Id level, and surprisingly, was not ablated by subsequent high-dose myeloablative chemotherapy.

Prostate cancer A clinical trial for prostate cancer has been performed at Stanford employing DCs purified from peripheral blood by conventional density gradient centrifugation and pulsed with prostatic

acid phosphatase (PAP) as the tumor antigen. Based upon preclinical data demonstrating that immunization with xenogeneic homologues can break tolerance to self antigens, this trial utilizes a xenogeneic PAP, derived from rodents, rather than human PAP as the immunogen (Fong et al., 1997, 1999). Twenty-one patients with recurrent or metastatic prostate cancer and measurable serum levels of PAP were immunized with two monthly injections of autologous PAP-pulsed DC. All patients developed T-cell proliferation to mouse PAP following vaccination. Ten of 21 patients also developed crossreactive T-cell proliferation against endogenous human PAP. Seven patients had stable disease following vaccination without any other treatment or intervention. In this trial the route of DC administration was investigated; it was found that DCs can prime immunity when administered intravenously, intradermally, or intralymphatically. In another trial performed at the Mayo Clinic 13 hormone refractory prostate cancer patients were immunized with two monthly infusions of DCs pulsed with a recombinant human PAP fused to human GM-CSF (Burch et al., 2000). DCs were obtained from PBMCs with the closed density centrifugation system. Patients also received three monthly subcutaneous injections of the fusion protein alone. T-cell proliferation was reported in a subset of patients to human PAP and/or human GM-CSF following DC vaccination, while induction of anti-PAP (five of 11 patients) and anti-GM-CSF (10 of 11 patients) antibodies was seen only following the subcutaneous ‘booster’ injections with soluble protein. Three of 11 evaluable patients had disease stabilization following vaccination for 206 to 274 days.

Colorectal cancer A clinical trial in colorectal cancer patients is ongoing at Stanford targeting an agonist HLA*A0201 restricted epitope of carcinoembryonic antigen (CEA) 605–613. The ability of growth factors to increase numbers of circulating DCs is also being investigated in that trial. Flt-3 ligand, which has been shown to

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preferentially expand DC precursors in mice in vivo, was administered to patients prior to leukapheresis. DCs were then enriched with a modified gradient approach and pulsed with peptide and KLH before being reinfused. In the initial six patients, Flt-3 ligand increased the mean number of DCs/leukapheresis from 3.1  106 to 230  106. Six of six patients immunized with Flt-3 ligand-expanded DCs developed KLHspecific T-cell proliferative responses, while five of the five evaluated to date also have evidence of expansion of CTLs not only specific for the agonist epitope, but also for tumor cells expressing the native form of CEA. One of the initial six patients had regression of pulmonary metastasis following vaccination without any other treatment (Fong et al., manuscript submitted).

MONOCYTE AND CD34 PROGENITOR-DERIVED DCs DCs with potent ability to stimulate naïve and memory T cells in vitro can be grown from peripheral blood monocytes with cytokines GMCSF and IL-4 (Romani et al.,1996, Thurner, 1999a). Bhardwaj and her colleagues performed a series of carefully constructed clinical trials in normal volunteers in order to establish that human monocyte-derived DCs could induce immune T-helper and cytolytic T-cell responses, assess their kinetics and lay the groundwork for subsequent trials in cancer patients. Nine healthy subjects received a single dose of subcutaneously injected unpulsed control DCs, followed a month later by another subcutaneous injection of 1.6 up to 4 million DCs pulsed with KLH as a priming antigen, tetanus toxoid as a recall antigen, and the HLA-A2.1-restricted influenza matrix peptide (Dhodapkar et al., 1999). Four additional subjects received the KLH and flu peptides alone without DCs. Nine of nine patients had increased proliferation of peripheral blood cells to KLH, and five of six had increased proliferation to tetanus toxoid. Six of six HLA A2.1 subjects had at least a 2-fold increase in ELISPOT gamma interferon assay to flu matrix peptide after DC vaccination; none of

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the subjects who had peptides alone or control DCs alone, or who were not HLA A2.1 positive showed evidence of increased CTL reactivity to the flu peptide. The ELISPOT assay, which measured single cell CD8 reactivity to antigen by the release of gamma interferon, indicated that up to 70 CTL precursors were detected per 200 000 fresh PBMCs 20 days after DC vaccination, suggesting that up to 0.17% of the circulating CD8 cells (assuming approximately 20% CD8 in total PBMC) were flu specific. When fresh PBMC were restimulated ex vivo with peptide, a tetramer assay 7 days after peptide stimulation showed that significant increases were observed in antigen-specific gamma interferon-secreting CD8 cells in DC-vaccinated subjects. The same group of investigators reinjected three of the HLA-A2.1 subjects 6 to 12 months later with monocyte-derived DC pulsed only with the flu matrix peptide (Dhodapkar et al., 2000). All three subjects had a rapid increase in CD8 T cells that were flu specific, and one of the three developed CTLs that were specifically lytic for peptide-expressing targets. Of note is that after rechallenge with peptide-pulsed DCs, CTL responses were more rapid, and of a greater magnitude. CTL reactivity to flu matrix rose by day 7 and reached higher levels than after the initial DC vaccination, after which ELISPOT assays peaked by day 30. T cells with greater avidity for the target were represented among the revaccinated responders, since lower concentrations of stimulating peptide were needed to detect a T-cell response. These data suggested that helper cell stimulation was not needed for a rechallenge, and that significant quantitative and qualitative changes in T-cell populations occurred after repeated antigen-pulsed DC vaccinations.

Prostate cancer Early clinical trials of monocyte-derived DCs included patients with prostate cancer who were vaccinated with DCs pulsed with two HLA-A2.1restricted peptides from prostate-specific membrane protein (PSMP) (Murphy et al., 1996; Salgaller 1998; Tjoa et al., 1998; Murphy et al.,

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1999a, 1999b). In a phase II trial of 37 patients, peptide-pulsed DCs were infused intravenously at 6-week intervals in patients who had locally recurrent or metastatic prostate cancer. Fourteen patients had a CR or PR by nontraditional criteria (AUA criteria) which included decline in PSA or tumor decrease on Prostascint scan among the response criteria (Lodge et al., 2000). However, the investigators could not conclude that there was an association between augmented CD8 responses to PSMP peptides and clinical benefit. In fact, there was an association between preexisting nonspecific DTH reaction of at least three of the antigens on the Pasteur–Merieux multitest indicator and clinical response on that trial, and between clinical response and anti-CD3-stimulated nonspecific PBMC proliferation. Of the 14 CRs  PRs on that trial, nine had three or more positive DTH tests, whereas only seven of 22 nonresponders were positive; the numbers were felt to be too small for the differences to be of any statistical significance.

Melanoma The first published clinical trial in melanoma patients of DCs grown ex vivo with cytokines utilized cells delivered intralymphnodally by injection. A series of 16 melanoma patients who received peptide-pulsed (12 patients) or tumor lysate-pulsed (four patients) DCs were described (Nestle et al., 1998). Four patients had prior chemotherapy, and four prior biologic therapy. One million DCs were injected into an inguinal lymph node twice over 2 weeks and then monthly thereafter. DCs were pulsed with KLH to act as a CD4-helper antigen and to act as an immunologic tracer molecule. Toxicity was modest, with grade I/II fever, with pain and swelling at the injection site as the predominant side-effects. No autoimmune toxicity was detected. All 16 patients had a DTH response to KLH, and 11/16 had a DTH response to peptidepulsed DCs, but none to the peptides themselves. The DTH sites had a predominant CD8 infiltrate by immunohistochemistry, and in one

case the infiltrating cells were extracted, expanded and shown to be reactive to the gp100 and MART-1/MelanA tumor antigens. No data on systemic induction of CTL immunity were presented. A metastasis was resected from a responding patient, showing a predominant CD45RO memory T-cell infiltration. Five patients had clinical responses, with the two CRs still in remission 15 months later. The CR patients both had peptide-specific DTH reactions, which the authors felt were predictive. The same group has gone on to treat at least 40 patients with lysate- and peptide-pulsed DCs injected intralymphnodally, with a response rate of 25% (Nestle et al., personal communication). In a subsequently published study, 11 patients with metastatic melanoma received DCs grown from peripheral blood mononuclear cells, pulsed with a single HLA-A1-restricted MAGE-3 peptide and tetanus or tuberculin and induced to mature with monocyte-conditioned media (MCM) (Thurner et al., 1999b). The DCs in this trial were injected by combined subcutaneous, intradermal and intravenous routes in all patients. Four patients’ DCs were incubated in TNFα as well as the above cytokines and MCM. Tetanus toxoid or tuberculin were added to DCs pulsing as CD4 recall antigens and as an immune control, as KLH was used in the Nestle trial described above. The 11 patients had been diagnosed as having stage IV metastatic melanoma a mean of 10 months prior to entering the trial, and all had received previous chemo-, immuno- or chemo–immunotherapy. The treatment was not toxic, with only grade I/II side-effects, consisting mostly of local reactions at the cutaneous injection sites and fevers, although the authors felt that the fevers were due to tetanus-pulsed DCs, since in at least one patient with fever to 40°C the fevers disappeared after removal of the tetanus from the DC pulsing. By standard oncologic criteria, none of the 11 patients had an objective clinical response, although in six patients regression of some lesions was observed. Five of nine patients tested had greater than a 3-fold increase to tetanus toxoid by proliferation

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reactivity after vaccination, and five of five patients who received tetanus had significant increases in ELISPOT reactivity to tetanus antigens with no increase to tuberculin as a negative control. In the ELISPOT recall assays, decreases were seen in all five patients after intravenous injections as opposed to after cutaneous injection. The authors used a semiquantitative CTL recall assay to detect induction of CTLs directed against MAGE-3; in eight of 11 patients, there appeared to be a significant increase after cutaneous injections, but a decrease after intravenous injections as seen in the recall ELISPOT responses. Surprisingly, no evidence of MAGE-3specific DTH was observed pre- or postvaccination using either peptides or peptidepulsed mature DCs, and no evidence of MAGE3-specific CTL ELISPOT reactivity was observed in fresh blood. In two patients with multiple cutaneous lesions, MAGE-3 expression, which had been present prior to treatment, was absent, suggesting to the authors that immunoselection may have occurred. Weber et al. have described a clinical trial of 16 patients with metastatic melanoma who received increasing doses of multiple peptidepulsed monocyte-derived DCs intravenously twice 2 weeks apart (Weber et al., 1999). Similar to the Nestle trial, and unlike the Thurner trial, the monocyte-derived DCs were not treated with MCM or other agents to induce maturity. The gp100 209–217 (210M)-and tyrosinase 368–376 (370D)-substituted peptides were used. As in the above trials, grade I/II toxicity was observed, and no evidence of clinical autoimmune reactions were seen. One patient with small-volume lung disease had a complete remission lasting 8 months, two previously untreated patients have had minimal tumor shrinkage followed by stable visceral disease lasting 18 months each, and two patients had transient minimal responses followed by progression. Two responders had evidence of peptide-specific DTH reactivity, and four of the five responding or stable patients had evidence of increased gp100-or tyrosinase-specific CTL reactivity shown by release of gamma interferon by multiple restimulated PBMC postvaccine

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compared to prevaccine. In patients who were retreated with a second course of two intravenousinfusionsofpeptide-pulsedDCs,peptidespecific reactivity decreased. None of the patients with progression in the absence of any tumor shrinkage had evidence of peptidespecific CTL reactivity. The same group has gone on to treat patients with additional peptides pulsed on to DCs infused intravenously, followed by the addition of low-dose IL-2 subcutaneously for 5 days after each DC infusion as an ‘immune reconstitution’ based on preclinical work in the mouse suggesting that low-dose subcutaneously administered IL-2 added to tumor lysate-pulsed DC injections synergistically augmented the treatment effect. Of the first four patients treated with that regimen, one patient has shown evidence of an autoimmune hemolytic anemia resulting in a hemoglobin of 6, profoundly elevated LDH and an unmeasurable haptoglobin which was sustained for 4 weeks after the last dose of DCs. When reports of a flare of rheumatoid arthritislike symptoms in a patient treated with peptidepulsed DCs are added to the aforementioned episode, great caution must be advised in screening patients for autoimmune diseases prior to and monitoring their development after DC vaccinations. Linette pulsed a modified gp100 peptide 280–288 (9V) with a valine substitution at the 288 position on to monocyte-derived DCs and treated metastatic melanoma patients with peptidepulsed DCs given intravenously every 3 weeks for six injections (Linette 2000). One of the first six patients has had a partial response, and two of six patients had augmented CTLs reactive with the 280–288 epitope. Faid and colleagues treated 22 patients with MAGE-1 and MAGE-3-expressing tumors with monocyte-derived DCs pulsed with peptides from those two antigens (Faid et al., 2000). Ten patients received peptide-pulsed DCs, and 16 received DCs with KLH added to the peptides. There was no difference in the level and duration of MAGE1/3-specific CTLs generated in the peripheral blood using a sensitive ELISPOT assay, throwing doubt on the concept that KLH as ‘nonspecific help’ would augment

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the CTL response directed against a class I epitope peptide on DCs. Most of the trials described in this chapter have employed DCs produced by leukopheresis followed by direct density gradient purification and maturation without cytokines or Ficoll purification and adherence to plastic followed by incubation in IL-4/GM-CSF to produce ‘monocyte-derived’ DCs. Faires et al., have used elutriation or counter current centrifugation to purify monocytes, followed by incubation with GM-CSF, and maturation with calcium ionophore, IL-2 and IL-12 (Faires et al., 2000). DCs were pulsed with multiple melanoma peptides and administered intradermally, intravenously or intralymphnodally to patients with metastatic melanoma. Interestingly, none of the patients who received DCs either intravenously or intradermally had a clinical response, but two of the five patients that received intralymphnodal DCs had a clinical response, and four of the five had augmented DTH to peptide-loaded DCs. DCs derived from CD34 progenitors, grown in the presence of GM-CSF and TNF then pulsed with multiple melanoma peptides and injected subcutaneously have been used to treat patients with metastatic melanoma (J. Fay, personal communication). Twenty-four patients have been treated so far, and immune response as well as clinical response data are available for the first 11 patients. ELISPOT showed evidence of boosted immunity in seven patients, and clinical regression was seen in some patients which appeared to correlate with immune responses. In at least two patients, antimelanoma reactivity by ELISPOT reached the level of FLU-specific cells with treatment continuing on that trial which is now a randomized trial of adherent monocyte-derived compared with CD34 progenitor-derived DCs. A large number of preclinical experiments in rodents have shown that gene therapy of DCs is feasible and results in high levels of antigen expression after plasmid or viral vectormediated gene transfer. Haluska and colleagues have performed a phase I trial in melanoma patients of DCs transduced with adenoviral vec-

tors encoding the gp100 and MART-1 tumor antigens (29) (Haluska et al., 2000). Monocytederived DCs were infected with engineered adenoviruses, and injected subcutaneously every 3 weeks, six times. Three of the first five patients developed vitiligo, which has not been observed on prior DC trials in melanoma. Stift et al. (Stift et al., 2000) pulsed matured DCs with TNF or CD40 ligand plus tumor lysate ex vivo then treated 13 patients with various malignancies by intralymphnodal injection. This was followed by daily subcutaneous IL-2 for 10 days; no responses were seen in this pilot trial but DTH to lysate was observed in two patients; six of 13 patients had stable disease. DCs have been pulsed with alternative antigens such as heat-shock proteins and KLH, which contain immunogenic peptides. Recently, six patients were treated both intradermally and intravenously in a pilot study (Riccobon et al., 2000); no responses have been observed. Morse et al. treated 14 patients with CEA expressing malignancies with a strongly HLA-A2 binding peptide (CAP-1) pulsed on to immature monocyte-derived DCs given intravenously (Morse et al., 1999a). Up to 100 million DCs with a purity of 66% were achieved with minimal toxicity; only one minimal responder and one stable patient were observed in a group that had been extensively previously treated with chemotherapy. Little evidence of immune response was observed in that trial in peripheral blood. The same investigators treated six patients in a pilot trial of Flt-3 ligand-primed DCs which were then harvested by pheresis and pulsed with the CAP-1 CEA peptide (Zaremba et al., 1997), KLH and tetanus toxoid for adoptive transfer in patients with CEA-expressing malignancies (Morse et al., 1999b). Interestingly, after 24 hours on plastic, the resulting DCs which had been characterized as ‘immature’ now became mature with increased expression of the CD83 and CD80 markers. One of the first six patients has had an immune response to CEA by tetramer analysis, and 8.5% of the circulating cells were shown to be CD11/CD14− DCs. All patients had increased helper cell reactivity to KLH or tetanus, and interestingly a cytolytic T-

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cell response to KLH was detected in a patient for the first time.

Renal cell cancer A novel technique for antigen-pulsing of DCs consists of fusing autologous tumor cells with autologous or allogeneic DCs by the use of electrofusion with a brief pulse of electrical current. Kugler and colleagues treated 17 patients with renal cell cancer with fused tumor-allogeneic monocyte-derived DCs (Kugler et al., 2000). The DCs for each injection were derived from different normal donors. All patients had visceral metastases, and all but one had received no prior systemic therapy; six patients had bony disease and six had liver disease. Ten to 15% of the total mixtures fused as determined by twocolor flow cytometry prior to treatment. The resulting cell mixture was irradiated to 2000 R, and a total of 50 million each of tumor and allogeneic DCs were fused, irradiated, then injected subcutaneously twice, 6 weeks apart, just distal to a draining inguinal lymph node. Modest to minimal toxicity was observed, with local injection site pain, swelling and fever, as the most common manifestations. Four CRs and two PRs were seen. Eleven of 17 patients developed tumor cell-specific DTH reactivity, including five of six responders. MUC-1 was found to be expressed on all 17 patient tumors, and two of the four HLA A2.1 patients developed MUC 1.2-specific CD8 T cells after vaccination. Intracellular cytokine assays from those two patients showed that 0.3 and 0.8%, respectively, of their CD8 cells specifically secreted gamma interferon after a 6-hour exposure to an HLA-A 2.1-restricted MUC-1.2 peptide, but not to an irrelevant peptide. The use of tumor lysates to pulse DCs was originally described by Nestle et al., and has been strengthened by preclinical work in mice by the groups of Mule (Fields et al., 1998) and Bhardwaj (Sauter et al., 2000). The advantage of tumor lysates as antigen is that all possible antigens expressed at the surface would be available, and the antigens would be ‘customized’ to the patient, but practical considerations dictate that

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most patients with metastatic cancer might not have easily surgically accessible tumor that could be recovered and used under sterile conditions. Nonetheless, Mule and colleagues have initiated a phase I clinical trial of tumor lysatepulsed monocyte-derived DCs in all histologies with metastatic disease, and have observed responses in patients with sarcoma and melanoma. In one clinically responding patient with melanoma, increases in melanoma antigen-specific CTLs to 1:2000 in the peripheral blood by tetramer assay were observed (J. Mule et al., personal communication). Stift et al. have employed DCs isolated by affinity bead purification, then propagated them in fetal calf serum with cytokines and induced maturation with CD40 ligand (Stift et al., 2000). These DCs were pulsed on day 5 with tumor lysate from patients with advanced malignancies, then injected intralymphnodally under ultrasound guidance in an inguinal node weekly for 3 weeks. IL-2 was then injected subcutaneously at 1 million U/day for 12 days each month. Two patients with medullary thyroid cancer showed drops in the calcitonin tumor marker.

TREATMENT WITH CYTOKINES IN VIVO An attractive strategy to augment DC yield and activity is to pretreat patients with cytokines that induce proliferation of DC precursors and/or augment their maturation and activation. One alternative to simply preparing DCs from peripheral blood monocytes or purified CD34 progenitors ex vivo would be to prime the patient with a cytokine(s) that augmented the proliferation/activation of DC precursors, then perform a leukopheresis which would provide DC-enriched cells. The use of FIL-3 ligand in this role has already been discussed. Another such maneuver was suggested by Roth et al. who performed a phase I clinical trial of fixed doses of GM-CSF at 2.5 µg/kg daily for 14 days of every 28-day cycle, with increasing doses of IL-4 starting at 0.5 and increasing to 6 µg/kg/day (Roth et al., 2000). The cytokine

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combination increased the number of circulating cells that expressed DC markers like CD83 and HLA-DR, but also increased the number of CD14 cells disproportionately. The combination of GM-CSF and IL-4 also augmented mixed lymphocyte reactivity (MLR) and resulted in increased endocytosis by PBMCs. Twenty-one patients were treated on this phase I study, including 16 patients with renal cell cancer. When cells that had greatly augmented CD83 expression after GM-CSF/IL-4 treatment in vivo were sorted and analyzed for CD14 expression, it was absent, suggesting that two populations of cells were augmented; a predominant CD14-positive cohort, and a smaller cohort of CD14-negative classic CD83-positive DCs. One patient with prostate cancer had a sustained partial response on that trial. Cajal et al. pretreated 24 advanced cancer patients with GM-CSF for 4 days, followed by harvest of monocyte-derived DCs which were grown in IL4/GM-CSF then pulsed on day 6 with tumor lysate (Cajal et al., 2000). Therapy was given subcutaneously every 21 days for three cycles. One melanoma patient had a partial response; six patients were stable, and nine patients developed evidence of tumor-specific DTH reactivity after the first DC vaccination.

DC TRAFFICKING An important study which was performed to shed light on optimal routes of administration for DCs was described by Morse and Lyerly, who Indium-111 labeled monocyte-derived DCs from seven patients on a clinical trial of CEA RNA-pulsed DCs and localized the labeled DCs over time by radioscintigraphy (Morse et al., 1999c). Intravenously administered DCs initially localized to lung, then spleen, liver and bone marrow, but not noticeably to tumors or lymph nodes. Localization to draining lymph nodes only occurred after intradermal immunization, not after subcutaneous injection. By 48 hours, 70% of the input radioactivity remained at the subcutaneous injection site, and 65% of the intradermal material. A maximum of 0.41% of

the labeled injectate localized to a draining inguinal lymph node after intradermal injection in the thigh. To the authors, these data suggested that the intradermal injection route was superior. In this chapter we have attempted to describe the large and growing variety of clinical trials, and include recent data that are as yet only presented in abstract form from the recent ASCO meeting in New Orleans. Important questions that need to be addressed in the field of clinical DC treatment include: (1) The relative merits of preparing DCs from fresh blood by affinity purification without cytokines compared with prolonged ex vivo growth in the presence of cytokines. (2) Should DCs be prepared from peripheral blood mononuclear cells compared with CD34 derived progenitors? (3) Are DCs more optimally delivered by intravenous, cutaneous or lymph nodal routes? (4) What is the optimal antigen delivery route, including but not limited to peptides, whole proteins, RNA, DNA plasmid, heat-shock fusion proteins, cellular heterokaryon fusions, tumor lysates and gene-engineered virally encoded antigens? (5) What are the appropriate co-stimulatory and activation molecules to expose DCs in order to optimize DCs function in vivo (Morse et al., 1999d, 2000)? These are issues of unquestioned importance for the clinical future of DCs, and by the time the next edition of this text is published, many of the answers to these questions will be available from ongoing phase II studies.

REFERENCES Burch, P.A., Breen, J.A., Buckner, J.C. et al. (2000). Clin. Cancer Res. 6, 2175–2182. Cajal, R., Mayordomo, J.I., Yubero, A. et al. (2000). Proc. Amer. Soc. Clin. Oncol. 19, abstr. 1780 454a. Dhodapkar, M.V., Steinman, R.M., Sapp, M. et al. (1999). J. Clin. Invest. 104, 173–180. Dhodapkar, M.V., Krasovsky, J., Steinman, R.M., Bhardwaj, N. (2000). J. Clin. Invest. 105, R9–R14. Faid, L., Toungouz, M., Linin, M. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1779 454a. Faires, M.B., Bedrosian, I., Xu, S. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1778 453a.

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Fields, R.C., Shimizu, K. and Mule, J.J. (1998). Proc. Natl Acad. Sci. USA 95, 9482–9487. Fong, L., Ruegg, C.L., Brockstedt, D., Engleman, E.G. and Laus, R. (1997). J. Immunol. 159, 3113–3117. Fong, L., Benike, C., Engleman, E.G. et al. (1999). Proc. Am. Assoc. Cancer Res. 40, 85. Haluska, F.G., Linette, G.P., Jonasch, E. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1777 453a. Hsu, F.J., Kwak, L., Campbell, M. et al. (1993). Ann. NY Acad. Sci. 690, 385–387. Hsu, F.J., Benike, C., Fagnoni, F. et al. (1996). Nat. Med. 2, 52–58. Kugler, A., Stuhler, G., Waldin, P. et al. (2000). Nat. Med. 6, 332–336. Kwak, L.W., Campbell, M.J., Czerwinski, D.K. et al. (1992). N. Engl. J. Med. 327, 1209–1215. Linette, G., Jonasch, E., Longerich, S. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1833 467a. Liso, A., Stockerl-Goldstein, K., AuffermannGretzinger, S. et al. (2000). Biol. Blood Marrow Transplant. 6, 621–627. Lodge, P.A., Jones, L.A., Bader, R.A., Murphy, G.P. and Salgaller, M.L. (2000). Cancer Res. 60, 829–833. Morse, M.A., Coleman, R.E., Akabani, G., Niehaus, N., Coleman, D. and Lyerly, H.K. (1999a). Cancer Res. 59, 56–58. Morse, M.A., Lyerly, H.K., Gilboa, E., Thomas, E. and Nair, S.K. (1999b). Cancer Res. 58, 2965–2968. Morse, M.A., Vredenburgh, J.J. and Lyerly, H.K. (1999c). J. Hematother. Stem. Cell Res. 8, 577–584. Morse, M.A., Deng, Y., Coleman, D. et al. (1999d). Clin. Cancer Res. 5, 1331–1338. Morse, M.A., Mosca, P.J., Hobeika, A. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1840 469a. Murphy, G., Tjoa, B., Ragde, H., Kenny, G. and Boynton, A. (1996). Prostate 29, 371–380.

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Murphy, G.P., Tjoa, B.A., Simmons, S.J. et al. (1999a). Prostate 39, 54–59. Murphy, G.P., Tjoa, B.A., Simmons, S.J. et al. (1999b). Prostate 38, 73–78. Nestle, F.O., Alijagic, S., Gilliet, M. et al. (1998). Nat. Med. 4, 328–332. Reichardt, V.L., Okada, C.Y. et al. (1999). Blood 93, 2411–2419. Riccobon, A., Ridolfi, R., De Paola, F. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1842. Romani, N., Reider, D., Heuer, M. et al. (1996). J. Immunol. Meth. 196, 137–151. Roth, M.D., Gitlitz, B.J., Kiertscher, S.M. et al. (2000). Cancer Res. 60, 1934–1941. Salgaller, M.L., Lodge, P.A., McLean, J.G. et al. (1998). Prostate 35, 144–151. Sauker, B., Albert, M.L., Francisco, L., Larsson, M., Somersan, S. and Bhardwaj, N. (2000). J. Exp. Med. 191, 423–434. Stift, A., Dubsky, P., Friedl, J.P. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1820 464a. Thurner, B., Roder, C., Dieckmann, D. et al. (1999a). J. Immunol. Meth. 223, 1–15. Thurner, B., Haendle, I., Roder, C. et al. (1999b). J. Exp. Med. 190, 1669–1678. Timmerman, J.M. and Levy, R. (2000). J. Immunol. 164, 4797–4803. Tjoa, B.A., Simmons, S.J., Bowes, V.A. et al. (1998). Prostate 36, 39–44. Valone, F., Lacy, M., Mackenzie, M. et al. (2000). Proc. Am. Soc. Clin. Oncol. 19, abstr. 1776 453a. Weber, J.S., Jeffery, G., Marty, V. et al. (1999). Proc. Am. Soc. Clin. Oncol. 18, abstr. 1664 432a. Wen,Y.J., Ling, M., Bailey-Wood, R. and Lim S.H. (1998). Clin. Cancer Res. 4, 957–962. Zaremba, S., Barzaga, E. et al. (1997).Cancer Res. 57, 4570–4577.

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41 Dendritic cell therapies for HIV-1 infection Cara C. Wilson University of Colorado Health Sciences Center, Denver, Colorado, USA

To understand and manipulate immune responsiveness, and many clinical areas involving the immune system, one should not restrict the analysis to antigens and lymphocytes. One must also consider DCs, the captivating controllers of immunity. Ralph Steinman Cell

INTRODUCTION

to the development of adjuvant therapeutic approaches designed to generate or enhance HIV-specific cell-mediated immunity through modulation of T-cell reactivity. Therapeutic approaches using HIV-1 antigen-pulsed DCs are attractive for several reasons. These strategies provide both functional APCs, thereby repleting DCs lost during infection, and HIV-1 antigens complexed with APCs in a form that is potent at stimulating both naïve and memory T cells. DCs can now be cultured in large numbers from either monocytes (Romani et al., 1994; Kiertscher and Roth, 1996) or bone marrow precursors in vitro (Caux et al., 1992), making DC-based therapies more feasible.It stands to reason that therapeutic immunization of HIV-infected individuals with autologous-cultured DCs, appropriately loaded with HIV antigens, might lead to the generation of a more effective cellular immune response to HIV-1 in the setting of effective antiretroviral therapy.Clinicalstudiesdesignedtoaddressthehypothesis that DCs derived from HIV-infected patientsonHAARThavethe potential to stimulate

HIV-1 infection is characterized by progressive immune dysfunction and CD4 T-lymphocyte depletion resulting in the acquired immunodeficiency syndrome (AIDS). Dendritic cells (DCs), the most potent antigen-presenting cells (APCs) known for priming MHC class I- and IIrestricted T-cell responses to antigen, have been shown to be both depleted in number and dysfunctional in the setting of untreated HIV-1 infection (reviewed in Knight and Patterson, 1997). Treatment of HIV-infected patients with highly active antiretroviral therapy (HAART) regimens has led to marked reductions in HIV-1 viral load and improvements in peripheral CD4 T-cell counts (Autran et al., 1997; Li et al., 1998). However, there is evidence that HAART alone may not result in complete return of normal lymphocyte number and function, and in particular, HIV-specific immune function (Autran et al., 1997; Pontesilli et al., 1999; Rinaldo et al., 1999). Recognition of these limitations has led Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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strong HIV-specific cellular immune responses in vivo are currently underway.

OVERVIEW OF HIV-1 INFECTION HIV-1 infection is marked by a progressive depletion of peripheral CD4 T cells, T-cell dysfunction, defects in both number and function of (APCs) such as DCs and monocytes, thymic dysfunction, lymphoid destruction, and pancellular defects attributed to stem cell dysfunction. In the majority of HIV-infected persons, the ultimate result of this immune dysfunction is development of the opportunistic infections and malignancies associated with AIDS. Traditional approaches for treating HIV-1 infection have focused on the use of antiretroviral agents, alone and in combination. Antiretroviral monotherapy with nucleoside analogues such as AZT has resulted in some clinical benefit in delaying the onset of HIV-related disease, although the clinical benefit as well as the ability to raise CD4 count and lower viral load has been temporary. Therapy with combinations of antiretroviral nucleoside analogs has had a more pronounced effect on suppressing viral load and raising peripheral CD4 counts in infected persons. However, despite the well-documented antiviral effects of nucleoside analogs and other agents used in combination, their ability to reliably and completely reverse the immune deterioration associated with HIV-1 infection has not been shown. Therefore, combining therapeutic modalities designed to restore immune function with those aimed at suppressing viral replication may prove to be the most effective method of treating HIV-1 infection.

CELLULAR IMMUNITY TO HIV-1 Role of cytotoxic T lymphocytes (CTL) The progressive immune dysfunction associated with untreated HIV-1 infection occurs, in

many cases, despite evidence of early humoral and cellular HIV-specific immune responses. Although the correlates of protective immunity to HIV-1 infection have not been precisely defined, there is increasing evidence that certain types of immune responses might more effectively control HIV-1 replication than others. Cytotoxic T lymphocytes (CTLs) have been shown to be important in controlling viral replication in a number of chronic viral infections, and there is growing evidence that HIV-specific CTLs are important in the suppression of HIV-1 replication during chronic infection (Yang et al., 1997). The role of CTLs in controlling HIV-1 replication is best illustrated in studies of immune control in several clinical settings: in acutely HIV-1-infected individuals, in chronically infected individuals who fail to progress to AIDS, and in HIV-1-exposed but uninfected individuals. Studies of acute HIV-1 infection in humans revealed an early expansion of CD8 T cells which contained HIV-specific CTLs (Pantaleo et al., 1994), and longitudinal data showed that expansion of a restricted HIV-specific T-cell repertoire was associated with viral persistence and correlated with more rapid disease progression. Circulating virus-specific CTLs could be readily detected following acute infection with HIV-1, and their rise was temporally associated with a decline in viremia (Borrow et al., 1994; Koup et al., 1994). There is also some evidence that CTLs are involved in controlling HIV replication and delaying disease progression in chronically infected individuals (Klein et al., 1995b). Although the typical disease course during HIV-1 infection is one of gradually diminishing peripheral CD4 T-lymphocyte counts, with the development of AIDS occurring within a median time of 10 years from initial infection, approximately 5–10% of infected persons maintain stable, normal peripheral CD4 T-cell counts and are clinically asymptomatic for >10 years after infection (Levy, 1993; Klein and Miedema, 1995a).This heterogeneous cohort of individuals, termed ‘long-term nonprogressors’, has been intensively investigated to elucidate the factors

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responsible for this attenuated disease course. The presence of low viral loads and strong HIV-specific CTL activity is characteristic of this cohort (Cao et al., 1995; Ferbas et al., 1995; Pantaleo et al., 1995; Rinaldo et al., 1995; Harrer et al., 1996). In addition to the presence of high levels of memory CTLs, nonprogressors tended to have a strong, broadly reactive CTL response, targeting multiple epitopes in conserved regions of several HIV-1 antigens (Harrer et al., 1996). Further evidence of the important role of CTLs in controlling or preventing infection with HIV-1 can be found in the evaluation of HIVexposed but uninfected individuals, who exhibit HIV-specific T-cell responses in the absence of evidence of specific humoral immunity. Studies of individuals who have been exposed to HIV-1 but who remain uninfected (by serology or HIV1 DNA PCR) have been carried out to identify the natural mechanisms of resistance to or clearance of HIV-1. Both HIV-specific T-helper responses (based on cytokine production or proliferation) and CTLs have been identified in these individuals, often in the absence of serum antibodies against HIV-1 gene products (reviewed in Shearer and Clerici, 1996). The failure to detect evidence of HIV-1 infection, despite repeated exposures to HIV-1 in many cases, has led to the belief that a coordinated cell-mediated immune response to HIV-1 consisting of antigen-specific T helper and cytotoxic T lymphocytes might protect against HIV-1 infection. Recent studies that utilize more sensitive techniques for identifying circulating HIV-specific CTLs have confirmed the role of cellular responses in suppression of HIV-1 replication. Using MHC class I–peptide tetrameric complexes to directly quantitate circulating HIV-specific CTLs in vitro, Ogg et al. observed a strong inverse correlation between HIV-1 plasma RNA and the frequency of HIV-specific CTLs in chronically infected individuals (Ogg et al., 1998). Direct support for the role of CD8 HIV-1 specific CTLs in controlling HIV-1 replication in vivo has also been derived from studies in the SIV rhesus macaque model in which elimination of CD8

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cells is accompanied by a brisk rise in SIV plasma RNA (Jin et al., 1999; Schmitz et al., 1999).

Role of helper T lymphocytes (HTL) Despite the documented ability of HIV-specific CTLs to kill infected target cells presenting the appropriate epitope in vitro (Yang et al., 1997), the presence of HIV-specific CTLs in vivo is not sufficient to protect infected individuals from disease progression. CD4 helper T lymphocytes (HTLs) also appear to play a critical role in the establishment of effective immunity to HIV-1 and other chronic viral infections (Kalams and Walker, 1998). Defects in the ability of CD4 T cells from most HIV-infected individuals to respond by proliferation or cytokine production to HIV-1 antigens, as well as other ‘recall’ antigens, have been documented for many years (Shearer and Clerici, 1991). This HTL dysfunction is likely to be important in the immunopathogenesis of HIV-1 infection, in that dysfunctional T helper cells would be unable to appropriately assist in the expansion, differentiation, and maintenance of HIV-specific CTL. Some have even hypothesized that the failure to mount an HIV-specific CD4 T-cell response, and thereby provide help to CTLs, may underlie the inability of most infected patients to control HIV-1 replication. Rosenberg et al. determined that the presence of strong HIV-1 p24-specific T-helper responses inversely correlated with HIV-1 viral load in chronically infected individuals (Rosenberg et al., 1997). Furthermore, CD4 T-helper proliferative responses to HIV-1 p24 protein were more frequently found in the longterm nonprogressor cohort of infected individuals than in infected individuals with more standard disease progression, suggesting a protective effect of these cells in delaying disease progression (Rosenberg et al., 1997). Using a novel method to quantitate HIV-specific CD4 T cells in the peripheral blood based on cytokine secretion, Pitcher et al. also found a higher frequency of gag-specific CD4 HTL in HIVinfected patients with nonprogressive disease (Pitcher et al., 1999). These studies suggest that HIV-specific HTL responses may play an

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important role in preventing infection in seronegative persons exposed to HIV-1 or in protecting against disease progression in HIV-infected individuals. Antigen-specific CD4 T helper lymphocytes play a critical role in orchestrating the immune response based on patterns of cytokine production. Multiple factors may be involved in determining whether a more or less effective cell-mediated immune response to HIV-1 is mounted. These include genetic host factors (including HLA type), pathogenicity of the infecting virus, and the route and inoculum of initial infection. During active viral replication, certain viral proteins may inhibit the generation or maintenance of HIV-specific T cells through the induction of apoptosis (i.e. nef, gp120) (Oyaizu and Pahwa, 1995) or by altering the cytokine environment. The HIV-1 Nef protein, for example, has been shown to downregulate MHC class I expression, upregulate production of IL-10, and downregulate production of IL-2 and IFNγ, all factors that could conceivably adversely impact the generation or maintenance of the cell-mediated immune response to HIV-1 (Brigino et al., 1997; Collins et al., 1998). Indeed, overall cytokine dysregulation, which is characterized by a defect in TH1-type cytokines, such as IL-2 and IFNγ, and a dominance of TH2-type cytokines, such as IL-10, has been observed during progressive HIV-1 infection (Clerici and Shearer, 1993b). This TH2-dominated cytokine milieu may make it more difficult to mount or maintain effective cell-mediated immune responses to HIV-1 as disease progresses (reviewed in Clerici and Shearer, 1994). Once an immune response against HIV-1 has been generated, HIV-1 may subsequently evade immune regulation via multiple ‘stealth’ mechanisms. Factors such as the diversity of the initial immune response and the ability of HIV-1specific T-cell clones to crossrecognize variant peptides are likely to be important (Wilson et al., 1997a). If the virus succeeds in escaping immune control, the loss of CD4 T cells (and decreased specific T-cell help), progressive APC dysfunction (resulting from monocyte and DC infection or loss), and a cumulatively suppres-

sive cytokine milieu (TH2  TH1), may all contribute to the difficulty in generating T cells that react against evolving viral proteins. In this setting, therapeutic approaches designed to identify and correct these defects by enhancing or broadening T-cell immunity, and by generating responses against conserved epitopes, are likely to have the greatest impact.

The fate of HIV-specific HTLs and CTLs during the course of HIV-1 disease Since CD4 T cells (in particular activated CD4 T cells) are one of the main targets of HIV-1, it has been postulated that HIV-specific T helper cells that are activated appropriately during acute HIV-1 infection are subsequently infected and deleted from the T-cell repertoire. Evidence to support this theory of ‘clonal deletion’ can be found in the work of Rosenberg et al. (Rosenberg et al., 1997), who reported that persons acutely infected with HIV-1 and immediately treated with HAART subsequently developed strong and persistent HIV-1 p24-specific proliferative responses. Since such strong proliferative responses are rarely seen in chronically HIV-1infected individuals, it is postulated that these strong MHC class II-restricted responses can only be generated or maintained in the absence of replicating virus. The fate of clonally expanded, HIV-specific CD4 T cells during the natural course of HIV infection remains unclear, because other studies have suggested that T-cell dysfunction, not clonal deletion, may be responsible for the lack of HIV-1 antigen-specific proliferation in chronically-infected individuals. Using a sensitive technique that does not rely on T-cell proliferation, Pitcher et al. found that chronically infected patients did have detectable frequencies of HIV-specific CD4 T cells in their peripheral blood, but that the frequencies were lower than those seen in nonprogressors (Pitcher, 1999). Unfortunately, antigen-specific proliferative responses in that study were not directly compared with frequencies of cytokinesecreting cells. Some investigators have noted expansions in the CD4 T cell repertoire in HIV1-infected patients (Roglic et al., 1997), and in

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one report found that these expansions were not associated with HIV-specific T-cell proliferation (Lederman et al., 1998). However, a gradual resolution of these CD4 T cell expansions following the institution of highly active antiretroviral therapy suggests the possibility of an HIV-specific component (Gorochov et al., 1998; Lederman et al., 1998). Other investigators have reported that the addition of exogenous IL-12, or the blocking of IL-10, could restore HIV-specific proliferation in vitro in many infected patients tested (Clerici et al., 1993a). These results suggest that HIV-specific CD4 T cells may be present in HIV-infected individuals, but are either anergic or lacking adequate signals or cytokines necessary to proliferate in vitro. HIV-specific CTLs have been identified in HIV-infected individuals with progressive HIV-1 infection, although in many cases, the frequency of HIV-specific CTLs declines in end-stage disease (Carmichael et al., 1993; Geretti et al., 1996). The presence of CTLs in infected subjects with poorly controlled HIV-1 infection has raised the question as to whether the HIV-specific CTLs identified in vitro are actually functional in vivo. Recent studies support the notion that, in the absence of adequate HTL function, HIV-specific CTLs may be present but rendered anergic or otherwise dysfunctional (Spiegel et al., 2000). These studies underscore the important role that HTLs play in the immunologic control of HIV-1 infection. It is likely that therapeutic strategies that reconstitute a strong HIV-specific immune response in both the T helper and cytotoxic T-cell effector arms in such individuals would lead to a more effective and longer lasting suppression of HIV-1.

