Molecular Biology of B Cells
Tasuku Honjo Frederick W. Alt Michael S. Neuberger Editors Elsevier Academic Press
Molecular Biology of B Cells
Molecular Biology of B Cells Edited by
Tasuku Honjo Department of Medical Chemistry Kyoto University Faculty of Medicine Kyoto, Japan
Frederick W. Alt Howard Hughes Medical Institute The Center for Blood Research The Children’s Hospital, Boston, Massachusetts
Michael S. Neuberger MRC Laboratory of Molecular Biology Protein and Nucleic Acid Chemistry Division Cambridge, United Kingdom
AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO
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Contributors
Dr. Frederick W. Alt Howard Hughes Medical Institute, The Children’s Hospital, The Center for Blood Research, Boston, MA, USA
Dr. Adolfo Ferrando Department of Pediatrics, Children’s Hospital, DanaFarber Cancer Institute, Boston, MA, USA
Dr. Barbara Birshtein Department of Cell Biology, Albert Einstein College of Medicine, Bronx, NY, USA
Dr. Martin F. Flajnik Department of Microbiology and Immunology, University of Maryland, Baltimore, MD, USA
Dr. Constantin A. Bona Department of Microbiology, The Mount Sinai School of Medicine, New York, NY, USA
Dr. Raif S. Geha Department of Pediatrics, Harvard Medical School, Boston, MA, USA
Dr. Francisco Bonilla Division of Immunology, Children’s Hospital, Boston, MA, USA
Dr. Deborah L Hardie Medical Research Council Centre for Immune Regulation, The University of Birmingham Medical School, Birmingham, England, UK
Dr. Per Brandtzaeg Laboratory of Immunohistochemistry and Immunopathology (LIIPAT), Institute of Pathology, University of Oslo, Rikshospitalet, Oslo, Norway
Dr. Linda Hendershot Tumor Cell Biology, St Jude Children’s Research Hospital, Memphis, TN, USA
Dr. Marianne Bruggemann Laboratory of Developmental Immunology, The Babraham Institute, Babraham Hall, Babraham, Cambridge, UK
Dr. Tasuku Honjo Department of Medical Chemistry, Kyoto University Faculty of Medicine, Yoshida, Sakyo-ku, Kyoto, Japan
Dr. Peter Burrows Department of Microbiology, University of Alabama at Birmingham, 383 Wallace Tumor Institute, Birmingham, USA
Dr. Ellen Hsu Department of Physiology & Pharmacology, The State University of New York Health Science Center at Brooklyn, Brooklyn, NY, USA
Dr. Kathryn Calame Departments of Microbiology and Biochemistry & Molecular Biophysics, Columbia Unversity, Collge of Physicians and Surgeons, New York, NY, USA
Dr. John F. Kearney Division of Developmental and Clinical Immunology, Department of Microbiology, University of Alabama at Birmingham, 6th Avenue South, Birmingham, AL, USA
Dr. Michael C. Carroll Department of Pediatrics, Harvard Medical School, The Center for Blood Research, Boston MA, USA
Dr. Paul W. Kincade Immunobiology and Cancer Research Program, Oklahoma Medical Research Foundation, Oklahoma City, OK, USA
Dr. Michel Cogné Laboratoire d’Immunologie, Faculte de Medecine, Limoges Cedex, France
Dr. Kazuo Kinoshita Department of Medical Chemistry, Graduate School of Medicine, Kyoto University, Yoshida-Konoe, Sakyo-ku, Kyoto, Japan
Dr. Max D. Cooper Howard Hughes Medical Institute, The University of Alabama at Birmingham, 378 Wallace Tumor Institute, Birmingham AL, USA
Dr. Katherine L. Knight Department of Microbiology and Immunology, Loyola University Stritch School of Medicine, Maywood, IL, USA
Dr. Jason Cyster Howard Hughes Medical Institute and Department of Microbiology and Immunology, University of California San Francisco, San Francisco, CA, U.S.A.
Dr. Michael Krangel Department of Immunology, Duke University Medical Center, Jones Bldg, Research Drive, Durham, NC, USA
Dr. Nadia Danilova Department of Biology, Massachusetts Institute of Technology, Cambridge, MA, USA
Dr. Michael E. Lamm Department of Pathology, Case Western Reserve University School of Medicine, Cleveland, OH, USA
Dr. Randall S. Davis Divisions of Developmental and Clinical Immunology and Hematology/Oncology, Department of Medicine, University of Alabama at Birmingham, Birmingham, AL, USA
Dr. Dennis Lanning Department of Microbiology and Immunology, Loyola University Stritch School of Medicine, Maywood, IL, USA
Dr. Douglas T. Fearon Department of Medicine, University of Cambridge School of Clinical Medicine_Addenbrookes Hospital, Cambridge, UK
Dr. Tucker W. LeBien University of Minnesota Cancer Center, Minneapolis, MN, USA
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Contributors
Dr. Gerard Lefranc Laboratoire d’Immunogenetique Moleculaire, Institut de Genetique Humaine, Universite Montpellier II, Montpellier Cedex 5, France
Dr. Michael Reth Department of Molecular Immunology, Faculty of Biology III, University of Freiburg and Max Planck Institute for Immunobiology, Freiburg, Germany
Dr. Marie-Paul Lefranc Laboratoire d’Immuno Genetique Moleculaire, LIGM, Universite Montpellier II, UPR CNRS Institut de Genetique Humaine, Montpellier Cedex 5, France
Dr. Roy Riblet Torrey Pines Institute for Molecular Studies, San Diego, CA, USA
Dr. Susanna Lewis Genetics and Genomic Biology, Hospital for Sick Children Research Institute, Toronto, Ontario, Canada Dr. Gary W. Litman Department of Molecular Genetics, All Children’s Hospital, St. Petersburg, FL, USA Dr. A. Thomas Look Department of Pediatrics, Children’s Hospital, DanaFarber Cancer Institute, Boston, MA, USA Dr. Ian C. M. MacLennan Medical Research Council Centre for Immune Regulation, The University of Birmingham Medical School, Birmingham, UK Dr. Nancy Maizels Departments of Immunology and Biochemistry, University of Washington Medical School, Seattle, WA, USA Dr. Roy Mariuzza Center for Advanced Research in Biotechnology, W. M. Keck Laboratory for Structural Biology, University of Maryland Biotechnology Institute, Rockville, MD, USA Dr. Jim Marks Department of Anesthesia, San Francisco General Hospital, San Francisco, CA, USA Dr. Fumihiko Matsuda Centre National de Genotypage, Evry Cedex, France Dr. Fritz Melchers Deptartmen of Cell Biology, Biozentrum, University of Basel, Basel, Switzerland Dr. Herbert C. Morse III Laboratory of Immunopathology, National Institutes of Health, Bethesda, MD, USA Dr. H. Craig Morton Laboratory of Immunohistochemistry and Immunopathology (LIIPAT), Institute of Pathology, University of Oslo, Rikshospitalet, Oslo, Norway Dr. Masamichi Muramatsu Department of Medical Chemistry, Graduate School of Medicine, Kyoto University, Yoshida-Konoe, Sakyo-ku, Kyoto, Japan Dr. Lars Nitschke Institute of Virology and Immunobiology, Wuerzburg, Germany Dr. Marjorie A. Oettinger Department of Molecular Biology, Massachusetts General Hospital, Boston, MA, USA Dr. Barbara A. Osborne Department of Veterinary and Animal Science, University of Massachusetts, Amherst, MA, USA