Immunologic recovery after HAART HAART, which usually includes potent protease inhibitors in combination with nucleoside analogue reverse transcriptase (RT) inhibitors, has been shown to effectively lower viral load and increase peripheral CD4 T-cell counts in a majority of HIV-infected patients. Inititation of HAART leads to the rapid reduction of viral replication in lymph nodes and peripheral blood,

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and there is mounting evidence that viral suppression resulting from HAART leads to significant immune reconstitution. Immunologic recovery in HIV-infected patients treated with HAART is characterized by an early rapid rise in peripheral CD4 cells (that is largely comprised of memory T cells) and a decrease in lymphocyte activation markers on both CD4 and CD8 T lymphocytes, followed by a more gradual increase in the naïve T-cell populations (Autran et al., 1997; Pakker et al., 1997, 1998; Li et al., 1998; Connick et al., 2000). Maximally suppressive antiretroviral therapy also results in improvements in the ability to mount delayedtype hypersensitivity responses and lymphoproliferative responses to some common recall antigens (Autran et al., 1997; Pontesilli et al., 1999; Connick et al., 2000). Recent studies have also shown that thymic function can be restored in some HIV-infected individuals treated with HAART (Douek et al., 1998).

Limitations of HAART Despite the initial success of HAART in controlling HIV-1 replication in many patients and restoring some level of general immune function, there is evidence that such therapeutic combinations have limitations. Although T-cell numbers increase in most HIV-infected patients treated with suppressive antiretroviral agents, the extent of reconstitution is limited in many cases, and peripheral CD4 T-cell numbers do not always return to normal. Even in cases of significant viral suppression and increases in CD4 counts, it is unclear whether complete immunologic recovery can occur in patients with more advanced disease. Autran et al. investigated the extent to which HAART could reverse the immunologic abnormalities associated with advanced HIV-1 infection (Autran et al., 1997). They reported a significant rise in peripheral CD4 T cells (memory, then later naïve) and a decrease in lymphocyte activation markers on both CD4 and CD8 T cells. The numbers of peripheral CD4 T cells and the CD4:CD8 T cell ratio, however, did not normalize over the period of study.

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Although significant increases in proliferative responses to recall antigens have been reported in HIV-1–infected patients following suppressive therapy, HIV-specific proliferative responses are not generally restored following the institution of HAART, and may actually decline. As discussed, the majority of chronically infected individuals have never developed or have lost HIV-1-specific T-helper cell responses over time, yet early intervention with HAART in persons with acute HIV-1 infection has been associated with enhanced HIV-1-specific T-helper cell function (Rosenberg et al., 1997). HIV-1-infected individuals treated with potent antiviral therapy prior to seroconversion developed HIV-1specific T-helper cell responses to HIV-1 Gag that were analogous to those seen in persons who spontaneously control viremia in the absence of antiviral therapy (Rosenberg et al., 1997). In contrast, HAART did not readily lead to augmentation of these responses in persons who were started on therapy later in infection. HIV-specific lymphocyte proliferative responses were not detected in the setting of HAART during chronic infection using conventional assays (Lederman et al., 1998; Plana et al., 1998; Pontesilli et al., 1999; Connick et al., 2000), and a decrease over time on HAART in the frequency of HIV-specific CD4 T cells has been reported using sensitive cytokine release assays (Pitcher et al., 1999). In the case of HIV-1-specific CTL responses, which many individuals possess prior to therapy, the frequency of CTLs tend to decline following the initiation of HAART in both the acute and chronic setting (Ogg et al., 1998, 1999; Gray et al., 1999; Rinaldo et al., 2000). HIV-specific CTL responses have been shown to increase if viral suppression is incomplete or HAART is stopped (Dalod et al., 1998; Ortiz et al., 1999). The basis for decreases in HIV-specific cellular immune responses is presumably related to lowered antigen concentration, although HIV-1 antigens have been shown to persist in lymphoid tissue despite HAART (Wong et al., 1997). These studies suggest that therapeutic immunization to boost HIV-1-specific cellular immune responses might be beneficial in per-

sons who have complete viral suppression on prolonged HAART. Whether preserving HIV-1specific cell-mediated immune responses in early infection or inducing them in more advanced infection will allow the host defense to enhance the durability of antiretroviral therapy or permit its withdrawal remains to be seen. Thus, despite the initial success of HAART in controlling HIV replication in many patients, there is increasing evidence that the limitations of antiretroviral therapy alone may necessitate adjunct therapies. In addition to failure of successful therapeutic regimens to completely restore immunity, there are multiple factors that prevent antiretroviral agents from successfully controlling viral replication. In a number of patients who have failed to respond or have only transiently responded to HAART with significant decreases in viral load, evidence of viral resistance to one or more of the drugs in the combination has been described (Young et al., 1998; Zhang et al., 1998; Lorenzi et al., 1999). Failure has also been attributed to lack of compliance due to drug toxicities as well as to pharmacologic factors such as inadequate drug levels or tissue penetration. Generating a broader, more potent HIV-specific cellular immune response may be critical in maintaining a low viral load in the event of drug resistance or noncompliance.

ROLE OF DCs IN HIV-1 IMMUNOPATHOGENESIS Dendritic cells are felt to play two important roles in the setting of HIV infection: that of capturing and transporting HIV-1 to susceptible CD4 T lymphocytes in the lymph nodes, and that of initiating an HIV-specific cellular immune response. There are several lines of evidence suggesting that DCs may be involved in the transmission and immunopathogenesis of HIV-1 (reviewed in Knight and Patterson, 1997). It has been hypothesized that during HIV-1 transmission, immature DCs at mucosal sites are first targeted by HIV-1. As DCs migrate to draining lymph nodes they undergo a matura-

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tion process, allowing them to become potent stimulators of resting T cells in the lymph nodes. Although controversy exists as to the extent to which DCs from blood and tissues are actually infected with HIV-1 in vivo, it has become clear that even with latent or low-level replication of HIV-1 in DCs, virus can be effectively transmitted during T-cell priming (Tsunetsugu-Yokota et al., 1995). The mechanism of this transmission of HIV-1 from DC to T cell is an area of intense investigation. A number of elegant studies have shown that HIV-1 can both enter and infect DCs as well as to bind to their surface via specific receptors (Ayehunie et al., 1997; Blauvelt et al., 1997; Geijtenbeek et al., 2000), but the result of either pathway is that infectious HIV-1 is transported into draining lymph nodes. Subsequent interactions between DCs and T lymphocytes lead to the spread of HIV-1 and replication within the lymphocyte population, resulting in a cytopathic effect on CD4 T cells (Cameron et al., 1992). Pope et al. found that DCs isolated from skin and subsequently infected with HIV-1 could efficiently transmit HIV-1 to and promote extensive viral replication in memory CD4 T cells through DC–T cell binding and syncytia formation (Pope et al., 1994). The ‘kiss of death’, as this interaction between DCs and T cells has been described, could be postulated to explain the conspicuous absence of HIV-specific T helper cells observed even early during HIV-1 infection. The interactions between DCs and HIV-1, and the role of DCs in HIV-1 pathogenesis, are described in more detail in a separate chapter. In addition to the impact that DCs may have on HIV-1 disease as a result of their ability to transmit HIV-1 to T cells, it has been suggested that DCs themselves are adversely effected by HIV-1, through direct or indirect means. Several investigators have reported decreased numbers of dermal or blood DCs in HIV-1-infected persons with advanced disease (Belsito et al., 1984; Eales et al., 1988; Macatonia et al., 1990; Grassi et al., 1999). Additionally, there is evidence that DCs freshly isolated from HIV-infected persons or infected with HIV-1 in vitro are dysfunctional in their ability to stimulate autologous and allo-

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geneic T-cell proliferation or to induce a primary immune response (Blauvelt et al., 1995, 1996; Roberts et al., 1994). Additionally, DCs from HIVinfected individuals may express lower levels of MHC molecules (Eales et al., 1988). Whether DC death and dysfunction are a result of direct infection with HIV-1, or result from the indirect effects of unchecked HIV-1 replication, is not entirely clear. However, it is likely that the loss of critical DC function may contribute to the global immunodeficiency associated with chronic, progressive HIV-1 infection.

Function of DCs cultured from HIV-1infected patients In order for autologous DC-based therapies to be successfully implemented in the setting of HIV-1 infection, it will be necessary to obtain adequate numbers of DCs from HIV-infected donors that are highly functional and free of replicating HIV-1 infection. Functional studies of blood DCs, which comprise less than 1% of PBMCs (Freudenthal and Steinman, 1990), have been limited by the lack of sufficient cell numbers. The ability now exists to culture large numbers of DCs from blood or bone marrow precursors using simple culture methods and recombinant cytokines (Romani et al., 1994). Cultured DCs may be derived from differentiation of CD34 DC precursors (obtained from mobilized peripheral blood or bone marrow) or from ‘conversion’ of blood monocytes (Romani et al., 1994; Kiertscher and Roth, 1996) (Caux et al., 1992). DCs may be readily obtained from monocytes or DC precursors in the peripheral blood using simple techniques, such as adherence or CD14 selection, and a short culture period in cytokines, such as GM-CSF and IL-4 (Romani et al., 1994; Kiertscher and Roth, 1996). In several studies, cultured peripheral bloodderived DCs from HIV-infected donors were found to be highly functional (Fan et al., 1997; Chougnet et al., 1999; Sapp et al., 1999). Fan et al. found that DCs cultured from blood precursors of HIV-infected donors had similar morphology and phenotype to those derived from HIV-1 seronegative donors, although in some cases

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lower yields of DCs were obtained from HIV donors with low peripheral CD4 T-cell counts or high viral loads (Fan et al., 1997). Chougnet et al. evaluated monocyte-derived DCs from both HIV-1-infected and uninfected donors. DCs obtained from these two donor groups displayed similar expression of costimulatory molecules and produced similar amounts of betachemokines, IL-12, and IL-10 following CD40 ligation (Chougnet et al., 1999). One benefit of utilizing DCs cultured from peripheral blood monocytes as a therapeutic modality for HIV-1 infection is the ease of obtaining the cells. However, a possible concern is the fact that peripheral blood monocytes are susceptible to HIV-1 infection and may even harbor HIV-1 in a latent form (Bagasra and Pomerantz, 1993; Peng et al., 1995). Sapp et al. evaluated cultured DCs derived from T celldepleted blood precursors of asymptomatic HIV-infected donors with varying HIV-1 viral loads and peripheral CD4 T cell counts (Sapp et al., 1999). In addition to finding that the DCs from HIV donors were highly functional, no evidence of HIV-1 DNA was found in these DCs by a semiquantitative PCR analysis.

Immunotherapy with DCs A number of investigators have shown that CD34 cells derived from bone marrow, mobilized peripheral blood, or cord blood develop into a heterogeneous population of DCs upon culture in GM-CSF and TNFα, with the addition of stem cell factor (SCF) or Flt-3 ligand (FL) markedly increasing the yield of DCs (Caux et al., 1992, 1996; Szabolcs et al., 1995; Rosenzwajg et al., 1996). One attractive feature of utilizing CD34 stem cell-derived DCs (CD34–DC) relative to monocyte-derived DCs (M-DC) for immunotherapeutic purposes is the greater proliferative potential, and therefore relatively higher DC yield, of progenitor cells. A study of DCs grown from the CD34 precursor cells of HIV-infected individuals has recently been completed (Wilson et al., 2000). In this study, peripheral blood CD34 cells obtained from both HIV and HIV donors enrolled in

ACTG 285, a study of G-CSF mobilization of stem cells (Schooley et al., 2000), were cultured in GMCSF, TNFα, and either SCF or FL. The yield of DCs per CD34 cell was low in some HIV donors, in particular those with higher viral loads or with the presence of HIV-1 DNA in purified CD34 preparations. On average, DCs from both HIV and HIV donors expressed similar levels of MHC and costimulatory molecules, were potent stimulators of allogeneic T cells, and matured similarly in response to CD40 ligation or LPS exposure. The majority of mobilized CD34 cells from the HIV donors were free of HIV-1, based on the absence of HIV-1 DNA by PCR (Campbell et al., 2000), and no HIV-1 p24 antigen was detected in cultures of resting or activated DCs grown in SCF, TNFα, and GM-CSF. Furthermore, using a sensitive nested PCR technique, HIV-1 DNA was not detected in the CD34-derived DCs grown under these conditions from any of the 16 HIV-infected donors, even those few whose purified CD34 cells contained HIV-1. Low levels of HIV-1 p24 antigen were detected in a single culture of activated DCs grown in FL, TNFα, and GM-CSF. These results suggest that DCs derived from HIV donor CD34 progenitors are highly functional and largely free of replicating HIV-1. CD34 cells from HIV-infected individuals represent an important source of DC precursors for use in immunotherapeutic strategies. As the T-cell stimulatory properties of DCs are likely to be a critical factor in determining the success of DC-based therapies, several studies have compared the functional capacities of DCs derived from either CD34 progenitors or monocytes (Mortarini, 1997; Triozzi and Aldrich, 1997; Meierhoff, 1998; Ferlazzo et al., 1999, 2000). In these comparative studies, some subtle differences have been noted in certain functional aspects of CD34-derived DCs versus monocyte-derived DCs. Triozzi and Aldrich compared CD34-DC cultured in GM-CSF/TNFα to M-DC cultured in GM-CSF/IL-4 with regard to surface phenotype and using a number of functional assays (Triozzi and Aldrich, 1997). In that study, DCs derived from both types of precursors presented peptide antigen similarly, however, M-DC presented soluble protein and stimulated

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allogeneic cells in a MLR better than did CD34DC. Differences in culture conditions or maturation stage, as opposed to intrinsic differences in the DCs themselves, might in part account for these observed functional disparities. Another study compared mature, CD1a M-DC and CD34-DC established from the same individuals and grown in the same cytokine combination for phenotypic and functional characteristics (Ferlazzo et al., 1999). Following a culture period in GM-CSF, TNFα, SCF, FL and IL-4, CD1a DCs from either precursor type expressed similar levels of the maturation marker, CD83, and were similarly potent in stimulating allogeneic T cells to proliferate. However, CD34-DC pulsed with a cognate epitope peptide were better able to active and expand antigen-specific CD8 T cells than were M-DCs. Similar functional comparisons using DCs derived from monocytes and CD34 progenitor cells of HIV-1-infected donors have not yet been performed. The results of these studies suggest that both M-DC and CD34-DC have potent immunostimulatory properties in vitro. Clinical trials utilizing DCs derived from different sources will be necessary to demostrate the clinical relevance of these in vitro findings.

Role of DCs in initiating and maintaining HIV-specific immunity DCs, considered to be ‘nature’s adjuvant’, are the most potent professional APC for priming T-cell responses. DCs are felt to be essential for stimulation of naïve resting T cells, but they are also potent at stimulating expansion of memory T cells (Knight and Stagg, 1993). The ability of DCs to prime MHC class I- and class II-restricted Tcell responses may result in large part from APC expression of high levels of class I and class II MHC molecules as well as costimulatory molecules, such as B7.1, B7.2 and CD40 (reviewed in Banchereau and Steinman, 1998). DCs preferentially induce the development of TH1type CD4 lymphocytes, but may also induce CD8 CTL responses in the absence of T helper cells or IL-2 (Bhardwaj et al., 1994; Macatonia et al., 1995). The preferential induction of TH1

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responses is felt in part to be due to secretion of IL-12 by DCs (Macatonia et al., 1995; Heufler et al., 1996), and the production of this critical immunostimulatory cytokine has been confirmed in appropriately stimulated human DC cultures (Cella et al., 1996; Wilson et al., 1997b; de Saint-Vis et al., 1998). This TH1-biasing effect of DCs on the immune system is likely to be important in the setting of HIV-1 infection, where the generation of a strong cell-mediated immune response against HIV-1 is felt to be critical in achieving control of viral replication. The therapeutic use of functional DCs might also help to overcome the inhibitory effects that a TH2 cytokine milieu, characteristic of progressive HIV-1 disease, could have on the generation of important TH1-type HIV-specific immune responses. Since DCs are likely to be the principal cells involved in priming HIV-specific T-cell responses in vivo, an understanding of the factors involved in DC presentation of HIV-1 antigens to naïve and memory T cells is crucial to the development of vaccines and immune-based therapies for HIV-1 infection. Whereas DC infectivity with HIV-1 and the ability of DCs to transmit HIV-1 to T cells has been an area of active investigation, less is known about the nature of the Tcell responses generated to HIV-1 antigens by DCs from HIV-infected or uninfected persons. DCs from normal donors have been shown to process and present antigens from a variety of sources and via a number of pathways (reviewed in Banchereau and Steinman, 1998). DCs have the ability to take up exogenous antigens by receptor-mediated endocytosis or sample their antigenic environment by fluid-phase macropinocytosis (Sallusto et al., 1995), and display the ability to present exogenous protein antigens in a MHC class I-restricted manner (Bohm et al., 1995). It has recently been shown that DCs may possess a unique pathway by which exogenous antigens are processed for presentation via MHC class I (Rodriguez et al., 1999). Studies to date suggest that DCs can process HIV-1 antigens, provided in multiple formats, and present them to both naïve and memory HIV-specific CD4 and CD8 T cells in vitro. DCs

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cultured from the peripheral blood of HIVnegative donors and pulsed with HIV proteins or transfected to express HIV-1 proteins have been shown to be potent stimulators of primary MHC class I- and II-restricted T-cell responses in vitro (Wilson et al., 1999; and unpublished observations). Several reports suggest that DCs cultured from the peripheral blood of HIV-1-infected patients have typical DC morphology and phenotype (Fan et al., 1997; Chougnet et al., 1999), and are also potent stimulators of memory HIV-specific CTLs in vitro. Autologous, cultured DCs from HIV donors that were pulsed with HIV-1 peptides or infected with recombinant vaccinia encoding HIV-1 genes were found to be better stimulators of memory HIV-specific CTLs than were similarly treated autologous B-lymphocyte cell lines (Fan et al., 1997). Peripheral bloodderived DCs cultured from HIV-infected patients and pulsed with HIV-1 proteins, especially when the HIV-1 proteins were complexed with lipids, were shown to be potent stimulators of MHC class I-restricted CD8 T cell responses in vitro (Zheng et al., 1999). Taken together, the results of the studies described above suggest that cultured DCs are potent stimulators of both naïve and memory HIV-specific T-cell responses. The factors governing the effectiveness of DC-based vaccine approaches in vivo, including such variables as DC source and antigen format, remain to be defined.

CLINICAL STUDIES DC therapies are currently under investigation in the setting of cancer, and they appear to be safe and to induce antitumor effects in vivo in preliminary studies (Hsu et al., 1996; Holtl et al., 1998; Nestle et al., 1998). The safety and immunogenicity of autologous mature DCs administered subcutaneously to nine healthy volunteers was investigated by Dhodapkar et al. (Dhodapkar et al., 1999). In this study, subjects received an initial injection of DCs without antigen, followed 4–6 weeeks later by a single injection of DCs loaded with antigen (KLH, influenza

matrix peptide, with or without tetanus toxoid). Four additional subjects were given antigens alone, without adjuvant. Although no local or systemic reactions were noted after the injection of DCs without antigen, local erythema without induration occurred in six of nine subjects after the injection of antigen-loaded DCs. The injections were generally well-tolerated with transient fatigue and low-grade fever reported in one subject each. Additionally, one subject developed a transient low-titer positive ANA test (1:40) at 60 days after immunization without any other evidence of autoimmunity. The injections of either antigen or DCs alone did not result in significant increases in antigen-specific T-cell proliferation. In contrast, the single injection of antigen-pulsed DCs resulted in priming of KLHspecific CD4 T cells in all nine subjects and boosting of tetanus-specific T-cell responses in five of six immunized. Increases in influenzaspecific CD8 T cells were measured as early as a week following DC immunization. This study definitively showed that mature, monocytederived DCs, loaded with either protein antigen or peptide, could serve to stimulate both naïve and memory CD4 and CD8 antigen-specific T cells in vivo following a single DC immunization. To date, a single pilot study involving the administration of antigen-loaded DCs to HIVinfected persons has been performed (Kundu et al., 1998), although several others are planned. This phase I clinical trial, carried out by Drs Kundu and Merigan at the Center for AIDS Research at Stanford University Medical Center, assessed the safety and antigen-presenting properties of allogeneic or autologous DCs pulsed with recombinant gp160 or synthetic peptides administered to six HIV-infected volunteers. DCs were isolated from leukapheresed PBMCs from the recipient (one case) or HLAmatched siblings (five cases) using stepwise centrifugation and a short culture period with antigen, without exogenous cytokines. DCs were incubated with HIV-1 gp160 or synthetic peptides corresponding to HLA-A2-restricted CTL epitopes (in env, gag, pol of HIV-1) and infused intravenously to HLA-A2 HIV-infected subjects six to nine times at monthly intervals. Study

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subjects had CD4 counts ranging from 300 to 700/mm3 at the initiation of the study and viral loads ranging from 3 to 5 log10 plasma HIV-1 RNA copies/ml. Following infusions, patients were monitored for changes in viral load and HIV-specific immune responses. No clinically significant adverse effects were seen in any patient, and plasma HIV-1 RNA and CD4 T cell numbers were stable during the study period. No enhancement in HIV-specific immunity was noted in the three subjects with starting CD4 counts of <410/mm3, but the three subjects with higher starting peripheral CD4 counts displayed evidence of enhanced HIV-specific CTLs and/or lymphoproliferative responses. The small size of this pilot study precluded any conclusions as to clinical benefit of this approach, but the infusions were well-tolerated and enhanced HIV-specific immune responses were seen in 50%. A number of DC-based strategies designed to promote or enhance HIV-specific immunity are currently under development. Investigators at Walter Reed Army Institute of Research plan to investigate the safety and immunogenicity of autologous DCs transfected with recombinant canarypox vector encoding HIV-1 env/gag/pol genes (ALVAC-HIV ) and administered subcutaneously to HIV-1 seronegative donors (Mary Marovich, personal communication). In this phase I study, 36 healthy adult volunteers will receive either intramuscular ALVAC-HIV, intradermal ALVAC-HIV, or ALVAC-HIV-transfected autologous DCs administered subcutaneously on the same vaccination schedule at months 0, 1, 3 and 6. DCs will be generated from the adherent fraction of peripheral blood mononuclear cells and cultured for 5 days in rHuGM-CSF and rHuIL-4, then matured with autologous monocyte-conditioned medium (MCM) and infected with the canarypox vector. Immunogenicity will be assessed using lymphocyte proliferative responses to vaccine component antigens, bulk chromium release assays for CTLs, and assessment of antigen-specific γ-interferon producing cells by ELISPOT and flow cytometric techniques. One goal of this study is to investigate whether differential targeting of DCs, via differ-

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ent modes or routes of vaccine administration, results in differences in the strength of cellular immune responses to the vaccine antigen. In this trial, it is expected that administration of transfected DCs will promote the greatest immune response, followed by the intradermal administration (which aims to target DCs directly in vivo), with the standard intramuscular approach being least effective. The results of this biological proof-of-concept study will help to determine how strongly vaccine strategies designed to target DCs in vivo should be emphasized clinically. Another clinical approach involves testing the hypothesis that in the setting of low-viral replication imposed by HAART in HIV-infected patients, cultured DCs may be safely administered as a therapeutic vaccine in order to generate and/or maintain a broad and strong HIV-specific cell-mediated immune response. Investigators at the University of Pittsburgh Medical Center are developing DC-based, therapeutic vaccination approaches that aim to generate and/or enhance HIV-specific cellmediated immune responses in HIV-1-infected subjects receiving highly active antiretroviral therapy (Sharon Riddler, personal communication). In a phase I/II trial still in the development stages, HLA-A2.1 HIV-infected patients with peripheral CD4 counts greater than 300 cells/mm3 who have undetectable HIV-1 viral loads (HIV-1 plasma RNA <400 copies/ml) for more than 6 months will be vaccinated with autologous DCs loaded with HIV-1 CTL epitope peptides. DCs obtained from adherent peripheral blood mononuclear cells will be grown for 7 days in rHuGM-CSF and rHuIL-4, then pulsed with conserved HIV-derived HLA-A2-restricted CTL epitope peptides and matured using CDYO ligation. Subjects will receive at least two DC immunizations, administered either subcutaneously or intravenously, at 3-week intervals. Subjects will be followed clinically for changes in peripheral CD4 count and plasma HIV-1 RNA (viral load). Immunogenicity of the DC vaccine will be assessed by evaluating peripheral blood T-cell responses to the vaccine antigen (proliferation, cytotoxicity, cytokine secretion, MHC

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class I tetramer staining). The primary objectives of this study are to determine the safety and tolerability of autologous, peptide-loaded DCs in HIV-1-infected subjects. This study will also help to determine whether DC immunization in the setting of HAART suppression of viral replication will allow for the generation of stronger HIV-specific immunity, as well as to compare the efficacy of DC vaccines administered by intramuscular or subcutaneous routes.

SUMMARY Given that overall immune function improves in HIV-infected patients on HAART, yet HIVspecific immune responses wane over time, it stands to reason that vaccination with HIV-1 antigens in this setting of viral suppression might result in enhanced HIV-specific immunity. Enhancing HIV-specific cellular immunity during viral suppression may ultimately allow for more complete immune control of viral replication in the event that antiretroviral therapy fails or can no longer be tolerated. As DCs are likely to be the principal cells involved in priming HIV-1-specific T-cell responses in vivo, an understanding of the factors involved in DC presentation of HIV-1 antigens to naïve and memory T cells is crucial to the development of vaccines and immune-based therapies for HIV-1 infection. Depletion and dysfunction of freshly isolated blood DCs have been reported in the setting of HIV-1 infection, a factor likely to contribute to the T-cell defects characteristic of HIV1 disease. Despite the adverse effects of HIV-1 replication on DCs, highly functional DCs that are free of replicating virus can be grown from precursors in the peripheral blood of HIV-infected individuals. These cultured DCs maintain their immunostimulatory characteristics and are capable of stimulating both naïve and memory HIV-specific T cells in vitro. It has been hypothesized here that DC dysfunction might be corrected, and a more effective cellular immune response to HIV-1 generated, through the therapeutic use of appropriately primed,

cultured autologous DCs in the setting of viral suppression with HAART. Future clinical studies will be necessary to determine the most effective source of DCs, dose, route of administration, antigen format, and when to best apply DC vaccines therapeutically during the course of HIV-1 disease.

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42 Dendritic cell tolerogenicity and prospects for dendritic cell-based therapy of allograft rejection and autoimmune disease Lina Lu and Angus W. Thomson Thomas E. Starzl Transplantation Institute, and Department of Surgery, University of Pittsburgh Medical Center, Pittsburgh, PA, USA

Emerging evidence points towards a role for DCs in tolerance, and it will therefore be imperative to pursue DCs more vigorously in allergy, transplantation and autoimmunity. Jacques Banchereau and Ralph M. Steinman Nature

INTRODUCTION

tolerance within the thymus (Jenkinson et al., 1985; Matzinger and Guerder, 1989). DCs, including myeloid DC progenitors (Thomson and Lu, 1999a), lymphoid DCs (Süss and Shortman, 1996) and even ‘classic’ myeloid DCs (Inaba et al., 1997), were subsequently shown to be able to induce peripheral, antigen (Ag)specific unresponsiveness in various experimental models, or implicated as having a role in self tolerance. Mechanisms whereby DCs accomplish this, include selective activation of the TH2 subset (Khoury et al., 1996), induction of regulatory T cells (Clare-Salzler et al., 1992), induction of T-cell anergy (Lu et al., 1995a), or promotion of T-cell apoptosis (Süss and Shortman, 1996; Lu et al., 1997a). Moreover, what is important is

Dendritic cells (DCs) constitute a complex system of bone marrow (BM)-derived antigenpresenting cells (APCs) that initiate and regulate immune responses. Extensive recent studies have improved our understanding of DC-lineage development, differentiation, activation and function. The conventional view of DCs is that they play an important role as immunologic adjuvants and offer potential for therapy of infectious disease and cancer. The concept of tolerogenic DCs however, is also now wellrecognized. A role for DCs in central tolerance induction was demonstrated initially in the context of self Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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evidence that tolerogenic APCs traffic from peripheral tissues to draining lymph nodes and to there initiate the proliferation of naïve T cells. The biologic characteristics of DCs, such as antigen capture, trafficking to T-cell areas, and constitutive expression of major histocompatibility complex (MHC) class II, make this cell a potential tolerance inducer. Current data support the hypothesis that low or absent expression of critical cell surface costimulatory molecules (CMs), concomitant with stable antigen presentation, can render responding T cells unable to enter the cell cycle, or to undergo apoptosis. It is also believed that DCs not only provide a common set of signals to initiate clonal expansion of T-cells, but also provide T cells with selective signals leading to differential T-cell subset expansion. An explanation for these different Tcell responses to DCs is provided by the study of Rissoan et al. (Rissoan et al., 1999), who proposed that DCs exist as distinct subsets that differ with respect to their lineage origin, surface molecule expression and biological function. The development of subsets of DCs and their function is affected by environmental factors, in particular the cytokines, IL-10 and transforming growth factor β (TGFβ), prostaglandin E2 (PGE2), and corticosteroids. These molecules downregulate CM or MHC class II antigen expression, or inhibit IL-12 production by DCs, giving rise to ‘tolerogenic’ DCs with a TH2-driving function (Steinbrink et al., 1997) or capacity to directly inhibit CD4 T-cell proliferation and the cytotoxic activity of CD8 T cells (Moore et al., 1993; Caux et al., 1994a). Many other molecules that influence DC function can potentiate their tolerogenicity. Thus, the chimeric fusion protein cytotoxic T lymphocyte antigen 4 (CTLA4)-Ig can markedly subvert DC-induced T-cell responses in vitro (Lu et al., 1997b). Fas ligand (L) (CD95 L), that is expressed on murine lymphoid (Süss and Shortman, 1996) or myeloid DCs (Lu et al., 1997b), and tumor necrosis factor (TNF) receptor apoptosis-inducing ligand (TRAIL) (Fanger et al., 1999), that is expressed on human CD11c blood DCs, may regulate or eliminate T cells responding to antigen presented by the DCs. Thus, genetically engineered DCs

expressing immunosuppressive molecules, such as viral (v) IL-10, TGFβ, Fas L or CTLA4-Ig may offer potential for the suppression/deletion of allo- or autoantigen-specific T cells, with implications for tolerance induction. One of the challenges of contemporary applied DC biology is to ascertain whether DCs can be utilized for the selective prevention/ therapy of undesired immune responses, as occur in autoimmune diseases, allograft rejection, and allergic hypersensitivity reactions.

DCs AND TOLERANCE INDUCTION Tolerance is a fundamental property of the immune system and underlies selective unresponsiveness to self autoantigens. It is the ultimate goal of transplant immunologists seeking permanent, drug-free, donor alloantigenspecific immunologic unresponsiveness in humans, and of those wishing to restore tolerance to the autoantigens implicated in the pathogenesis of autoimmune diseases. The principal mechanisms by which tolerance is thought to be induced and/or maintained are clonal deletion, anergy and suppression by regulatory cells. One of the principal determining factors in these processes is the function of the APCs that are critical for initiation of the primary immune response. DCs can present antigen in either a tolerogenic or immunogenic fashion (Finkelman et al., 1996) depending on the inoculation regimen used. Recent studies in the human and mouse demonstrate that subsets of DCs from different lineages or different tissues may determine the type of immune response induced. Freshly isolated Peyer’s patch, but not spleen DCs produce IL-10, and induce the differentiation of TH2 cells (Iwaski and Kelsall 1999) that may account for oral tolerance. TH2 cytokine-secreting cells were detected in mice injected with liver-derived DC progenitors. By contrast, comparatively high numbers of interferon (IFN)-γ cells were induced by injection of BM-derived DCs (Khanna et al., 2000). DCs from fms-like tyrosine

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kinase 3 (Flt-3) ligand (Flt-3L)-treated mouse liver on the other hand, induce strong TH1 responses (Morelli et al., 2000). A subset of murine DCs (lymphoid DCs) regulates the responses of naïve CD8 T cells by limiting their IL-2 production (Kronin et al., 1996). Observations of the tolerogenic capacity of DC in many diverse experimental systems are summarized in Table 42.1, and include reports of a role for DCs both in central and peripheral tolerance. In organ transplantation, DCs have consistently been identified as prominent donor cells in the multilineage hematopoietic cell microchimerism that is observed in long-term surviving rodent and human allograft recipients (Starzl et al., 1992, 1993, 1996; Demetris et al., 1993; Qian et al., 1994), whether or not immunosuppressive therapy is used to promote graft survival. This finding, together with the well-known properties of DCs as highly effective APCs (Steinman, 1991), prompted our investigation of the significance and possible mechanistic role of donor-derived DCs in organ transplant tolerance. The possible role of DCs in tolerance induction was further evaluated in liver transplantation studies. Compared with the heart or kidney, the liver is comparatively rich in total number of DCs, yet is generally regarded as the least immunogenic of transplanted whole organs. In the mouse, liver transplant tolerance is achieved in virtually all MHC-incompatible mouse strain combinations, without the need for immunosuppression (Qian et al., 1994). Using techniques established to propagate DC progenitors from normal mouse liver (Lu et al., 1994), it was found that DC progenitors of donor origin could be propagated from the BM of mice that spontaneously accepted MHC-mismatched liver allografts. Donor-derived DCs could not however, be propagated from the marrow of mice that acutely rejected heart grafts from the same donor strain (Lu et al., 1995b; Thomson et al., 1995a). Donor-derived DC progenitors can also be propagated from the blood of human liver allograft recipients given adjunctive donor BM (Thomson et al., 1995b; Rugeles et al., 1997).

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Evidence from recent immunohistochemical studies of Demetris et al. (1997) further suggests that persistence of donor-derived DCs may be important in the long-term maintenance of transplantation tolerance and the prevention of (chronic) organ allograft rejection. Growth, maturation and function of interstitial DCs in the liver are affected by microenvironmental factors, in particular cytokines, that are influenced by the process of liver transplantation. The interstitial DCs within normal liver allografts that are accepted without immunosuppressive therapy in mice, are predominantly functionally immature, with tolerogenic potential. Treatment of liver donors with Flt-3L strikingly augments the number of potential stimulatory DCs, leading to rapid induction of host antidonor immune responses and acute liver graft rejection, an effect that can be reversed by blocking the allo-stimulatory function of the DCs with CTLA4-Ig or anti-CD40L mAb (Thomson and Lu, 1999b). Each of these observations suggests that allogeneic DCs might play a role in the induction of tolerance, rather than simply be a consequence of it.

Central tolerance induced by DCs The thymus plays a major role in immune homeostasis by the deletion of self-reactive T cells, thus contributing to the maintenance of self tolerance. A number of cell types within the thymus, including DCs, provide signals responsible for the negative selection of T cells. Matzinger and Guerder (1989) outlined the critical role of DCs in central tolerance by demonstrating that the tolerogenic properties of the APC-depleted mouse thymus could be restored by reconstitution with purified splenic DCs. Intrathymic injection of Mls-incompatible spleen or thymic DCs can induce tolerance via clonal anergy (Inaba et al. 1991). Similar results have been reported in parent → F1 BM chimeras, and in transgenic mice (Widera et al., 1987; Gao et al., 1990). Strong T-cell tolerance in parent →F1 BM chimeras, prepared with supralethal irradiation, was evident at the level of mature thymocytes, and presumably occurred in the

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Observations of the tolerogenic capacity of dendritic cells (DCs)

Tolerogenic effect reported

Reference

Thymic DCs

Encephalitogen pulsing and adoptive transfer prevents EAE Tolerance acquired to host MHC in chimeric thymus Deletion enhances effector phase of CH Transfer reduces incidence of diabetes in NOD mice in vivo Diabetogenic peptide-pulsed/unpulsed DCs prevent diabetes in NOD mice IFN-γ-stimulated NOD mouse DCs exert antidiabetogenic effects Reconstitution of DC-depleted thymuses restores ability to delete thymocytes in vivo

Khoury et al., 1995 Jenkinson et al., 1985 Grabbe et al., 1995 Clare-Salzler et al., 1995

Langerhans cells Pancreatic lymph node DCs BM-derived DCs Splenic DC

Co-stimulator-deficient DC progenitors

CD40-deficient BM DCs Renal interstitial DCs Cervical lymph node DCs Mesenteric lymph node DCs Peyer’s patch DCs Mouse DCs

Large numbers of DCs reduce local HVG reaction in vivo Large numbers of DCs/high antigen load inhibit antitumor immunity CD8/Fas-L DCs induce apoptosis in activated T cells in vitro CD8 DCs regulate the response of naïve CD8 T cells by limiting their IL-2 production Induction of alloantigen-specific T cell unresponsiveness in vitro Adoptive transfer prolongs allograft survival Induction of IL-10-producing T cells and downregulation of CD8 responses Induction of Tr1 cells that suppress responses of activated T cells Induction of alloantigen-specific T-cell hyporesponsiveness Discordant expression of MHC class II and invariant chain may be a regulatory mechanism for peripheral T-cell tolerance Adoptive transfer delays the onset of lymphocytic thyroiditis Uptake of dying cells by DCs for maintenance of tolerance to self-Ag Induce the differentiation of TH2 cells, oral tolerance Injection of rat anti-DC mAbs induces rat Ig-specific tolerance Induction of T-cell anergy by tumor-associated Ag

Feili-Hariri et al., 1999 Shinomita et al., 1999 Matzinger and Guerder, 1989 Knight et al., 1983a Knight et al., 1983a Süss and Shortman, 1996 Kronin et al., 1996 Lu et al., 1996 Fu et al., 1996; Rastellini et al., 1995 Dhodapkar et al., 2001 Jonuleit et al., 2000 Gao et al., 1999 Saleem et al.,1997 Delemarre et al., 1995 Huang et al., 2000 Iwasaki and Kelsall, 1999 Finkelman et al., 1996 Grohmann et al., 1997

EAE, Experimental allergic encephalomyelitis, CH, contact hypersensitivity, NOD, non-obese diabetic, HVG, host-versus-graft.

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TABLE 42.1

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thymus itself. It was demonstrated that T-cell contact with thymic epithelial cells (including DCs) induced clonal deletion of most of the host-reactive T cells, but also spared a proportion of low-affinity cells. These findings indicate that the role of DCs in deletion of autoreactive T cells within the thymus is not dependent upon unique characteristics of thymic DCs, but may also be mediated by signaling provided by DCs from other tissues. Since the thymus is the primary site for central tolerance induction, a direct approach to tolerance by clonal deletion is intrathymic antigen injection. Acquired thymic tolerance, induced by intrathymic injection of antigen has been demonstrated in a number of autoimmune diseases (Koevary and Blomberg, 1992; Gerling et al., 1992; Khoury et al., 1993; Goss et al., 1994). Thus, injection of the immunodominant peptide of myelin basic protein (MBP) into the thymus of Lewis rats, protects the animals from experimental allergic encephalomyelitis (EAE) (Khoury et al., 1993). Thymic DCs isolated from these animals can transfer protection to naïve recipients, suggesting that DCs mediate the effects of acquired thymic tolerance (Khoury et al., 1995). Moreover, thymic but not splenic DCs pulsed ex vivo with the encephalitogenic peptide protect naïve recipients from EAE when injected systemically (Khoury et al., 1996). Interestingly, animals thymectomized before systemic injection of the ex vivo pulsed APCs were not protected, raising the question of sitespecific homing of DCs. The capacity of intrathymic injection of donor spleen cells (Ohzato and Monaco, 1992), BM cells, renal glomeruli (Remuzzi et al., 1991), pancreatic islets (Posselt et al., 1990) or DCs (Ridge and Matzinger, 1996) to induce tolerance has been reported, indicating that this effect is not restricted to hematopoietic cells. While intrathymic injection has been very effective for tolerance induction in rodents, successful large animal studies have not been reported. In humans, the disadvantages of this approach are that (1) there is no effective method to eliminate mature donor-reactive T cells in large animals, and (2) the thymus of

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adult large animals, including humans, is involuted, making it difficult to determine whether it still functions.

Peripheral tolerance induced by DCs It is now evident that DCs can provide both stimulatory and downregulatory signals for immune reactions. Recent studies suggest that in vitro generated myeloid DCs with an ‘appropriate’ surface phenotype (‘immature’ or ‘costimulatory molecule deficient’), or CD8α lymphoid DCs (in mice) can subvert allogeneic T-cell responses. In addition, adoptive transfer of autoantigen-pulsed thymic DCs (Khoury et al., 1995) or splenic DCs whose co-stimulatory molecule expression has been blocked by the chimeric fusion protein CTLA4-Ig (Khoury et al., 1996), can inhibit development of autoimmune disease (EAE) in rodents. Possible mechanisms involved in DC tolerogenicity can be summarized as follows. Selective activation of TH 2-like T-cell subsets Splenic APCs, particularly DCs, can mediate protection from EAE if they are pulsed with peptide in vitro in the presence of CTLA4-Ig (Khoury et al., 1996). The mechanism of protection is probably a TH2 switch, since immunohistology of the CNS demonstrated almost complete inhibition of the TH1 cytokines, IL-2 and IFN-γ, with upregulation of the TH2 cytokines, IL-4 and IL-13. Splenic DCs pretreated with TGFβ and peptide were also tolerogenic (S.J. Khoury, personal communication). Thus, nontolerogenic DCs may become tolerogenic by blocking their ability to provide costimulatory signals, or by modification with suppressive cytokines (Liu et al., 1998). Other investigators have observed that Ag-specific suppression of delayed-type hypersensitivity responses can be achieved by i.v. administration of Ag-pulsed Langerhans cells (LCs) (Morikawa et al., 1992) or splenic DC (Morikawa et al., 1993), possibly via a mechanism involving selective activation of TH2-like T-cell subsets (Morikawa et al., 1995).

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It appears that under steady-state conditions, resident (rat) respiratory tract DCs are programmed for the induction of TH2-biased responses. A possible explanation for this finding is the presence of gene transcripts encoding IL-10 within the DCs, suggesting that IL-10 production by these cells might contribute toward the TH2 bias operative in mucosal immune responses to inert protein antigens (Stumbles et al., 1998). In addition, freshly isolated Peyer’s patch DCs produce IL-10 and induce differentiation of TH2 cells (Iwaski and Kelsall, 1999). Evidence that the comparative immunologic privilege of hepatic allografts might be due, in part, to induction of TH2 responses by liverderived DC progenitors was provided by Khanna et al. (2000) who showed that these cells induce clusters of IL-10- and IL-4-secreting MNCs in allogeneic lymphoid tissue in vivo, whereas much reduced IFNγ responses were observed compared with those induced by BM-derived DCs. Induction of regulatory T cells T regulatory (Tr) cells are a variety of cells that display regulatory functions in vitro or in vivo. They can be subdivided into a number of subsets based on expression of cell surface markers, production of cytokines, and mechanisms of action. Recent work by Jonuleit et al. (2000) and Dhodapkar et al. (2001) strongly suggests that immature DC may control peripheral tolerance by inducing the differentiation of human Tr cells. Thus, immunization with immature DC results in downregulation of CD8 T-cell responses to recall antigen and induction of IL-10producing T cells. In addition, immature DC-induced Tr1 cells can suppress responses of activated T cells. These observations open new therapeutic perspectives for the use of immature DC in autoimmune diseases and allogeneic cell or organ transplantation, and also imply that use of immature DC for anti-cancer vaccination should be avoided. Protection of prediabetic nonobese diabetic (NOD) mice from development of type-1 diabetes by transfer of DCs isolated from pancreatic

lymph nodes of diabetic NOD mice (ClareSalzler et al., 1992), may possibly be mediated by a mechanism involving the enhanced induction of regulatory T cells. Similarly, Delemarre et al. (1995) reported that transferring cervical node DCs from autoimmune lymphocytic thyroiditis (LT)-resistant BB/Wor rats to LT-prone BB/Wor recipients delayed the onset of LT. Induction of T-cell anergy Selective blockade of costimulatory signals is a promising strategy for therapeutic immunosuppression (Larsen et al., 1996; Kirk et al., 1997). We and others have demonstrated that DCs whose allostimulatory function is impaired, either by incomplete maturation, selective blockade of B7 family costimulatory molecules (using CTLA4-Ig), the influence of specific cytokines (i.e. IL-10) or ultraviolet B irradiation can either induce alloantigen-specific hyporesponsiveness or apoptosis in vitro, or suppress in vivo immune reactivity (Table 42.1). It has also been reported that retroviral delivery of vIL10 to replicating murine myeloid DC progenitors renders these cells capable of inducing alloantigen-specific hyporesponsiveness in vitro (Takayama et al., 1999). Grohmann et al. (1997) reported that a tumor-associated and self antigen peptide presented by DC could induce T-cell anergy in vivo, but that IL-12 could prevent or revert the anergic state. Intrathymic injection of antigen may mediate peripheral tolerance through interaction of thymic DCs with activated peripheral T cells that circulate to the thymus. It has been suggested that this interaction leads to peripheral T-cell anergy (Chen et al. 1997). Induction of activated T-cell apoptosis The expression by DCs of molecules associated with the inhibition of T-cell growth or the induction of T-cell apoptosis (i.e. nitric oxide (NO), or FasL) may render DCs capable of subverting Tcell responses. Mouse splenic CD8α lymphoid DCs, that express FasL, induce apoptosis in activated allogeneic CD4 T cells, resulting in

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diminished T-cell proliferation in mixed leukocyte reactions (MLRs) (Süss and Shortman, 1996). These DCs can also impair IL-2 production by CD8 T cells. DCs exposed to IFNγ, bacterial endotoxin ( lipopolysaccharide; LPS), or allogeneic T cells can synthesize NO synthase (NOS). NO production suppresses allogeneic Tcell proliferation in MLRs, and also induces apoptosis, both in activated T cells and in the DCs themselves (Lu et al., 1996; Bonham et al., 1996a). Highly purified myeloid DCs grown from mouse BM in GM-CSF  IL-4 have been found to express FasL mRNA by reverse transcriptase polymerase chain reaction (RT-PCR), and to uniformly express surface FasL by both flow cytometric and immunocytochemical analysis. These cells, but not DC propagated from FasLdeficient (gld) mice, induced dose-dependent increases in DNA fragmentation in Fas Jurkat T cells. The same DCs also induced apoptosis of alloactivated T cells, once the CD28/B7 pathway was blocked using CTLA4-Ig (Lu et al., 1997a). It is interesting to speculate that FasL donor liverderived DCs might be responsible for host T-cell deletion observed in murine spontaneous liver allograft acceptance (Qian et al., 1997a). Cell interactions resulting in the apoptosis of liver graft-infiltrating T cells might be mediated by FasL donor DCs located in portal areas (Woo et al., 1994). It has yet to be determined, however, whether there is selective deletion of specific, donor-reactive T cells in the mouse orthotopic liver transplant model (Qian et al., 1997b) in which graft acceptance occurs in most strain combinations without immunosuppression. Apoptosis of immune-reactive host cells is also associated with the marked prolongation of heart allograft survival observed in mice preconditioned by systemic infusion of MHC class IIhi, costimulatory molecule-deficient donor DC, in combination with anti-CD40L mAb (Lu et al., 1999a). Another important apoptosis-inducing ligand, TRAIL, is expressed on human CD11c blood DCs after stimulation with either IFNγ or α, and confers the ability to kill TRAIL-sensitive targets, including activated T cells (Fanger et al., 1999). TRAIL may serve as an innate effector

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molecule on CD11c DCs for the regulation or elimination of DC-activated T cells. Downregulation of the clonal expansion of B cells by follicular DC In germinal centers, B lymphocytes are intimately associated with follicular dendritic cells (FDCs). It has been hypothesized that FDCs are involved in the regulation of B-cell growth and differentiation through cell–cell interaction. Highly enriched preparations of human FDCs were cultured with mitogen-stimulated B cells in the absence of T cells, and B cell [3H]-TdR uptake was inhibited by up to 80% (Freedman et al., 1992). Furthermore, supernatants from cultured FDCs were also able to inhibit B-cell proliferation. These results demonstrated that FDCs may downregulate the clonal expansion of B cells that occurs within lymphoid follicles as part of the normal physiologic immune response. Significance of the route of DC administration, cell number and antigen load Under experimental conditions, DCs exhibit a dichotomous potential to regulate immune responsiveness, depending on the route of in vivo administration, number of cells injected, the amount of antigen presented, and the source or population of DCs. Thus, administration of DCs loaded with low doses of tumor antigen can enhance tumor rejection in mice, while administration of DCs loaded with high doses of tumor antigen, or administration of large numbers of tumor antigen-pulsed DCs inhibits the antitumor effect of DCs (Knight et al., 1983a). BM-derived DC propagated in GM-CSF  IL-4 accelerate donor-specific kidney allograft rejection if delivered i.v., but can prolong pancreatic graft survival when administrated via the portal vein (Gorczynski et al., 1996). Liver-derived DC progenitors have been shown to prolong pancreatic islet allograft survival. By contrast, splenic DCs propagated from the same mice, using the same technique, induce rejection of islet allografts in diabetic recipients (Rastellini et al., 1995). There is also considerable evidence

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that specific components of the local tissue microenvironment, e.g. the cytokine TGFβ, may be responsible for imparting tolerogenic activity to DCs that migrate from the anterior chamber of the eye (an immunologically privileged site) to regional lymphoid tissue (Streilein et al., 1992). In summary, DCs are capable of eliciting positive (sensitizing) and negative (tolerizing) responses in the immune system. This reflects molecular regulation of the immunological activity of DC subsets, including expression of first and second signals for T-helper cell activation, stimulatory or suppressive cytokines (IL-6, IL-10, IL-12), and death-inducing ligands by these important APCs (reviewed later). The ability to generate immunoregulatory DCs in sufficient number, under well-defined conditions, has considerable implications for the development of effective, DC-based therapy of cancer, infectious disease, autoimmunity and allograft rejection.

MOLECULAR BASIS OF DC TOLEROGENICITY Costimulatory molecule deficiency The potent capacity of DCs to activate immunologically naïve (resting) T cells is related to their high constitutive expression of MHC class II and T-cell costimulatory molecules, such as CD40, CD80, CD86 and OX40L. Deficiency or absence of expression of these molecules reduces the APC function of DCs, and may render them potentially tolerogenic. Thus, DCs residing in peripheral nonlymphoid tissues (e.g. skin) express relatively low levels of co-stimulatory molecules, and are poor T-cell stimulators when freshly isolated (Larsen et al., 1994). Others have observed that LC precursors in the epidermis function poorly as T-cell stimulators (Romani et al., 1986), and there is even evidence that LCs may provide downregulatory signals during elicitation of cutaneous (contact sensitivity) inflammatory reactions (Grabbe et al., 1995). MHC class IIlo DC precursors in the airway epithelium of rats (Nelson and Holt, 1995;

Stumbles et al., 1998), freshly isolated heart or kidney DCs (Austyn et al., 1994), and DC progenitors propagated from the mouse liver (Lu et al., 1994; Khanna et al., 2000) are also poor allostimulators. Absence of expression of invariant chain by interstitial DCs within nonlymphoid tissue (Saleem et al., 1997) may also be a significant factor in the regulation of self tolerance. Presentation of antigen to T cells by APCs in the absence of costimulatory molecules induces T-cell anergy (Schwartz, 1990). The importance of costimulatory molecule deficiency on DCs in peripheral tolerance induction has been emphasized by the fact that CTLA4-Ig, a potent blocker of the costimulatory B7–CD28 pathway, significantly prolongs allograft survival in various organ transplantation models (Lin et al., 1993; Sayegh et al., 1995). Studies in our laboratory have demonstrated that DCs propagated in vitro from mouse BM in the presence of suboptimal concentrations of GM-CSF express only low levels of costimulatory molecules, i.e. MHC class II, CD80lo, CD86. These cells not only induce donor-specific T-cell unresponsiveness in vitro (Lu et al., 1995a), but also prolong donor strain cardiac (Fu et al., 1996) or pancreatic islet allograft survival (Rastellini et al., 1995), when administered systemically 1 week before transplantation. Likewise, in an autoimmune disease model, MBP-pulsed DCs on which CD80 and CD86 expression has been blocked by CTLA4-Ig protect rats from the induction of EAE (Khoury et al., 1996). It has also been suggested that rhesus monkey BM-derived DC progenitors (MHC class II/dim) may exert tolerance-promoting activity, both in vivo and in vitro (Thomas et al., 1994). These observations have led to speculation that costimulatory molecule-deficient DCs could play a role in peripheral T-cell tolerance induction.

Expression of death-inducing ligands It has been proposed that expression of CD8 by the lymphoid DC subpopulation of mouse thymus and spleen (Welsh and Kripke, 1990; Vremec et al., 1992) might enable these cells to

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exhibit a ‘veto’ function (Hambor et al., 1990; Sambhara and Miller, 1991). Further work demonstrated that CD8α DCs isolated from the spleen expressed FasL and were capable of killing activated CD4 T cells via the Fas/FasL pathway (Süss and Shortman, 1996). However, more recent evidence indicates that CD8 is not important in the ‘veto’ function of these lymphoid DCs (Kronin et al., 1997). In a recent study we have also demonstrated (Lu et al. 1997a) that DCs propagated from mouse BM in GM-CSF  IL-4 (myeloid DC; MHC class IIhi, CD80hi, CD86hi) express functional FasL, but induce only a low level of apoptosis in alloantigen-activated T cells. Once the B7–CD28 pathway is blocked with CTLA4-Ig, however, significantly increased T-cell apoptosis can be detected, indicating that FasL and B7 molecules on DC may play counterregulatory roles in proliferation and survival of T cells following DC–T cell interaction. It appears that Fas–FasL interaction is probably not the only death-inducing pathway that exists in DC–T cell interaction, because a high level of Tcell apoptosis can also be induced by DCs propagated from the BM of FasL-deficient (gld ) mice in the presence of CTLA4-Ig (Lu et al., 1997a). These studies suggest that other molecules expressed by DCs may contribute to DC-induced apoptosis. Candidate molecular pathways include TNF/TNF-R, and TRAIL/ TRAIL-R (Lynch et al., 1995; Wiley et al., 1995). There is recent direct evidence that activated human blood CD11c DCs express TRAIL and kill TRAIL-sensitive target cells. Moreover, soluble TRAIL-R blocked target cell death, suggesting that DC-mediated apoptosis was TRAIL specific. By contrast, the human IL-3Rαpre-DC subset displayed minimal cytotoxicity toward the same targets, demonstrating a molecular basis for functional differences between the human CD11c (DC1) and IL-3Rα (DC2) DC subsets (Fanger et al., 1999).

Production of NO Studies of LPS-treated mice have suggested that NO production by presumptive intrathymic DCs may be responsible for thymocyte apoptosis and

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possibly self tolerance (Fehsel et al., 1995). These findings are consistent with our observation of NO production by a subpopulation of MDCs following treatment in vitro with IFNγ or LPS, or interaction with allogeneic T cells (Bonham et al., 1996a; Lu et al., 1996).

NF-κB inhibition The active dimer of nuclear factor (NF)κB is a rapid response transcription factor that plays a central role in immunological processes by regulating genes involved in immune and inflammatory responses (Rescigno et al., 1998). There is now considerable evidence that NFκB activation is responsible for DC maturation and activation. Thus, mature DCs express high levels of NFκB family proteins (Granelli-Piperno et al., 1995), whereas signaling by members of the TNFα R family, such as CD40 and RANK, results in activation of NFκB in DCs (Banchereau and Steinman, 1998). Rescigno et al. (1998) reported that TPCk, an NFκB inhibitor, effectively blocked LPS-induced nuclear translocation of p65, associated with inhibition of costimulatory molecule expression on DCs (Bielinska et al., 1990). Studies performed in Rel B knockout mice have suggested that Rel B, a member of the NFκB family, is a key molecule that determines the constitutive expression of κB-containing housekeeping genes in DCs (Lee and Burckart, 1998). Our recent studies show that inhibition of NFκB translocation with cyclosporine (CsA) is linked to decreased costimulatory molecule expression and allostimulatory activity of DCs. This effect could be partially overcome by IL-4, or potentiated by TGFβ (Lee et al., 1999). The double-stranded phosphorothioate oligodeoxynucleotide, NFκB ODN, that specifically binds NFκB, inhibits costimulatory molecule expression on DCs, without or with only minimal effect on MHC antigen expression that is necessary for the induction of antigen-specific T cells (Giannoukakis et al., 2000). Moreover, inhibition of NFκB translocation in DCs with NFκB ODNs potentiates the tolerogenicity of costimulator-deficient DC, both in vitro and in vivo (Giannoukakis et al., 2000).

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DC subsets/microenvironmental factors Evidence has accumulated that DC subsets can determine the type of immune response by providing T cells with polarizing signals (Kapsenberg and Kalinski, 1999). It remains unclear what the signals are and how they work. Most reports have demonstrated that the cytokines produced by DCs are important factors in directing T-cell proliferation at an early stage of DC–T cell interaction. Tissue environmental factors and the nature of invading pathogens appear to affect the development of different DC subsets. For instance, viruses may induce the development of type 1 DCs with high IL-12 production, whereas helminths may induce type 2 DCs, as a result of their PGE2producing capacity (Kalinski et al., 1999). Thus the special properties of DCs (surface expression of costimulatory/regulatory molecules, responsiveness to immunosuppressive cytokines, T-cell interaction, and in vivo homing to T-dependent areas of lymphoid tissues (Austyn and Larsen, 1990) make their manipulation to maximize their tolerogenic potential an attractive approach for cell-based therapy of immune-mediated disorders (allograft rejection and autoimmune disease). These manipulations will now be considered in more detail.

PROPAGATION OF TOLEROGENIC DCs FOR THERAPEUTIC APPLICATION – THE INFLUENCE OF CYTOKINES, TRANSCRIPTION AND MICROENVIRONMENTAL FACTORS Even with extensive processing procedures, only trace numbers of DCs can be obtained from normal tissue. Since Inaba et al. (1992a, 1992b) first described a method to propagate DC progenitors from normal mouse BM or blood with GM-CSF, the availability of large numbers of myeloid DCs has opened up possibilities for immunotherapy using DCs as vaccines. Several

groups have subsequently described methods for culture of large numbers of highly stimulatory human, nonhuman primate, mouse or rat DCs derived from different tissues using GMCSF, most often in combination with IL-4. More limited studies have reported the propagation of DCs with tolerogenic properties. It has been well-demonstrated that cytokines play a key role in the regulation of DC maturation and activation. TNFα enhances the development of mature human DCs from progenitors (Caux et al., 1992; Szabolcs et al., 1995), whereas IL-1β inhibits the function of LC (Grabbe et al., 1994a). Monocytes represent an abundant source of precursors that can polarize towards either DCs or macrophages, depending on the external stimuli. This polarization can be driven in vitro by the addition of appropriate cytokines (GM-CSF  IL-4 or macrophage (M)-CSF  colony-stimulating factor-1). Monocyte-derived DCs generated in culture with GM-CSF and IL-4 have a very high level of cell surface MHC class II and costimulatory molecules. In addition, they express the low-affinity receptor for Fcγ, FcγRII (CD32) (Sallusto et al., 1995). Rather conflicting results have been obtained concerning the effects of IFNγ on the antigenpresenting function of DCs, and on the expression of costimulatory or MHC class II molecules by DCs (Grabbe et al., 1994b; Lutz et al., 1996). IL10 has been shown to reduce the expression of HLA-DR or CD86 molecules on human peripheral blood DCs (Buelens et al., 1995), and murine LCs (Enk et al., 1993; Kaeamura and Furue, 1995). It inhibits the upregulation of CD58, CD83 and CD86 on in vitro generated human DCs (Steinbrink et al., 1997). There is also evidence that IL-10 pretreatment strongly inhibits the DCinduced responses of naïve and primed CD4 T cells in allogeneic MLR and anti-CD3 assays, and induces alloantigen- or peptide-specific anergy in CD4 T cells (Steinbrink et al., 1997). Moreover, IL-10 suppresses production of IL-8 and TNFα by DCs activated by LPS (Buelens et al., 1997) and skews towards TH2 differentiation by blocking IL12 synthesis by DCs (Buelens et al., 1997; De Smedt et al., 1997). It has also been reported that IL-10 accelerates murine LC apoptosis in culture

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TABLE 42.2 Experimental manipulations rendering tolerogenicity to DC Dendritic cell type

Tolerogenic effect reported

Reference

CTLA4-Ig-treated splenic DCs IL-10 treated Langerhans cells IL-10 treated splenic DCs Immature DCs exposed to IL-10

Protection from EAE Ag-specific anergy cells induction in vitro Induce TH2 response in vivo Induction of alloantigen- or peptide-specific anergy in T cells Promote TH2 development Promote TH2 responses

Verberg et al., 1996 Enk et al., 1993 De Smedt et al., 1997 Steinbrink et al., 1997

DCs grown in PGE2 Corticosteroid-induced monocyte-derived DCs (DC3) Ultraviolet B-irradiated Langerhans cells DCs grown in CsA NFκB-specific decoy-treated DCs

Ag-specific unresponsiveness of TH1 cells in vitro Induction of Ag-specific unresponsiveness Induction of Ag-specific unresponsiveness in vitro, and prolongation of graft survival

Kalinski et al., 1997 De Jong et al., 1999 Simon et al., 1991 Lee et al., 1999 Giannoukakis et al., 2000

EAE, experimental allergic encephalomyelitis, CsA, cyclosporine.

(Ludewig et al., 1995). Interestingly, Kalinski et al. have reported that DCs grown in the presence of prostaglandin(PG) E2 are unable to secrete IL-12, and when they present antigen to T helper cells, they promote the development of TH2 cells (Kalinski et al., 1997). DCs propagated from different tissues may exhibit different phenotypes and functions. Experiments using adult mice have shown that the liver harbors stem cells and DC progenitors that can be propagated using GM-CSF, but that only reach functional maturation when exposed to extracellular matrix protein (collagen type-1) with which DCs are spatially associated in normal liver (Lu et al., 1994). Donor-derived liver DC progenitors prolong pancreatic islet allograft survival (Rastellini et al., 1995). Clearly, it is possible to propagate potentially tolerogenic DCs from different sources of precursors under welldefined laboratory conditions. In vitro manipulations that may confer tolerogenic properties on DCs are summarized in Table 42.2. Here we review several methods for obtaining ‘tolerogenic’ DCs.

Propagation of ‘tolerogenic’ DCs using suboptimal concentrations of GM-CSF GM-CSF is a key cytokine for myeloid DC propagation from proliferating precursors or

hematopoietic stem cells, and its effect on the maturation of DCs appears to be dose-related. DC progenitors deficient in cell surface costimulatory molecules can be propagated from the BM of normal or NOD mice using an appropriate low concentration of GM-CSF, with removal of contaminating granulocytes by antibody  complement depletion or gradient separation. The cells are characterized as DEC205, MHC class II, CD80dim, CD86/dim, CD40dim, poor stimulators of naïve allogenic T cells in MLR, and endocytosishi. These ‘immature’ DCs induce alloantigen-specific hyporesponsiveness in allogeneic T cells (Lu et al., 1995a), and prolong heart allograft survival if given i.v. (2  106) 7 days before organ transplantation (Fu et al., 1996; Lutz et al., 2000). Both costimulatory molecule expression on the surface of these DCs and their allostimulatory function are upregulated, and phagocytic activity (a property of immature DCs) lost upon maturation following further culture in GM-CSF  IL-4.

Propagation of ‘tolerogenic’ DCs from mouse BM with GM-CSF  TGFβ1 TGFβ1 is a potent immunosuppressive cytokine (Derynck, 1994) that has been reported to affect the proliferation and differentiation of hematopoietic progenitors,

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including DCs. TGFβ has been strongly implicated as a key microenvironmental factor, acting on APCs, in the generation of immune deviation and immunological privilege, such as occurs in the anterior chamber of the eye (Streilein et al., 1992; Wilbanks and Streilein, 1992).Yasunori et al. (1997) showed that TGFβ1 almost totally blocked mouse DC maturation in vitro. This effect was thought to be mediated via Fc-receptor (CD32)-bearing suppressor cells, presumably macrophages. However, TGFβ1 did not inhibit the production of immature, MHC class II, CD86 DCs, and also did not suppress the differentiation of these cells. Strobl et al. (1996) reported that TGFβ1 promoted GM-CSF, TNFα and c-kit ligand-induced DC development from CD34 hematopoietic progenitors in human cord blood. The human cord blood-derived DCs that developed in the presence of additional TGFβ1 were CD1a, and did not have strong allo-MLR stimulatory activity. Their surface phenotype and biological function were similar to that of freshly isolated epidermal LCs (immature DCs). We have used TGFβ1 in conjunction with GM-CSF to facilitate the generation of mouse BM-derived DCs deficient in costimulatory molecule expression and with potential tolerogenic activity. Compared with propagation of DCs in low concentrations of GM-CSF, relatively high yields and purity of potentially tolerogenic cells can be obtained from normal mouse BM using GM-CSF plus TGFβ1. The blocking effect of TGFβ on DC maturation can, however, be reversed either by adding TNFα or IL-4, or by removing inhibitory cells (CD32 adherent cells). DCs propagated in the presence of GM-CSF  TGFβ1 can significantly prolong survival of vascularized heart allografts from the same donor strain (Lu et al., 1997b), if given 7 days before organ (cardiac) transplantation. There is also evidence that TGFβ1 impairs the antigen presenting function of human BM-derived APCs (Bonham et al., 1996b).

Propagation of ‘tolerogenic’ DCs from mouse BM with GM-CSF  NFκBspecific ‘decoy’ oligodeoxynucleotides (ODNs) Much evidence suggests that NFκB activation is responsible for DC maturation and activation. NFκB activation can be inhibited by various pharmacological agents at each activation step, including glucocorticoid-dependent upregulation of IκB, CsA-dependent calcineurin inhibition, and deoxyspergualin (DSG)-dependent inhibition of nuclear translocation (Lee and Burckart, et al., 1998). The therapeutic use of these agents may be limited by weak NFκB inhibition, or by undesirable side-effects. A recently

FIGURE 42.1 (A) NFκB ODNs specifically bind NFκB extracted from DCs. Nuclear proteins extracted from DCs propagated from mouse BM with GM-CSF  IL-4 were assessed for the presence of NFκB by electrophoretic mobility shift assay (EMSA). Specificity of the NF-κB ODNs for NFκB binding was demonstrated by utilizing unlabeled NFκB ODN as a competitor to a consensus NFκB probe. The addition of excess unlabeled NFκB ODN or consensus NFκB probe to the binding reaction resulted in the disappearance of the NFκB band. (B) Treatment of DCs in culture with NFκB ODN inhibits NFκB DNA-binding activity. Nuclear NFκB binding activity of DCs propagated from mouse BM with GM-CSF  IL-4 in the presence or absence of NFκB ODN decoys or control ODN2 was determined by EMSA. NFκB binding activity was detected in nuclear extracts of GM-CSF  IL-4 DCs. The addition of NFκB ODN to these cultured DCs completely inhibited NFκB binding. Addition of control ODN2 had no effect.

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599

104

50 0

101

102

103

104

MHC class II

0

MHC class I

100

101

102

103

104

100

100

101

50

103

CD 86

102

103

104

CD 11c

0

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0 100

CD 80

50

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50

CD 40 NF-κB ODN PBS

0

isotype

0

50

A

101

102

103

104

100

101

102

103

104

B

3

[H]TdR incorporation (CPM  103)

C3H Spleen

B10 Spleen

control ODN2 DC

IL-4 DC

control ODN1 DC

FIGURE 42.3 Prolongation of B10 (H2b) vascularized cardiac allograft survival in C3H (H2k) recipients by NFκB ODN-treated donor DCs. C3H (H2k) recipients were given 2  106 unmodified or NFκB ODN- or control ODN-treated DCs propagated from B10 mice i.v. 7 days before B10 heterotopic cardiac transplantation. NFκB ODN DC treatment markedly prolonged heart graft survival.

NF-κB ODN DC

30 25 20 15 10 5 0

0.5 1 3 5 10 20 50 100200 Stimulator cell number (103)

FIGURE 42.2 (A) Inhibition of costimulatory molecule expression on NFκB ODN DCs. DCs were propagated from B10 (H-2b, IAb ) BM in GM-CSF  IL4, and in the presence of NFκB ODN or control ODN1. CD80, CD86, but not CD11c, or MHC class I or II expression was significantly impaired in NFκB ODN DCs (shaded profiles) compared with untreated DCs (open profiles), or control ODN1 DCs (data not shown). (B) Markedly inhibited allostimulatory function of NFκB ODN DCs. Irradiated B10 BM DCs propagated with GM-CSF  IL-4 in the presence of NFκB ODN or control ODN1, 2 were used as stimulators and C3H (H-2k, IEk) spleen T cells as responders. Cultures were maintained for 72 hours. 3[H]TdR was added 18 hours before harvesting. Reproduced with permission. Copyright of The American Society of Gene Therapy, 2000.

developed immunosuppressive strategy exploits the natural binding sites of NFκB to create decoy ODN DNA molecules. Short, double-stranded, phosphorothioate-modified ODNs containing consensus NFκB binding sites, were able to inhibit NFκB transactivation of HIV LTR-based

reporter genes in vitro, and significantly reduced the degree of inflammation in arthritic rat joints (Miagkov et al., 1998). We have demonstrated that NFκB ODNs are incorporated efficiently by BM DCs, and specifically inhibit NFκBdependent transcription of a reporter gene (Giannoukakis et al., 2000) (Figure 42.1). Moreover, DCs propagated in GM-CSF and NFκB ODNs selectively suppressed costimulatory molecule expression without interfering with MHC class I or II expression (Figure 42.2). DCs propagated with GM-CSF and NFκB ODNs induced allogeneic donor-specific T-cell hyporesponsiveness in mixed leukocyte cultures, associated with inhibition of TH1-type cytokine production. Furthermore, administration of NFκB ODN-modified donor-type DCs resulted in significant prolongation of organ allograft survival in the absence of immunosuppression (Figure 42.3). Our results suggest that interference with NFκB and other transcriptional pathways involved in immune stimulation by DCs using ODN approaches could be a novel means to promote tolerance induction in organ transplantation.

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THERAPEUTIC APPLICATION OF DCs IN EXPERIMENTAL ORGAN TRANSPLANTATION Immature donor DCs prolong allograft survival Significant prolongation of mouse heart allograft survival can be achieved by pretreatment of recipients with immature donor-derived DCs propagated from mouse BM, using either GMCSF (Fu et al., 1996; Lutz et al., 2000) or GM-CSF  TGFβ1 (TGFβ-DCs) (Lu et al., 1997b). One to 8  106 immature donor (B10; H2b) DCs were injected i.v. into C3H (H2k) recipients, 7 days before B10 heart transplantation. DCs propagated using GM-CSF  IL-4 (mature DCs; IL-4 -DCs) accelerated rejection of subsequent heart grafts from B10 donors. Median graft survival time was reduced strikingly from 10 to 5 days in the IL-4 DC pretreatment group. By contrast, TGFβ-DCs (2  106 i.v., 7 days before organ transplantation) significantly prolonged median heart graft survival time (from 10 to 26 days). This therapeutic effect was correlated with suppression of donor antigen-induced T-cell proliferation and antidonor target cell CTL responses. TGFβ-DC of the donor strain were unable to prolong BALB/c (third party) heart graft survival, indicating that the effect of the TGFβ-DC was antigen specific. The inhibition of cardiac allograft rejection by TGFβ-DC was also dependent on the timing and the amount of antigen delivered by DC administration.

Blockade of costimulation potentiates the therapeutic effect of donor DCs Although costimulatory molecule-deficient donor-derived DCs can prolong allograft survival, most published observations to date indicate that they do not induce permanent allograft acceptance in nonimmunosuppressed recipients. Failure of these cells to induce allograft tolerance may be due to the ‘late’ upregulation of cost-imulatorymoleculesonthesedonorDCsfollowing their interaction with host T cells. This hypothesis is supported by the observation that

in vitro exposure of B10 TGFβ-DC to C3H spleen T cells for 18 hours induces CD40 and B7 molecule expression on the DC (Lu et al., 1999a). To achieve long-term graft acceptance, it appears important to prevent ‘late’ upregulation of costimulatory molecules, or to block upregulated costimulatory signal expression with agents such as CTLA4-Ig. The CD40/CD40L (gp 39) pathway has been shown to play an important role in DC activation, and is crucial for the establishment of primary and secondary responses to T-dependent antigens (Armitage et al., 1992; Yang and Wilson 1996). Thus, CD40 engagement upregulates the expression of CD80 and CD86 on DCs, providing a link between the CD40/CD40L and CD80: CD86/CD28:CTLA4 pathways (Caux et al., 1994b; McLellan et al., 1996). Ligation of CD40 on DCs also induces production/secretion of IL-12, an important TH1 cell-activating cytokine. Combination of allogeneic immature DC injection with blockade of the CD40/CD40L pathway using anti-CD40L mAb, 7 days before cardiac transplantation, results in long-term graft survival, with indefinite (>100 days) graft acceptance in 40% of recipients (Lu et al., 1999a). A single injection of anti-CD40L mAb alone failed to prolong heart graft survival. This therapeutic effect is not confined to use of DCs as the donor cell population. Thus, Parker et al. (1995) have shown in a different model, that anti-CD40L mAb combined with donor small lymphocytes blocks rejection of pancreatic islet allografts. Moreover, Hancock et al. (1996) found that treatment of recipients with donor spleen cells and anti-CD40L mAb led to indefinite survival of mouse cardiac allografts, although repeated administration of the Ab was required. The mechanism(s) by which administration of anti-CD40L mAb together with either costimulator-deficient donor DCs (Lu et al., 1997b) or resting B cells (Niimi et al., 1998) induces long-term graft survival is not clear. It was noted in the resting B-cell studies, that longterm graft survival was associated with striking inhibition of intragraft TH1 cytokine and IL-12 expression, and with reciprocal upregulation of TH2 cytokines. Combined immature donor DC and anti-CD40L mAb therapy markedly inhib-

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ited CTL responses in both graft-infiltrating cells and recipient spleen cells. More significantly, increased apoptosis of graft-infiltrating cells and spleen cells (Plate 42.4) is associated with comparatively high levels of donor cell microchimerism found in the combined treatment group, compared with untreated animals, or those given either immature donor DC or mAb treatment alone (Lu et al., 1999a). In conclusion, the manner in which antigen is presented by DCs plays an important role in the regulation of allogeneic DC–T cell interactions in vivo. The critical role of CD40 expression on donor DCs, and of CD40L in T-cell activation, provides a rational basis for blockade of this pathway to augment the tolerogenic potential of donor costimulator-deficient DCs in the longterm suppression of organ allograft rejection.

Immunosuppressive drugs inhibit DC maturation, and may promote DC tolerogenicity in vivo Evidence has accumulated that various immuno-suppressive drugs, including corticosteroids, CsA and DSG, can inhibit DC maturation, either in vitro or in vivo. Corticosteroids inhibit the maturation of immature, human monocyte-derived DCs, as assessed by lack of expression of CD83, and prevention of loss of antigen-uptake capacity. These corticosteroidinduced DCs (termed ‘tolerogenic DC3’) that express low levels of costimulatory molecules, produce reduced levels of IL-12p 70, and induce the development of TH2 cells (De Jong et al., 1999). Studies by Lee et al. (1999) have shown that CsA also inhibits (murine) DC maturation in vitro, and that this effect is associated with inhibition of the nuclear translocation of the gene regulatory protein NFκB, that has been shown to play a significant role in DC maturation. Recent findings of Thomas et al. (1999) suggest that DSG inhibits the maturation of rhesus monkey DC, both in vitro and in vivo. Thus, administration of DSG to anti-CD3 immunotoxintreated recipients of renal allografts inhibited DC maturation, i.e. expression of CD83, CD86 and rel B (an NFκB family member) within lym-

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phoid tissue, in association with the induction of transplant tolerance. We have also reported that exposure of BM DCs to NFκB-specific ‘decoy’ ODNs potentiates the tolerogenicity of DC in vitro and in vivo (Figures 42.2 and 42.3). Taken together, these findings indicate that one mechanism underlying the immunosuppressive/ tolerogenic properties of diversely acting immunosuppressive drugs may be inhibition of DC maturation.

Other forms of combination immunosuppressive therapy using DCs In addition to costimulation blockade or the use of immunosuppressive drugs, other approaches to tolerance induction using donor DCs in experimental organ transplantation have been explored. These include a combination of DCs with anti-T-cell mAbs, or other immunosuppressive agents. Administration of anti-CD4 mAb potentiates the capacity of immature donor DCs to prolong heart allograft survival in mice (Gao et al., 1999), whereas anti-CD4  antiCD8 mAb enhances the ability of donor DCs to prolong mouse skin graft survival. In the rat, total lymphoid irradiation  antithymocyte globulin in combination with immature donor DCs facilitate the long-term acceptance of heart allografts. Strategies using DCs to prolong allograft survival are summarized in Table 42.3. These include a recent report of the use of allopeptide-pulsed host DCs to induce donorspecific tolerance following their intrathymic injection in antilymphocyte serum (ALS)-treated, cardiac-allografted rats (Garrovillo et al., 1999).

THERAPEUTIC APPLICATION OF DCs IN EXPERIMENTAL AUTOIMMUNE DISEASE Evidence that DCs can be used or manipulated to exhibit ‘tolerogenic’ effects in autoimmune disease has come predominantly from studies in experimental models of type-1 diabetes or multiple sclerosis. Clare-Salzler et al. (1992) showed that pancreatic lymph node DCs from diabetic

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TABLE 42.3

Prolongation of allograft survival by DCs

Treatment

Graft

Refernce

Costimulator-deficient DCs Immature DCs

Mouse cardiac Mouse cardiac

Liver-derived DC progenitors CD40-deficient DCs NFκB specific decoy-treated DCs Allopeptide-pulsed host DCs CD8αDCs

Mouse pancreatic islet Mouse cardiac Mouse cardiac Rat cardiac Mouse cardiac

Fu et al., 1996 Suri et al., 1998; Lutz et al., 2000 Rastellini et al., 1995 Gao et al., 1999 Giannoukakis et al., 2000 Garrovillo et al., 1999 O’Connell et al., 2001

Mouse cardiac Mouse skin Mouse cardiac Rat cardiac Mouse skin

Lu et al., 1997b, 1999 Markees et al., 1999 Gao et al., 1999 Hayamizu et al., 1998 Gozzo et al., 1999

DCs alonea

DCs in combination with: Anti-CD40L mAb Anti-CD4 mAb TLIATG Anti-CD4  CD8 mAbs a

Donor-derived DCs unless specified. TLI, total lymphoid irradiation; ATG, antilymphocyte globulin.

NOD mice could inhibit the development of disease when adoptively transferred to prediabetic NOD recipients. More recently, Feili-Hariri et al. (1999) have reported that BM-derived DCs from NOD mice pulsed with pancreatic islet antigenderived peptides and injected into prediabetic NOD recipients could prevent diabetes development. Moreover, it appeared on the basis of IgG1 Ab responses in the DC-treated mice, that a TH2 response was generated, suggesting that the balancebetweenregulatoryTH2andeffectorTH1cells may have been altered. It has also been reported that IFNγ-stimulated NOD mouse splenic DCs exert antidiabetogenic effects (Shinomiya et al., 1999). Adoptive transfer of autoAg (MBP)-pulsed thymic DCs or splenic DCs (treated with CTLA4-Ig) prevents the development of EAE in the rat (Khoury et al., 1995; Khoury et al., 1996), also associated with a TH2 switch.

GENETIC ENGINEERING OF TOLEROGENIC DCs The foregoing information has shown that the antigen-presenting function of DCs is impaired

and their potential tolerogenicity enhanced, either by their incomplete maturation, the influence of anti-inflammatory cytokines (IL-10, viral IL-10, or TGFβ), blockade of surface costimulatory molecules (CTLA4-Ig or anti-CD40L mAb) or immunosuppressive drugs. Since DC maturation is linked to nuclear translocation of NFκB, antagonism of this gene transcription regulatory protein by specific antagonist molecules may also enhance DC tolerogenicity. In addition, the expression by DCs of key molecules associated with inhibition of T-cell growth (NO), or with induction of T-cell apoptosis (FasL, or other death-inducing ligands) suggests that transfer of genes encoding one or more of these molecules (summarized in Table 42.4) either alone or in combination, may render DCs tolerogenic in models of transplantation or autoimmune disease. DCs can be genetically modified using retroviral or adenoviral vectors (Song et al., 1997; Specht et al., 1997) or transfected with particulate/plasmid-based vectors that take advantage of the highly phagocytic nature of immature DC/DC progenitors. Thus, Song et al. (1997), Specht et al. (1997) and Wan et al. (1997) have

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TABLE 42.4

603

Candidate genes as modulators of DC tolerogenicity

Gene

Possible effect

vIL-10

Inhibition of expression of CD80, CD86 and MHC II; suppression of IL-12; induction of TH2 response Inhibition of T cell proliferation Neutralizes IFNγ Competitive intracellular binder of IL-12-p35 Apoptosis-inducing activity for T cells Blockade of B7 costimulatory signal Blockade of CD40–CD40L pathway Apoptosis inducing activity for T cells Apoptosis inducing activity for T cells Inhibition of NFκB nuclear translocation

TGFβ IFN-γdR EBI-3 INOS CTLA4-Ig CD40-Ig Fas ligand TRAIL (TNF family) IκB

INOS, inducible nitric oxide synthase; IFN-γdR, defective IFN-γ receptor.

demonstrated that transfer of genes encoding model antigens renders mouse BM-derived DCs capable of inducing protective and/or therapeutic antitumor immunity. A variety of techniques have been employed to engineer DCs genetically to express immunosuppressive molecules. These include vIL-10 (Takayama et al., 1998, 1999), TGFβ (Lee et al., 1998), CTLA4-Ig (Lu et al., 1999b) and FasL (Matsue et al., 1999; Min et al., 2000) (Table 42.5). DC progenitors from mouse BM transfected with vIL-10-encoding plasmids via gene gun delivery synthesize vIL-10 and exhibit substantially reduced capacity for the induction of allogeneic T-cell proliferation (W. Storkus and T. Tueting, personal communication). vIL-10 production by DCs is associated with significant reductions in cell surface expression of MHC class II and costimulatory molecules (CD80, CD86). We have also reported (Takayama et al., 1998) that retroviral transduction of mouse BMderived myeloid DCs to produce vIL-10 substantially impairs their allostimulatory activity. Moreover, it renders the DCs able to induce alloAgspecific T-cell hyporesponsiveness. Production of a ‘sorting’ retroviral vector, encoding both vIL10 and enhanced green fluorescent protein, permits selection (purification) of positive transfectants with substantially reduced T-cell stimulatory activity (potentially tolerogenic, vIL-10secreting DC) (Figure 42.5) (Takayamaetal.,1999).