Dr. Matthew Scharff Department of Cell Biology, Albert Einstein College of Medicine, Bronx, NY, USA Dr. Mark S. Schlissel Department of Molecular and Cellular Biology, Division of Immunology, University of California-Berkeley, Berkeley, CA, USA Dr. JoAnn Sekiguchi Howard Hughes Medical Institute, The Children’s Hospital, The Center for Blood Research, Boston, MA, USA Dr. Ranjan Sen Department of Biology, Brandeis University, Waltham, MA, USA Dr. Mark Shlomchik Section of Immunobiology, Yale University School of Medicine, New Haven, CT, USA Dr. Robero Sitia Department of Molecular Pathology and Medicine, Universita Vita-Salute San Raffaele, DIBIT-HSR Scientific Institute, Milan, Italy Dr. Janet M. Stavnezer Department of Molecular Genetics and Microbiology, University of Massachusetts Medical School, Worcester, MA, USA Dr. Lisa A. Steiner Department of Biology, Massachusetts Institute of Technology, Cambridge, MA, USA Dr. Freda Stevenson Molecular Immunolgy Group, Tenovus Laboratory, Southampton University Hospitals Trust, Southampton, UK Dr. Eric Sundberg Center for Advanced Research in Biotechnology, W. M. Keck Laboratory for Structural Biology, University of Maryland Biotechnology Institute, Rockville, MD, USA Dr. Naoya Tsurushita Protein Design Labs, Inc., Fremont, CA, USA Dr. Maximiliano Vásquez Protein Design Labs, Inc., Fremont, CA, USA Dr. Ulrich H. von Andrian The Center for Blood Research and the Department of Pathology, Harvard Medical School, Boston, MA, USA Dr. Urich H. von Andrian Department of Pathology, The Center for Blood Research, Boston, MA, USA Dr. Gregory W. Warr Department of Biochemistry, and Center for Marine Biomedicine and Environmental Sciences, Medical University of South Carolina, Charleston, SC, USA Dr. Jurgen Wienands Department of Biochemistry & Molecular Immunology, Universität Bielefeld, Abteilung Biochemie I, Bielefeld, Germany Dr. Catherine Willett Phylonix Pharmaceuticals, Inc., USA
Dr. Andreas Radbruch Deutsches Rheumaforschungszentrum Berlin, 10117 Berlin, Germany
Dr. Gillean Wu Dean, Faculty of Pure and Applied Science, York University, Toronto, Ontario, Canada
Dr. Klaus Rajewsky Harvard Medical School, Center for Blood Research, 200, Longwood Avenue, Boston, MA, USA
Dr. Hans G. Zachau Adolf-Butenandt-Institut Molekularbiologie, Muenchen, Germany
Dr. Jeffrey V. Ravetch Laboratory of Molecualr Genetics and Immunology, The Rockefeller University, New York, NY, USA
Dr. Zhixin Zhang Howard Hughes Medical Institute, University of Alabama at Birmingham, Birmingham, AL, USA
Contents
5. The Mechanisms of V(D)J Recombination
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JOANN SEKIGUCHI, FREDERICK W. ALT, AND MARJORIE OETTINGER
1. Human Immunoglobulin Heavy Chain Locus
Antigen Receptor Gene Assembly 62 Mutational Analyses of Recombination Signal Sequences 64 “Beyond 12/23” Restriction of V(D)J Rearrangements 64 Influence of Coding Flanks 65 The Biochemistry of V(D)J Cleavage 65 RAG1/2-RSS Binding 66 RAAG1/2 Post-Cleavage Complex 67 A Role for HMG1 (or HMG2) in V(D)J Recombination 67 A Closer Look at RAAG1 and RAG2 68 Colding and Signal Joint Formation Requires the NHEJ Pathway 71
FUMIHIKO MATSUDA
Organization of the Human VH Locus 2 Analysis of Human VH Segments 7 Evolution of the Human VH Locus 10 Human CH Locus 12
2. Immunoglobulin Heavy Chain Genes of Mouse ROY RIBLET
Igh-V or VH Genes of the Ighb Haplotype 19 Polymorphism in VH Genes 20 Evolution 24 Genomic Considerations 24
6. Transcription of Immunoglobulin Genes KATHRYN CALAME AND RANJAN SEN
3. Immunoglobulin K Genes of Human and Mouse
Transcriptional Regulatory Elements in Immunoglobulin Heavy and Light Chain Genes 83 Proteins Binding in Ig Transcriptional Regulatory Elements 86 Areas of Current Research 89 Discoveries Resulting from the Study of Ig Gene Transcription 93
HANS G. ZACHAU
General Features of Human and Mouse K Genes 27 Human Immunoglobulin K Genes 27 Mouse Immunoglobulin K Genes 30 Aspects of Evolution of the K Genes 33
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