Adenoviral delivery of CTLA4-Ig to myeloid DC (Plate 42.6) results in secretion of the transgene product, blockade of B7 molecule expression, and the capacity of the genetically modified DCs to induce alloAg-specific T-cell hyporesponsiveness (Lu et al., 1999b). These CTLA4Ig-transduced DCs traffic in vivo to T-cell

FIGURE 42.5 Genetic modification of DC with a ‘sorting’ vector encoding an immunosuppressive molecule. Flow-sorted murine myeloid DCs retrovirally transduced with a vector encoding vIL-10 and enhanced green fluorescent protein (‘sorted vIL-10 Tr DC’) show markedly reduced allostimulatory activity in (A) MLR and (B) CTL assays. Control gene (Zeo)- or vIL-10-EGFP-) transduced C57BL/10 (H2b) DCs were used as stimulators of naïve allogeneic C3H (H2k) T cells. Reproduced with permission. Copyright of Lippincott Williams & Wilkins, Inc., 1999.

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TABLE 42.5

Genetic manipulation of DCs to display immunosuppressive effects

Gene transferred

Delivery vector

Effects observed

Reference

vIL-10

Ecotropic retrovirus/ centrifugal transduction Ecotropic retrovirus encoding vIL-10 and EGFP

Alloantigen-specific T-cell hyporesponsiveness Highly selected transduced DCs markedly impaired T cell activity

Takayama et al., 1998

TGFβ

Adenovirus (E1/E3 deleted)

Impaired T cell function Enhanced in vivo survival

Lee et al., 1998

CTLA4-Ig

Electroporation (cDNA)

Prolongation of islet allograft survival Alloantigen-specific hyporesponsiveness; enhanced in vivo survival Alloantigen-specific hyporesponsiveness; inhibition of TH1/ augmentation of TH2

O’Rourke et al., 1999

Adenovirus (E1/E3 deleted)

Ecotropic retrovirus/ centrifugal transduction

Fas L (CD95L)

Particle-mediated cDNA PBK-CMV phagemid

In vivo antigen-specific immunosuppression Alloantigen-specific hyporesponsiveness; prolongation of heart allograft survival

Takayama et al., 1999

Lu et al., 1999

Takayama et al., 2000

Matsue et al., 1999 Min et al., 2000

EGFP, enhanced green fluorescent protein.

areas of recipient secondary lymphoid tissue, where they survive in enhanced numbers compared with control gene-transduced DCs (Plate 42.7). Interestingly, retrovirally transduced myeloid DCs expressing CTLA4-Ig show markedly inhibited ability to prime allogeneic T cells in vivo, and skew towards a predominant TH2-cell response (Takayama et al., 2000). Electroporation of cDNA encoding immunosuppressive molecules into a mouse DC line renders the cells capable of prolonging pancreatic islet allograft survival (O’Rourke et al., 1999). Particle-mediated delivery of cDNA encoding FasL into a mature murine DC line confers the capacity to induce antigen-specific immunosuppression in vivo (Matsue et al., 1999). In a recent study, Min et al. (2000) have reported that transfer of cDNA encoding human FasL to murine BM-derived DCs by lipofection renders the DCs capable of inducing apoptosis in Fas target T cells and promoting alloAg-specific hyporesponsiveness in vivo.

Significantly, repeated i.p. injection of these FasL-transduced DCs prolonged vascularized cardiac allograft survival in fully MHCmismatched recipients. Collectively, these studies indicate the potential of genetically engineered DCs for the regulation of undesired allo- or autoimmune responses.

ACKNOWLEDGMENTS Our work is supported by National Institutes of Health grants DK 49745 and AI 41011, and by grants from the Roche Organ Transplantation Research Foundation, and the Juvenile Diabetes Foundation International. We thank our many colleagues who have made significant contributions to these studies, and Ms. Shelly Lynn Conklin for typing the manuscript.

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Lu, L., Bonham, C.A., Chambers, F.D. et al. (1996). J. Immunol. 157, 3577–3585. Lu, L., Qian, S., Hershberger, P.A. et al. (1997a). J. Immunol. 158, 5676–5684. Lu, L., Li, W., Fu, F. et al. (1997b). Transplantation 64, 1808–1815. Lu, L., Li, W., Zhong, C. et al. (1999a). Transplantation 68, 747–757. Lu, L., Gambotto, A., Lee, W.-C., Bonham, C.A., Robbins, P.D. and Thomson, A.W. (1999b). Gene Ther. 6, 554–563. Ludewig, B., Graf, D., Gelderblom, H.R., Becker, Y., Kroczek, R.A., and Pauli, G. (1995). Eur. J. Immunol. 25, 1943–1950. Lutz,M.B., Assmann,C.U.,Girolomoni,G.andRicciardiCastagnoli,P.(1996).Eur.J.Immunol.26,586–594. Lutz, M.B., Suri, R.M., Niimi, M., Ogilvie, A.L.J., Kukutsch, N.A., Robner, S., Schuler, G. and Austyn, J.M. (2000). Eur. J. Immunol. 30, 1813–1822. Lynch, D.H., Ramsdell, F. and Alderson, M.R. (1995). Immunol. Today 16, 569–574. Markees, T.G., Philips, N.E., Gordon, E.J. et al. (1999). Transplant. Proc. 31, 884–885. Matsue, H., Matsue, K., Walters, M., Okumura, K., Yagita, H. and Takashima, A. (1999). Nat. Med. 5, 930–937. Matzinger, P. and Guerder, S. (1989). Nature 338, 74–76. McLellan, A.D., Sorg, R.V., Williams, L.A. and Hart, D.N.J. (1996). Eur. J. Immunol. 26, 1204–1210. Miagkov, A.V., Kovalenko, D.V., Brown, C.E. et al. (1998). Proc. Natl Acad. Sci. USA 95, 13859–13864. Min, W.P., Gorczynski, R., Huang, X.Y. et al. (2000). J. Immunol. 164, 161–167. Moore, K.W., O’Garra, A., Malefyt, D.E.W., Vieram, P. and Mosman, T.R. (1993). Annu. Rev. Immunol. 11, 165–190. Morelli, A.E., O’Connell, P.J., Khanna, A., Logar, A., Lu, L. and Thomson, A.W. (2000). Transplantation 69, 2647–2657. Morikawa, Y., Furotani, M., Kuribayashi, K., Marsuura, N. and Kakudo, K. (1992). Immunology 77, 81–87. Morikawa, Y., Furotani, M., Mastsuura, N. and Kakudo, K. (1993). Cell Immunol. 152, 200–210. Morikawa, Y., Tohya, K., Matsuura, N. and Kakudo, K. (1995). Immunology 85, 575–581. Nelson, D.J. and Holt, P.G. (1995). J. Immunol. 155, 3517–3524. Niimi, M., Pearson, T.C., Larsen, C.P et al. (1998). J. Immunol. 161, 5331–5337. O’Connell, P.J., Li, W., Takayama, T., Logar, A.J., Qian, S. and Thomson, A.W. (2001) Transplant Proc. 33, 94. Ohzato, H. and Monaco, A.P. (1992). Transplantation 54, 1090–1095. O’Rourke, R., Lower, J., Kang, S.M. et al. (1999).

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PLATE 42.4 (A) Pre-administration of donor immature DC  anti-CD40L mAb inhibits the antidonor CTL response, both within the graft and systemically. Cytotoxic activity of freshly isolated normal (C3H) heart nonparenchymal cells, heart graft-infiltrating cells (GIC), and recipient spleen cells against target cells (EL4) expressing donor (B10) alloantigen (H-2b) was determined by 4-hour 51Cr-release assay. Cytotoxicity was profoundly suppressed 14 days after heart transplantation. (B) A comparatively high incidence of apoptotic cells is observed in the heart GIC of mice pretreated with immature donor DC  anti-CD40L mAb. Apoptotic cells within B10 heart grafts of untreated C3H mice or mice pretreated (day7) with either immature B10 DC  anti-CD40L mAb, or mature B10 DC were detected by in situ TUNEL staining. Results were obtained on day 7 posttransplantaion, with the exception of the bottom right (day14). Reproduced with permission. Copyright of Lippincott Williams & Wilkins, Inc., 1999.

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PLATE 42.6 DCs are efficiently transduced with AdEGFP. (A) Flow cytometric analysis of mouse BM DCs propagated in GM-CSF  IL-4 and transduced with Ad-EGFP; 85% cells are EGFP. (B) Fluorescent AdEGFP-transduced cell exhibiting dendritic morphology (400 original magnification). (C) Ad-CTLA4Ig transduction markedly inhibits the allostimulatory function of DCs. BM DCs (B10) were transduced with either Ad-LacZ or CTLA4-Ig, then tested as stimulators of naïve allogeneic (C3H) T cells in 3-day MLRs. Reproduced with permission. Copyright of Stockton Press 1999.

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PLATE 42.7 (A–C) (Above) Ad-CTLA4Ig transduction of DCs does not affect their migratory activity and enhances their survival in unmodified allogeneic recipients. B10 BM DCs transduced with either Ad-LacZ or Ad-CTLA4-Ig were injected s.c. (2  105) into the hind footpad of C3H recipients. Seven days later, donor-derived cells were detected in the spleen by immunohistochemical staining for IAb. (Below) The incidences of donor-derived DCs (IAb) in spleens of recipients were determined by the mean number of positive cells per 100 low power fields (LPF). Reproduced with permission. Copyright of Stockton Press, 1999.

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43 Genetic engineering of dendritic cells Andrea Gambotto, Vito R. Cicinnati and Paul D. Robbins University of Pittsburgh School of Medicine, Pittsburgh, PA, USA

Fatti non foste a viver come bruti, ma per seguir virtute e canoscenza [You were not made to live as brutes, but to follow virtue and knowledge] Divina Commedia, Inferno Dante Alighieri

INTRODUCTION

GENE TRANSFER METHODS

The ability to manipulate dendritic cell (DC) function by gene transfer of cytokines, costimulatory molecules or antigens represents an attractive strategy to enable some DCbased therapies. Available vectors include retroviruses, adenoviruses, lentiviruses, adenoassociated virus, herpes simplex virus, cationic liposomes, naked DNA and DNA-coated gold beads (Table 43.1). Several of these methods have been shown to be effective in genetic engineering of DCs. Due to better in vitro culture methods based on expansion of DC progenitors with appropriate growth factors and cytokines, DCs generated from peripheral blood have been available in large numbers only during the last few years (Romani et al., 1994, 1996; Strunk et al., 1996; Thurner et al., 1999). This increase in the ability to generate large numbers of DCs in culture has resulted in more widespread application of DCs to treating disease, including the use of gene transfer strategies (Table 43.1) (Austyn, 1998; Giannoukakis et al., 1999a; Kirk and Mule, 2000). Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

Murine leukemia virus (MLV) Retroviruses are RNA viruses that replicate through a double-stranded DNA intermediate, integrating into the host DNA where they are able to express viral RNA for the life of the cell. Thus, retroviruses have the advantage of permanently modifying cells. The MLV genome encodes for three polyproteins, gag, pol and env, that are required in trans for viral replication and packaging. However, all that is required for viral replication in cis are the 5 and 3 longterminal repeats (LTRs) that contain promoter, polyadenylation and integration sequences, a packaging site termed psi, and a tRNA binding site, as well as several additional sequences involved in reverse transcription. The genes encoding these three viral proteins can be removed and heterologous genes and transcriptional regulatory sequences inserted. In order to produce infectious, replication-defective retroviral vectors, cell lines that express the three viral genes in trans have been generated, and are

609

Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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Gene delivery to Dendritic Cells

Viral-Vectors

Transferred Genes Cytokine

Retrovirus

TAA*

References Viral

Various

Modelantigen

Marker

IFN-γ IL10 IL-12

63 64, 65 66 67

EBV Influenza CTLA4Ig

EGFP EGFP GM-CSF IL-2 IL-4 IL-7 IL-10 IL-12 TGF-β

84–89 126, 127 92 92

MART-1 gp100 TRP2 MUC1 p53 α-FP Bcl-xL

95–97 98

EBV

99 94 94

B7 CTLA4Ig Lymphotactin CD40L Fas-L OX-2 PymT β-gal OVA EGFP GM-CSF

β-gal

140 128 137, 138, 139 129, 130, 138 135, 138 111, 112 115, 116 114, 117 118 119 120 121 136, 137 122, 123 131, 132 140 138 106 107 108, 109

93 141 142

MART-1 MAGE-A1 gp100 tyrosinase

146–148, 150 149 151 152 145 154

HIV EBV Vaccinia Virus

81 75, 76

83 77 78, 79 80, 82

β-gal OVA

Lentivirus

AdenoAssociated-Virus

Murine

63 MART-1

Adenovirus

Human

B7.1 ICAM-1 LFA-3

159 159 159 158 108, 144

β-gal OVA 1acZ AP IacZ EGFP

Herpes Simplex Virus

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143 153 160 160

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TABLE 43.1 (cont.) Viral-Vectors

Transferred Genes Cytokine

Liposomes

TAA*

References Viral

Various

Modelantigen

Marker

MART-1 HIV HSV β-gal CAT EGFP

Gene Gun

IL-12 IFNβ-

MART-1 gp100 tyrosinase Mage-1 Mage p53

Human

Murine

166 167 163

162

164 164 92, 167 165 165 165 165 165 178 178 177 181 183 182 180 175 176 184 185

HPV Influenza Naked DNA

GM-CSF IL-4 gp100 p53 HSV Influenza FasL

Electroporation CaPO4 mRNA

Luciferase EGFP EGFP CEA PSA TERT

92, 168, 169 92 170 173 183

*  Tumor-associated antigen

termed packaging cells. The vector, in the proviral form of double-stranded DNA, can then be introduced into the packaging cells either transiently or stably by transfection, resulting in the production of infectious virus. Following infection of a target cell and integration of the vector into the host DNA, the virus is unable to replicate due to the absence of the viral transacting proteins (Miller, 1992; Coffin, 1996). The majority of gene therapy trials to date have utilized the extensively characterized murine retroviral vectors based on MLV since it has no homology with human retroviruses and causes no direct pathology in mice. Retroviral vectors have the additional advantage of stably integrating into the host DNA in the infected cell and expressing the transgene for the life of that cell. Packaging lines that can produce moderate

titers of the replication-defective vectors have been developed that do not give rise to replication-competent helper virus through recombination (Kim et al., 1998a). The dis-advantage of the MLV-based retroviral vectors is that they require cell division, in particular mitosis, for integration (Miller et al., 1990). Thus, the current retroviral vectors are better suited for ex vivo gene therapy where isolated cells can be propagated in culture and modified by retroviral infection prior to transplantation into the recipient (Gambotto et al., 2000).

Lentiviruses Lentiviruses are a class of retroviruses that includes human immunodeficiency virus (HIV), simian immunodeficiency virus (SIV ) and

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equine infectious anemia virus (EIAV) that are able to infect nondividing cells (Haase, 1986). Recently, vectors based on these lentiviruses have been developed that are able to infect nondividing cell types including β-cells from the pancreatic islets as well as neurons (Naldini et al., 1996; Glannoukakis et al., 1999b). The further development of lentiviral vectors including methods for producing the vectors should allow for their general application to in vivo gene therapy (Akkina et al., 1996).

Adenoviral vectors Adenoviruses are double-stranded DNA viruses that cause respiratory and eye infections in a number of mammalian species, incuding humans. Although human adenovirus serotype 5 is the best characterized and used predominantly as a vector for gene transfer, vectors based on other serotypes as well as other species of adenoviruses are also being developed (Prevec et al., 1989; Bautista et al., 1991). To date, 51 human Ad serotypes, Ad1 to Ad51, have been reported and subdivided into six subspecies, namely A to F, based mainly on similarities in DNA structure, replication and transcriptional organization, rather than on similar DNA sequences (De Jong et al., 1999). Adenoviruses contain four early genes that encode multiple polypeptides important for the virus life cycle. The proteins encoded by the E1 region, the first gene expressed following infection, are essential to activate transcription of the E2, E3 and E4 genes that encode for additional regulatory proteins as well as polypeptides important for viral replication. The major late promoter (MLP) is activated late in infection, resulting in expression of proteins important for packaging of the viral DNA (Graham and Prevec, 1992). In order to generate a replication-defective virus from Ad-5, the E1 gene is deleted from the viral genome and an expression cassette carrying the gene of interest inserted in place of E1. The ∆E1 virus is unable to replicate unless the E1 polypeptides E1A and E1B, are provided in trans. The virus is then replicated in the human embryonic kidney cell line 293 that stably

expresses E1A and E1B. The ∆E1 virus is replication defective since the necessary E1 proteins are not provided in trans. Adenoviral vectors can be grown to higher titer and are able to infect a wide variety of both dividing and nondividing cell types. However, the episomal viral DNA that is unable to replicate in the infected cell is eventually degraded, resulting in loss of gene expression. In addition, the ‘first-generation’ ∆E1 vectors still express a low level of viral proteins, resulting in continued stimulation of an immune response to the virally infected cell in vivo (Graham and Prevec, 1992; Graham, 2000).

Herpes simplex virus vectors Herpes simplex viruses (HSVs) are large linear DNA viruses of approximately 150 kb that can either infect cells lytically or latently, depending upon the cell type (Roizman and Sears, 1996). One advantage of HSV vectors is that they are able to incorporate exogenous DNA of up to 35 kilobases, allowing for insertion of large fragments of DNA. The normal life cycle of HSV following uptake of the virus into cells involves expression of a set of immediate–early genes including ICP0, ICP4, ICP22, ICP27 and ICP47. In order to generate a vector based on HSV, the immediate–early genes are important for subsequent activation of the early and late genes which are deleted or inactivated. The resulting defective virus can be grown on complementing cell lines expressing the appropriate immediate–early genes, similar to adenoviral vectors. The first HSV vector that was developed had a deletion in the ICP4 gene (DeLuca et al., 1985), but was still able to express the other immediate–early gene products that are toxic to certain cell types. Recently, further deletion of ICP0, ICP22 and ICP27 has resulted in a virus that is nontoxic and can express a transgene for extended periods in culture and in vivo (Marconi et al., 1996). Alternatively, it is possible to package plasmid vectors, termed amplicons, using defective HSV helper virus or plasmids encoding helper functions. To generate helperfree amplicons, a plasmid carrying the HSV origin of replication and packaging signals is co-

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transfected with a series of plasmids that contain all of the individual HSV genes missing only the packaging sequence (Glorioso et al., 1995).

induced by DNA-damaging agents, incuding UV light (Yakobson et al., 1989; Russell et al., 1995).

Adeno-associated virus

Vaccinia virus vectors

Adeno-associated virus (AAV) is a member of the parvovirus family of single-stranded small DNA viruses that require a helper virus such as adenovirus or herpes simplex virus for replication (Siegl et al., 1985). The virus contains two genes, rep and cap, that encode polypeptides important for viral replication and packaging. Similar to retroviruses, rep and cap can be supplied in trans with only the inverted terminal repeat (ITR) sequences required in cis for the virus life cycle (Berns and Bohenzyky, 1987; Berns and Giraud, 1996). Thus, therapeutic genes can be inserted between the ITRs and infectious virus generated by co-transfection of the 293 cells with the vector and a Rep and Cap expression plasmid followed by infection with a first-generation adenoviral vector (Fan and Dong, 1997; Li et al., 1997). The resulting virus can be purified away from the contaminating adenovirus by centrifugation. However, to avoid possible adenoviral contamination, infectious recombinant AAV can also be generated by cotransfection of a plasmid expressing the complementing adenoviral genes (Muzyczka, 1992; Kotin, 1994). Following infection with wild-type AAV, the virus forms a doublestranded DNA intermediate that is able to integrate site specifically into a region of chromosome 19 through a rep-dependent mechanism (Kotin et al., 1992). However, recombinant AAV vectors integrate randomly and may actually persist as episomal DNA in nondividing cells (Flotte et al., 1994; Kearns et al., 1996). One of the rate-limiting steps in the virus life cycle is the rate of second-strand synthesis, necessary for expression of the transgene (Ferrari et al., 1996). Normally, adenoviral gene products are able to stimulate the rate of secondstrand synthesis, but in the absence of adenoviral helper functions, AAV requires the presence of cellular factors. These unknown factors are either high in certain cell types or can be

Recombinant vaccinia virus vectors are members of the Poxviridae family of viruses. These viruses replicate and transcribe their genome in the cytosal of infected cells. Recominant of viral DNA into the host’s genome is thus not of concern. Vaccinia virus infects most mammalian cell types (Moss, 1996) and is thus the vector of choice for transient gene expression. Its substantial genome can be replaced, thereby allowing insertion of large DNA fragments up to 25 kb in size (Smith and Moss, 1983). The infectious cycle of this virus is divided into three phases. Early-phase genes encode enzyme proteins, whereas intermediate-phase genes drive the expression of the late-phase genes encoding structural proteins (Baldick et al., 1992). The 7.5 kDa protein encoding the promoter gene is active during early and late phases of infection and can be used to drive expression of the inserted gene of interest (Taylor and Paoletti, 1988; Merchlinsky and Moss, 1992). Wild-type vaccinia virus is cytolytic and less virulent poxvirus vectors, such as modified vaccinia ankara (MVA) or fowlpox virus, which do not repicate completely in mammalian cells, are used mostly for transduction of cells (Sutter and Moss, 1992; Somogyi et al., 1993; Carroll and Moss, 1997). Inhibition of vaccinia replication can also be achieved by psoralen or UV-light treatment, prior to use for transduction (Paoletti, 1996).

Nonviral vectors Although nonviral gene transfer is less efficient than virus-mediated DNA delivery, it has several advantages. It is considerably cheaper to produce large amounts of DNA than certified pathogenfree viruses; there is no possibiity of recombination giving rise to replication-competent virus; and the plasmid-based delivery systems express no immunogenic proteins. For these reasons, many laboratories have focused on improving

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nonviral gene delivery methods, particularly for vaccination strategies. The mechanism by which DNA vaccination is faciliated consists of a combination of at least three distinct means: (1) antigen synthesis and presentation by myocytes (Wolff et al., 1992); (2) direct plasmid transfection of APCs with subsequent antigen presentation (Casares et al., 1997; Torres et al., 1997); and (3) antigen synthesis by myocytes and transfer to APCs for presentation (Ertl and Xiang, 1996; Fu et al., 1997). Although uptake of DNA by cells occurs with modest efficiency, complexing of the DNA to various chemical formulations increases DNA uptake and subsequent gene expression (Perales et al., 1994). In particular, condensation of the DNA using different polycations is able to improve DNA uptake and prevent degradation (Felgner and Ringold, 1989). Polycationic lipids, such as cationic liposomes, mixed with plasmid DNA are thought to fuse with the cell membrane of targeted cells thereby improving gene delivery efficiency in vitro and in vivo (Felgner et al., 1995). It is also possible to couple the DNA to polypeptides to target DNA to specific cell surface proteins (Wu et al., 1991) or to coat the DNA on to gold beads for delivery using bioballistic strategies (Yang and Sun, 1995). Electroporation is another method used for delivery of plasmid DNA into the nucleus of cells. It takes advantage of the interaction of an appied electric field with the lipid dipoles of a pore within the cell membrane that induces and stabilizes the permeation sites and thus enhances cross-membrane transport of molecules (Neumann et al., 1982).

GENE TRANSFER TO DCs Retroviral vectors Human DCs The first viral vectors used for gene transfer to DCs were recominant replication-defective retroviral vectors based on the Moloney murine leukemia virus. As these retroviral vectors preferentially infect dividing cells, DCs or DC

precursors that are actively dividing are necessary for targeting viral infection. Since CD34 DC precursors derived from bone marrow, cord blood or peripheral blood cells proliferate actively (in contrast to terminally differentiated cultured DCs) they have been used as targets for retrovirus-mediated transduction. Bregni et al. (1992) reported for the first time the successful retroviral transfer of the neomycin resistance gene into human peripheral blood hematopoietic progenitors, whereas Cassel et al. (1993) transduced the neomyicin resistance gene into a human CD34-enriched precursor stem cell population from bone marrow and peripheral blood. Similarly, efficient retroviral transduction of sorted CD34+ cells from cord blood with the thymidine kinase (TK)-neo gene has been observed (Lu et al., 1993). After transduction, CD34+ cells can be easily expanded to mature DCs expressing the gene of interest. Based on this concept, DCs were engineered to secrete functional IFNγ and L-12 by retroviral infection, resulting in genetically modified DCs inducing more highly TH1-biasing T-cell proliferation (Ahuja et al., 1998). Reeves et al. (1996) induced antimelanoma CTLs in vitro with DCs derived from CD34+ cells transduced (22–28%) with the melanoma tumor-associated antigen (TAA) MART-1 gene. Similarly, Scabolcs et al. (1997) achieved transduction rates of between 11.5 and 21.5% and showed that retrovirally transduced DCs were phenotpically unmodified and potent T-cell stimulators. There are also several reports of successfully inducing antigen-specific CTLs from healthy donors after in vitro stimulation with DCs transduced with EBV (LMP2 gene) and influenze virus genes, respectively (Rooney et al., 1998; Heemskerk et al., 1999). Retroviral transduction rates can be enhanced with multiple infection cycles. In addition, Elwood et al. (1996); Bello-Fernandez et al. (1997); Dao et al. (1997) and Movassagh et al. (1999) have all reported the beneficial use of Flt-3 ligand during differentiation of DCs in order to enhance the transduction rate. By using a semi-solid methylcellulose assay, Movassagh et al. (1999) were able to transduce more than 90% of CD34+-derived DCs. Finally, there have

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been several reports of successful transduction of peripheral blood monocyte-derived DCs (moDCs) with retroviral vectors (Aicher et al., 1997; Westerman et al., 1998), despite evidence that moDCs, in contrast to CD34-derived DCs, do not proliferate during differentiation into DCs (Ardesha et al., 2000). Murine DCs The inability of retroviral vectors to transduce nonproliferating cells, such as moDCs, does not represent a particular problem for the typical application of murine-derived DCs. Protocols for the generation of murine DCs usually include use of bone marrow-derived stem cells that have a high proliferation rate. Protocols for obtaining proliferating DCs have been developed that involve generation of bone marrowderived DCs in culture with GM-CSF and IL-4 for 7 days. Zitvogel et al. (1996) and Nishioka et al. (1999) reported the successful transduction of murine DCs with the two IL-12 subunit genes and showed that these genetically engineered DCs were able to induce CTL-mediated tumor resistance. Specht et al. (1997) co-cultured DCs with ecotropic retroviral producer cell lines and delivered the model antigen β-gal with a transduction rate of up to 72%. Moreover, they were able to induce β-gal-specific CTLs, indicating that the transduced retroviral gene product could be processed and presented within DCs. Similarly, it was demonstrated that resistance against tumors expressing the model antigen ovalbumin (OVA) could be generated by retroviral gene transfer of OVA to DCs and that induction of antigen-specific CTLs was dependent on CD4 T-cell help (De Veerman et al., 1999; Schnell et al., 2000). Gasperi et al. (1999) transduced immature DCs with a retroviral vector containing the reporter gene enhanced green fluorescent protein (EGFP) and were able to generate a highly homogeneous cell population after cell sorting. These transduced DCs were able to mature fully under LPS stimulation, as indicated by a highly mature phenotype and a strong allostimulatory function, and thus may be suitable for further functional studies.

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Takayama et al. (1998) showed that IL-10-transduced DCs have impaired allostimulatory capacity, leading to T-cell hyporesponsiveness. Using a repeated centrifugal method for transduction of the DC culture, they were able to achieve a retroviral transduction efficiency of 35–40%. In addition, they sorted DCs by means of co-delivery of the EGFP gene, yielding a transduced DC population of 95% purity (Takayama et al., 1999). The same group recently showed that the transduction of the fusion protein CTL antigen 4 immunoglobulin (CTLA4-Ig) into DCs induced alloantigen-specific T-cell hyporesponsiveness (Takayama et al., 2000). The transduction efficiency was 62% as defined by antigen expression by immunocyto or histochemistry. Taken together, these results demonstrate the ability to stably transduce both murine and human DCs, resulting in induction or subversion of specific immune responses.

Lentivirus vectors Lentiviral vectors are derived from the HIV virus with a particular tropism for lymphocytes. Although they belong to the family of retroviruses, they seem able to infect cells in the resting G0 cell cycle phase. Thus they appear suitable for gene delivery to nondividing human DCs. Li et al. (1998) were able to transduce CD34 cells with an env- and nef-deleted HIV1DeltaEN vector, pseudotyped with vesicular stomach virus G (VSV-G), yielding stable gene expression as measured by GFP reporter gene expression. Gruber et al. (2000) from the same group constructed a HIV-1DeltaEN V(3) vector deleted in the remaining accessory HIV genes (vif, vpr and vpu), which did not impair transduction efficiency of moDCs. Muthumani et al. (2000) developed a lentiviral vector containing the virion-associated protein (vpr), essentially for replication in monocytes and macrophages, fused with the marker gene EGFP and showed that it was capable of transducing various leukocytes including DCs. Chinnasamy et al. (2000) transduced moDCs, derived from both healthy volunteers and melanoma patients, with a VSV-G-pseudotyped lentiviral vector. The

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transduction efficiency, as shown by EGFP expression, ranged from 70 to 90%. Transduced DCs showed strong allostimulatory capacity and maintained the ability to respond to activation signals, such as CD40L and LPS. Mangeot et al. (2000) constructed a lentiviral vector derived from the simian immunodeficiency virus (sivMAC251) that was able to transduce CD34 cell-derived as well as moDCs efficiently with a transduction rate of 40%. Recently, Negrè et al. (2000) from the same group showed that CD40Linduced maturation of moDCs enhances the transduction rate, as shown by GFP expression. Schroers et al. (2000) transduced PBMC-derived DCs with a transduction efficiency of 25–30% that was decreased when mature DCs were used. In addition, they showed integration of HIV-1 into the host cell genome. Similarly, Curran et al. (2000) used a feline immunodeficiency virus (FIV) for stable gene delivery into DCs with high efficiency.

Adenoviral vectors Human DCs In contrast to retroviral vectors, Ad-based vectors are able to accept large DNA inserts, are not host specific and are able to infect nondividing, resting cells. These advantages led to the development of replication-incompetent Ad5 vector for gene transfer purposes, bypassing in this way the need to isolate C34 cells for the generation of DCs. Arthur et al. (1997) compared adenoviral gene transfer with nonviral methods, such as DNA/liposome complexes, electroporation and CaPO4 precipitation and found that the adenoviral transduction efficiency exceeded 95% and was up to 1000-fold higher than nonviral methods. Moreover, transduction by adenoviral vectors requires a high multiplicity of infection (MOI), because of the low level of adenoviral-specific receptors on the cell surface of DCs. Similarly, Dietz and VukPavlovic (1998) demonstrated expression of the marker gene EGFP in 90% of adenovirustransduced CD83 DCs. Sonderbye et al. (1998) showed the immunomodulatory potential of

Ad-transduced human umbilical cord DC precursors and mature blood DCs. We have demonstrated the efficiency of adenoviral gene transfer to DCs using three different fluorescent marker genes. As shown in Plate 43.1, human DCs can be infected efficiently with up to three different vectors simultaneously. The transgenic expression of the co-stimulatory molecule B7 in DCs enhanced MLR stimulatory capacity, whereas the transduction of CTLA4-Ig reduced MLR stimulation capacity of DCs. Butterfield et al. (1998), Perez-Diez et al. (1998) and Philip et al. (2000) showed that melanoma-specific CTLs can be generated by DCs adenovirally transduced with MART-1 genes, whereas Linette et al. (2000) have been able to induce CTL responses to gp100. Ranieri et al. (1999) were able to raise HLA-A*0201-restricted CTL responses against EBV antigens (LMP2B) in vitro following adenovirus-mediated gene transfer. Diebold et al. (1999a) transfected DCs with complexes consisting of adenovirus mannose polyethylenimine DNA. The gene delivery rate was enhanced using these transfection complexes since they are taken up via the highly expressed mannose receptor on DCs. Similarly, Tillman et al. (1999) were able to enhance adenoviral gene transfer into DCs by a CD40-targeted adenoviral vector and induce maturation of DCs. There is some question regarding the effect of infection of DCs with adenoviral vectors at high MOI. Zhong et al. (1999) reported that adenoviral vectors are nonperturbing genetic vectors for human DCs, whereas Jonuleit et al. (2000) found that the MLR stimulation capacity of DCs was suppressed in a dose-dependent manner following transduction. Rea et al. (1999) and Rouard et al. (2000), on the other hand, showed that infection with adenovirus human DCs, rendering them fully mature with high expression of CD40, CD80, CD83 and CD86 thereby enhancing their allostimulatory potential. The differences in results between these groups may reflect the types of DCs used as well as the purity of the adenoviruses. It is likely that infection itself or the delivery of large amounts of viral proteins to DCs in the form of viral particles will result in DC maturation.

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Murine DC Adenovirus-mediated gene delivery into DCs has been studied extensively in murine models since 1997 and consists mainly of transfer of model- and tumor-associated antigens (TAA) as well as of cytokines and other immunomodulatory molecules. Wan et al. (1997) and Song et al. (1997) transduced DCs with genes encoding the model TAA poyoma middle T and beta-galactosidase (β-gal) genes, respectively, resulting in strong CTL-mediated immunity. Moreover, they showed that direct intravenous injection of adenovirus-transduced DCs did not show virally mediated side-effects such as hepatic toxicity, in contrast to immunizations performed with direct injection of recombinant adenovirus. In this context, Brossart et al. (1997), and more recently, Wan et al. (1999a), have reported that DCs transduced with adenovirus can be administered repeatedly, resulting in only a low-level induction of anti-Ad antibodies. Preexisting high titers of anti-Ad neutralizing Abs does not negatively affect DC-mediated antigen delivery and immunization. Moreover, Brossart et al. (1997) showed the rejection of tumor cells, expressing the model antigen OVA, with adenovirus-infected DCs but not with adenovirus alone in immunized animals. The strong induction of antitumor CTLs by DCs adeno-virally transduced with TAA is related to predominant T helper 1 cytokine production, that favors a cellular-mediated immune response, as shown by Inoue et al. (1999). A large number of TAAs, derived from melanoma, have been examined in adenovirally transduced DC-mediated immunization studies. Ribas et al. (1997) stably transfected a syngeneic murine fibrosarcoma line with the melanoma antigen MART-1/Melan-A and showed partial and complete protection with i.v. administration of MART-1-transduced DCs (Ribas et al., 1999). Depending on the mouse strain, multiple immunizations with DCs expressing the TAA showed decreased protection that seemed to be Fas mediated (Ribas et al., 2000). Kaplan et al. (1999) showed both protection and inhibition of tumor growth in tumorbearing mice after immunization with gp100 and

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tyrosinase-related protein 2 (TRP2)-transduced DCs. The antitumor activity was dependent on CD4+ T-cell help and improved by the simultaneous immunization of the two antigens. The importance of CD4+ T-cell help has been also reported with DCs transduced with human gp100, where antitumor immunity was induced dependent on the direct priming of naïve CD4+ T cells (Wan et al., 1999b). Yang et al. (2000a) demonstrated in a transgenic murine model that DCs transduced with human gp100 process the same A2.1-restricted peptide epitopes as human DCs. Similarly, mice immunized with TRP2transduced DCs were capable of inducing protection against mouse melanoma-induced lung metastasis (Tüting et al., 1999) and the transduced DCs showed 30–60% TAA expression as determined by cytoplasmic coexpression of EGFP. The injection of MUC1-transduced DCs was also shown to be capable of inhibiting growth of MUC1-expressing tumors (Gong et al., 1997). Ishida et al. (1999) genetically engineered DCs to express human p53, a tumor antigen, resulting in a significant reduction in growth of established tumors. Finally, the adenoviral gene transfer of alpha-fetoprotein showed inhibition of tumor growth in a murine hepatocellular tumor model by immunization with DCs genetically engineered to express alpha-fetoprotein (Vollmer et al., 1999). Another approach has involved the adenoviral gene transfer of the antiapoptotic gene Bcl-xL to DCs, increasing their surviral following in vivo delivery. This approach has resulted in significant inhibition of tumor growth in a murine prostate carcinoma (Pirtskhalaishvili et al., 2000). The usefulness of combining the potent antigen presentation capacities of DCs with cytokine-mediated antitumor activity has been demonstrated in several studies. Cao et al. (1998) studied the efficacy of tumor peptidepulsed DCs transduced with the lymphotactin gene, whereas Zhang et al. (1999) showed the efficacy of tumor RNA and lymphotactintransduced DCs. Fushimi et al. (2000) intratumorally injected adenovirus engineered to produce the human CC chemokine macrophage

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inflammatory protein 3 alpha (MIP-3α) that is chemotactic for DCs. They demonstrated that the antitumor response was CD8+ T-cell dependent, since in CD8-deficient mice this effect was abrogated. Similarly, Wang et al. (2000) showed that the adenovirus-mediated transgene expression of granulocyte–monocyte colonystimulating factor (GM-CSF) in the lung airways caused accumulation of DCs concomitantly with a high number of CD4 and CD8 cells and increased levels of IFNγ. Ozawa et al. (1999) delivered GM-CSF into bone marrow-derived DCs and Langerhans cells. This increased MHC class II and CD86 expression leading to enhanced antigen presentation capacity. Cao et al. (1999) fused tumor cells with GM-CSFtransduced DCs and induced a strong specific CD8 and CD4 T cell-mediated antitumor response that induced tumor protection and extended survival in tumor-bearing mice. Miller et al. (2000) administered IL-7-transduced DCs intratumorally in a lung cancer model and induced complete tumor regression. Various controls consisting of IL-7, AdIL-7 vector alone, IL-7-transduced fibroblasts, or unmodified DCs, were inferior in inducing tumor resistance, suggesting the important role of IL-7-DCs in the antitumor response. There have been two reports of potent therapeutic and long-lasting CD8 and CD4 mediated antitumor response in distinct tumor models by intratumor injection of DCs adenovirally transduced with IL-12 (Melero et al., 1999; Furumoto et al., 2000). In addition, Kikuchi et al. (2000a) modified DCs to express CD40L, resulting in sustained inhibition of tumor growth. Moreover, in a murine model, they could induce a humoral response against microbial agents in a CD4-independent manner (Kikuchi et al., 2000b). Several groups have examined the effects of adenoviral infection of DC maturation and have shown that DC infection with adeno-viral vectors results in DC maturation through a NFκB-dependent mechanism. In particular, Hirschowitz et al. (2000) and Morelli et al. (2000) reported that DCs transduced with adenoviral vectors show signs of activation, as demonstrated by upreguation of CD86, CD54, MHC class II, and increased production of IL-12 p40

protein in culture supernatants. Many investigators have observed modest therapeutic effects of injection of DCs infected with control vectors compared with uninfected controls. Thus, although adenoviral vectors appear to infect human and murine DCs efficiently, they may also have effects on DC function in vivo. In addition to stimulating an immune response by adenoviral transfer of genes encoding immunostimulatory cytokines and molecules, DCs can also be modified to inhibit the immune response. Lee et al. (1998) showed that transforming growth factor beta (TGFβ)transduced DCS exhibited very poor allostimulatory capacity with unmodified migration characteristics in vivo. Similarly, Lu et al. (1999a, 1999b) transduced the immunosuppressive CTLA4-Ig into DCs, resulting in blockade of CD80/CD86-mediated CD28 ligation on T cells. These CTLA4-Ig-expressing DCs had poor allostimulatory capacities and induced allogenspecific T-cell hyporesponsiveness. Gorcynski et al. (2000) studied the effect of transducing TGFβ, IL-10, IL-12 and the regulatory molecule OX-2 into DCs. They found that co-expression of IL-10 and TGFβ prolonged graft survival that was enhanced by additional delivery of OX-2. Genetically modified DCs have also been shown to be therapeutic in animal models of autoimmune disease. Ad-IL-10 transduced DCs, somewhat unsuspectedly, enhanced allostimulatory activity of DCs in mixed lymphocyte reactions, CTL assays, increased organ graft rejection and augmented antidonor alloantibody (Lee et al., 2000). Injection of DCs infected with an adenoviral vector expressing either IL-4 or FasL resulted in significant inhibition of established collagen-induced arthritis in the mouse (Kim et al., 2000; Kim and Robbins, unpubished).

Adeno-associated virus vectors (AAV) Only recently have recombinant AAV vectors been explored for genetic manipulation of DCs. AAVs have a broad host range and are capable of inducing persistent transgene expression. Liu et al. (2000) transduced monocytes with a AAV/GM-CSF/Neo vector and delayed the

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IL-4 supplementation for 2 days showing that transduced monocytes were able to proliferate. After addition of recombinant IL-4 the GM-CSFtransduced monocytes differentiated into DCS. GM-CSF gene integration into the host’s genome was demonstrated in a sorted CD83+ DC population. The transduction of the GM-CSF gene may substitute for exogenous GM-CSF in DC cultures. Infection of DCs by AAV may be enhanced by methods to increase second-strand synthesis such as low-dose irradiation. Zhang et al. (2000) were able to transduce murine DCs with the marker gene β-gal and showed that immature DCs in contrast to mature DCs were able to induce CD40L-dependent T-cell immunity in mice previously immunized i.m. with AAV-lacZ.

Vaccinia virus vectors Human DC Recombinant vaccinia virus vectors have a wide range of tropism and infect most mammalian cell types making them useful for transient gene expression. Brown et al. (1999) studied the transduction of moDCs with a host range-restricted avipoxvirus, fowlpoxvirus (FWPV), and showed over 80% expression of the marker gene lacZ. Moreover, they showed that FWPVs are able to infect both immature and mature cord blood CD34+ progenitor-derived DCs. Recently, the same group reported that immunization with fowlpox-transduced DCs induces a potent and long lasting transgene-specific class I-restricted T-cell immunity (Brown et al. 2000). Fan et al. (1997) transduced moDCs from HIV-1-infected patients with a recombinant vaccinia virus, expressing HIV antigens gag, pol and env and were able to induce high levels of MHC class Irestricted anti-HIV-1 memory CTLs. Kim et al. (1997, 1998b) and Zajiac et al. (1997) delivered melanoma-associated antigens MARTI1/Melan A into DCs that served for in vitro stimulation of autologous, antigen-specific CTLs. Similarly, Chaux et al. (1999) were able to identify HLAA*0201-restricted epitopes for MAGE-A1 with vaccinia virus encoding MAGE-A1-transduced DCs for in vitro stimulation of T cells. Bettinotti

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et al. (1998) showed in addition, that antigen processing and presentation is restricted to a unique allele/ligand combination. Yang et al. (2000b) generated CTIs induced by DCs transduced with multiple epitopes from the gp100 melanoma TAA, that efficiently lysed target cells expressing individual epitopes. Drexler et al. (1999) transduced DCs with tyrosinase gene, encoding a melanoma-specific differentiation antigen, incorporated into a modified vaccinia virus Ankara (MVA). They were able to induce antimelanoma CTL that could be augmented with repeated immunization, despite the fact that the patients had been vaccinated previously against smallpox. Di Nicola et al. (1998) compared adenoviral and vaccinia virus-mediated gene transfer into CD34+-derived DCs and showed that transduction with either viral vector resulted in efficient gene expression of the modified membrane-exposed alkaline phosphatase (AP) reporter gene. Because of sustained cytopathic effects of vaccinia infection on DCs, Subklewe et al. (1999) used psoralen and UVinactivated recombinant vaccinia virus encoding EBNA-3A for transduction of DCs. Nevertheless, they were able to achieve a strong anti-EBV CTL response in vitro. Several groups have studied the effects of vaccinia virus infection on DCs and found that the virus inhibits DC maturation as defined by low expression of co-stimulatory molecules and HLA molecules, as well as low T-cell activation capacity (Engelmayer et al., 1999; Drillien et al., 2000; Jenne et al., 2000), making it less suitable as a vehicle for immunization. Murine DCs Bronte et al. (1997) delivered the model TAA βgal into DCs capable of inducing prolonged survival in tumor-bearing mice. Brossart et al. (1997) compared distinct viral vectors (recombinant vaccinia or adenovirus) and peptide/ protein pulsing for DC-mediated antigen delivery DCs. The strongest CTL induction could be achieved with virally transduced DCs. Hodge et al. (2000) were able to enhance the T-cell stimulatory capacity of DCs by delivery of the

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costimulatory molecules CD80, intercellular adhesion molecule-1 (ICAM-1) and leukocyte function-associated antigen-3 (LFA-3), by means of either a replication-defective avipox or a replication-competent vaccinia virus and showed in either case enhanced T-cell activation and CTL induction.

Herpes simplex virus vectors HSV-1 vectors are also able to transfer large genes efficiently to a wide range of both dividing and nondividing cells. Given that HSV is neurotropic and capable of long-term persistence, it is the vector of choice for gene transfer into the central nervous system. To date, there have been only a few reports of HSV-based DC gene delivery. Coffin et al. (1998) transduced CD34+ cells and DCs with GFP and lacZ and obtained 100% transduced cells after cell sorting by flow cytometry. Stevenson et al. (2000) constructed a herpesvirus saimiri-based vector and were able to transduce immature DCs at a rate of 25–35%. Salio et al. (1999) reported that HSV infection mediated an inhibitory effect on DC maturation.

NONVIRAL GENE TRANSFER Human DCs Nonviralgenetransferisfarlessefficientthanviral gene transfer, but has the advantages of being less expensive and nominally associated with less pathogenicity/immunogenicity. Also, it is possible that nonviral gene transfer will have less of an effect on DC function or maturation than viral infection. DCs can be transfected in vitro using cationic lipids to deliver plasmid DNA encoding HSV proteins into DCs, resulting in induction of antigen-specific CD8+ (Rouse et al., 1994). Similarly, Alijagic et al. (1995) reported the ability to transfect, PBMC-derived-DCs with two reporter genes (CAT and β-gal) using a liposome-based transfection technique. Tüting et al. (1998) transiently transfected moDCs with plasmid DNA encoding human melanoma TAA-derived MART-1/Melan-A, pMel-17/gp100,

tyrosinase, Mage-1 or Mage-3 by particle bombardment in order to stimulate autologous PBMC responder T cells. Co-transfection of genes coding for IL-12 or IFNa enhanced the antigen-specific CTL response. Similarly, moDCs transiently transfected using cationic liposomes with a MART-1-encoding plasmid were able to induce CTLs able to lyse HLA-matched donor cells expressing the antigen in vitro (Philip et al., 1998). PBMC-derived DCs were also transiently transfected with the HIV-derived nef gene and the expression and subsequent processing of nef by transfected DCs did not alter the presentation of an immunodominant nef epitope to CTLs of HIV+ patients (Chassin et al., 1999). Lohmann et al. (2000) demonstrated that CD83-derived DCs could be transfected by electroporation with cDNA-encoding tumor peptides. Transfection efficiency of up to 10% was obtained as measured by expression of the reporter EGFP gene. Van Tendeloo et al. (1998) compared distinct types of DCs for their susceptibility to nonviral gene transfer mediated by electroporation.They found that lipofection was unsuccessful in CD34derived DCs, Langerhans cells and moDCs, whereas electroporation achieved gene transfer in CD34-derived DCS and Langerhans cells with a transfection efficiency of 16% and 12%, respectively. Transfection efficiency of moDCs by electroporation was very low (only 2%). Nair et al. (1998) transfected PBMC-derived DCs with carcinoembryonic antigen (CEA) mRNA linked to the LAMP-1 lysosomal targeting sequence enhancing a CD4 T-cell response necessary for optimal induction of CD8 CTLs. Morse et al. (1998) showed that optimal antitumor activity of DCs occurs when they are transfected with RNA encoding TAA before maturation with CD40L. Similarly, whole mRNA from tumor cells (Bozkowski et al., 2000) as well as mRNA specifically encoding PSA (Heiser et al., 2000) have been used for transfection of DCs, resulting in induction of CTL-mediated antitumor immunity.

Murine DCs Condon et al. (1996) demonstrated in vivo transfection of DCs by means of cutaneous genetic

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immunization following particle bombardment (gene gun). A similar approach was performed where naked plasmid DNA encoding HSV antigens was delivered intramuscularly and induced mainly a CD4 TH1 response (Marickan et al., 1997). DCs isolated from the skin and lymph nodes after i.m. injection of naked DNA encoding influenza polypeptides, presented antigen to T cells (Casares et al., 1997). Similarly, Porgador et al. (1998) showed that gene gun immunization with infuenza antigens directly targeted DCs in the skin and were responsible for specific CD8+ T-cell priming. Particle-mediated transfer of bone marrow-derived DCs ex vivo with DNA encoding the tumor-associated antigen E7 derived from the human papilloma virus (HPV)16 as well as a ‘self’ epitope from the tumor suppressor gene p53 could induce effective antitumor activity (Tüting et al., 1997). Interestingly, it appears as if only a relatively small number of genetically engineered DCs are required to induce a potent-immune response. Akbari et al. (1999) showed that DNA vaccination resulted in transfection of a small proportion of DCs that led to general activation of all DCs and effective T-cell activation due to crosspriming by death of keratinocytes expressing the antigen. Petersen et al. (1999) transfected DCs with either the full-length p53 gene or a minigene encoding an immunodominant epitope in a HLA-A2 transgenic mouse model and found that specific CTLs could be generated with both gene constructs. In addition to naked DNA, liposomes and particle-mediated delivery of plasmids, DNA can also be delivered to DCs using various conjugates. Diebold et al. (1999b) linked plasmid DNA to mannose polyethylenimine (ManPEI) conjugates for targeted DNA delivery into DCs. These ManPEI/DNA complexes are taken up efficiently by DCs via endocytosis. The co-transfection of cytokines also appears to enhance the immune stimulatory capacity of DCs in that immature bone marrow-derived DCS were modified to express GM-CSF, enhancing the in vivo antigen-presenting capacity of DCs due to better migratory capacity. Yang et al. (1999) transfected murine DCs with human gp100 and elicited better antigen-specific CD8

and CD4 T-cell responses as compared with naked DNA or peptide-pulsed DC immunizations. The immune function of DCs can also be altered following nonviral gene deivery. Recently, Nair et al. (2000) showed induction of CTLs against melanoma and thymoma cells using telomerase reverse transcriptase (TERT) RNA transfected DCs. Min et al. (2000) transfected bone marrow-derived DCs with Fas ligand (FasL) and showed that they were capable of inducing Fas-mediated apoptosis of Fas cells, inhibiting allogeneic MLR in vitro and prolonging survival of mismatched allografts. Relatively small numbers of mIL-4-transfected DCs were sufficient to modify the immune response to induce higher IgE and lower IgG2a levels. Wei et al. (2000) increased the transfection rate of bone marrow-derived DCs four times by the use of a cytoplasmic T7 vector compared to pCMV plasmid, as measured by expression of the firefly luciferase gene.

Future prospects We now are poised to see a variety of gene transfer strategies be tested in clinical studies. Although considerable complexity is associated with the use of exogenous DC isolation, culture and transfection, the ability to derive comparable DCs and effectiveness in various models not requiring exogenous culture have yet to be demonstrated. In the interim, such studies appear to be sound and may ultimately be the major means to immunize, particularly in the setting of chronic diseases such as cancer, allograft rejection, chronic infectious disease and autoimmunity.

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PLATE 43.1 Triple adenoviral co-infection of human DCs. Human DCs can be efficiently infected with up to three different adenoviral vectors. Panels A–D show a typical fluorescent human DC after co-infection with AdEYFP, Ad-EGFP and Ad-EBFP. Figure shows fluorescence with (A) 550 nM emission filter for EYFP detection, (B) 510 nM emission filter for EGFP detection, (C) 440 nM emission filter for EBFP detection and (D) triple band pass filter for simultaneous detection of all three colors.

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44 Phagocytosis of apoptotic cells Matthew L. Albert The Rockefeller University, New York, USA

To examine the causes of life, we must first have recourse to death. Frankenstein Mary Shelley

(Bevan, 1976b). More recently, this exogenous pathway for the generation of MHC I/peptide complexes has also been shown to result in the tolerization of T cells specific for tissuerestricted antigen, a phenomenon called ‘crosstolerance’ (Kurts et al., 1996; Heath et al., 1998). As it is now believed that the activation of naïve T cells is a property restricted to APCs, these in vivo observations offer a potential solution to the question of how CD8 T cells are activated for the targeting of cells that express antigen not found in APCs. Examples of such antigens include tumor-restricted proteins and viruses that do not infect professional APCs (e.g. human papillomavirus, HPV ) (von Knebel Doeberitz, 1992). Additionally, this exogenous pathway provides a potential pathway by which tissue-restricted antigen may access an APC for the tolerization of self-reactive T cells in the periphery (Heath et al., 1998). While these observations indicate that the

INTRODUCING THE DEAD An exogenous pathway for MHC I antigen presentation Significant evidence for an indirect pathway for the loading of MHC class I molecules has emerged over the last 25 years. The most compelling data come from in vivo experiments in mouse models demonstrating that viral, tumor and histocompatibility antigens can be transferred from MHC-mismatched donor cells to host bone marrow-derived antigen-presenting cells (APCs) to elicit antigen-specific CTL responses that are restricted to self MHC molecules (Bevan, 1976a, 1976b; Huang et al., 1994; Sigal et al., 1999). Michael Bevan originally coined this phenomenon ‘cross-priming,’ as antigen is ‘crossing the MHC barrier,’ initially invoked in the generation of MHC class I peptide epitopes and the activation of CD8+ T cells Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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immune system possesses a natural mechanism by which exogenous antigens may access MHC I molecules of APCs, there were two undefined features: the mechanism of antigen transfer (whole cells versus free protein or peptide); and the identity of the APC. In vitro studies have now defined that exosomes, heat-shock proteins, immune complexes and apoptotic cells can all serve as vehicles for the delivery of antigen to the DC system in a manner that permits the cross-presentation of antigen (Suto and Srivastava, 1995; Albert et al., 1998a; Zitvogel, et al., 1998; Regnault et al., 1999). In all in vitro systems in which a direct comparison has been make, dendritic cells (DCs), but not macrophages (MΦs) are capable of crosspresenting antigen (Albert et al., 1998a; Rodriguez et al., 1999; Yrlid and Wick, 2000). In this chapter, we will focus on apoptotic cells as a source of antigen for this indirect pathway. Specifically, I will follow the circuitous route antigens within apoptotic cells must course in order to activate versus tolerize CD8 T cells. In vitro studies will be used to outline defined and potential pathways, and where possible, correlations will be made to in vivo experiments. I will begin by reviewing the antigens which have been cross-presented via this pathway. The discussion will then focus on the dying cell, specifically identifying unique features of apoptotic death that make this exogenous pathway efficient. Next, the interaction between surface ligands on the apoptotic cell and the various receptors on the immature DC will be examined. I then outline three possible routes whereby antigen derived from the apoptotic cell may access the DC’s class I molecules: a recycling pathway for MHC I in which antigen is loaded in the endosome; a pathway whereby retrograde transport of the antigen from the endosome to the ER facilitates entry into the ‘classical’ class I antigen presentation pathway; and an endosome to the cytosol transporter, again allowing antigen processing via the ‘classical’ MHC I pathway. Finally, I speculate on how the microenvironment in which the apoptotic cell is captured may influence the immunologic outcome – priming versus tolerization of the CTL.With a better understand-

ing of the complexities of this apoptosis-dependent pathway for the generation of MHC I/peptide complexes, it may be exploited for DC-based therapies for the activation or tolerization of antigen-specific T cells.

ANTIGENS FROM THE DEAD Dendritic cells engulf apoptotic cells and cross-present viral, tumor and bacterial antigen In our attempts to understand immunity to tissue-restricted antigen, we tested the role of dying cells in the delivery of antigen to APCs. In doing so, we developed a physiologically relevant model whereby human DCs may acquire relevant antigens and stimulate MHC class Irestricted CTLs by phagocytosing apoptotic cells. Our studies provided the first evidence that apoptotic cells are a critical trigger for the crosspresentation of exogenous antigen (Albert et al., 1998b). Apoptotic cells are an attractive source of antigen for a number of reasons. When cells die an apoptotic death, antigen within the cell is concentrated within discrete lipid membranebound vesicles called apoptotic bodies and blebs (Rosen et al., 1995). In addition to being concentrated sources of antigen, vesicles derived from apoptotic cells contain ‘eat-me’ signals, which help facilitate the acquisition of the apoptotic material by phagocytes (Savill, 1996). Other surface or intravesicular proteins, still unknown, may also serve to help facilitate the cross-presentation antigen within the apoptotic cell. Possibly, heat-shock proteins within the apoptotic material serve to chaperone peptide epitopes into the MHC I presentation pathway (Srivastava et al., 1998). To date, in vitro experiments have defined three classes of antigen (viral, bacterial and tumor antigens) which may be cross-primed via this apoptosis-dependent pathway (Plate 44.1). Our studies have defined an experimental system in which apoptotic cells expressing influenza antigen can be phagocytosed by human monocyte-derived DCs, in turn permitting the

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generation of influenza epitopes for the generation of MHCI/peptide complexes and the activation of potent influenza-specific CTLs (Albert et al., 1998b). The influenza antigen may be derived from influenza-infected cells or cell lines transfected with the gene encoding the influenza matrix protein (Albert and Bhardwaj, 1998; Albert et al., 1998b). The apoptotic cell may be syngeneic, allogeneic or xenogeneic, examples of which include primary monocytes, human tumor cell lines (HeLa and 293T cells), as well as various murine cell lines (RAW, EL4 and L929 cells) (Albert et al., 1998b; Albert submitted). The mechanism of triggering apoptotic death have included viral-induced death, UV-B, UV-C and gamma-irradiation (Albert et al., 1998a, 1998b) and unpublished data). We have also demonstrated that apoptotic tumor cell lines expressing a defined tumor antigen, may be captured by DCs for the generation of tumor-specific killer T cells (Albert et al., 1998c). These studies were performed on patients with paraneoplastic cerebellar degeneration (PCD), an important and unique instance of naturally occurring tumor immunity and autoimmune neuronal degeneration in humans (reviewed in Darnell, 1996). The target antigen in this disorder, cdr2, has been cloned and characterized as a Purkinje cell-specific protein which is aberrantly expressed by breast and ovarian tumor cells. While there are cdr2-specific antibodies (termed Yo) present in patients with PCD, no role for these antibodies in the pathogenesis of the disease has been defined as (1) treatment that reduces the titer of PND antibodies does not alter the clinical course of the disease; (2) passive transfer of antibodies to animals does not reproduce the disorder; and (3) the cdr2 antigen is intracellular, suggesting that antibodies may not be sufficient to cause disease (Albert et al., 1998c). We therefore hypothesized that CTLs play an important role in the tumor immunity and possibly the neuronal degeneration evident in these patients (Darnell, 1996). Using apoptotic HeLa cells as a source of cdr2, we have demonstrated that DCs derived from PCD patients can cross-prime cdr2-specific T cells; and that the presence of such an expanded pop-

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ulation of T cells is specific to patients with PCD (Albert et al., 1998c). These data suggest the following model for naturally occurring tumor immunity in these patients: apoptotic tumor cells phagocytosed by peripheral tissue DCs cross-present epitopes derived from cdr2 on class I MHC molecules; and after migrating to a draining lymph node, these DCs might activate cdr2-specific T cells that return to the tumor site, targeting cdr2-expressing cells. The observation that tumor antigens derived from engulfed apoptotic cells can be employed for the activation of tumor-specific T cells was recently confirmed by Russo et al. (2000). In their studies, MAGE-3 epitopes were generated by DCs which had phagocytosed apoptotic cells expressing the MAGE-3 antigen, thus facilitating the activation of class I-restricted tumor infiltrating lymphocytes derived from patients with melanoma. This is an encouraging result, as it suggests that this apoptosis-dependent crosspriming pathway can be employed for immunotherapy in cancer patients. However, it is unclear how these authors resolve their findings with those of Lee et al., who have demonstrated that while melanoma patients indeed have expanded populations of tumor-specific T cells, these cells are functionally unresponsive – unable to directly lyse melanoma target cells or produce cytokines in response to mitogens (Lee et al., 1999). It has also been demonstrated that DCs can engulf apoptotic Salmonella-infected macrophages, and in turn generate both MHC I and MHC II epitopes, derived from bacteriaexpressed antigen (Yrlid and Wick, 2000). In these studies, the bacterium itself triggered apoptotic death. In contrast, when the MΦ were infected with a Salmonella mutant that does not induce death, cross-presentation was not observed, unless the infected MΦ are induced to undergo apoptotic death by other means. Importantly, the critical features of this pathway included: DC as the requisite APC, for while uninfected MΦs were capable of efficiently phagocytosing the apoptotic cells, they could not cross-present the bacterial antigen; and the apoptotic cell as the means of antigen transfer.

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This latter feature was demonstrated using culture conditions that led to Salmonellainduced necrotic death of the infected MΦ, demonstrating a drastic reduction of the presentation of bacterial-encoded antigen on the MHC I of the DC (Yrlid and Wick, 2000). While our in vitro studies have now been confirmed by others (Russo et al., 2000; Yrlid and Wick, 2000), there continues to be some controversy in the literature. In contrast to the critical role of apoptotic cells others have defined a system in which the killing of tumor cells by non-apoptotic pathways is associated with ‘high immunogenicity’ of the expressed tumor antigens. This observation was correlated with the induction of HSP expression in tumor cells (Melcher et al., 1998). I believe that the difference between their study and our findings reflects an unfortunate semantic distinction regarding the definition of aptosis. The tumor cells used in the study of Melcher et al. were transfected with the herpes simplex virus (HSV) gene expressing thymidine kinase (tk), and death was achieved by treating the cells with ganciclovir. Classically, this procedure leads to the induction of apoptotic death (Hamel et al., 1996). To prevent the induction of apoptosis, the authors introduced bcl-2 into their tumor cells; and when treated with ganciclovir, their HSV-tk, bcl-2-expressing tumor cells did not become TUNEL positive. Still, HSPs within the tumor cell were upregulated. According to most definitions, necrotic death is considered nonenergy dependent, as compared to apoptotic death, which requires ATP. Thus, it is confusing what Melcher et al. consider to be necrotic death in light of the fact that they report protein upregulation in response to ganciclovir. It should also be noted that death in the presence of various caspase inhibitors leads to a lack of some of the classical features associated with apoptosis yet the cells die an energy- dependent programmed cell death (McCarthy et al., 1997; Miller et al., 1998; Mills et al., 1998). Thus, their studies may in fact be consistent with the pathway described here. In re-interpreting this data, it is intriguing to consider that the specific apoptotic cascade

involved in triggering cell death may result in distinct immunologic outcomes.

PACKAGING THE DEAD Apoptotic cells efficiently concentrate and deliver antigen to dendritic cells To appreciate the efficiency of the antigen transfer in this apoptosis-dependent exogenous pathway, it is important to review some characteristic features of apoptotic cells. In all cases of programmed cell death, an orderly and energydependent process occurs, resulting in morphological disintegration. This disintegration includes: chromatin margination, condensation and fragmentation; cytoplasmic condensation and vacuolization; and maintenance of cell membrane integrity, often involving extensive cell surface blebbing (Wyllie et al., 1980, 1984). Additionally, the dying cell exposes novel and altered surface markers on its plasma membrane, which facilitates the recognition and rapid clearance of the dying cells by phagocytes and neighboring cells, thereby preventing adverse inflammatory reactions might occur secondary to leakage of the cellular contents (Savill, 1997). Three aspects of this process warrant further discussion: the packaging of antigen into discrete vesicles; the nature of the packaged antigen; and the exposure of novel ligands on the surface of the dying cell.

Concentration of antigen in apoptotic bodies and blebs Alterations in the cytoskeleton of the dying cell are an early feature characteristic of apoptosis. In many cell types, these changes lead to the formation of apoptotic bodies and blebs – discrete surface structures that may be critical for facilitating the efficient transfer of antigen to the DC. It has been demonstrated that apoptotic bodies and blebs contain concentrated viral and self antigen as compared to the apoptotic corpse (Casciola-Rosen et al., 1994a; Rosen et al., 1995). Apoptotic bodies range in size from 0.8 to 6 µm,

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and selectively package the nuclear contents of the dying cell. They have been shown to include concentrated levels of the ribonucleoproteins Ro and La, as well as the small nuclear ribonucleoproteins, or snRNPs – all are known autoantigens in systemic lupus erythematosus (SLE) (Casciola-Rosen et al., 1994a, 1994b). In contrast, apoptotic blebs are small, 0.2-1 µm, and contain contents of the rough endoplasmic reticulum (ER). Sindbis virus-infected cells undergo apoptotic death, and it has been shown that the resulting surface blebs contain abundant viral glycoprotein in their surface membranes (Rosen et al., 1995). Notably, both apoptotic bodies and blebs have phosphatidylserine (PS) on their surface and, as discussed below, this might facilitate recognition and uptake of these vesicular structures by phagocytes (Fadok et al., 1992). Based on these observations, it has been suggested that apoptotic blebs and bodies may serve as vehicles for the delivery of antigen to an APC for the initiation of immune responses. While this remains an intriguing possibility for the transport of antigenic material, it is unclear if this has physiologic relevance. In situ, it is extremely difficult to observe TUNEL-positive apoptotic cells, and neither apoptotic blebs nor bodies have been found in nonpathologic situations. Given the three-dimensional interactions between cells in tissues (as opposed to tissue culture), and the rapidity of engulfment of early apoptotic cells, it is no surprise that corpses are not visualized histologically – presumably this means that the dying cell will be internalized prior to pinching off into blebs and bodies. As such, it is still unclear if the formation of apoptotic blebs and bodies has relevance to immunity.

HSPs within the apoptotic cell may chaperone peptides to DCs Whether the means of antigen delivery is apoptotic fragments or the entire apoptotic cell itself, the nature of the antigen transferred within the apoptotic material is of great interest. Possibilities include: whole or partially degraded

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protein, thus requiring further processing by the DC; peptide epitopes, pre-processed within the apoptotic cell and chaperoned to the DC via binding to a heat shock protein (HSP). HSP/peptide complexes may be concentrated into apoptotic bodies/blebs as a way for antigen to be ‘pre-processed, packaged and delivered,’ thus facilitating the rapid loading of MHC I and MHC II molecules in the DC. This is an appealing hypothesis as HSP/peptide complexes have been shown to transfer peptide epitopes to APCs for the ‘cross-priming’ of T cells (Suto and Srivastava, 1995). HSPs are a highly conserved family of proteins that serve a variety of cellular functions including molecular chaperones for protein folding and protein transport. A role for HSPs as chaperones of peptides into the class I MHC antigen presentation pathway of APCs has also been described. Early experiments indicated that HSPs isolated from tumor cells or virus-infected cells elicit protective immunity to the cognate tumor or viral antigen in various in vivo murine models (Suto and Srivastava, 1995). As HSP genes do not show polymorphism (Maki et al., 1990), it was suggested that the HSPs themselves are not immunogenic, but instead serve as chaperones for tumor or viral peptides (reviewed in Srivastava et al., 1998). It is now known that four HSPs possess the ability to chaperone peptides into the MHC I antigen presentation pathway of an APC – two members of the HSP90 family, GP96 in the ER and the cytosolic HSP84/86 molecules; the cytosolic HSP70s; and Calreticulin, an ER chaperone involved in the generation of stable MHC I/peptide complexes (Udono and Srivastava, 1993, 1994, 1995; Arnold et al., 1995; Basu and Srivastava, 1999). We have begun to examine the role for HSPs in dying cells and preliminary data suggest that GP96 is concentrated in apoptotic blebs while cytosolic HSP70s can be found in the apoptotic bodies (unpublished data). Additionally, it should be pointed out that HSPs are stressinduced proteins and many of the same triggers of apoptotic death (e.g. viral infection) will also result in the upregulation of HSP family members, thus making them excellent candidates for

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chaperoning peptide epitopes to the DC (Wu, 1995). Other surface or intravesicular proteins, still unknown, may also serve to help facilitate the cross-presentation of antigen.

Surface ligands on apoptotic cells While much is known regarding induction and signal transduction cascades responsible for the triggering of apoptosis, less is known about changes in the plasma membrane of the dying cell. Information regarding the alterations that occur is essential in order to understand the mechanism of recognition and engagement of the apoptotic cell by the phagocyte. Clearly, an appropriate ligand for such an interaction is one that is exposed on the outer leaflet of the apoptotic cell’s plasma membrane, but not present on a living cell. Investigations employing a variety of cell types imply that changes may include carbohydrate motifs (e.g. N-acetyl glucosamine), distinct patterns of surface phospholipid exposure, as well as subtle structural changes (Savill, 1997; Platt et al., 1998). One of the best-characterized changes in the plasma membrane occurs during early apoptotic death (Bruckheimer and Schroit, 1996). Effector proteins in the caspase cascade inactivate a flippase or lipid translocase that is responsible for maintaining anionic phospholipids exclusively within the inner leaflet of the plasma membrane (Connor and Schroit 1991; Connor et al., 1994; Diaz and Schroit, 1996). Additionally, there is the activation of a lipid scramblase, which effectively scrambles the lipid bilayers of the dying cell. Together, these two events result in loss of membrane asymmetry throughout the cell, including the plasma membrane. One of the newly exposed phospholipids is phosphatidylserine (PS), and it has been proposed that this may serve as a ligand for engagement with surface receptors on the phagocyte (Fadok et al., 1992; Zwaal and Schroit, 1997). Experiments involving competing apoptotic cell ingestion with either liposomes containing PS or soluble phosphoserine salts suggest that this indeed constitutes a means of phagocyte recognition of the dying cell. Supporting these data

are the observation that externalized PS may bind soluble proteins to form a molecular complex that is recognized by relevant phagocyte receptors (Savill, 1996). There is also evidence that suggests a role for newly exposed sugar residues on the surface of an apoptotic cell. Seminal studies by Wyllie and colleagues implicated macrophage lectins, selective for N-acetyl glucosamine, Nacetyl galactosamine and galactose, in lowtemperature binding of apoptotic thymocytes by resident peritoneal macrophages (Duvall et al., 1985). Furthermore, there is evidence that mannose/fucose carbohydrate motifs may be involved in the recognition and uptake of apoptotic cells via still undefined lectin receptors (Hall et al., 1994). Recently, it was demonstrated that ICAM-3 (CD50) undergoes a putative conformational shift which allows for the most distal extracellular domain of the protein (domain 1) to interact with CD14, a receptor on macrophages which mediates recognition of apoptotic cells (Devitt et al., 1998; Moffatt et al., 1999). While this finding is intriguing, as it offers a new paradigm for the generation of novel epitopes on the surface of the apoptotic cell, it is unlikely that ICAM-3 plays a role in DC-mediated phagocytosis of apoptotic cells as they do not express the CD14 receptor.

EATING THE DEAD Receptor utilization may distinguish the immunologic outcome of capturing an apoptotic cell Until recently, very little was known about the molecular mechanisms responsible for the engagement and internalization of apoptotic cells by phagocytes, but within the past few years there have been a number of molecules implicated in this process. Surface receptors involved in the phagocytosis of apoptotic cells now include: CD14; several members of the scavenger receptor family; the ABC1 transporter (a putative chloride channel and human

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homolog of CED-7); the complement receptors CR3 and CR4 (also known as αMβ2 and αXβ2, respectively); the Fc receptor (FcR); the αvβ3/CD36 complex, bridged by the soluble protein thromospondin-1 (TSP-1); and the αvβ5/CD36 receptor complex, possibly bridged by the soluble protein lactadherin (reviewed in Platt et al., 1998). Additionally, putative receptors include the macrophage mannose receptor (MR) and a still undefined phosphatidylserine receptor (Fadok et al., 1992; Savill, 1996). With the exception of CD14 (not present) and αvβ3 (not active, Albert et al., 1998b) all of these receptors are potentially involved in DCmediated uptake of apoptotic cells. As an exhaustive discussion of these molecules is beyond the scope of this chapter, I will focus on the three receptor systems reported to play a role in the DC’s ability to capture apoptotic cells: the αvβ5/CD36 receptor complex, the complement receptors and the Fc receptor.

The αvβ5/CD36 receptor complex The first demonstration that integrin receptors are involved in the capture of apoptotic cells was offered by Savill and colleagues, who demonstrated that the vitronectin receptor αvβ3 mediates phagocytosis in macrophages (Savill et al., 1990). It is now known that αvβ3 forms a receptor complex with CD36, which is bridged by the soluble protein Thrombospondin-1(TSP-1), thus facilitating binding of the apoptotic cell. TSP-1 is perfectly suited to act as a ‘molecular bridge’ as it possess an Arg-Gly-Asp (RGD) domain, permitting interaction with the αvβ3 integrin; a thrombospondin domain, allowing binding to CD36; and two factor VIII-like domains, which are known to bind anionic phospholipids (Roberts et al., 1985; Robson et al., 1988; Silverstein et al., 1992; Kosfeld and Frazier, 1993). In contrast to macrophages, immature DCs express little αvβ3; instead they employ an alternate αv heterodimer, the αvβ5 integrin receptor, for the efficient internalization of apoptotic cells. As is the case in the macrophage receptor complex, αvβ5 functions in concert with CD36. Importantly, the expression of this receptor pair

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is restricted to the immature DC as compared to macrophages which lack β5. Notably, this distinction in receptor utilization constitutes the first known distinction between macrophages and DCs in their handling of apoptotic material (Albert et al., 1998b). With respect to a bridging molecule, preliminary data suggest that DCs do not produce TSP-1; and that TSP-1 inhibitors, which block phagocytosis in macrophages, do not affect αvβ5-mediated phagocytosis of apoptotic cells in DCs (unpublished data). Studying other phagocyte systems, an interesting candidate protein has been identified. Lactadherin (LA), also known as MFG-E8, was originally defined for its role in breast milk secretion (Couto et al., 1996; Aoki et al., 1997). The ductal epithelial cells in the breast secrete plasma membrane-bound-milk fat globulins; these lipid droplets expose PS on its outer leaflet and are coated by the soluble factor LA, which acts analogous to an opson in facilitating receptor recognition by the target cell (Newburg et al., 1998; Peterson et al., 1998a, 1998b). This receptor system has been defined in bovine and turns out to be αvβ5/CD36 (Hvarregaard et al., 1996; Andersen et al., 1997). There has also been a suggestion that LA may interact with αvβ3. Although unrelated structurally, LA, like TSP-1, is a multidomain protein which contains an EGF-like domain, an RGD domain (αvβ5-interacting domain), and two factor V/VIII-like C domains (C1 and C2) in its C-termini that mediate binding to anionic phospholipids (Couto et al., 1996). Thus, LA is the ideal candidate to act as a ‘molecular bridge,’ facilitating DC capture of apoptotic cell (Plate 44.2). Of particular interest, it has been recently shown that the dominant protein which coats the outer leaflet of exosomes is MFG-E8/LA (Thery et al., 1999). Future studies will be required to determine if the utilization of the αvβ5 receptor is directly responsible in mediating the cross-presentation of antigen by DCs.

The microenvironment in which cells die determines receptor utilization With respect to other receptor systems involved in the phagocytosis of apoptotic cells by DCs,

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two others warrant discussion, as they help illustrate the possibility that receptor utilization influences the immunologic outcome. The complement receptors efficiently mediate capture of apoptotic cells in both macrophages and DCs (Mevorach et al., 1998; Albert and Mevorach unpublished data). These receptors probably bind C3bi, which in turn is capable of opsonizing apoptotic cells (Plate 44.2) (Mevorach et al., 1998). Interestingly, phagocytosis via CR3 does not lead to the release of inflammatory cytokines, and in fact, in the absence of FcR activation, may actually be immunosuppressive (Marth and Kelsall, 1997). While these observations have not been directly tested in DCs, it offers the possibility that the capture of antigen via the CRs might lead to cross-tolerization of antigen-specific T cells. There is also evidence that DCs can capture apoptotic cells via the FcR. Immature DCs express several Fc receptors, but it is unclear which ones will be critical for the phagocytosis of antibody-apsonized apoptotic cells (Plate 44.2). Still, it should be pointed out that such an event has the possibility to result in both the maturation of the DC as well as the cross-presentation of the internalized antigen (Manfredi et al., 1998; Regnault et al., 1999). In this way, antigen derived from the apoptotic cell could be used for the priming of antigen-specific T cells. It is important to understanding the full repertoire of receptors employed by DCs for the binding and internalization of apoptotic cells, as the data available suggests that receptor utilization may play a role in determining the immunologic outcome of the phagocytic event. Interestingly, in each of the three receptor systems described, a soluble factor is required to bridge the DC and the apoptotic cell (Plate 44.2). Thus, it is possible that the microenvironment in which cells die may determine receptor utilization, in turn affecting the potential fate of cross-presentation – activation or tolerization of antigen-specific T cells. According to this hypothesis, the presence of an antibody specific for a surface antigen on the apoptotic cell would allow for capture via the FcR and the activation of CTLs (e.g. anti-PS antibodies present in SLE

where apoptotic death is thought to be a trigger of lupus flares, Casciola-Rosen and Rosen, 1997). In contrast, the complement cascade may be activated by normal cell turnover, as it has been suggested that negatively charged phospholipids such as cardiolopin and PS activate C1 in the absence of antibody (Kovacsovics et al., 1985; Mevorach et al., 1998). In this way tissue-restricted self antigen may be handled in a manner that results in the tolerization of self-reactive T cells. With respect to the use of the αvβ5 integrin receptor, uptake of apoptotic cells via this receptor is apparently immunologically quiescent (Sauter et al., 2000), and therefore the cross-presentation of antigen has the potential to be directed for either the cross-priming or the cross-tolerization of CTLs, in part depending on additional criteria such as the presence of absence of antigen-specific CD4 helper T cells (discussed below).

PROCESSING THE DEAD Potential paths for MHC I/peptide complex formation It is believed that all entries into the cell converge at the endosome. In general, the endocytic system is comprised of pleiomorphic interconnected arrays of vesicles, larger vacuoles and tubules which communicate with the plasma membrane and the trans-Golgi network (TGN) (Geuze, 1998). The phagolysosome is the first stopping point for the engulfed apoptotic cells and within 30–60 minutes, LAMP vesicles begin to fuse with it, presumably aiding in the initial processing of the internalized antigen (Albert and Kleijmeer, unpublished data). Of great interest is the mechanism by which the engulfed antigen finds its way from the phagolysosome to the MHC I of the DC. Possibilities include: (1) loading of MHC I within the endosome, presumably within the late endosomal compartments, as is the case in MHC II antigen presentation; (2) retrograde transport of internalized antigen via the Golgi complex to the ER, thus delivering antigen to the biosynthetic

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MHC I processing pathway; and (3) a mechanism by which antigen may exit the endosome and enter the cytosol, thus entering the ‘classical’ class I MHC antigen presentation pathway of the DC (Plate 44.3). Note, all three of these pathways may be operational simultaneously; in fact, different antigens may access the MHC I of the DC via different routes.

Pre-processed epitopes may be loaded on to MHC I within the late endosome In a number of recent studies, MHC I was detected within all of the described endocytic compartments (Gromme et al., 1999; Kleijmeer et al., in press). Early endosomes (EEs), characterized by the presence of the transferrin receptor (TfR), were shown to contain 17% of the class I heavy chain (HC) within the endocytic compartments. The multivesicular late endosomes (LEs) and lysosomes, identified by the presence of LAMP-2 and CD63, contained 56% and 27% of endocytic class I HC labeling, respectively. Importantly, β2m could also be detected within each of these endocytic compartments offering the possibility that stable MHC I complexes exist in these compartments. Cyclohexamide treatment, which blocks the biosynthetic pathway of MHC I, indicated that the class I present in the endocytic compartments was derived from the cell surface (Monique Kleijmeer, personal communication). Our recent findings have confirmed the presence of MHC I in the endocytic compartment of DCs; furthermore, we have demonstrated co-localization of the phagocytosed apoptotic material with MHC I of the DC. This finding may simply reflect the internalization of surface MHC I molecules for purposes of degradation in the lumen of the lysosomes, however, there remains the possibility of a functional role for this observation. Supporting this possibility is the fact that much of the MHC I found within the multivesicular LEs is concentrated on the internal membranes. Notably, the internal membranes make up the vesicles which are released into the extracellular medium upon LE fusion with the cell surface; in other words, MHC I localizes on the surface of what will

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become exosomes (note, exosomes have the ability to initiate a class I-restricted immune response; Raposo et al., 1996; Zitvogel et al., 1998; Kleijmeer et al., submitted). Apart from the exocytosis of LEs, other mechanisms have been described whereby class I MHC that is associated with the limiting membrane of the LE may be inserted into the plasma membrane (Reid and Watts 1990). These observations offer the possibility that antigen derived from the phagocytosed apoptotic cells may be loaded on to recycling MHC I and subsequently the MHC I/peptide complex transported to the cell surface. This model suggests that a ‘pre-processed antigen’ would be required as it would be inefficient for whole protein to be degraded nonspecifically within the LE by resident catabolic enzymes. Not only would this be a slow process, it would be by ‘chance’ alone that an appropriate nine amino acid peptide capable of binding the MHC I groove would be generated. Notably, none of the requisite molecules involved in class I antigen presentation are found in the endocytic compartment. The notion of such a preprocessed antigen fits well with a role for HSPs as chaperones for the peptide antigen (see ‘Packaging the Dead’). As discussed, protein synthesized within the donor cell would be processed within the endogenous class I antigen presentation pathway leading to the generation of HSP/peptide complexes (Srivastava et al., 1998). These complexes are ideal chaperones of antigenic peptides for the transfer of antigen to the DC for a number of reasons. Once generated, the HSP/peptide complexes might be concentrated within apoptotic bodies or blebs that are engulfed by the immature DC. The HSP would also serve to protect the peptide antigen from degradation upon entry into the endocytic compartment of the DC. Finally, it has been suggested that some of the HSP family members may be capable of facilitating the loading of MHC I (Li and Srivastava, 1993). Alternatively, an as yet uncharacterized lysosomal enzyme may play a role in the processing of internalized antigen for the generation of MHC I epitopes. Possibly, DC-restricted

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cathepsin molecules will play a role in the processing of antigen for the generation of MHC I antigenic determinants in the endocytic system (Villadangos et al., 1999).