MARIE-PAULE LEFRANC AND GÉRARD LEFRANC
FRITZ MELCHERS AND PAUL KINCADE
IGL Genes and IMGT-ONCOLOGY 37 The Human IGL Genes 40 The Mouse IGL Genes 50
Three Waves of Hematopoiesis During Embryonic Development 101
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Pluripotent Hematopoietic Stem Cells 103 Pathways of Hematopoietic Progenitor Cells Toward B Lymphocyte Lineage Commitment and Differentiation 106 Control of Lymphoid Cell Development by Transcription Factors 107 Plasticity if PAX-5–Deficient Pre-B Cells 109 The Surrogate Light Chain 110 Pre-B Cells and Their Differentiation to More Mature B Lineage Cells 112 Rearrangements at the L Chain Loci at the Transition from Large to Small Pre-B-II Cells 114 Immature B Cells 116 Selections of Immature B Cells 117
8. Allelic Exclusion, Isotypic Exclusion, and the Developmental Regulation of V(D)J Recombination
10. Development and Function of B Cell Subsets JOHN T. KEARNEY
Selection and Differential Survival Mechanisms—B Cell Receptor Signaling 156 Compartmentalization of B Cell Subsets 157 Other Factors Involved in Formation of B Cell Subsets 157
11. Structure and Function of B Cell Antigen Receptor Complexes MICHAEL RETH AND JURGEN WIENANDS
Structure of the BCR Complex 161 Coupling Between the BCR and SYK 162 Redox Regulation of BCR Signaling 162 ITAM- and Non-ITAM-Controlled Signaling Pathways to SLP-65 163 ITAM-Independent Signaling and Fine-Tuning 165
MICHAEL S. KRANGEL AND MARK S. SCHLISSEL
Rag Expression 127 The 12/23 Rule 128 Accessibility Hypothesis 128 Enhancer and Promoter Control of V(D)J Recombination 128 Trans-Acting Factors 130 Chromatin Dynamics and V(D)J Recombination 130 Ordered Rearrangement Within Ig and TCR Loci 132 Allelic Exclusion at Ig and TCR Loci 133 Ig Light Chain Isotypic Exclusion 136 Future Directions 136
9. The Development of Human B Lymphocytes PETER D. BURROWS, TUCKER LEBIEN, ZHIXIN ZHANG, RANDALL S. DAVIS, AND MAX D. COOPER
Stages of Human B Cell Differentiation 141 Sites of Human B Cell Development 143 Human Immunoglobulin Genes 143 The Role of Surrogate Light Chains in Human B Cell Development 144 Repertoire Diversification via Receptor Editing and VH Replacement 145 Regulation of Antibody Production by B Cell Receptors 147 Immunodeficiency Diseases 148 B Lineage Leukemia 149
12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22 LARS NITSCHKE AND DOUGLAS T. FEARON
CD19 171 Inhibitory Co-Receptors on B Cells 177
13. The Dynamic Structure of Antibody Responses IAN C. MACLENNAN AND DEBORAH L. HARDIE
Three Routes to Antibody Production 187 Stages of Adaptive Antibody Responses 187 How and Where B Cells Encounter Antigen 188 Primary Cognate Interaction of B Cells with Primed T Cells 189 Exponential Growth of Activated B Cells 190 Proliferation, Hypermutation, and Selection in GC 192 Sustained Survival of Memory B Cell Clones and Plasma Cells 197
14. Dynamics of B Cell Migration to and within Secondary Lymphoid Organs JASON G. CYSTER AND ULRICH H. VON ANDRIAN
Lymphoid Organ Entry 203 Compartmentalization of Mature B Cells 209 B Cells at Sites of Inflammation 213 Homing of Antibody Secreting Cells (ACSs) 213
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15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System PER BRANDTZAEG, H. CRAIG MORTON, AND MICHAEL E. LAMM
Immune-Inductive Tissue Compartments 223 Characteristics of B Cells in Secretory Effector Tissues 226 B-Cell Stimulation in MALT Structures 229 Class Switch and Ig A Isotype Promotion 232 Mechanisms Directing Homing and Retention of Mucosal B Cells 234 What Is Actually Known About Human Mucosal B Cells? 238