Retrograde transport of exogenous antigen to the ER It is also possible to envisage a route whereby antigen derived from the engulfed apoptotic cell is trafficked to the ER for entry into the classical MHC I presentation pathway of the DC. Studies focused on the mechanism of entry of bacterial protein toxins have helped to define one pathway by which this might occur. Toxins such as cholera toxin, Shiga toxin and the plant toxin ricin, all find entry into the ER via retrograde transport through the biosynthetic/secretory pathway (Sandvig et al., 1994; Majoul et al., 1996). Cholera toxin (CT), like other toxins in this family, have a classical AB5 subunit composition (reviewed in Johannes and Groud, 1998). The A fragment is the toxin and in the case of CT, the subunit is a Gsα ribosylating enzyme that leads to an increase in the rate of cAMP synthesis. While the A fragment contains all of the effector function, it cannot itself enter the cells. Entry requires the non-covalently linked homopentemer B fragments, and occurs via different mechanisms for the different family members. Importantly, both Shiga toxin (ST) and CT have been shown to co-localize with the TfR in the EEs (see above). Once within the endosome the toxin complex disassociates: in pulse chase experiments, the B subunits can be localized in LEs and eventually lysosomes where they are degraded; and, the A subunit is transported from the EE to the TGN (Sandvig et al., 1994; Cieplak et al., 1995; Majoul et al., 1996, 1998). While endosome to TGN transport is still poorly characterized, this pathway is not unique to the bacterial toxins. In fact, a number of housekeeping gene products such as the mannose 6-phosphate receptor (MPR), TGN38 and furin, all traffic from the plasma membrane to the TGN via the endosome – some via the EE while others require a slightly acidic pH and probably exit from the LE (Demaurex et al.,

1998). Of note, MPR transport to the TGN is regulated by the small GTPase Rab9 (Diaz et al., 1997). By employing a dominant negative Rab9 mutant, it might be possible to explore the role of endosome to TGN transport in the crosspresentation of antigen by DCs. Once within the TGN, it is possible to invoke a well defined pathway for the trafficking of proteins between the Golgi apparatus and the ER (Pelham, 1991; Miesenbock and Rothman, 1995). Proteins that follow such a route often contain an ER retention sequence as is the case for CT, a COOH-terminal Lys-Asp-Glu-Leu (KDEL) motif. Proteins that contain such a motif are known to bind ERD2, the KDEL receptor, which exists mainly in Golgi-like structures and binds with increased affinity under the slightly acidic condition found in the Golgi cisternae (Lewis et al., 1990). In this way, proteins possessing a KDEL motif are captured in the Golgi and transported to the ER; the ERD2 receptor exhibits decreased affinity for the KDEL motif with increasing pH, thus in the neutral environment of the ER, bound proteins are released. Indeed, the KDEL sequence is less of an ER retention signal and instead is considered more as a retrieval signal important for mediating this Golgi to ER pathway (Pelham, 1991). The relevance of this pathway to the crosspresentation of antigen derived from apoptotic cells becomes clearer when we again consider a possible role for HSPs in the trafficking of antigen from the dying cell to the MHC I of the DC. GP96 possesses an NH2-terminal KDEL motif, and normally resides within the ER where it plays a role in chaperoning peptide antigen to MHC I (see above) (Altmeyer et al., 1996). If GP96 (or other ER chaperones such as calreticulin) were involved in chaperoning antigen in this apoptosis-dependent exogenous pathway, it is conceivable that endosome to TGN trafficking, followed by retrograde transport, would provide the exogenous HSP peptide complex access to biosynthetic MHC I within the ER. Our observation that lactacystin only partially inhibited cross-presentation suggests that a proteasome-independent pathway exists for the loading of MHC I in the DC (Albert et al.,

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1998a). Both the loading of the MHC I/peptide complex within the endosome, and an endosome to ER (via the TGN) pathway offers the possibility of such a proteasome-independent mechanism.

trafficking antigen derived from apoptotic cells in a manner which permits the formation of MHC I/peptide complexes.

PRESENTING THE DEAD A ‘phagosome-to-cytosol’ pathway may permit access to the classical MHC I pathway Phagocytosed antigen may also exit the endosome and enter the cytosol whereupon it can be ubiquinated and processed by the proteasome, thus entering the ‘classical’ MHC I antigen presentation pathway. This is supported by the recent work of Rodriguez et al., who suggest that DCs can traffic exogenous antigen derived from immune complexes from endocytic compartments to the cytosol (Rodriguez et al., 1999). Interestingly, this transport pathway is restricted to DCs, again demonstrating a distinction between DCs and MΦ in their handling of exogenous antigen. This work suggests that cytosolic transport does not occur as a result of nonspecific leakage, but instead results from the existence of a novel membrane-transport pathway. Bioactive horseradish peroxidase (HRP), derived from internalized HRP-immune complexes, could be isolated from the cytosol of the DC within 2–4 hours; in contrast, resident lysosomal proteins such as cathepsin D and β-hexosaminidase remained vesicular. Consistent with the ability of the antigen to access the MHC I presentation pathway, treatment of DCs with proteasome inhibitor resulted in enhanced recovery of bioactive HRP (Rodriguez et al., 1999). Possible mechanisms by which antigen is transported to the cytosol include the expression of a dendritic cell-restricted transporter; or a putative dendritic cell-specific endocytic compartment, in which antigen may efficiently escape to the cytosol (Rodriguez et al., 1999). In either case, the existence of this endosomecytosol transport pathway accounts for the unique ability of DCs to present internalized antigens on MHC class I molecules. Future studies will be required in order to determine if this pathway also accounts for the DC’s ability to

Distinct outcomes of engulfing apoptotic cells Until recently, phagocytosis of apoptotic cells was not considered to induce an immune response. However, our data and data from other laboratories suggest that apoptotic cells are not simply degraded for purposes of clearance. Instead, they are acting to specifically trigger suppression, stimulation and possibly tolerization of T cells. These distinct outcomes are in part dependent on APCs available for the phagocytosis of the dying cell; the microenvironment in which the apoptotic cell dies; and the presence or absence of antigen-specific CD4+ T cells.

Macrophage capture of apoptotic cells may result in immunosuppression Macrophages efficiently engulf apoptotic cells and were originally thought to clear dying cells in an immunologically quiescent manner (Savill, 1996). It has recently been demonstrated, however, that phagocytosis of apoptotic cells actively inhibited the production of pro-inflammatory cytokines such as IL-1β, IL-12 and TNFα (Voll et al., 1997; Fadok et al., 1998). In contrast, production of TGFβ1 and platelet-activating factor (PAF) was increased. Both are considered to be immunosuppressive as addition of exogenous TGFβ1 or PAF results in the inhibition of LPSstimulated cytokine production (Fadok et al., 1998). Interestingly, these studies indicated that receptor utilization by the macrophage for the binding and internalization of the apoptotic cell is critical. While crosslinking CD36 on the macrophage (a co-receptor for αvβ3, see ‘Eating the Dead’) mimicked the anti-inflammatory effect of the apoptotic cell (Voll et al., 1997), the

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phagocytosis of antibody-opsonized apoptotic cells abrogated this immunosuppressive effect (Fadok et al., 1998). This may reflect an important distinction between the way macrophages versus DCs behave postengulfment of an apoptotic cell, and lends further evidence that receptor usage may influence the immunologic outcome of capturing an apoptotic cell.

DC maturation and CD4 T-cell help are critical for the ‘cross-priming’ of CTLs As discussed above, immature DCs are capable of capturing apoptotic cells and cross-presenting antigen derived from these dying cells to activate class-I restricted CTLs. In situ, the capturing of apoptotic cells by DCs is likely to occur in the periphery, as this is where immature DCs are situated. As such, a maturation signal is probably required for the DCs to migrate to the draining lymph organ for the initiation of T-cell responses. However, DC maturation is not triggered by the uptake of the apoptotic cells themselves (Sauter et al., 2000). It has therefore been proposed that either an additional signal is required to trigger DC migration, or a constitutive pathway exists for DCs to traffic to the draining lymph node (Huang et al., 2000). In addition to the possibility that the maturation state of DCs might affect the outcome of the cross-presentation of antigen, there is likely to be an important role for the CD4 helper T cell. In our current studies, we have discovered that the presence or absence of CD4 helper cells modulates the apoptosis-dependent exogenous pathway for MHC I antigen presentation (Albert et al., submitted). This finding is in accordance with described in vivo models (Bennett et al., 1998; Green et al., 1998; Ridge et al., 1998; Schoenberger et al., 1998). It is important to consider that in some cases, this requirement for CD4 help distinguishes the exogenous pathway from the classical MHC I antigen presentation pathway. For example, DCs directly infected with influenza virus generate endogenously synthesized influenza antigen, and are capable of activating CTLs in the absence of helper T cells. In contrast, DCs cross-presenting influenza

antigen derived from phagocytosed apoptotic cells require cognate help from CD4 T cells. In the absence of such help, the CD8 T cells are either ignored or tolerized (Albert et al., submitted), possibly depending on antigen dose (Kurts et al., 1999a). These observations suggest that we must next address the question of CD4 tolerance versus activation in order to understand the regulation of CTL responses specific for tissue-restricted antigen.

The exogenous pathway may also result in cross-tolerization of CD8T cells specific for tissue-restricted self antigen The uptake of apoptotic cells by DCs may also prove to be important for the induction of ‘crosstolerance,’ an in vivo phenomenon first described by Kurts et al., (1996). In their experiments, the OVA protein was expressed under the tissue-restricted, insulin promoter in transgenic mice (mRIP-OVA). When OVA-specific, MHC Irestricted TCR transgenic T cells were adoptively transferred into these mice, the T cells accumulated and expanded in the draining lymph node. These T cells were responding to antigen that had been cross-presented by bone marrowderived cells and not to the islet cells themselves; and the proliferative response was followed by deletion (or tolerance) via a Fasdependent mechanism (Kurts et al., 1996, 1998a, 1999b; Heath et al., 1998). In contrast, when OVA-specific MHC II-restricted TCR transgenic T cells were co-injected, now the CD8 T cells expanded, became effector cells and lysed the OVA-expressing islet cells (Bennett et al., 1997; Kurts et al., 1997). It was therefore proposed that a bone marrow-derived cell can capture exogenous antigens for both class I and class II presentation, transport these antigens to the draining lymph nodes and present the antigen to naïve CD4 and CD8 T cells. Possibly, the key presenting cell is the DC given its capacity to capture antigens in the periphery and migrate to the T-cell areas of the lymphoid organ. Furthermore, it is likely that apoptotic islet cells are responsible for transferring antigen to the

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DC, as the cross-presentation of OVA antigen in this model system was enhanced when the OVA-expressing cells were targeted in situ by adoptively transferred activated CTLs (Heath et al., 1998; Kurts et al., 1998b). It will be important to define an in vitro system to model the cross-tolerization of CD8T cells to establish that indeed apoptotic cells may transfer antigen to DCs for the deletion of selfreactive T cells.

LESSONS FROM THE DEAD Cross-presentation of apoptotic cells in physiologic and pathologic settings Much work is still required to sort out what factors determine which cell gets to ‘eat’ the apoptotic cell, and in what context this event might occur. Clearly, there is a sharp distinction between the fate of the apoptotic material engulfed by the macrophage versus the DC. Receptor utilization may also play a role in directing antigen for different immunologic outcomes. Additionally, there is likely to be a differential effect if the material is cross-presented by the DC in the presence or absence of an inflammatory/maturation signal. Finally, there is a role for the CD4 T cell in distinguishing crosspriming from cross-tolerance, thus focusing the attention on the T-cell repertoire and the tolerization of CD4 T cells, both centrally and peripherally. In this final section, I offer three in vivo situations in which the apoptosis-dependent cross-presentation pathway may be critical in mediating the observed immunologic responses.

Viral immunity in the absence of direct infection of an APC Based on the modulating factors described, and our knowledge of the distinct localization of macrophages versus DCs in normal tissue, it is possible to propose a model for what might occur in various settings of antigen challenge. Let us first consider an apoptotic keratinocyte

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(KC) in the epidermal skin. KCs may undergo apoptotic death during normal cell turnover, exposure to UV irradiation (e.g. the sun), or as a consequence of viral infection (Synder, 1975; Kane and Maytin, 1995; Chrysomali et al., 1996; Pei, 1996). In normal skin, there are an abundance of Langerhans cells (LCs), a myeloidderived subset of immature DCs found in the epidermis (Stingl et al., 1987). LCs are thought to form a continuous network within the epidermis, and serve as ‘sentinels;’ they are responsible for capturing antigen and relaying the signal to the T cells in the draining lymph node (Shelley and Juhlin, 1976a, 1976b). In contrast, there are no macrophages in the normal epidermis. Thus, in the absence of inflammation (or a ‘danger signal’, Matzinger, 1994, 1998), either a neighboring KC or an LC would capture the dying cell. In the case of the latter, the LC might traffic to the draining lymph node and given the lack of a maturation signal and/or the lack of CD4 T-cell help, the result might be the cross-tolerization of any skin-specific T cells it engages (Plate 44.4a). Indeed, this apoptosis-dependent exogenous pathway for MHC I antigen presentation might represent a critical pathway for the induction of self-reactive CD8 T cells. Let us now consider the situation when the KC is infected with a virus, for example, human papillomavirus (HPV). Papillomaviruses have a high degree of species and tissue specificity. With regard to cellular tropism, HPVs infect only surface squamous epithelium of the skin or mucosa, and to my knowledge, papillomaviruses have not been shown to infect LCs or DCs (Shah and Howley, 1996). As is the case with EBV, even in the absence of direct infection of a DC, there are profound CTL responses detectable in individuals infected with HPV. Indeed, it is thought that CD8+ T cells are responsible for controlling the infection. Additionally, it is known that many strains of HPV induce apoptotic death in KCs. It is therefore tempting to speculate that the activation of HPV-specific T cells occurs via the cross-presentation of antigen derived from the apoptotic KCs. Given the presence of a virus, a putative danger signal, the LCs might capture the

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apoptotic HPV-infected KC, become activated and migrate to the draining lymph node whereby it could induce HPV-specific T cells (Plate 44.4b). It is also intriguing to reflect on what might occur if cross-presentation of antigen in blocked or in some still undefined manner abrogated by the virus. In the case of papillomaviruses, let us now consider the highly pathogenic HPV-16 and HPV-18 strains, which express E6 and E7, early gene products with high affinity for p53 and retinoblastoma (Rb), respectively. Their ability to efficiently inhibit apoptotic death has been correlated with the fact that these viral strains transform epithelial cells and are the causative agents of cervical carcinoma. While it has yet to be demonstrated, I propose that the inhibition of P53-mediated apoptotic death may lead to a defect in the cross-priming of HPV-16- and HPV18- specific T cells, resulting in poor control of the virus and the possibility for these viral strains to promote tumorigenesis.

Inappropriate cross-priming – a possible trigger for autoimmunity There is indication that a delicate balance exists between macrophage-mediated immunosuppression and DC cross-presentation of antigen derived from apoptotic cells. In the event of an imbalance, autoimmunity might occur. One relevant example of this is SLE. It is known that sun exposure is a trigger for lupus flares, suggesting that apoptotic cells might be a source of antigenic material for the exacerbation and possibly even the initiation of SLE (Casciola-Rosen and Rosen, 1997). It has been described that patients with SLE have a defect in their ability to phagocytose apoptotic material. This has been demonstrated both in vitro by studying macrophage function as well as in vivo by visualizing lesional skin of lupus patients (Mir and Balsalobre, 1994; Hermann et al., 1998; Pablos and Gomez-Reino, 1998). Apoptotic corpses were detected in the UV-irradiated skin of SLE individuals; in contrast, normal individuals clear apoptotic cells in the skin with such rapidity that they cannot be detected. This defect in phagocy-

tosis can be mapped to the molecular level in a subset of patients who lack C1q, a protein involved in the complement pathway (Walport et al., 1998). It should be noted that the CR3 (aMβ2) and CR4 (αXβ2) complement receptors are involved in the clearance of apoptotic cells, and that signaling through the complement receptors is thought to be immunosuppressive (see ‘Eating the Dead’). As a result of this phagocytic defect, the quantity of apoptotic material apparently overwhelms the normal clearance capacity of macrophages leading to the possible inappropriate cross-presentation of antigen by DCs. Still, it is unclear why in the case of SLE patients the cross-priming of self-reactive T cells prevails over cross-tolerization. As SLE is a polygenetic disorder with gene susceptibility loci that map to the MHC II gene complex, it is intriguing to propose that the availability of a self-reactive CD4 T cell might explain the skewing of antigen cross-presentation toward priming.

Tumor immunity versus tumormediated immunosuppression The last example we will consider focuses on the different consequences resulting from the capture of apoptotic tumor cells. There have been considerable attempts at demonstrating tumorspecific cellular immune responses in patients with cancer, to no avail. Expanded populations of functionally active tumor-specific CTLs have not been previously found in humans (Schuler and Steinman, 1997). This is the case even when the tumors express known tumor-specific antigens such as the melanoma MAGE/MART antigens (Boon and Old, 1997). At first, this might appear surprising given what we now know about the cross-presentation of antigen. Indeed, the cell proliferative cycle is intertwined on a molecular level with the cell death cycle, so tumor cells have an even higher likelihood of undergoing apoptotic death than normal tissue. In fact, it is possible to characterize tumors by their apoptotic index with more aggressive tumors having a higher propensity to undergo apoptosis

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(King and Cidlowski, 1995). So why don’t tumorspecific antigens get cross-presented for the initiation of tumor-reactive CTLs? A number of answers to this question can be supported by existing data, thus offering an explanation for the establishment of a bona fide cancer. If the cross-priming of tumor-specific T cells did occur, the tumor might still evade immune attack if it possessed a way to immunosuppress the tumor-specific T cell (e.g. expression of FasL, Hahne et al., 1996). Alternatively, an absence of CD4 T cell help for the tumorspecific CD8 killer cell might lead to crosstolerance as opposed to cross-priming. Finally, the apoptotic tumor cells might be sequestered (captured) by macrophages, thus leading to an immunosuppressive outcome. Notably, it has been reported that up to 50% of tumor bulk can be macrophages; additionally, there are reports that tumor cells can induce DC apoptosis (Esche et al., 1999). Indeed, both possibilities would result in a deficiency in the crosspresentation of tumor cell-derived antigen. There are, however, documented cases of tumor regression and tumor immunity. As discussed, we have revealed that paraneoplastic cerebellar degeneration (PCD), an example of naturally occurring tumor immunity, correlates with the cross-priming of PCD-specific T cells. With such a model, it is now possible to define the factors which determine if the tumor cells will be handled in a way that leads to the crosspriming of tumor-specific T cells as occurs in the case of PCD patients (Albert et al., 1998c), or the cross-tolerization and/or immunosuppression that occurs in individuals with cancer (Lee et al., 1999). Parameters to consider include the tissuerestriction of the tumor antigen – in other words, in the case of PCD pathogenesis, is it critical that the associated tumor antigen, cdr2, is normally brain-specific? While the neuronal expression of cdr2 is what establishes the Purkinje cell as a target for the immune system, our recent finding that 60% of ovarian tumors from neurologically normal individuals suggests that alone (Darnell et al., 2000), the ectopic expression of an ‘immune-privileged’ antigen is not sufficient to trigger tumor immunity. These ovarian cancer

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patients need to be evaluated for the presence of cdr2-specific CD8T cells in order to fully resolve this issue. Additionally, the presence of CD4 helper T cells needs to be studied, as this might explain the distinct outcomes in the two populations of patients – cross-priming in he case of PCD and cross-tolerance in neurologically normal women harboring cdr2 tumors. If PCD patients uniquely cross-prime their cdr2-specific T cells, then possibly the meaning of ‘immune-privilege’ has more bearing on the peripheral tolerization of self-reactive CD4 T cell responses. Alternatively, we may need to consider the possibility that susceptibility to PCD is linked to the class II MHC gene complex as is true in many autoimmune disorders, including those thought to be mediated by CD8 T cells in their effector phase, e.g. type I diabetes.

Who gets to ‘eat’ . . . and when I have spent much time considering the role of DCs in uncovering this exogenous pathway for MHC I antigen presentation and CTL activation. Ironically, in considering the in vivo relevance of this pathway, it has become apparent that CD4 T cell help, and the macrophage’s access to the apoptotic material will both critically influence the outcome of apoptotic death in situ. With respect to the role of macrophages, I am intrigued by their ability to capture the apoptotic material five to ten times more efficiently than DCs. Based on the distinct localization of DCs versus macrophages within the body, it is possible to appreciate how the decision of ‘who gets to eat’ might be temporally regulated. Returning to the example of HPV infection in the skin, it is clear that the LCs get to ‘eat’ early. And once it had captured an apoptotic cell, it would exit the skin, traffic to the draining lymph node and engage any HPV-specific T cell it might encounter. In this way, an immune response would be initiated. Meanwhile, back in the peripheral tissue, as a result of an inflammatory signal, macrophages will be recruited from the dermis and peripheral blood. Given that the LC has already captured the needed antigen for

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the activation of T cells, it is now appropriate for the macrophage to enter the tissue and clear the site of inflammation. The presence of apoptotic cells in the context of a danger signal leads to immunosuppression – all immune responses must be regulated and the macrophage may offer the requisite negative feedback to regulate the immune stimulation. Indeed, the two cells work in concert to both defend the infected tissue as well as clear the inflammatory site for future tissue remodeling. This same interplay is apparent in the example of SLE, as it is likely the skewing toward DC engulfment of the apoptotic cell that triggers the activation of self-reactive T cells. And in the case of tumor-mediated immunosuppression, the balance tips in the other direction. By sequestering the apoptotic tumor cells, the macrophage prevents the DC from capturing antigen, in essence, preventing the activation of tumor-specific CTLs, thus offering one mechanism by which the tumor manages to evade the immune system. With a better understanding of the complexities of this apoptosis-dependent pathway for the generation of MHC I/peptide complexes, it may be possible to develop DC-based therapies for the activation or tolerization of antigen-specific T cells.

ACKNOWLEDGMENTS I would like to thank Drs Sebastian Amigorena, Monique Kleijmeer, Nathalie Blachere, Raymond Birge, Drar Mevorach, Roy Silverstein and Ralph Steinman for helpful conversations and fruitful collaborations. I also wish to recognize Drs Nina Bhardwaj and Robert Darnell who were integral contributors to many of the experiments and hypotheses outlined in this chapter.

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PLATE 44.1 Apoptotic cells are capable of transferring antigen to dendritic cells for activation of CD8 and CD4 T cells. DCs are capable of acquiring antigen from apoptotic cells, cross-presenting antigen on class I and class II MHC, and inducting antigen-specific CD8 and CD4 T cell responses. The DC / CD4 T cell interaction is likely a cognate, antigen-specific engagement. This engagement may be critical for the cross-priming of the CD8 T cell. Possibly, the absence of CD4 T cell help leads to the cross-tolerization of the CD8 T cell.

PLATE 44.2 Receptor utilization may predict the immunologic outcome of antigen cross-presentation. DCs utilize various receptors for the recognition and internalization of apoptotic cells. It is intriguing to consider how receptor utilization may influence the immunologic outcome of antigen cross-presentation. Possibly, capture via the FcR results in DC activation and the cross-priming of antigen-specific T cells. Complement receptor (CR) utilization, in contrast, may result in the tolerization of T cells. With respect to the utilization of the αvβ5 / CD36 receptor complex, other immunomodulatory signals will determine the immunologic outcome. Interestingly, the soluble factors that bridge these receptors to the apoptotic cells are distinct, thus suggesting that the microenvironment in which apoptotic cell death occurs may determine receptor utilization and the subsequent immune response.

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PLATE 44.3 Schematic outline of potential pathways for trafficking of antigen in the apoptosis-dependant exogenous pathway. Shown here is a schematic representation of how antigen from an apoptotic cell may be trafficked for the generation of MHC I / peptide complexes. Possibilities include (i) a recycling pathway in which peptide epitopes are loaded on MHC I within the endocytic compartment; (ii) an endosome to trans-golgi pathway that permits ‘preprocessed’ antigen to access the ER by retrograde transport; and (iii) a DC-restricted transporter allowing antigen to exit the endosome and enter the cytosol.

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PLATE 44.4 Dendritic cells cross-presenting antigen derived from apoptotic cells may initiate T cell tolerance or immunity. This illustration outlines the proposed in vivo pathways by which immature DCs may capture antigen from dying cells in the periphery and induce tolerization (a) or the activation of CD8 T cells (b). The tolerogenic response may depend on the microenvironment in the lymph node and / or on the absence of antigen-specific CD4 T cell help (a). In contrast, the ability to activate CTLs depends on the presence of a maturation signal and the presence of antigen-specific CD4 T cell help. Once activated, these CD8 T cells may return to the site of infection and lyse virally-infected cells or cells on their way toward malignant transformation (b).

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45 Dendritic cells and the biology of parasitism Karen A. Norris School of Medicine, Pittsburgh, Pennsylvania, USA

Thunder is good, thunder is impressive; but it is lightning that does the work. Mark Twain

INTRODUCTION

refer to microorganisms, particularly protozoa and helminths, that require a host for some aspect of their survival and propagation. The scientific interest in parasites mainly derives from the enormous medical and economic impact of parasitic infections worldwide and the biological complexity of the host-parasite interaction. One important aspect of many parasitic infections is an early and sometimes persistent immunosuppression, which may lead to chronic persistence of the parasites in the mammalian host. A better understanding of the immune responses to parasites and the pathology associated with failure to clear the infection may eventually lead to successful vaccine development or immunotherapy for these devastating diseases.

The results of a large body of research in recent years have established the importance of the DC as a key cell heralding the presence of microbial pathogens in the tissues and directing the type and extent of the acquired immune responses. It is this early interaction as well as priming and memory T-cell activation, that places DCs as the ‘lightning’ that sparks the interaction between innate and acquired immunity. Only recently has the interaction between protozoan parasites and DCs been investigated. Not surprisingly, these complex microorganisms have developed unique ways to thwart the immune responses at the early stages of antigen presentation and lymphocyte priming. The focus of this chapter will be three protozoan parasites and their interaction with DCs: Leishmania sp., Trypanosoma cruzi and Plasmodium falciparum. The word ‘parasite’ is defined as one who exploits the hospitality of the rich and earns welcome by flattery. In scientific terms, parasites Dendritic Cells: Biology and Clinical Applications ISBN 0–12–455851–8

LEISHMANIA Leismania species cause a complex spectrum of diseases that include limited cutaneous

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Copyright © 2001 Academic Press. All rights of reproduction in any form reserved.

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Leishmaniasis, a more extensive mucocutaneous, and a systemic, life-threatening disease, kala azar (visceral Leishmaniasis). The broad range of clinical manifestations is due in part to differences in pathogenic characteristics among different Leishmania species, as well as differences in host immune responses. Leishmania sp. are transmitted by the bite of phlebotomine insects (sandflies) which have a wide distribution in the tropics and subtropics. The agent of visceral leishmaniasis is L. donovani with subspecies identified by geographic regions. Several distinct species and subspecies are responsible for cutaneous leishmaniasis, and they also have specific geographic distribution. Old World species include L. tropica, L. major and New World species are L. mexicana and L. braziliensis. In addition to the clinical importance of leishmaniasis, experimental murine leishmaniasis has been a particularly important model of host–pathogen interaction that has provided much information regarding the development and significance of T-helper dichotomy. In experimental murine models of leishmaniasis, disease susceptibility or resistance has been clearly shown to be associated with the development of a T helper 1 (TH1) or a T helper 2 (TH2) response, respectively (for review see Reiner and Locksley, 1995). The transmission of Leishmania occurs via the bite of sandflies and infectious metacyclic promastigote forms initiate infection of phagocytic cells. Parasites replicate intracellularly as amastigotes and sustain the infection when the cell ruptures and the released parasites invade neighboring phagocytic cells, particularly macrophages, monocytes and Kupffer cells. Several studies have established that Leishmania are present in dermal DCs and epidermal Langerhans cells (LCs) taken from cutaneous lesions in human leishmaniasis (El Hassan et al., 1995; Isaza et al., 1996) and in experimental murine cutaneous leishmaniasis (Blank et al., 1993; Moll et al., 1993; Von Stebut et al., 1998; Konecny et al., 1999; Marovich et al., 2000). In addition, a number of groups have shown that metacyclic promastigotes can readily infect human (Marovich et al., 2000) and murine DCs,

in vitro (Von Stebut et al., 1998; Konecny et al., 1999). Likewise, amastigotes are capable of infecting and replicating within fetal skinderived DCs (FSDDC) (Von Stebut et al., 1998) and splenic DCs (Konecny et al., 1999). Experimental infection of DCs with Leishmania promastigotes and amastigotes results in increased surface expression of CD40, CD86 and HLA-DR, indicating the initiation of the maturation process (Von Stebut et al., 1998; Marovich et al., 2000). In contrast, other studies using bulk murine FSDDC (Von Stebut et al., 1998) and spleen-derived DCs did not detect significant surface changes in expression when exposed to metacyclic promastigotes (Konecny et al., 1999). These discrepancies may reflect different culture conditions of both parasites and DCs, as well as the likelihood that with low infection rates, changes in expression of cell surface markers may be too small to detect in bulk cultures and must be considered at a cellular level. Indeed, although Marovich et al. reported that L. major-infected human DCs demonstrated a mature phenotype, based on surface expression of CD40, CD86 and HLA-DR, they were unable to detect production of IL-12p70, IL-1b, IL-6, IL-10 and TNFα in culture supernatants (Marovich et al., 2000). In contrast, single-cell analysis using intracellular cytokine staining and flow cytometry revealed that L. majorinfected DCs are a major source of IL-12p70 in a CD40/CD40 ligand-dependent process (Marovich et al., 2000). To address the question of whether infection with Leishmania impaired the antigen processing and presentation of parasite antigens to T cells, Marovich et al. recovered monocytederived DCs from patients infected with L. major, and exposed the cells to L. major promastigotes in vitro. When the infected DCs were subsequently exposed to autologous T cells, significant proliferation of the T cells was observed, along with increased IFNγ production, compared to uninfected DCs (Marovich et al., 2000). Interestingly, a number of studies have shown that macrophage IL-12 production is inhibited by infection with L. major (Kaye et al., 1994; Carrera et al., 1996; Belkaid et al., 1998;

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Weinheber et al., 1998), although infection of mice with L. major clearly drives protective IL-12 and IFNγ responses in vivo (Vieira et al., 1994). In vitro studies point to promastigote-infected DCs as a major contributor of IL-12 production provided there is CD40 ligand co-stimulation (Marovich et al., 2000). Taken together, these results indicate that DCs are the likely site of initiation of infection of the vertebrate host with Leishmania, and that upon infection, the cells undergo maturation and are a significant source of early IL-12 production. Furthermore, antigenic processing and presentation functions of infected human DCs are apparently unimpaired. This situation is in contrast to the results of studies with the closely related protozoan parasite, Trypanosoma cruzi and with Plasmodium falciparum (discussed below), which apparently alter the key functions of DCs as a potential immune evasion strategy.

TRYPANOSOMA CRUZI Trypanosoma cruzi, the causative agent of Chagas’ disease, is a protozoan parasite that currently infects nearly 20 million people in Latin America (Kirchhoff, 1993). The parasite is transmitted to the human host by bloodsucking reduviid bugs and blood transfusion from infected individuals. Infection begins with the introduction of parasites into the bite wound or direct infection of mucous membranes by trypomastigotes. Trypomastigotes infect tissue macrophages and DCs, and replicate freely in the cytoplasm as intracellular amastigotes. After several rounds of replication, amastigotes convert to trypomastigotes that are released when the cell ruptures. Trypomastigotes gain access to the bloodstream where they disseminate, and eventually target heart muscle cells, tissue macrophages and autonomic neurons of the digestive system. In humans and in experimental murine models, the initial acute stage of infection is accompanied by profound immunologic disturbances, with eventual control of parasitemia several weeks after infection. By 1 to 2 months after infection, parasites are

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present at very low levels in the bloodstream and in the tissues, however, they are rarely, if ever completely cleared by the host. The life-long infection results in an asymptomatic state in most individuals, although approximately 30% of chronically infected patients progress to symptomatic disease with increased inflammatory damage of the heart, digestive, and/or peripheral nervous systems. Experimental murine models of Chagas’ disease have revealed that early control of T. cruzi infection depends in part upon (IFNγ) produced by NK cells and T cells (DosReis, 1997). Parasites multiply readily in resting macrophages, but are killed by IFNγ-activated macrophages (Gazzinelli et al., 1992; Munoz-Fernandez et al., 1992). IFNγ is also a key cytokine for the induction of an adaptive response to intracellular pathogens such as T. cruzi. The role of DCs has only recently been investigated in the context of T. cruzi infection. Van Overtvelt and colleagues have shown that T. cruzi trypomastigotes infect and replicate within human DCs, and that shortly after entry into the cell, the parasites escape the phagocytic vacuole and replicate in the cell cytoplasm cells (Van Overtvelt et al., 1999). These observations are consistent with the infection of human and murine macrophages by T. cruzi, where it has been shown that parasites replicate in resting macrophages (de Araujo-Jorge, 1989). In contrast to other pathogens such as Mycobacterium tuberculosis, Borrelia burgdorferi and others (Guzman et al., 1994; Filgueira et al., 1996; Ghanekar et al., 1996; Henderson et al., 1997), infection of immature DCs with T. cruzi does not stimulate their maturation (Van Overtvelt et al., 1999). In fact, a decrease in basal levels of IL-12 and TNFα was observed in T. cruzi-infected immature DCs (Van Overtvelt et al., 1999). In addition, LPS-induced DC maturation was significantly inhibited by T. cruzi infection as IL-12, IL-6 and TNFα levels were all reduced compared to uninfected, LPS-treated DCs. In contrast to the dramatic reduction in IL-12 and TNFα production, infection of DCs with T. cruzi did not have an appreciable effect on basal levels of HLA-DR, CD40, CD54, CD80 and CD86, all of

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which are involved in the antigen-presenting functions of DCs. The LPS-induced upregulation of HLA-DR and CD40 was significantly reduced, implicating infection with diminished maturation of DCs and antigen-presenting capabilities; however, this was not directly tested (Van Overtvelt et al., 1999). Similarly, infection of macrophages with T. cruzi results in defective MHC class II presentation, perhaps suggesting a common pathway of inference with antigenprocessing and presentation in the two cell types (La Flamme et al., 1997). Most interestingly, in experiments conducted in trans-wells, Van Overtvelt and colleagues found soluble factors derived from T. cruzi trypomastigotes that were capable of similar inhibitory effects on DC cytokine production and maturation as infection with live parasites (Van Overtvelt et al., 1999). The inhibitory factor(s), was derived from spent media of T. cruzi trypomastigote cultures and has been shown to have profound T cell inhibitory activity, although the specific component responsible for the biologic activity has not been identified. Preliminary evidence suggests that the biologic activity found in culture supernatants may be a small molecular weight, heat stable factor (Van Overtvelt et al., 1999). These results are similar to the observation of a T. cruzi-derived immunosuppressive factor that was found to inhibit Tcell proliferation and IL-2 receptor expression on human T lymphocytes (Kierszenbaum et al., 1990; Kierszenbaum et al., 1994; Kierszenbaum et al., 1998). Downregulation of T-cell proliferation has also been associated with T. cruziderived glycosylinositolphospholipids (Gomes et al., 1996), thus suggesting that the parasites produce multiple factors that may result in immune dysregulation of T-cell activation. Although it remains to be shown whether T. cruzi infection of DCs affects T-cell priming functions, the results of Van Overtvelt and colleagues suggest that T. cruzi can infect and replicate within cells that are critical to the generation of protective immunity, and alter the expression of key cytokines involved in the protective immune response: IL-12 and TNFα (Aliberti et al., 1996; Hunter et al., 1996; DosReis,

1997). In most cases of human Chagas’ disease, the infection is controlled during the acute phase, although parasites persist for the life time of the host. The impairment of DC function by parasite infection or parasite-derived factors probably contributes to the sophisticated immune evasion mechanisms associated with the persistence of the parasite within the human host, thus leading to the pathology of the chronic disease state (Tarleton and Zhang, 1999).

PLASMODIUM FALCIPARUM Similar to T. cruzi, Plasmodium falciparum also inhibits DC maturation (Urban et al., 1999). Plasmodium sp. are the etiologic agents of malaria, with P. falciparum responsible for most of the morbidity and nearly all of the mortality associated with the disease. Malaria is responsible for approximately 1.5 to 2 million deaths per year with the majority of deaths occurring in Sub-Saharan Africa in children under 5 years of age. As with Leishmania and T. cruzi, P. falciparum is transmitted to the human host by the bite of an insect, in this case the Anopheles mosquito. Parasites enter the host when the mosquito takes a bloodmeal and the sporozoite stage quickly reaches the liver, where it enters and replicates within hepatocytes. Merozoite stage parasites are released from hepatocytes and invade and replicate with erythrocytes. During the erythrocytic stage, parasites express antigens on the surface of erythrocytes that mediate adherence of the infected erythrocytes to endothelial cells. Adherent, infected erythrocytes are sequestered in capillary beds, thus avoiding splenic clearance. Specific immunity to Plasmodium has been attributed to cytotoxic T cells directed against the liver-stage parasites, and antibodies directed against antigens associated with the erythrocytic stage (Miller et al., 1994). Although antimalaria immunity develops slowly during childhood, protection is poorly developed and generally does not protect against symptomatic disease. In addition, malaria induces profound immune dysregulation and suppression during

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REFERENCES

clinical disease (Williamson and Greenwood, 1978; Scorza et al., 1999). Pathogenesis is associated with adherence of infected erythrocytes to endothelial cells via Plasmodium-encoded glycoproteins (PfEMP-1) that are expressed on the red cell surface (Miller et al., 1994). The PfEMP-1 protein undergoes antigenic variation and thus contributes to immune evasion associated with malaria (Smith et al., 1995). Infected erythrocytes adhere to host cells via a number of receptors, particularly thrombospondin, CD36 and CD54 (ICAM-1). Since both CD36 and CD54 are expressed on immature DCs (Zhou and Tedder, 1995), Urban and colleagues addressed the issue of adherence of infected erythrocytes to DCs (Urban et al., 1999). In these studies, RBCs infected with parasite lines expressing PfEMP-1 adhered to immature DCs and inhibited LPSinduced maturation. In addition, erythrocytes infected with a P. falciparum line defective in PfEMP-1 expression did not adhere to DCs and did not inhibit maturation. No difference in the proportion of DCs undergoing apoptosis or necrosis was observed between cells treated with infected or uninfected erythrocytes, which indicated that the inhibition of maturation was not due to direct parasite-induced cytopathic effects. In addition, DCs exposed to parasiteinfected erythrocytes were shown to be poor stimulators of T-cell responses in vitro (Urban et al., 1999), indicating that the interaction of parasite-infected cells alters DC maturation and antigen presentation and probably contributes to the immunosuppression observed clinically. The studies of the interaction of P. falciparum and T. cruzi are prime examples of the complex interaction between host and parasite, with immune responses impaired at a key point: DC maturation and priming of the T-cell responses. Clearly, these infections result in immune dysregulation at multiple levels, but the demonstration that specific parasite products can mediate DC impairment could provide specific targets for vaccine development and novel immunotherapeutics.