16. The Cellular Basis of B Cell Memory KLAUS RAJEWSKY AND ANDREAS RADBRUCH
Generation of B Cell memory and Memory B Cells in T Cell-Dependent Antibody Responses 247 Memory Plasma Cells 252 Adaptive B Cell Memory 254
17. Immunoglobulin Assembly and Secretion
19. Regulation of Class Switch Recombination MICHEL COGNÉ AND BARBARA K. BIRSHTEIN
CSR Requires Specific Stimuli Occurring in a Defined Germinal Center (GC) Microenvironment 289 Proximal CIS Regulatory Elements for GT 291 Distant Regulatory Region for GT and CSR: The 3¢ IGH Enhancers 293 Mechanisms for 3¢ IGH Regulatory Region-Mediated Regulation of GT 295 Coordinated Regulation of Transcription, Recombination, and Replication 300
20. Molecular Mechanisms of Class Switch Recombination JANET STAVNEZER, KAZUO KINOSHITA, MASAMICHI MURAMATSU, AND TASUKU HONJO
Outline of Mechanisms for CSR 307 Isotype Specificity of CSR 312 AID, The Sole B Cell-Specific Factor Required for CSR 313 Cleavage of the S Region 314 Processing and Joining of DNA Ends After Cleavage 315 Comparison of CSR with SHM 319
LINDA M. HENDERSHOT AND ROBERTO SITIA
Mechanisms of IG Synthesis and Assembly 261 Multiple Layers of Quality Control Exist to Aid and Monitor the Assembly of Functional IGs 264 Transport of Assembled IG Molecules to the Golgi 267 Degradation of Misfolded and Unassembled IG Subunits 267 Differentiation to Plasma Cell 268
18. Fc and Complement Responses JEFFREY V. RAVETCH AND MICHAEL C. CARROLL
Consequences of FCgRIIB Deficiency 275 Consequences of Complement and Complement Receptor Deficiencies 276 Fc Receptors 2276 Complement Receptors 280 Co-Receptor Signaling Versus Antigen Localization to FDC 281 Frontiers: Complement Versus Fc Receptors 285
21. Molecular Mechanisms of Hypermutation NANCY MAIZELS AND MATTHEW D. SCHARFF
Characteristics of Somatic Hypermutation of Immunoglobulin Variable Regions 327 Activation and Targeting of Hypermutation by Transcription and CIS-Elements 329 Hypermutation Occurs Within a Limited Window of B Cell development 330 The AID Gene Is Critical for Hypermutation 331 Phase One of Hypermutation: C Æ U Deamination and Base Excision Repair 332 Mismatch Repair Factors in Phase Two of Hypermutation 332 DNA Breaks in Hypermutation 334 Competing Pathways of Repair: Error-Prone DNA Synthesis or Strand Transfer 335 Evolution and Hypermutation 335
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22. Selection During Antigen-Driven B Cell Immune Responses: The Basis for High Affinity Antibody MARK J. SHLOMCHIK
Overview of the B-Cell Immune Response 339 Affinity Maturation in the Early Stages of the B-Cell Immune Response 341 The CG Is a Second Major Site for Affinity Maturation 342 Affinity-Based Selection Continues After the GC Reaction Has Ended 344 An Integrated View of the Strategic Design of the B-Cell Immune Response: Future Directions 344
23. Chromosomal Translocations in B Cell Leukemias and Lymphomas A. THOMAS LOOK AND ADOLFO FERRANDO
Translocations Associated with a Block in Lymphoid Differentiation 349 Translocations Associated with Suppression of Apoptosis During Lymphoid Development 352 Translocations Associated with Increased Proliferation in Lymphoid Precursors 354 Activation Cell Cycle Regulation in Mantle Cell Lymphoma and Myeloma 355
24. Classification and Characteristics of Mouse B Cell–Lineage Lymphomas
26. Immunodeficiencies Caused by B Cell Defects FRANCISCO A. BONILLA AND RAIF S. GEHA
Clinical Features of the Agammaglobulinemias 403 Autosomal Recessive Hyper-IGM Syndrome 410 Murine Models of Human B-Cell Deficiency 411