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2000 (cont.) Wilson, C.C., Martin, K.S., Campbell, T.B. et al. Peripheral blood CD34+ cells from HIV-infected donors differentiate in vitro into functional dendritic cells free of replicating HIV-1. Abstract B139 in Program 39.14. American Association of Immunologists Meeting, Seattle, Washington, May 12–16, 2000. [41] Wolfers, J., Andr‚ F., Ruegg, C. et al. DC derived-exosomes are effective cancer vaccines: preclinical data towards a clinical trial. Proc. Amer. Assoc. Cancer Res. 41 (Abstr. 5556) 874. [13] Womer, K.L., Vella, J.P. and Sayegh, M.H. Chronic allograft dysfunction: mechanisms and new approaches to therapy. Semin. Nephrol. 20, 126–147. [32] Woods, G.M., Doherty, K.V., Malley, R.C., Rist, M.J. and Muller, H.K. Carcinogen-modified dendritic cells induce immunosuppression by incomplete T-cell activation resulting from impaired antigen uptake and reduced CD86 expression. Immunology 99, 16–22. [5] Wykes, M. and MacPherson, G. Dendritic cell–B-cell interaction: dendritic cells provide B cells with CD40-independent proliferation signals and CD40-dependent survival signals. Immunology 100, 1–3. [6] Wykes, M., MacDonald, K.P.A., Quin, R., Hart, D.N.J. and McGuckin, M.A. Activated dendritic cells express MUC1. 6th International Symposium on Dendritic Cells. Port Douglas. [8] Yang, S., Kittlesen, D., Slinghuff, C.L., Jr., Vervaert, C.E., Seigler, H.F. and Darrow, T.L. Dendritic cells infected with vaccinia vector carrying the human gp100 gene simultaneously present multiple specificities and elicit high-affinity T cells reactive to multiple epitopes and restricted by HLA-A2 and -A3. J. Immunol. 164, 4204–4211. [43] Yang, S., Linette, G.P., Longerich, S., Roberts, B.L. and Haluska, F.G. HLA-A2.1/K(b) transgenic murine dendritic cells transduced with adenovirus encoding human gp100 process the same A2.1-restricted peptide epitopes as human antigen-presenting cells and elicit A2.1-restricted peptide specific CTL. Cell Immunol. 204, 29–37. [43] Yokoyama, H., Kuwao, S., Kohno, K., Suzuki, K., Kameya, T. and Izumi, T. Cardiac dendritic cells and acute myocarditis in the human heart. Jpn. Circ. J. 64, 57–64. [26]

Yrlid, B.U. and Wick, M.J. Salmonella-induced apoptosis of infected macrophages results in presentation of a bacteria-encoded antigen after uptake by bystander dendritic cells. J. Exp. Med. 191, 613–624. [20, 44] Zhang, Y., Chirmule, N., Gao, G.P. and Wilson. J. CD40 ligand-dependent activation of cytotoxic T lymphocytes by adeno-associated virus vectors in vivo: role of immature dendritic cells. J. Virol. 74, 8003–8010. [43] Zhang, Y., Wang, Y., Ogata, M., Hashimoto, S., Onai, N. and Matsushima, K. Development of dendritic cells in vitro from murine fetal liver-derived lineage phenotype-negative c-kit(+) hematopoietic progenitor cells. Blood 95, 138–146. [7] Zhong, L., Granelli-Piperno, A., Pope, M. et al. Presentation of SIVgag to monkey T cells using dendritic cells transfected with a recombinant adenovirus. Eur. J. Immunol. 30, 3281–3290. [35] Zlotnik A. and Yoshie, O. Chemokines, a new classification system and their role in immunity. Immunity 12, 121–127. [5, 37] 2001 Darnell, J.C., Albert, M.L. and Darnell, R.D. cdr2, a target antigen of naturally occurring human tumor immunity, is widely expressed in gynecologic tumors. Cancer Res. 60, 21–36. [44] Kleijmeer, M., Escola, J.M., Uydtehaag, F.G.C.M. et al. MHC class I and II molecules co-localize to endocytic compartments and exosomes of dendritic cells and B-lymphocytes. in press. [44] Kurts, C., Cannarile, M., Klebba, I. and Brocker, T. Cutting edge: dendritic cells are sufficient to crosspresent self-antigens to CD8 T cells in vivo. J. Immunol. 166, 1439–1442. [Preface] Moldenhauer, A., Nociari, M., Rafii, S. and Moore, M.A.S. Generation of human dendritic cells from CD34+ cells in coculture with tumor necrosis factor alpha-stimulated endothelial cells. submitted. [21] Tone, M., Tone, Y., Fairchild, P.J., Wykes, M. and Waldmann, H. Regulation of CD40 function by its isoforms generated through alternative splicing. Proc. Natl Acad. Sci. USA 98, 1751–1756. [Preface]

ANNOTATED REFERENCES BY YEAR

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AG see arabinogalactan ageing, wound healing and 543–4 AIA see antigen-induced arthritis Ailos gene, functions 122 Alexa dyes, light microscopy 236–7 allergy 523–38 allergen sensitisation, T cells 525–6 allergic disease, developmental progression 534–5 allergic responses, early/late phase 523–4 atopic allergic disease, T cell activation 525–8 developmental progression 534–5 future directions 535 type 2 T helper cells 524–5 allografts acute rejection 442–5 CD40 ligation 446 cellular rejection 442–5 chronic rejection 445–8 DC-lymphocyte interactions 446–7 DCs therapeutic application 600–1 hematolymphoid cells 440, 442–8 liver, irradiation 444–5 liver sinusoidal endothelial cells 444 MHC class II antigens 441 myeloid DC transduction 446 normal physiology, re-establishing 448–9 organ-based immunity, overview 440–2 rejection, DC-based therapy 587–607 solid organ, DCs in rejection and acceptance 439–57 survival prolongation, by DCs 600–2 T cells, allogeneic 446 therapy, DC-based 587–607 tolerance induction 449–53 _v_5/CD36 receptor complex, apoptotic cells 633 ALPS see autoimmune lymphoproliferative syndrome

4–1BBB ligation 60–1 55kDa actin-bundling protein, arterial wall 549 80/1, dendritic cell line 167–72 AAV see adeno-associated virus ACAID see anterior chamber-associated immune deviation actin, phagocytosis 213–16 activated T-cell apoptosis, induction 592–3 activation B cells 333 DCs 196, 207 autoimmune diseases 461–4 chemokines 207 T cells 21–8 activation markers, LCs 303 adaptive immune responses 507–14 DCs as sentinels 473–4 influenza virus 507–11 adaptive immunity, DCs 252–3, 473–4, 507–14 adeno-associated virus (AAV) gene transfer 610, 613 vectors, gene transfer to DCs 618–19 adenoviral vectors, gene transfer to DCs 616–18 adenoviruses 519–20 DC interactions 519–20 gene transfer 610, 612 adherent cells, progenitors 87–8 adhesion molecules DC populations 107–8 DC-T cells interaction 319–20 LCs 38, 303 adhesion and signalling, DCs 108–9 adjuvants, immune, DCs v monocyte/macrophages 271–2 adoptive immunotherapy, DCs for 93–4 adrenergic agonists, IL-12 production suppression 534

771

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772

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Page 772

INDEX

alveolar macrophages (AMs), and DCs 321–2 AMLR see autologous mixed lymphocyte response AMs see alveolar macrophages anergy T-cell, induction 592 tolerance induction method 588 angiogenesis, wound healing 542 anterior chamber-associated immune deviation (ACAID) 392, 404 anti-HLA II antibodies 428 anti-influenza immune responses, DCs induction 508–11 antigen exogenous, cross-presentation 627–44 oral 331–2 antigen capture LCs 37, 304–5 regulation 151–63 antigen concentration, apoptotic cells 630–1 antigen dose, and peptide affinity 532 antigen load, peripheral tolerance 593–4 antigen packaging 630–2 antigen persistence, autoimmune diseases 461–2 antigen presentation CD1 molecules 160 packaging the dead 630–2 antigen presenting functions, DCs v microglia 378–9 antigen presenting molecules, LCs 303, 304–5 antigen processing dendritic cell lines 169–70 Leishmania 646–7 subsets, DCs 160–1 antigen transport, and reproductive tract DCs 414 antigen uptake CD8_ DCs 362 intestinal DCs 330–1 monocyte/macrophage interaction 269–70 antigen uptake/processing, LCs 303 antigen-coated particles, phagocytosis 153 antigen-induced arthritis (AIA) 461–2, 463 antigen-presentation assays, DCs endocytic capacities 222–3 antigen-presenting cells (APCs) EBV interactions 515–16 endocytic/exocytic pathways 180 functions, DCs 97–8 HSV interactions 516–17 microglia as 396–7 vaccinia virus interactions 517–18 antigen-processing capabilities DCs 270 macrophages 270 antigen-pulsed DCs antigen transfer 293 lymph nodes 144, 293 antigen-responding T cells, augmented virus growth 495–6

antigenic complexity, bacteria 475 antigenic peptide-MHC complexes, immune response regulation 52, 53, 55 antigens cell surface, DCs 99–100 cell surface loading 158 from the dead 628–30 presentation 106–7 processing 106–7 T cell responses 294 transfer 290–6 transfer in vitro 293–4 transfer in vivo 294–6 uptake 106–7 uptake mechanisms 151 aorta, DC clustering 551 APCs see antigen-presenting cells apoptotic cells _v_5/CD36 receptor complex 633 antigen concentration 630–1 cross-presentation pathologic settings 639 physiologic settings 639 immature DCs 634 immunologic outcomes 632–4 LA 633 macrophage capture 637–8 phagocytosis 627–44 activated T-cell 592–3 antigens from the dead 628–30 eating the dead 632–4 introducing the dead 627–8 lessons from the dead 639–42 outcomes 637 packaging the dead 630–2 presenting the dead 637–9 processing the dead 634–7 receptor utilization 632–3 surface ligands 632 TSP-1 633 TUNEL-positive 630–1 virus-infected, CD8+ T cells 509–10 apoptotic debris, v necrotic debris 270–1 aquaporins, DCs fluid uptake, macropinocytosis 152 arabinogalactan (AG), bacterial cell walls 475 arsenical skin tumors 427 arterial wall, DCs identification 547–9 ascites, malignant 434 macrophages from 85, 92 astrocytes, antigen-specific T cell responses 378 atherosclerosis 547–57 animal models 555 atherosclerotic lesions 552–4 clustering of DCs 551 DC clustering, aorta 551 DCs identification, arterial wall 547–50 distribution of DCs 552–4

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Page 773

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atherosclerotic lesions (cont.) foam cells 555 heterogeneity of DCs 554–5 immunohistochemical DCs identification 549–50 vascular DCs heterogeneity 554–5 vascular DCs origin 550–1 vascular pathology 547 atherosclerotic lesions DCs in 552–4 T cells 552–4 atopic allergic disease, T cell activation 525–8 attenuated vaccines, DCs involvement 500–1 augmented virus growth, antigen-responding T cells 495–6 autocrine pathways, DCs 296–7 autoimmune, ocular 402–4 autoimmune diseases antigen persistence 461–2 CNS 379–81 cryptic self-epitopes 464 DCs activation 461–4 DCs priming 461–4 homeostatic cytokines 464–5 IL-10 465 immunostimulatory CPG motifs 464 polygenicity 460–1 regulatory T cells 465–7 subdominant self-epitopes 464 systemic autoimmunity 462–3 therapy, DC-based 587–607 tolerance maintenance 464–7 viral infections 463–4 autoimmune encephalomyelitis (EAE) 373–4, 379–81, 382–4 autoimmune lymphoproliferative syndrome (ALPS) 460 self-antigen persistent priming 467 autoimmune responses, perpetuation of initial 467–9 autoimmune uveoretinitis (EAU) 394, 403–4, 406 autoimmunity DCs and 459–71 systemic 462–3 autoimmunity amplification, B cells 468–9 autoimmunity trigger, cross-priming, inappropriate 640 autologous mixed lymphocyte response (AMLR) 462 B7 family co-stimulatory/polarizing factors 55 costimulation 532 expression 57 ligands for CD28, CTLA-4, ICOS 54–7 B cell areas DC cell subsets 493–4 HIV-1/SIV infection 495 B cell functions, DC subsets 257–8 B cell non-Hodgkin’s lymphoma 562–3

773 B cells activated 255–6 activation 333 autoimmunity amplification 468–9 CD4+ T helper cells 258 clonal expansion downregulation 593 DC interactions 258–9 differentiation 256 and FDCs 31–2, 593 growth enhancement 255–6 IgA isotype switching 256–7 IL-12 256 immune response regulation 52–3 maturation, FDCs 143 OX40–OX40L interactions 259 priming 367 thymus 13–14 transmission EM 237–9 bacteria antigenic complexity 475 cell wall models 474 complexity 474–5 DC suface molecules 478–80 DCs interaction 473–86 evasion strategies, immune responses 474–5 growth, intracellular 480–2 survival, intracellular 480–2 bacteria-induced upregulation, co-stimulatory molecules 483–4 bacterial antigen, cross-presentation 628–30 bacterial infection, DCs 482–4 BAFF, DC modulatory factor 62 BALT see bronchus-associated lymphoid tissue basal cell tumors 427 BBB see blood-brain barrier _2 integrins 306 _-adrenergic receptor agonists, immunosuppressive activity 374–5 biomedical research, dendritic cell lines uses 172–5 biotin, labeling enhancement 236 Birbeck granules (BGs) arterial walls 548–9 LCH 432 LCs 36, 37, 38, 225–6, 269, 302, 350 bladder carcinoma 434 blood DC migration from 140–1, 275–7 DC migration via 138–40, 141 blood clotting 541 blood DCs 101–3 cord 88–90, 105 cultured 102–3 fresh monocyte subsets 102 peripheral, gradient enrichment 562–5 blood-brain barrier (BBB) 372–4, 393 blood-cerebrospinal fluid (CSF) barrier 373 blood-ocular barrier (BOB) 390, 391

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774

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Page 774

INDEX

BM-derived DCs, tolerogenic capacity 590 BOB see blood-ocular barrier bone marrow precursors, DCs 100–1 bone marrow transplantation, hematopoietic DC precursors 433 bone marrow-derived DCs, isolation, culture, propagation 78–9, 88 breast tumors 427 ‘bridge’ subsets, DCs 142 bronchial DCs, migration 318 bronchoalveolar tumors 427 bronchus-associated lymphoid tissue (BALT) 315–16, 318 c-kit ligand, DCs generation in vitro 188–90 C-type lectin receptors, receptor-mediated endocytosis 155–6 calcineurin-inhibiting drugs, allografts 445 calcitonin gene-related peptide (CGRP) 311 calcium ionophores, cytokine-independent DC differentiation 92–3 canarypox 518–19 DC interactions 518–19 cancer cervical 427 DCs in HPV-associated 415–16 clinical trials, DCs 561–71 head/neck, DC infiltration 426 identifying DCs in 425–37 carbocyanine dyes, light microscopy 236–7 Castleman’s disease 427 caveolar membrane system, potocytosis 217 CB1, dendritic cell line 167–72 CCR1, lung DCs 317 CCR5, lung DCs 317 CCR5–using HIV-1 isolates replication 489–90 CCR6 chemokine receptors 204–5 DC migration 267–8 defensins 205 CCR7 DC migration 267–8, 280 LCs 46 TCA4/SLC binding 125 CD1 family, DCs 99–100 CD1 molecules antigen presentation 160 trafficking 160 CD1a+ cells CNS 382 emigration and maturation 44–7 CD1aCD3CD4+ thymic pDCs, developmental origin 17–18 CD2–LFA-3 (CD58) interaction 58 CD2+ cells, progenitors 88 CD4, and HIV-1 infection 490–2 CD4+ T cells, HIV-1 infection 491–3

CD4+ T helper cells B cells 258 cellular immunity to HIV-1 576 chronic allergic inflammatory response 524–8 cross-priming CTLs 638 fate 576–7 tumor immunity 641 CD8_ DCs 121–2, 123 antigen uptake 362 cytokine production 363 liver 340–2, 344 lymph nodes 361–3 spleen 361–3 T cells priming 363 CD8+ T cells cross-tolerization 638–9 HIV-1 resistance mediation 499–500 influenza antigen presentation 509–10, 511 CD11c+ cells, DCs subsets 363–4 CD11c expression, DC precursors defined by 87 CD11c cells cord blood 89–90 innate immune responses 506–7 CD14, phagocytosis mediation 153 CD14+ cells, DCs subsets 363–4 CD14+ monocytes, DC migration 276–7 CD15s, skin DCs 305–6, 307 CD28, ligation, B7 family members 54–7 CD28–mediated signalling, CXCR5 54 CD34 progenitor-derived DCs, cancer clinical trials 565–9 CD34+ cells bone-marrow 88 cord blood 88–9 hematopoietic precursors, LCs 40–2 peripherally circulating progenitors 86 CD34+ progenitor-derived DCs, for immunotherapeutic purposes 580 CD34+CD1a- thymic precursors 19 CD36 avb5/CD36 receptor complex, apoptotic cells 633 phagocytosis mediation 153 CD40 chemokine production by DCs 207 lung DCs 316, 317 T. cruzi infected DCs 648 CD40 ligation, allografts 446 CD40–CD40L interaction cytokine-independent differentiation 92–3 lung DCs 319–20 maturation signal, DCs 61 CD40–deficient BM DCs, tolerogenic capacity 590 CD40L DC activation 196–7 IL-12 secretion 196 CD40L-activated DC1s/DC2s 22–3, 25

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Page 775

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CD41 effector cells, EBV cellular immune responses 515 CD44, DCs migration 309 CD68 428 CD70, DC modulatory factor 62 CD80, lung DCs 316, 317 CD81 effector cells, EBV, cellular immune responses 514–15 CD83, emigration and maturation 44–7 CD86, lung DCs 316 cell adhesion method, light microscopy preparation 234 cell biology, dendritic 119–29 cell contact-dependent activation, NK cells 250 cell surface antigens, DCs 99–100 cell surface loading, antigens 158 cell surface marker expression, characteristics 78 cell suspensions, cryosectioning 234 cellular debris, apoptotic v necrotic 270–1 cellular immunity, immune response component 53–4 cellular rejection, allografts 442–5 central nervous system (CNS) autoimmune diseases, DCs in 379–81 CD1a+ cells 382 DCs 371–87 compartmentalization 375–6 demyelination 382 HIV infection 381 immunosuppressive environment 374–5 infectious diseases 381–2 inflamed, DCs recruitment 382–4 inflammation 382 lymphatic drainage 374 parenchyma, DCs absence 375–6 pathology, DCs in 379–82 sentinels, microglia 377–9 central tolerance, DC-induced 589–91 cervical canal 412 cervical cancer 427 DCs in HPV-associated 415–16 cervical DCs, STDs 414–16 cervical intraepithelial neoplasia (CIN) 415 cervical lymph node DCs, tolerogenic capacity 590 CGRP see calcitonin gene-related peptide Chagas’ disease, biology of parasitism 647–8 chaperoning DC-lymphocyte interactions, transplantation 446–7 peptides 631–2, 635 character regulation, immune response 53–4, 55 checkpoints, developmental 4–7 chemoattractant receptors, DCs 108 chemokine receptors CCR6 204–5 DC-produced 134 expression, inflamed CNS 383 HIV infection 205

775 immature DCs 204–5 LCs 42–4, 303, 305 lymphocyte-produced 134 nonlymphoid tissues 204–5 responsiveness 204–5 resting DCs 204–5 switch paradigm 205–6 wound healing 542 chemokines DC activation 207 DC differentiation 207 DC recruitment, tumors 207–9 DC-derived, efferent responses 138 DC-produced 66–7, 134 DCs and 203–11 DCs secreting 144–5 expression, inflamed CNS 383 inflammatory 205–6 leukocyte recruitment 382–4 lymphocyte-produced 134 microglia 396 monocyte/macrophage interaction mediation 267–8 phenotypic characterization, DCs 110 production by DCs 206–7 responsiveness, MCP-1 205 role 136 wound healing 542 chemokines receptors, monocyte/macrophage interaction mediation 267–8 chemotaxis, leukocytes 132–3 chimerism, allograft acceptance/rejection 450–4 chlamydia 416 choroid DCs in 402–4 macrophages 398–9 choroid plexuses, DCs localization 376–7 chronic allergic inflammatory response, type 2 T helper cells 524–8 chronic myelogenous leukemia (CML) 166 chronic rejection, allografts 445–8 chronic venous insufficiency (CVI), wound healing 543 CIIVs, transport vesicles 225 ciliary body, macrophages 398–9 CIN see cervical intraepithelial neoplasia circumventricular organs (CVOs), BBB 372–3 CLA see cutaneous lymphocyte-associated protein classical germinal centers, FDCs 32 clathrin coat-mediated endocytosis 215 clinical studies, HIV-1 infection, DC therapies 582–4 clinical trials, DCs for cancer 561–71 clonal deletion, tolerance induction method 588 Clostridium, survival strategy 481 CML see chronic myelogenous leukemia CMRF-44 429 CMRF-44 mAb 99–100

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776

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Page 776

INDEX

CMRF-56 mAb 99–100 CMRF-58 mAb 100 CNS see central nervous system co-simulator-deficient DC progenitors, tolerogenic capacity 590 co-stimulation blockading, DCs therapeutic effect 600–1 co-stimulator-deficient DC progenitors, tolerogenic capacity 590 co-stimulatory factors DC-produced 55–6 immune response regulation 53, 55 co-stimulatory molecules 109 bacteria-induced upregulation 483–4 coagulation, blood 541 coiling phagocytosis mechanism 478 morphological appearance 479 colorectal cancer, clinical trials 564–5 colorimetric labeling methods, light microscopy 235–6 combination immunosuppressive therapy 601 complement (C’) receptors 218, 305 antigens 106–7 phagocytosis 151, 153, 214 confocal microscopy, fluorescence 236–7 constitutive chemokines 144–5 constitutive recruitment, DC progenitor cells 135 conventional phagocytosis, model 478, 480 cord blood DCs 88–90, 105 costimulation, B7 family 532 costimulatory molecule deficiency, tolerogenicity 594 costimulatory molecules DCs 362–3 expression, inhibition 599 LCs 38, 303 cross-presentation, exogenous antigen 627–44 cross-priming CTLs 634 DC endosomes and 225 inappropriate 640 T cells 631 cross-tolerization, CD8+ T cells 638–9 crosslinking, Fc_RI 528–30 crosspresentation, dying cells to T cells 506 cryosectioning, light microscopy preparation 233, 234 cryptic self-epitopes, autoimmune diseases 464 CTLA-4 ligation, B7 family members 54–7 lung DCs 319 T cell inhibition 54–7 CTLs see cytotoxic T lymphocytes culture guide, DCs 77–96 cultured DC-like cells, in vitro 105–6 cultured DCs, fluorescent immunolabeling 233–4 cutaneous lymphocyte-associated protein (CLA), skin DCs 305–6, 307

cutaneous nerves, immune-associated function 302 CVI see chronic venous insufficiency CVOs see circumventricular organs CXCR5 CD28–mediated signalling 54 FDCs 32 cyclosporine, allografts 445 cytokine administration, DC development 197–200 cytokine production bacterial infection 482–4 CD8_ DCs 363 cytokine profiles, dendritic cell lines 170–1 cytokine receptor profiles, dendritic cell lines 170–1 cytokine receptors LCs 303, 305 monocyte/macrophage interaction mediation 268 cytokine signals, DCs as recipients of 187–202 cytokine-independent differentiation, DCs 92–3 cytokines administration 197–200 cancer treatment 569–70 DC1 effector function 23–5 DC2 effector function 25–6 DCs activation regulation 194–7 DCs development regulation 359–61 DCs generation in vitro 188–90 DCs maturation regulation 194–7 DCs regulation 189–97 exogenous 192 expression, intracellular 23–5 gene transfer studies 192 homeostatic 464–5 immunosuppressive activity 374–5 inhibiting DC differentiation/maturation 196 knockout studies 191–2 LCs selective tissue-homing 42–4 microglia 396 monocyte/macrophage interaction mediation 268 phenotypic characterization, DCs 110 production, DCs 62–7 T cell activation 320 tolerogenic DCs propagation 596–9 transgenic studies 192 cytolytic activity, NK cells 248–50 cytoplasmic markers, phenotypic characterization, DCs 111–12 cytoskeletal markers, phenotypic characterization, DCs 111 cytosol, phagosome-to-cytosol pathway, MHC 1 pathway 637 cytosolic proteins, exosomes 183 cytospins, light microscopy preparation 233–4, 235 cytotoxic activity, NK cells 245–6 cytotoxic T lymphocytes (CTLs) 318 cellular immunity to HIV-1 574–5 cross-priming 634, 638 fate 576–7

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Page 777

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D1, dendritic cell line 167–72 D2SC/1, dendritic cell line 167–72, 173–4, 175 DC1 effector function cytokines 23–5 pathogens 23–5 DC1s, TH1 differentiation 22–3 DC1s/DC2s, CD40L-activated 22–3, 25 DC2 effector function cytokines 25–6 pathogens 25–6 DC function, mice 21–2 tolerogenic DCs 21–2 DC activation, intracellular pathways 484 DC cell subsets, B cell areas and T cell areas 493–4 DC clustering, aorta 551 DC effectors, tissue microenvironments 26 DC endosomes, and cross-priming 225 DC interactions, influenza virus 507 DC lineage LCs 5–9 mice 21–2 monocyte/macrophage lineage 263–74 myeloid-related 264–5 DC maturation cross-priming CTLs 638 parasitic inhibition 645–9 regulation, cytokines 194–7 DC therapies, clinical studies, HIV-1 infection 582–4 DC trafficking cancer clinical trials 570 endothelial cells 284–5, 393 DC types, thymus 18–19 DC-CK1 chemokine, DC-produced 66–7 DC-lymphocyte interactions allografts 446–7 chaperoning, transplantation 446–7 DC-mediated triggering, NK cells 246–50 DC-T cell mixtures, immunodeficiency virus growth 490–2 DC-T cells interaction, adhesion molecules 319–20 DCs see dendritic cells DDCs see dermal dendritic cells death-inducing ligands, expression 594–5 debris, apoptotic v necrotic 270–1 DEC-205 EAE 380 receptor-mediated endocytosis 156 DEC-205 receptor 217–18 defensins, CCR6 205 dendritic cell biology 119–29 dendritic cell lines antigen processing 169–70 biomedical research, uses in 172–5 cytokine profiles 170–1 cytokine receptor profiles 170–1 development 165–77

777 endocytic potential 169–70 establishment of stable 166–8 functional properties 168–72 growth factor requirement 169 growth factor-dependent 166–8 growth-regulatory factors 170 immunotherapies development 174–5 maturation 173–4 phenotypic properties 168–72 surface phenotypes 168–9 T cell-stimulatory capacity 171–2 testing 165–77 dendritic cell sarcomas 432 dendritic cell-regulated immunoregulation 51–74 dendritic cells (DCs) activation, cytokine 196 adaptive immunity 252–3, 473–4, 507–14 adhesion 107–8 adhesion and signalling 108–9 adoptive immunotherapy 93–4 in allergy 523–38 allografts 439–57 rejection therapy 587–607 alveolar macrophages 321–2 antigen presenting functions 378–9 antigen-processing capabilities 270 APC functions 97–8 atherosclerosis 547–57 autocrine pathways 296–7 autoimmune diseases therapy 587–607 autoimmunity 459–71 B cells differentiation 256 growth enhancement 255–6 interactions 258–9 bacteria interaction 473–86 bacterial infection 482–4 biochemical analyses 172–3 blood DCs 88–90, 101–3 bone marrow precursors 100–1 bone marrow-derived 78–9, 88 in cancer 425–37, 561–71 CD1 family 99–100 CD40–CD40L interaction 61 cell surface antigens 99–100 characterizing 429–30 chemoattractant receptors 108 chemokine production 206–7 and chemokines 203–11 CNS 371–87 cord blood 88–90, 105 costimulatory molecules 362–3 culture guide 77–96 cytokine production 62–7 cytokine signal recipients 187–202 DC interaction 289–98 defined 97–8

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Page 778

778 dendritic cells (cont.) development 358–61 and secondary lymphoid tissue 124–6 differentiation, cytokine-independent 92–3 differentiation pathways 98, 442 disorders 429–30 DC-related 430–4 endocytic capacities 220–3 and endothelium 275–87 eye 389–409 fetal liver-derived 82, 90 fetal thymus-derived 90 fetal tissue 90 FITC-labelled 139 from diseased tissue 106 functional criteria 97–8 functional phenotype 106–12 genetic engineering 602–4, 609–25 heart 349–51 hematopoietic precursors 3–11 HIV-1 disease synopsis 501–2 HIV-1 infection 487–504, 573–86 HIV-infected individuals, culturing from 94 homing 307–9 human, isolation, culture, propagation 82–94, 105 humoral response regulation 255–6 IFN_ production 248–50 imaging 231–42 immortalization, oncogenes 166 immune responses mucosal 257 regulation 366–7 immunisation targets, mucosal vaccines 484 immunostimulatory functions 67–9 in vitro cultured 105–6 innate immunity 252–3, 473–4, 506–7, 510 interdigitating 104–5 interstitial 103–4 intestinal 325–36 intestinal immune responses 325–6 isolation guide 77–96 kidney 347–9 liver 338–47 localization 131–49 long-term cultures 79 lung-derived 80 lymph node-derived 80, 90–1 lymph nodes 357–70 lymphoid 22, 145–6 lymphoid microenvironments 361–6 lymphoid tissue-derived 104–5 lymphoid-related 265 lymphotoxin ligands 123–4 lysis 78 macrophage interaction 321–2 macrophage-derived 84–5 markers 99–100

INDEX

maturation 58, 61, 68, 78–9, 132 bacterial infection 482–4 cross-priming CTLs 638 mechanisms 173–4 parasitic inhibition 645–9 stages 265–6 migration 98, 107–8, 131–49, 203, 307–9 lymphoid microenvironments 365–6 mobilization 131–49 modulation immunostimulatory functions 67–9 polarizing functions 67–9 modulatory factors, membrane-bound 54–62 molecular analyses 172–3 monocyte-derived 80–2, 82–3 mouse, isolation, culture, propagation 78–82, 105 mucosal immune responses 257 MV interactions 512 NK cell cytolytic activity 248–50 nonlymphoid tissue 103–4, 133–5 nonlymphoid tissue see nonlymphoid tissues ontogenic heterogeneity 358 ontogeny 306–9, 401–2 pancreas 351–3 paracrine effects 296–7 paradoxes, phenotypic 119–20 parenchymal organs 337–56 pathogen-modulated immunostimulatory and polarizing functions 67–9 peritoneal cell-derived 82 perivasculature localization 278–9 Peyer’s patch-derived 80 phenotype application 112 phenotype paradoxes 119–20 phenotypic characterization 97–117 plasticity 187, 368 polarizing factors 55–6 polarizing functions 67–9 propagation guide 77–96 regulation, cytokine 189–97 respiratory tract 315–23 retina 399–404 sex hormones and 413 short-term cultures 79 signal 1/2 carriers 52–3, 55 signal 3 carriers 53–4, 55 SIV infection 487–504 skin entrance 307–9 skin-derived 80, 91 soluble factors 533 sources 263–4 species issues 98 spleen 142–3, 357–70 spleen migration 138–40, 140–1 spleen-derived 79, 90–1, 126 subset paradoxes 119–20 subsets see subsets, DCs

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Page 779

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dendritic cells (cont.) surface molecules 478–80 surface phenotypes 100–6 switch recombination 256–7 T cell activation 318, 320–1, 588 T cell mutual activation 196–7 targeted knockouts 119–29 therapy, allograft rejection and autoimmune diseases 587–607 thymus 13–20, 365 thymus-derived 80 TNF_ 67–8, 78–9 tolerance and autoimmunity balance 460–1 tolerance induction 588–94 tonsil-derived 90–1, 104–5 tumor-derived 82, 91–2 uveal tract 399–404 VEGF 196, 264, 279 virus interactions 505–22 virus transmission 496–7 wound healing 539–45 see also Langerhans cells (LCs) dendrocytes, immune-associated function 302 dermal dendritic cells (DDCs) functional aspects 309–12 functional features 37–40 `gatekeeper’ function 40 identifying molecules 304 immune responses 35 immunostimulatory function 309–12 maturation function 309–12 phenotype 302–6 phenotypic features 37–40 dermal microvascular unit, immune-associated function 302 development, dendritic cell lines 165–77 development, DCs 358–61 checkpoints 4–7 lymphoid lineages 358–9 myeloid lineages 358–9 regulation, cytokines 359–61 and secondary lymphoid tissue 124–6 dextrans, endocytosis measurement 220 differentiation DCs chemokines 207 cytokine-independent 92–3 inhibition, DCs 196 pathways, DCs 98, 442 T cells 332–3 diseased tissue, DCs from 106 disorders DC-related dysfunction 432–4 proliferation 430–2 DMG36 multipotential leukemic cell line 166 DNA

779 naked DCs 311 gene transfer 611, 620–1 donor leukocytes 345 donor-derived DCs, transplantation tolerance 589 dyes, fluorescence imaging 236–7 dying cells to T cells, crosspresentation 506 dynamin-2, phagocytosis 214–15 E-cadherin 306, 307 EAE see autoimmune encephalomyelitis early stages DC development 14–16 T cell development 14–16 EAU see autoimmune uveoretinitis EBV see Epstein-Barr virus EBV-transformed B (EBV-B) lymphocytes 179, 184 efferent responses, role of DC-derived chemokines 138 ELC/MIP-3_ chemokine 144–5, 280 electron microscopy (EM) 237–9 DCs endocytic capacities 221 LHs 36 materials sources 242 electroporation CaPO4 mRNA, gene transfer 611 ELISPOT reactivity, melanoma 567–8 EM see electron microscopy emigration DC precursors 275–8 LCs 44–7 endocytic capacities, DCs 220–3 endocytic pathway, DC, specializations 224–6 endocytic potential, dendritic cell lines 169–70 endocytic receptors 217–19 complement receptor 218 DEC-205 receptor 217–18 FcR 218 mannose/_ glucan receptor 217 scavenger receptor 218–19 endocytosis antigen transfer 291 DC endocytic capacities 220–3 DC endocytic pathway specializations 224–6 developmental regulation 223 endocytic capacities, DCs 220–3 endocytic pathway specializations 224–6 endocytic receptors 217–19 measurement 219–20 phagocytosis 213–15 pinocytosis 215–16 potocytosis 216–17 receptor-mediated 154–7 studies 213–29 types 213–17 endogenously-processed self-antigen, tolerance and immunity 466 endometrial tumors 427

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780

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Page 780

INDEX

endothelial cells DC trafficking 284–5, 393 liver sinusoidal, allografts 444 lymphoid, immune-associated function 302 transendothelial migration, DCs 132, 279–84 endothelium, DCs and 275–87 DC precursor emigration 275–7 DC trafficking 284–5 perivascular DC localization 277–9 TEM 279–84 epithelia DCs mobilization 137 vulvo-vaginal 412 wound healing 542 epithelial sites DCs at 488–93 DCs migration 138–40 epitopes cryptic self-epitopes 464 pre-processed epitopes, MHC 1 635–6 subdominant self-epitopes 464 Epstein-Barr virus (EBV) 514–16 APC interactions 515–16 CD41 effector cells 515 CD81 effector cells 514–15 cellular immune responses 514–15 EBV-transformed B (EBV-B) lymphocytes 179, 184 host immune responses 514 esophageal tumors 427, 433–4 exogenous antigen, retrograde transport to ER 636–7 exogenous cytokine 192 exosomes cytosolic proteins 183 DC-derived 179–85 hsc73 183 MHC class I proteins 181–2 protein composition 182–3 protein structure model 183 tumor peptide-pulsed 180–1 experimental organ transplantation, DCs therapeutic application 600–1 eye DCs 389–409 DCs function regulation 404–5 DCs recruitment 405–6 immune cells entry 392–4 immune priveleged status 391–2 inflammation 405–6 macrophage populations 398 microglia 394–7 mononuclear cell recruitment 405–6 perivascular cells 397 retinal DCs 399–404 FACS (fluorescence-activated cell sorting), DC purification 78, 79, 80, 84

Fas ligand, DC modulatory factor 62 fascin, and cancers 429 Fc receptors antigens 106–7 LCs 303 phagocytosis 151, 214 receptor-mediated endocytosis 154–5 Fc-type phagocytosis 480 Fc_RI crosslinking 528–30 DC expression 528–9 disease chronicity mediation 528–9 emigration and maturation 44–7 expression regulation 531 function 529–30 IgE 528–30 IL-4 529 mast cells 529 MHC class II molecules 529 structure 528 FcR 218 immune complex uptake 151 FcR-associated phagocytosis 151, 214 FDCs see follicular dendritic cells female genital tract DCs 412–14 fetal tissue DCs 82, 90 liver 82, 90 thymus 90 fibroblasts, immune-associated function 302 FITC-labelled DCs 139 fixation methods, light microscopy 234–5 fixed tissues, identifying DCs in 426–9 flow cytometry DCs endocytic capacities 222 PBMC DCs 434 Flt-3 receptor mRNA, DC hematopoiesis 8 Flt-3L DC growth factor 198–9 DCs development regulation 360 DCs generation in vitro 188–90, 198–9 liver DCs 340 NK cells 247–8 tumor regression 247–8 fluoresceinated dextrans (F-DXs), endocytosis measurement 220 fluorescence confocal microscopy 236–7 fluorescence immunohistochemical labeling methods, light microscopy 235–6 fluorescence-activated cell sorting (FACS), DC purification 78, 79, 80, 84 fluorescent immunolabeling, cultured DCs 233–4 fluorochromes, light microscopy 236 FM 3–25, endocytosis measurement 219 foam cells, atherosclerosis 555 follicular dendritic cells (FDCs) 29–34 B cell maturation 143 B cells 31–2, 593

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follicular dendritic cells (FDCs) (cont.) in cancers 429 classical germinal centers 32 CXCR5 32 germinal center reactions 31–2 IL-6 32 IL-13R_1 expression 32–3 networks 30 origin 30–1 perspectives 33 production, LT 190–3 PrP-deficient mice 31 SCID mice 31 T cells 31–2 forced endocytosis, phagocytosis mechanism 478 fractalkine 207 free radicals, microglia 396 freezing, light microscopy preparation 232–3 FSDC, dendritic cell line 167–72 full chimerism, allograft acceptance/rejection 450–4 function regulation, DCs 404–5 functional criteria, DCs 97–8 functional heterogeneity 367–8 functional phenotype, DCs 106–12 G-CSF CD34+ cells mobilization 86 DCs development regulation 360–1 gangliosides, immunosuppressive activity 374–5 gastric tumors 427 gastrointestinal epithelia, DCs migration 140 GCDCs see germinal center DCs gelvatol, light microscopy preparation 236 gene gun, gene transfer 611, 620–1 gene transfer future prospects 621 methods, genetic engineering 609–14 non-viral 620–1 studies, cytokines 192 to DCs 614–20 genetic engineering DCs 609–25 gene transfer methods 609–14 tolerogenic DCs 602–4 genital tract see reproductive tract germinal center DCs (GCDCs) 31–2, 143, 258 GFP see green fluorescent protein glutaraldehyde fixation IEM 241 SEM 239–40 GM-CSF adenoviral vectors, gene transfer 618 DC maturation 67–8, 279 DCs development regulation 360 DCs generation in vitro 188–90, 268 DCs regulation 193 hematopoietic progenitor cells differentiation 199

781 IL-4 combined 200 immune response enhancement 199 LCs 36, 37–9, 41–2 RPE cells 405 tolerogenic DCs propagation 597–9 gradient enrichment, peripheral blood DCs 562–5 granulation phase, wound healing 540 granulocyte-macrophage colony-stimulating factor see GM-CSF green fluorescent protein (GFP) labelled DCs 317–18 phagocytosis 476 growth factor receptors 305 growth factor requirement, dendritic cell lines 169 growth factor-dependent dendritic cell lines 166–8 growth-regulatory factors, dendritic cell lines 170 HAART see highly active anitretroviral therapy Hashimoto’s thyroiditis 459 DC infiltration 468 self-antigen priming 467 head cancers, DC infiltration 426 heart DCs 349–51 disease 351 function 351 histological location 349–50 ontogenic development 350–1 phenotype 349–50 population dynamics 350 heat shock proteins (HSPs) DC maturation 68 peptide chaperoning 631–2, 635 helper T lymphocytes (HTL) cellular immunity to HIV-1 575–6 fate 576–7 hematolymphoid cells, allografts 440, 442–8 hematopoietic development, signals 7–9 hematopoietic precursors, DCs 3–11 hematopoietic progenitor cells (HPCs) DCs 4 differentiation, GM-CSF 199 myeloid lineage 358 hematopoietin/activin family, co-stimulatory/polarizing factors 55 hepatitis, viral 347 hepatoma 427 herpes 416 herpes simplex virus vectors, gene transfer to DCs 620 herpes simplex virus-1 (HSV-1) 516–17 APC interactions 516–17 DCs role in vivo 517 immune responses evasion 516 herpes simplex viruses, gene transfer 610, 612–13 herpes viruses, human 514–17 highly active anitretroviral therapy (HAART) 573, 576 clinical studies 583–4

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782

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INDEX

highly active anitretroviral therapy (cont.) immunologic recovery after 577 limitations 577–8 histamine, IL-12 production suppression 534 histiocytomas, non-Langerhans 431–2 histiocytosis, Langerhans cell 382, 430–1 histiocytosis X 322 HIV, DCs binding 418 HIV infection chemokine receptor expression 205 CNS 381 intestinal DCs 333–4 HIV transmission, reproductive tract 416–18 HIV-1 cellular immunity 574–8 FDC network 30 HIV-1 disease, DCs contribution synopsis 501–2 HIV-1 immunopathogenesis, DCs role 578–82 HIV-1 infection CD4 490–2 CD4+ T cells 491–3 cultured DCs function 579–81 DC therapies 573–86 clinical studies 582–4 DCs during 487–504 overview 574 HIV-1/SIV, sexual transmission 488–93 HIV-infected individuals, DCs culturing from 94 HIV-specific immunity, DCs role 581–2 HLA II molecules 428 HLA-DR arterial wall 549 T. cruzi infected DCs 648 HLA-DR-positive cells 315 Hodgkin’s lymphoma 427, 428 B cell non-Hodgkin’s lymphoma 562–3 IL-13 33 homeostatic cytokines, autoimmune diseases 464–5 horseradish peroxidase (HRP) endocytosis measurement 220 MHC 1 pathway 637 HPCs see hematopoietic progenitor cells HPV see human papillomavirus HRP see horseradish peroxidase hsc73, exosomes 183 HSPs see heat shock proteins HSV-1 see herpes simplex virus-1 HTL see helper T lymphocytes human DCs isolation, culture, propagation 82–94, 105 markers 99–100 human herpes viruses 514–17 human papillomavirus (HPV) 414–16, 639 humoral immunity, immune response component 53–4 humoral response, regulation, DCs role 255–61