27. Diverse Forms of Immunoglobulin Genes in Lower Vertebrates GARY W. LITMAN, MARTIN F. FLAJNIK, AND GREGORY W. WARR
Cartilaginous Fish: An Unusual Example of Gene Multiplicity 417 Bony Fish: IG Heavy Chain Genes Resemble IgM and IgD 419 Lobe-Finned Fish: A “Transitional” Arrangement of Recombining Elements 422 Fleshy-Finned Fish: An Ancient Origin for Isotype Diversity 422 Amphibians and Reptiles: The Possible Origins of Class Switching 422 Light Chain Genes: Diverse Structures and Organization in Lower Vertebrates 423 Transcriptional Control of IG Genes in Lower Vertebrates 425 A Unifying Hypothesis to Explain the Origins of the Adaptive Immune Receptor 427 Immune Molecules in Jawless Vertebrates 427 Protochordates: Different Contexts for Diversified B Regions 428
HERBERT C. MORSE III
Comparative Classification of Mouse and Human B Cell–Lineage Neoplasms 366 Characteristics of Mouse B Cell–Lineage Lymphomas 368 Pathogenesis 373
25. B Cells Producing Pathogenic Autoantibodies
28. Immunoglobulin Genes and Generation of Antibody Repertoires in Higher Vertebrates: A Key Role for GALT DENNIS LANNING, BARBARA A. OSBORNE, AND KATHERINE L. KNIGHT
Avians 433 Lagomorphs 436 Artiodactyls 440 Other Mammals 444
CONSTANTIN A. BONA AND FREDA K. STEVENSON
Subsets of Autoantibodies 382 Criteria to Define Pathogenic Autoantibodies 383 Genetics of Autoantibodies 384 Molecular and Immunochemical Characteristics of Human Pathogenic Autoantibodies 388 Human Pathogenic Autoantibodies with Murine Counterparts 392
29. The Zebrafish Immune System LISA A. STEINER, CATHERINE E. WILLETT, AND NADIA DANILOVA
Hematopoiesis 450 Adaptive Immunity in Zebrafish: Organs and Molecules 452
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Genetic Approaches 457 Major Histocompatibility Complex (MHC) 460 Innate Immunity 460 Infection 464
30. The Origin of V(D)J Diversification SUSANNA M. LEWIS, GILLIAN E. WU, AND ELLEN HSU
The Alien Seed 473 The Evolution of BCR and TCR Loci 481 Considerations on the UR-V Gene 483
Antibody Phage Display 516 Use of Phage Display to Bypass Hybridoma Technology 519 Use of Phage Display to Bypass Immunization 520 A Comparison of Different Phage Antibody Library Types and Applications 521 Strategies for Selection of Phage Antibodies 522 Increasing Antibody Affinity Using Phage Display 522 Alternative Antibody Display Technologies 524
33. Humanization of Monoclonal Antibodies NAOYA TSURUSHITA AND MAXIMILIANO VASQUEZ
31. Antibody Structure and Recognition of Antigen ERIC J. SUNDBERG AND ROY A. MARIUZZA
A Structural Framework for Molecular Recognition 491
Murine, Chimeric, and Humanized Antibodies 533 Computer-Guided Design of Humanized V Regions 534 Other Humanization Methods 537 Immunogenicity of Humanized Antibodies 540 Humanized Antibodies Approved for Clinical Use 540
32. Monoclonal Antibodies from Display Libraries
34. Human Monoclonal Antibodies from Translocus Mice
JIM MARKS
MARIANNE BRÜGGEMANN
Overview of Antibody Phage Display 513 Prokaryotic Expression of Antibody Fragments 514 Generation of Antibody Gene Repertoires Using the Polymerase Chain Reaction 515
Human IG Transloci 547 The Mouse Strains 552 Index 563
Preface