HUVEC monolayers 283, 284–5 hyalocytes 398 ICAM-1 arterial wall 549 lung DCs 319 reverse transmigration mediation 284 ICAM-1 (CD54),-2,-3, ligands for LFA-1 (CD11a/CD18) 57–8 ICAM-3, apoptotic cell surface ligands 632 ICOS, ligation, B7 family members 54–7 IDCs see interdigitating DCs IDDM see insulin-dependent diabetes mellitus identifying DCs arterial wall 547–50 in fixed tissues 426–9 immunohistochemical identification, arterial wall 549–50 in tumors 432–3 ultrastructural criteria, arterial wall 547–9 IEM see immunoelectron microscopy IFN_ DC generation in vitro 188–90 DC maturation 195 DC-produced 66 IFN_ DCs maturation 195 MDDCs 253 production, DCs 248–50 IgA isotype switching, B cells 256–7 IgE, Fc_RI 528–30 Ikaros gene family DC hematopoiesis 7–8 functions 122 immune system effects 126 phenotypic characterization, DCs 111 IL-1, DC-produced 63 IL-1 family, co-stimulatory/polarizing factors 55 IL-1_ DCs generation in vitro 188–90 DCs migration 308 IL-3, DCs generation in vitro 188–90 IL-4 autocrine pathways, DCs 296–7 chronic allergic inflammatory response 524–5 DCs generation in vitro 188–90, 200 DCs regulation 193 Fc_RI 529 GM-CSF combined 200 sources 533 TH2 polarisation 533 IL-5, chronic allergic inflammatory response 524–5 IL-6 blood DCs 320 DC-produced 63 DCs generation in vitro 188–90 FDCs 32

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IL-7, DCs generation in vitro 188–90 IL-10 320–1 autoimmune diseases 465 DC-produced 63–4 DCs generation in vitro 188–90 IgA isotype switching 256–7 sources 64 IL-12 autocrine pathways, DCs 296–7 B cell differentiation 256 DC-produced 64–5, 320 mucosal immune responses 257 production suppression 533–4 secretion, CD40L 196 T helper cell-polarizing factor 64–5 IL-12p70, Leishmania-infected DCs 646 IL-13 chronic allergic inflammatory response 524–5 DCs generation in vitro 188–90 IL-12 production suppression 534 IL-13R_1 expression, FDCs 32–3 IL-15, DC-produced 65 IL-18 320 DC-produced 65–6 ILT3/1 expression, DC precursors defined by 87 imaging DCs 231–42 electron microscopy 237–9 fluorescence and confocal microscopy 236–7 immunoelectron microscopy 240–1 light microscopy 232–6 scanning electron microscopy 239–40 immature DCs allograft survival prolongation 600 apoptotic cells 634 CCR5–using HIV-1 isolates replication 489–90 chemokine receptors 204–5 influenza virus 507–8 lung 316–17 phagocytosis 475–8 immortalization of DCs, oncogenes 166 immune adjuvants DCs v monocyte/macrophages 271–2 peptide-pulsed PBMCs 271–2 immune cells entry, eye 392–4 nonparenchymal 394–9 through BBB 373–4 immune complex uptake, FcR 151 immune priveleged status, eye 391–2 immune responses character regulation 53–4, 55 components 53–4 enhancement, GM-CSF 199 innate 506–7 intestinal, and DCs 325–6 magnitude regulation 52–3

783 microorganisms evasion strategies 474–5 mucosal, DCs 257 regulation B cells 52–3 DCs 366–7 intestinal DCs 331–3 T cells 21–6, 52–4 tolerance and immunity 366–7 immune system effects, Ikaros gene family 126 immunisation targets, DCs, mucosal vaccines 484 immunity adaptive, DCs 252–3, 473–4, 507–14 HIV-specific, DCs role 581–2 induction, stress response-induced proteins 156–7 innate, DCs 252–3, 473–4, 510 organ-based, overview 440–2 skin 301–14 viral, infection absence 639–40 immunocytochemistry, cultured DCs preparation 233–4 immunodeficiency virus growth DC-T cell mixtures 490–2 see also HIV immunodeficiency virus production, tonsils 498 immunodeficiency viruses immunity, potential DCs role 499–501 MALT 497–9 immunoelectron microscopy (IEM) 240–1 immunogenicity, switch from tolerance 332 immunologic outcomes, apoptotic cells 632–4 immunoregulation, dendritic cell-regulated 51–74 immunostimulatory CPG motifs, autoimmune diseases 464 immunosuppression LCs 40 macrophage capture, apoptotic cells 637–8 and parasitism 645–9 solid organ allografts 439–40, 449 immunosuppressive drugs DC maturation inhibition 601 DC tolerogenicity promotion 601 immunosuppressive environment, CNS 374–5 immunosuppressive therapy, combination 601 immunotherapies adoptive 93–4 development, DC-based 174–5 in vitro cultured DC-like cells 105–6 induction of lymphoid tissue, T cells 126–7 infection, DC progenitor recruitment 135 infectious diseases, CNS 381–2 infectious influenza virus, CD8+ T cells 509 inflammation chronic, allergy 523–8 DC progenitor recruitment 135 inflammatory chemokines 205–6 inflammatory phase, wound healing 540

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784

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influenza antigen presentation 628–9 CD8+ T cells 509–10, 511 influenza virus adaptive immune responses 507–11 anti-influenza immune responses, DCs induction 508–11 DC interactions 507 DC subsets 508 immature DCs 507–8 mature DCs 508 monocytes 508 inhibition of DC function 109–10 innate immune responses 506–7, 510 CD11c- cells 506–7 innate immunity, DCs 252–3, 473–4, 506–7, 510 insulin-dependent diabetes mellitus (IDDM) 459 DC suppression 465–6 perpetuation 467 integrins _2 306 phagocytosis mediation 153 interaction, DC/DC 289–98 intercellular adhesion molecule-1 see ICAM-1 interdigitating DCs (IDCs) 104–5, 441 liver 347 interferon family co-stimulatory/polarizing factors 55 type 1 interferons, DC maturation 195 interstitial DCs 103–4 intestinal DCs 325–36 antigen uptake 330–1 exit 327, 328–9 functional properties 329–30 future directions 334 HIV 333–4 immune response regulation 331–3 in situ identification 326–7 isolation 329 life history 327–9 migratory properties 327–9 recruitment 327–8 retroviral infection 333–4 steady state 327 stimulated migration 327–8 TNF_ 328–9 intestinal immune responses, and DCs 325–6 intracellular pathways, DC activation 484 intracellular proteases, MHC class II antigens 158 iris, macrophages 398–9 irradiation, liver allografts 444–5 isolation, intestinal DCs 329 isolation guide, DCs 77–96 isopentane, light microscopy preparation 232–3 isotype switching, IgA, B cells 256–7 juvenile xanthogranuloma 431

KCs see keratinocytes keratinocytes, immune-associated function 302 keratinocytes (KCs), LCs 36 kidney DCs 347–9 function 349 histological location 347–8 ontogenic development 349 phenotype 347–8 population dynamics 348–9 knockout studies, cytokines 191–2 knockouts, targeted 119–29 Kupffer cells DCs migration 141, 338–9 cf liver DCs 338–9 LA see lactadherin labeling methods, light microscopy 235–6 lactadherin (LA), apoptotic cells 633 Lag-antigen, arterial wall 549, 551 LAM see lipoarabinomannan lamina propria (LP), intestinal immune responses 325–6 Langerhans cell granules (LCGs) 36–8, 225–6, 269, 302, 350, 432, 548–9 Langerhans cell histiocytosis (LCH) 382, 430–1 Langerhans cells (LCs) 35–50 activation markers 38, 303 adhesion molecules 38, 303 antigen capture 37, 304–5 antigen presenting molecules 303, 304–5 antigen uptake/processing 303 antigenic profiles 38 CCR7 46 characteristics 37–40 chemokine receptors 42–4, 303 chemotactic factor receptors 39 costimulatory molecules 38, 303 culture 83 cytokine receptors 39, 303 cytological characteristics 302 cytology 303 DC lineage 5–9 development 40–2 emigration 44–7 enzyme profiles 38 Fc receptors 303 functional aspects 309–12 functional features 37–40 future directions 47 general aspects 302 GM-CSF 36, 37–9, 41–2 identifying molecules 304 immune-associated function 302 immunostimulatory function 309–12 immunosuppression 40 maturation 37–9, 44–7 maturation function 309–12

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Langerhans cells (LCs) (cont.) mature cell characteristics 303 migration 44–7, 139–40, 277 MIP-3_ expression 43–4 morphology 38 phenotype 302–6 phenotypic changes 44–5 phenotypic features 37–40 resting cell characteristics 303 T cells 45–7 TGF_1 40, 41–2 tissue-homing properties 42–4 tolerogenic capacity 590 viral infections 40 see also dendritic cells (DCs) langerin 37, 304 LBDC see low buoyant density cells LCGs see Langerhans cell granules LCH see Langerhans cell histiocytosis LCs see Langerhans cells LCs, CD34+ hematopoietic precursors 40–2 lectin receptors, C-type, receptor-mediated endocytosis 155–6 Leishmania, biology of parasitism 645–7 lentivirus vectors, gene transfer to DCs 615–16 lentiviruses, gene transfer 610, 611–12 leukemias, DCs from 91–2 leukemic DC cultures, generation 166 leukocyte recruitment, chemokines 382–4 leukocytes chemotaxis 132–3 donor 345 migration 132–3 LFA-1 (CD11a/CD18), ligation, ICAM-1 (CD54),-2,-3 57–8 LFA-3, lung DCs 319 LFA-3 (CD58)-CD2 interaction 58 ligands uptake, by MDDCs 155 LIGHT DC modulatory factor 62 LT_-like activity 124 light microscopy DC imaging 231–6 materials sources 242 Lin-CD11c+ cells, DC migration 277 Lin-CD11c- plasmacytoid monocytes, DC migration 277 lipid scramblase, apoptotic cell surface ligands 632 lipoarabinomannan (LAM), receptor-mediated endocytosis 155–6 lipopolysaccharides (LPS), bacterial cell walls 474–5 liposomes, gene transfer 611 lipotechoic acids (LTA), bacterial cell walls 475 liquid propane, light microscopy preparation 232–3 Listeria monocytogenes, survival strategy 481 liver DCs 338–47 disease 347

785 Flt-3L 340 function 343–7 histological location 338–42 ontogenic development 343 phenotype 338–42 population dynamics 342–3 liver sinusoidal endothelial cells, allografts 444 liver sinusoids, DCs migration via 141–2 localization, DCs 131–49 choroid plexuses 376–7 lymph nodes 143–4 meninges 376–7 perivasculature 278–9 secondary lymphoid tissues 142–5 spleen 142–3 localization signals, secondary lymphoid tissues DCs 144–5 long-term cultures, DCs 79 low buoyant density cells (LBDC), liver DCs 343 LP see lamina propria LPS see lipopolysaccharides LT see lymphotoxin LTA see lipotechoic acids LT_-like activity, LIGHT 124 LT_R, FDC networks 30 lucifer yellow, endocytosis measurement 219–20 lung tumors 427, 433–4 lung-derived DCs 315–23 fate 317–18 immature DCs 316–17 isolation, culture, propagation 80 origin 317–18 lymph, DC migration via 138–40 lymph DCs 330 lymph node DCs cervical, tolerogenic capacity 590 mesenteric, tolerogenic capacity 590 pancreatic, tolerogenic capacity 590 lymph node-derived DCs, isolation, culture, propagation 80, 90–1 lymph nodes antigen-pulsed DCs 144 B cell areas 493–4, 495 DC subsets 143, 361–4, 493–4 DCs in 143–4, 357–70 DCs migration 138–40 DCs and virus load 493–7 LCs migration 139–40 localization in 143–4 peptide-pulsed DCs 144 regional, DCs migration 141–2 T cell areas 493–5 lymphatic endothelium, DCs reverse transmigration 279–84 lymphatic vessels, DCs migration to 279–84 lymphocyte-DC interactions, allografts 446–7 lymphocyte-produced chemokines 134

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786

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Page 786

INDEX

lymphocytes, development 5–7 lymphoid DC progenitors 145 lymphoid DCs 145–6 evidence 22 secondary lymphoid tissues 145–6 lymphoid endothelial cells, immune-associated function 302 lymphoid lineages, DCs development 358–9 lymphoid microenvironments DCs 361–6 DCs migration 365–6 lymphoid organs, DCs migration 205–6 lymphoid precursors, DCs from 359 lymphoid tissue development 119–29 induction, T cells 126–7 secondary DC development 124–6 DCs localization 142–5 lymphoid tissue-derived DCs 104–5 lymphoid tissues, mucosal-associated, immunodeficiency viruses 497–9 lymphoid-related DCs 265 lymphotoxin ligands, DC development 123–4 lymphotoxin (LT) FDC networks role 30 FDC production 190–3 lymphotoxin-mediated events 126–7 lysis, DCs 78 M5/114 (anti-MHC class II) 400 MA see mycolic acid macaque DCs, phenotype 488 macrophage capture, apoptotic cells 637–8 macrophage migration inhibitory factor (MIF), wound healing 541 macrophage populations, eye 398 macrophage-derived DCs 84–5 macrophage/monocyte lineage 263–74 macrophages antigen-processing capabilities 270 choroid 398–9 ciliary body 398–9 DC maturation 265–6 DCs interaction 321 iris 398–9 peritoneal-derived DCs 82 thymus 13–14 macropinocytosis 215–16 antigen uptake mechanism 151–2 cf micropinocytosis 215 model 478 monocyte/macrophage interaction 269–70 morphological appearance 479 MALDI-TOF-MS see matrix-assisted laser desorption ionization time-of-flight mass spectrometry male reproductive tract DCs 414

malignant ascites 434 macrophages from 85, 92 MALT see mucosal-associated lymphoid tissues mannose, phagocytosis 214, 478–9 mannose receptors antigen uptake mechanism 151 lung DCs 317 MDDCs 217 pinocytosis 216 receptor-mediated endocytosis 155–6 mannose/_ glucan receptor 217 marginal zone DCs (MZDCs) 142, 143 markers, DCs 99–100 mast cells Fc_RI 529 immune-associated function 302 matrix-assisted laser desorption ionization time-offlight mass spectrometry (MALDI-TOF-MS), exosomes protein composition 182–3 maturation DCs 58, 61, 68, 78–9, 132, 173–4 LCs 37–9, 44–7 maturation characteristics, DCs at body surfaces 488–9 maturation inhibition, DCs 196 maturation stages, DC 265–6 mature DCs, influenza virus 508 MCM see monocyte-conditioned medium MCP-1 chemokine responsiveness 205 EAE 406 MCP-4, DC-produced 66–7 MDDCs see monocyte-derived DCs measles virus (MV) 512–13 cell adverse effects 513 DCs interactions 512 DCs role in vivo 513–14 T cell interactions 512–13 mediators, dendritic cell-regulated immunoregulation 51–74 melanoma 427 clinical trials 566–9 membrane-bound modulatory factors, DCs 54–62 membrane-bound TNF-TNFR family members 59–62 meninges, DCs localization 376–7 Meningococci, immune response evasion 475 mesenteric lymph node DCs, tolerogenic capacity 590 mesenteric lymph nodes (MLN), intestinal immune responses 325–6 metastatic melanoma, clinical trials 567 MFG-E8, apoptotic cells 633 MHC 1, pre-processed epitopes 635–6 MHC 1 antigen presentation, exogenous pathway 627–8 MHC 1 pathway, phagosome-to-cytosol pathway 637 MHC 1/peptide complex formation, potential paths 634–5

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MHC biosynthesis regulation 151–63 degradation regulation 151–63 MHC class I expression, eye 392 MHC class I molecules, NK cells 246 MHC class I proteins antigens proteolysis 159–60 cross-priming 159 exosomes bearing 181–2 synthesis 158–60 trafficking 158–60 MHC class II+ DCs eye 392, 399 pancreas 351–2 parenchymal organs 339–40 MHC class II antigens allografts 441 cell surface loading 158 DC types, thymus 18–19 intracellular proteases 158 macropinocytosis 152 MHC class II compartments (MIICs), APCs 180 MHC class II expression, eye 392, 399, 404 MHC class II markers, intestinal DCs 326 MHC class II molecules BBB 372 Fc_RI 529 lung DCs 316 transfer between allogeneic DCs 292 MHC class II pathway, regulation 224–5 MHC class II proteins recycling pathways 157–8 synthesis 157–8 trafficking 157–8 MHC molecules, transfer between DCs 291–3 microchimerism allograft acceptance/rejection 450–4 tolerance induction, allografts 450–1 microenvironmental factors, tolerogenic DCs propagation 596–9 microglia antigen presenting functions 378–9 as APCs 396–7 chemokines 396 CNS sentinels 377–9 cytokines 396 distribution 394–7 free radicals 396 functions 377–8, 396 in vitro 396–7 NO 396 ontogeny 394–7 OX2 395–6 perivascular cells 397 phenotype 394–6 pinocytosis 396 proteases 396

787 retinal 394–7, 399–404 microorganisms complexity 474–5 evasion strategies, immune responses 474–5 TLR recognition 481 micropinocytosis, cf macropinocytosis 215 microscopy, light, DC imaging 231–6 MIF see macrophage migration inhibitory factor migration DCs 98, 107–8, 131–49, 203, 307–9 lymphoid microenvironments 365–6 lymphoid organs 205–6 LCs 44–7, 139–40 leukocytes 132–3 migratory cell cycle 305–6 migratory characteristics, DCs at body surfaces 488–9 MIICs see MHC class II compartments MIP-1_ expression, liver 346 MIP-3_ DCs migration 366 expression, LCs 43–4, 204–5 mixed chimerism, allograft acceptance/rejection 450–4 mixed leukocyte reaction (MLR) 289–90 liver DCs 343 stimulation 292 MLN see mesenteric lymph nodes MLR see mixed leukocyte reaction MLV see murine leukemia virus mobilization DCs 131–49 nonlymphoid sites 136–8 modulatory factors, membrane-bound, immune response 54–62 molecular basis, DC tolerogenicity 594–6 molviol, light microscopy preparation 236 monocyte chemotactic protein-1 see MCP-1 monocyte progenitor-derived DCs, cancer clinical trials 565–9 monocyte-conditioned medium (MCM) 266 monocyte-derived DCs (MDDCs) culture 80–2, 82–3 generation 133–5, 263–4 IFN_ production 253 isolation 80–2, 82–3 ligands uptake, fluorescent 155 mannose receptor expression 217 NK cell cytolytic activity 253 propagation 80–2, 82–3 monocyte/macrophage interaction direct 269–71 mediators 266–9 monocyte/macrophage lineage 263–74 monocytes DC maturation 265–6 DC progenitors 4–5, 22, 133–6 differentiation 280–2

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788

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monocytes (cont.) influenza virus 508 plasmacytoid 22 subsets 282–4 transmigration 281–2 transmission EM 237–9 wound healing 541 monocytic leukemias 432 mouse DCs isolation, culture, propagation 78–82, 105 markers 99 tolerogenic capacity 590 mucosal immune responses, DCs 257 mucosal sites, DCs at 418–19 mucosal vaccines 484 mucosal-associated lymphoid tissues (MALT) 488 chronic virus production, tonsils 498 immuno-deficiency viruses 497–9 SIV infection rapid spread 498–9 multiple myeloma, clinical trials 563–4 multivesicular endosomes (MVBs) 179, 184 murine leukemia virus (MLV ), gene transfer 609–11 MV see measles virus MVBs see multivesicular endosomes mycolic acid (MA), bacterial cell walls 474–5 mycosis fungoides/SS 427 myeloid DC transduction, allografts 446 myeloid DCs, evidence 22 myeloid lineages, DCs development 358–9 myeloid origin, DCs 4–5 myeloid-related DC lineage 264–5 myeloma, multiple 563–4 myelomonocytic leukemias 432 MZDCs see marginal zone DCs naked DNA, gene transfer 611, 620–1 natural interferon-producing cells (NIPCs) 463–4 natural killer cells see NK cells neck cancers, DC infiltration 426 necrotic debris, v apoptotic debris 270–1 negative selection, T cells 366–7 `nest’ subsets, DCs 142 neurons, immunosuppressive activity 374–5 neuropeptides, immunosuppressive activity 374–5 neutrophils, wound healing 541 NF_B inhibition 595 NF_B-specific ‘decoy’ ODNs 598–9 NIPCs see natural interferon-producing cells nitric oxide (NO) microglia 396 monocyte/macrophage interaction mediation 268–9 production 595 synthesis 172 NK cells 245–54 cell contact-dependent activation 250

cytolytic activity 248–50 MDDCs 253 cytotoxic activity 245–6 DC-mediated triggering 246–50 Flt-3L 247–8 MHC class I molecules 246 T cells/DCs 15–16 thymus 13–14, 15–16 tumor regression 247–8 NO see nitric oxide NOD mice see prediabetic nonobese diabetic mice non-opsonic phagocytosis 480 non-replicating virus, CD8+ T cells 509 non-viral gene transfer 620–1 non-viral vectors, gene transfer 613–14 nonclathrin-dependent phagocytosis 152 nonenzymatic probes, endocytosis measurement 220 nonfluorescent probes, endocytosis measurement 220 nonlymphoid tissues chemokine receptors 204–5 DCs in 103–4, 133–5 non epithelial DC populations 103–4 superficial epithelial DC populations 103 nonparenchymal immune cells 394–9 nonresponsiveness, DC induced 290 ocular autoimmune 402–4 ODNs see oligodeoxynucleotides oligodeoxynucleotides (ODNs), NF_B 598–9 oncogenes, immortalization of DCs 166 ontogenic heterogeneity, DCs 358 ontogeny, skin DCs 306–9 opsonic phagocytosis 480 oral antigen 331–2 organ transplantation, DCs therapeutic application 600–1 organ-based immunity, overview 440–2 OX2, microglia 395–6 OX40 ligation 59–60 OX40–OX40L interactions, B cells 259 PAF see platelet-activating factor Paget’s disease 427 PAMPs see pathogen-associated molecular patterns pancreas DCs 351–3 disease 353 function 352–3 histological location 351–2 ontogenic development 352 phenotype 351–2 population dynamics 352 pancreatic lymph node DCs, tolerogenic capacity 590 papillomavirus, human 414–16, 639 paracrine effects, DCs 296–7 paradoxes, DC phenotypes 119–20

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paraformaldehyde fixation IEM 241 light microscopy 232, 234–5 paraneoplastic cerebellar degeneration (PCD), antigen presentation 629 parasitism biology of 645–50 Leishmania 645–7 Plasmodium falciparum 648–9 Trypanosoma cruzi 647–8 PARC/DC-CK1 chemokine 144–5 parenchymal organs DCs in 337–56 MHC class II+ DCs 339–40 passenger leukocytes 345 pathogen-associated molecular patterns (PAMPS) 463 pathogens DC1 effector function 23–5 DC2 effector function 25–6 pattern-recognition receptors, antigens 106–7 PBC see primary biliary cirrhosis PBMC see peripheral blood mononuclear cells PCD see paraneoplastic cerebellar degeneration PECAM-1 see platelet endothelial cell adhesion molecule peptide affinity, and antigen dose 532 peptide chaperoning, HSPs 631–2, 635 peptide-pulsed DCs lymph nodes 144 melanoma 566–7 peptide-pulsed PBMCs, immune adjuvants 271–2 peptidoglycan (PG) cell wall, bacterial 474–5 perfusion fixation, light microscopy 232 peripheral blood DCs, gradient enrichment 562–5 peripheral blood mononuclear cells (PBMC) 434 peripheral lymph nodes, DCs and virus load 493–7 peripheral tissues DC precursor migration 275–8 DCs migration from 279–84 peripheral tolerance DC-induced 591–4 self-antigen specific 466–7 peripherally circulating progenitor cells 85–8 peritoneal cell-derived DCs, isolation, culture, propagation 81, 82 perivascular cells 397 microglia 397 perivasculature localization, DCs 278–9 perpetuation of initial autoimmune responses 467–9 Peyer’s patch DCs, tolerogenic capacity 590 Peyer’s patch-derived DCs, isolation, culture, propagation 80 Peyer’s patches (PP) DC subsets 144, 365 functional heterogeneity 367–8 intestinal immune responses 325–6 phenotypic heterogeneity 367–8

789 PG see peptidoglycan cell wall PGE2 320–1 PGL see phenolic glycolipids phagocytic receptors, DCs 153–4 phagocytosis 213–15 actin 213–16 antigen transfer 291 antigen uptake mechanism 151, 152–4 antigen-coated particles 153 apoptotic cells 627–44 antigens from the dead 628–30 eating the dead 632–4 introducing the dead 627–8 lessons from the dead 639–42 outcomes 637 packaging the dead 630–2 presenting the dead 637–9 processing the dead 634–7 complement receptors 151, 214 defined 152–3 dynamin-2 214–15 FcR-associated 151, 214 GFP 476 immature DCs 475–8 mannose 214 mechanisms 478 monocyte/macrophage interaction 269–70 Rho family GTPases 214 types 152–3 phagosome-to-cytosol pathway, MHC 1 pathway 637 phenolic glycolipids (PGL), bacterial cell walls 474–5 phenotype, application, DCs 112 phenotypic characterization DCs 97–117 application of phenotype 112 cell surface antigens 99–100 functional phenotype 106–12 future 112–13 surface phenotype 100–6 phenotypic heterogeneity 367–8 phosphatidylserine (PS), apoptotic cell surface ligands 632 phosphatidylserine receptors, phagocytosis mediation 153 pilosebaceous unit, immune-associated function 302 pinocytosis 215–16 antigen transfer 291 macropinocytosis 151–2, 215–16 microglia 396 placental DCs 413 plasmacytoid T cells/monocytes 22, 146, 198 Plasmodium falciparum, biology of parasitism 648–9 plasticity DC phenotypes/function 368 dendritic cells 187 FDC network 30

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platelet endothelial cell adhesion molecule (PECAM1) 308 platelet-activating factor (PAF) 204 PM see protoplasmic membrane Pneumococci, immune response evasion 475 polarization, T cells 21–8 polarizing factors, DC-produced 55–6 polarizing signals, immune response character 53–4, 55 POMC see proopiomelanocortin potocytosis 216–17 potocytosis (cont) caveolar membrane system 217 poxviruses 517–18 vaccinia virus 517–18 PP see Peyer’s patches pre-processed epitopes, MHC 1 635–6 precursor migration, DC, peripheral tissues 275–8 prediabetic nonobese diabetic (NOD) mice autoimmune response perpetuation 467 pancreas DCs 353 preparation methods, light microscopy 232–5 primary biliary cirrhosis (PBC) 347 priming, self-antigen 467–8 prion protein (PrP)-deficient mice, FDC networks 31 progenitor cells DCs constitutive recruitment 135 nonlymphoid sites 133–4 propagation 346–7 hematopoietic 4 monocytes 4–5, 22 peripherally circulating 85–8 progenitors, lymphoid DC 145 progesterone, viral transport facilitation 418 proopiomelanocortin (POMC) 311 propagation guide, DCs 77–96 propane, light microscopy preparation 232–3 prostaglandin E2, immunosuppressive activity 374–5 prostate cancer, clinical trials 564, 565–6 prostate tumors 427 proteases intracellular, MHC class II antigens 158 microglia 396 protoplasmic membrane (PM), bacterial 474–5 PrP-deficient mice see prion protein-deficient mice PS see phosphatidylserine Pseudomonas, immune response evasion 475 psoriasis, wound healing 544 PU.1 transcription factor, DC development 9 pulmonary DCs, distribution 315–16

receptor utilization apoptotic cells 632–3 microenvironment, immunologic outcome 633–4 receptor-mediated antigen uptake 151 receptor-mediated endocytosis 154–7 receptors, LCs selective tissue-homing 42–4 recruitment, DCs, inflamed CNS 382–4 regional lymph nodes, DCs migration 141–2 regulation DCs development, cytokines 359–61 DCs function 109–10 cytokine 189–97 regulatory T cells autoimmune diseases 465–7 induction 592 Reiter’s syndrome 462 rejection, allograft 442–8 RelB transcription factor DC development 8–9 expression 429 targeted knockouts 120–3 remodeling phase, wound healing 540 renal cell cancer, clinical trials 569 renal interstitial DCs, tolerogenic capacity 590 reproductive tract antigen transport, and DCs 414 DCs in 411–21 female genital tract DCs 412–14 HIV transmission 416–18 male reproductive tract DCs 414 STDs 414–18 respiratory tract, DCs in 315–23 responsiveness, DC induced 290 resting DCs, chemokine receptors 204–5 retina DCs 399–404 layout 389–91 retinal pigment epithelium (RPE) 389–91, 405 GM-CSF cells 405 retroviral infection, intestinal DCs 333–4 retroviral vectors, gene transfer to DCs 614–15 retrovirus, gene transfer 610 reverse transendothelial migration (TEM) DCs 93, 132, 279–84 molecular mediators 284 rheumatoid arthritis, DCs in synovial fluid 278 Rho family GTPases macropinocytosis regulation 152 phagocytosis 214 Rickettsia, survival strategy 481 RPE see retinal pigment epithelium

RANK (TRANCE ligand), DC modulatory factor 61 RANTES, eye 406 Rauscher leukaemia virus, DC autocrine pathways 296 re-epithelialization, wound healing 540

S100 intracytoplasmic calcium binding proteins 428 S-100 protein 306 arterial wall 549, 551 Salmonella typhimurium, survival strategy 481

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Salmonella-infected macrophages, antigen presentation 629–30 SALT see skin-associated immune system sarcoidosis 322 sarcomas, dendritic cell 432 scanning electron microscopy (SEM) 239–40 scavenger receptors 218–19 antigen uptake mechanism 151 phagocytosis mediation 153 receptor-mediated endocytosis 156 SCID mice see severe combined immunodeficiency mice secondary DC reactions 431 secondary lymphoid chemokine (SLC) 280 secondary lymphoid tissues DC development 124–6 DCs localization 142–5 DCs localization signals 144–5 lymphoid DCs 145–6 self-antigen specific peripheral tolerance 466–7 self-antigens, priming 467–8 SEM see scanning electron microscopy severe combined immunodeficiency (SCID) mice, FDC networks origin 31 sex hormones, and DCs 413 sexual transmission, HIV-1/SIV 488–93 sexually transmitted diseases (STDs) 411 cervical DCs 414–16 Shigella, survival strategy 481 short-term cultures, DCs 79 Siayl Lewis X, skin DCs 305–6, 307 signalling and adhesion, DCs 108–9 signals DCs localization, secondary lymphoid tissues 144–5 dendritic cell-regulated immunoregulation 51–74 hematopoietic development 7–9 signal 1/2 carriers, DCs 52–3, 55 signal 3 carriers, DCs 53–4, 55 SIS see skin immune system SIV infection DCs during 487–504 rapid spread, MALT 498–9 Sjogren’s syndrome B cell autoimmunity amplification 468–9 DC infiltration 468 self-antigen priming 467 skin immunity 301–12 role 35 tumors 427 see also wound healing skin immune system (SIS) 301 skin immunity 301–14 DDCs functional aspects 309–12 DDCs phenotype 302–6 LHs functional aspects 309–12 LHs phenotype 302–6

791 skin DCs ontogeny 306–9 skin-associated immune system (SALT) 301 skin-derived DCs HIV-1/SIV infection 488–9 isolation, culture, propagation 80, 91 SLC see secondary lymphoid chemokine SLE see systemic lupus erythematosus SLE nephritis, DC infiltration 468 slide mounting technique, light microscopy 236 solid organ allografts, DCs in rejection and acceptance 439–57 soluble factors, DC-produced 533 species issues, DCs 98 spleen DC subsets 142 DCs in 142–3, 357–70 DCs migration 138–40, 140–1 DCs subsets 361–4 spleen DCs, tolerogenic capacity 590 spleen-derived DCs 126 isolation, culture, propagation 79, 90–1 splenic APCs, peripheral tolerance 591–2 splenic DCs, tolerogenic capacity 590 spondyloarthritis, DC infiltration 468 Staphylococci, immune response evasion 475 STCP-1, DC-produced 66–7 STDs see sexually transmitted diseases streptavidin, labeling enhancement 236 Streptococcus, survival strategy 481 stress response-induced proteins, immunity induction 156–7 subdominant self-epitopes, autoimmune diseases 464 subsets, DCs 131–2, 364 antigen processing 160–1 B cell areas 493–4 B cell functions 257–8 CD11c+ cells 363–4 CD14+ cells 363–4 immune response determination 596 influenza virus 508 lymph nodes 143, 361–4 mobilization by Flt-3L 198 paradoxes 119–20 Peyer’s patches 144, 365 spleen 142, 361–4 T cell areas 493–4 thymus 18–19 subsets, monocytes 282–4 sucrose infusion, light microscopy 232 suppression by regulatory cells, tolerance induction method 588 surface ligands, apoptotic cells 632 surface molecules, DC 478–80 surface phenotype, DC populations 100–6 switch paradigm, chemokine receptors 205–6 switch recombination, DCs 256–7

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792

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systemic autoimmunity, autoimmune diseases 462–3 systemic lupus erythematosus (SLE) 459, 640, 642 nephritis, DC infiltration 468 T1/ST2L-T1/ST2 interaction 58–9 T cell activation, atopic allergic disease 525–8 T cell areas acute SIV infection 494–5 DC cell subsets 493–4 T cell receptors (TCRs), tolerance and immunity 460–1 T cell-mediated immune responses 21–6, 52–4 T cell-stimulatory capacity, dendritic cell lines 171–2 T cells activation 21–8 DCs 318, 320–1, 588 allergen sensitisation 525–6 allogeneic, allografts 446 antigen responses 294 atherosclerotic lesions 552–4 canarypox 519 cross-priming 631 DC cells interaction, adhesion molecules 319–20 DCs mutual activation 196–7 development, early stages 14–16 differentiation regulation 332–3 dying cells crosspresentation 506 FDCs 31–2 immune response regulation 21–6, 52–4 induction of lymphoid tissue 126–7 inhibition, CTLA-4 54–7 inhibitory activity, T. cruzi infected DCs 648 LCs 45–7 MV interactions 512–13 negative selection 366–7 plasmacytoid 22 polarization 21–8 priming, CD8_ DCs 363 proliferation, TNF_ 320 proliferation downregulation, T. cruzi-derived glycosylinositolphospholipids 648 regulatory, autoimmune diseases 465–7 thymus 13–14 tissue inflammation 525–8 transmission EM 237–9 virus transmission 496–7 T helper cell-polarizing factor, IL-12 64–5 T lymphocytes DC recruitment 125–6 immune-associated function 302 T-cell anergy, induction 592 T-cell apoptosis, activated, induction 592–3 T-cell futile cycles 426 tacrolimus, allografts 445 TADCs see tumor-associated DCs TAMs see tumor-associated macrophages

TARC (thymus and activation-regulated chemokine), DC-produced 66–7 targeted knockouts 119–29 DC development, secondary lymphoid tissue 124–6 lymphotoxin ligands 123–4 RelB 120–3 T-cell induction, lymphoid tissue 126–7 TCA4/SLC CCR7 binding 125 expression 125–6 TCRs see T cell receptors TECK (thymus-expressed chemokine), DC-produced 66–7, 145 telomerase reverse transcriptase (TERT) RNA transfected DCs 621 TEM see reverse transendothelial migration; transendothelial migration; transmission electron microscopy TERT see telomerase reverse transcriptase RNA transfected DCs testing, dendritic cell lines 165–77 TGF_1 DCs generation in vitro 188–90 DCs regulation 189, 190 immunomodulatory effect 464–5 LCs 40, 41–2, 464–5 tolerogenic DCs propagation 597–9 TGF_ IgA isotype switching 256–7 immunosuppressive activity 374–5 wound healing 542–3 TH1 differentiation, DC1s 22–3 TH1/TH2 regulation 366 TH2 polarisation IL-4 533 theoretical aspects 530–4 skewing, T cell priming 531 TH2–like T-cell subsets, selective activation 591–2 therapeutic application of DCs, experimental organ transplantation 600–1 therapy, DC-based, allograft rejection and autoimmune diseases 587–607 thrombospondin-1 (TSP-1), apoptotic cells 633 thymic DCs characteristics 16–17 developmental origin 17–18 tolerogenic capacity 590 thymocyte development, early stages 14–16 thymus DC types 18–19in 13–20, 365 DCs subsets 18–19 thymic DCs characteristics 16–17 thymus and activation-regulated chemokine (TARC), DC-produced 66–7 thymus-derived DCs, isolation, culture, propagation 80

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thymus-expressed chemokine (TECK), DC-produced 66–7, 145 thyroid tumors 427 tissue macrophages, immune-associated function 302 tissue microenvironments, DC effectors 26 TLR see Toll-like receptors TNF, FDC networks role 30 TNF family, co-stimulatory/polarizing factors 55–6 TNF-TNFR family, membrane-bound 59–62 TNF_ autoimmune response perpetuation 468 bacterial infection, DCs 483 DC maturation 67–8, 78–9, 483 DC-produced 62–3 DCs generation in vitro 188–90 DCs migration 308 DCs regulation 193 intestinal DCs 328–9 T cell proliferation 320 tolerance and immunity balance 460–1 endogenously-processed self-antigen 466 immune responses 366–7 peripheral tolerance 466–7 tolerance induction allografts 449–53 central tolerance, DC-induced 589–91 DCs and 588–94 peripheral tolerance, DC-induced 591–4 tolerance maintenance, autoimmune diseases 464–7 tolerance mechanisms, allografts 451 tolerogenicity DCs 21–2, 587–607 capacity 590 genetic engineering, tolerogenic DCs 602–4 immunosuppressive effects 604 modulation 603 molecular basis of tolerogenicity 594–6 promotion, immunosuppressive drugs 601 propagation for therapeutic application 596–9 therapeutic application, autoimmune disease 601–2 tolerance induction 588–94 Toll-like receptors (TLR) 479–80 microorganisms recognised by 481 tonsil-derived DCs 90–1, 104–5 tonsils immunodeficiency virus production, MALT 498 virus isolation 499 TRAIL DC modulatory factor 61–2 peripheral tolerance 593, 595 transcription factors phenotypic characterization, DCs 111 tolerogenic DCs propagation 596–9 transendothelial migration (TEM), DCs 132, 279–84

transgenic studies, cytokines 192 transmission electron microscopy (TEM) 237–9 protocol 238–9 transplantation DC-lymphocyte interactions, chaperoning 446–7 solid organ, DCs in rejection and acceptance 439–57 triggered-membrane ruffling, phagocytosis mechanism 478 Trypanosoma cruzi, biology of parasitism 647–8 TSP-1 see thrombospondin-1 tumor antigen, cross-presentation 628–30 tumor growth suppression, DC-derived exosomes 180–1 tumor immunity CD4+ T helper cells 641 v tumor-mediated immunosuppression 640–1 tumor infiltration, DC 427 tumor peptide-pulsed exosomes 180–1 tumor regression Flt-3L 247–8 NK cells 247–8 tumor-associated DCs (TADCs) 207–9 tumor-associated macrophages (TAMs) 208 tumor-derived DCs 82, 91–2 tumor-mediated immunosuppression, v tumor immunity 640–1 tumors chemokine recruitment of DCs 207–9 DCs adoptive transfer 250 identifying DCs in 432–3 TUNEL-positive apoptotic cells 630–1 type 1 interferons, DC maturation 195 type 2 T helper cells, chronic allergic inflammatory response 524–8 ulcerative colitis, DC infiltration 468 uterus 413 uveal tract DCs in 399–404 function, DCs 401–2 layout 389–91 migration capacity, DCs 401–2 ontogeny, DCs 401–2 vaccines attenuated, DCs involvement 500–1 mucosal 484 vaccinia virus 517–18 APC interactions 517–18 DCs role in vivo 518 gene transfer 610, 613 vaccinia virus vectors, gene transfer to DCs 619–20 VALT see vascular-associated lymphoid tissue vascular DCs heterogeneity 554–5

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Vascular DCs (cont.) origin 550–1 phenotypes 554 vascular endothelial cells, immune-associated function 302 vascular endothelial growth factor (VEGF) DC differentiation 196 DC maturation inhibition 264, 279 wound healing 542–3 vascular endothelium, DCs reverse transmigration 279–84 vascular pathology 547 vascular-associated lymphoid tissue (VALT) 551 vascularized tissue, DCs mobilization 137–8, 140–1 VEGF see vascular endothelial growth factor video microscopy, DCs endocytic capacities 221–2 viral antigen, cross-presentation 628–30 viral hepatitis 347 viral immunity, infection absence 639–40 viral infections autoimmune diseases 463–4 LCs 40

viral vectors gene transfer 609–13 non-viral vectors, gene transfer 613–14 virus growth, augmented, antigen-responding T cells 495–6 virus transmission DCs 496–7 T cells 496–7 virus-infected apoptotic cells, CD8+ T cells 509–10 viruses, DCs interactions 505–22 vulvo-vaginal epithelia 412 Wiskott-Aldrich syndrome 432 wound healing 539–45 abnormal 543–4 ageing and 543–4 normal 540–3 overview 539–40 xanthogranuloma, juvenile 431 XS52, dendritic cell line 167–72, 173–4 XS106, dendritic cell line 167–72, 175