bly was tied to the developmental programs of B and T lymphocytes. Moreover, we had begun to get a glimpse of how the expression of Ig receptors and other molecules on the surface of particular types of B lineage cells was linked to aspects of B cell development and function. The second edition of Immunoglobulin Genes appeared in 1995. Over the intervening six years, tremendous progress had been made on several fronts. Substantial organizational information had been obtained with respect to the IgH and Igk loci in humans and mice. In fact, the complete IgH V region locus had been isolated on overlapping cosmids and YACs. A huge breakthrough came from the identification of the developing lymphocyte-specific RAG gene products, which are the specific components of the site-specific VDJ recombinase required to assemble Ig and TCR variable region genes in B and T cells, respectively. The availability of the RAG products allowed dissection of the VDJ recombination mechanism in detail and also facilitated identification of the generally expressed nonhomologous DNA end-joining proteins, which are co-opted by the VDJ reaction to complete the joining phase of V(D)J recombination. Gene targeted mutational studies had also begun to be employed to test the function of Ig genes and some of their regulatory sequences as well as that of other molecules that function in developing and mature B lymphocytes. In the eight years that have passed since the publication of the second edition of Immunoglobulin Genes, we have witnessed remarkable progress on several fronts. First, as anticipated, the IgH and IgL loci have now been fully sequenced in humans and mice. In fact, the sequences of the entire genome of human, mouse, and many other organisms have now been obtained. Another formidable advance has been the elucidation of the basic mechanisms underlying CSR and SHM that are critical to generation of antigenspecific antibodies. The explosion of information on these processes was stimulated by the discovery of AID, an enzyme that is fundamental to both CSR and SHM, as well as to the gene conversion process that diversifies chicken Ig
Studies of immunoglobulin (Ig) genes have been one of the major focal points in modern biology. The apparently unique property of Ig gene loci, and their related T cell receptor loci, to be somatically assembled from germline gene segments has long fascinated biologists. Moreover, studies of the mechanisms by which Ig genes are expressed, and how this relates to the development of B lymphocytes, have contributed much to our understanding of fundamental genetic and cellular processes, ranging from transcription, differential RNA processing, site- and region-specific recombination, general DNA repair, and cellular signaling mechanisms. Because of the unique insights and wealth of instructive materials that derived from these studies, 14 years ago we decided to cover the field and its advances in a book on Immunoglobulin Genes. Eight years ago, the second edition of Immunoglobulin Genes continued to track this progress, and now we have continued and expanded this coverage in the third edition of Immunoglobulin Genes, which we have entitled Molecular Biology of B Cells. The change in title reflects the increased scope of the current volume, which covers the elucidation of the intimate links between Ig genes and many of the fundamental processes involved in generating and affecting the humoral arm of the immune response. The first edition of Immunoglobulin Genes appeared in 1989. At that time, the advent of DNA cloning and molecular biology had allowed a relatively full elucidation of the dynamic mechanisms involved in the somatic assembly of Ig variable (V) region gene segments. We knew then, at least in general terms, about some of the basic aspects of the V(D)J recombination, IgH class switch recombination (CSR), and somatic hypermutation (SHM) processes that form the fundamental basis for the diverse humoral immune response. In particular, molecular genetic approaches had provided an enormous amount of information on the structure, organization, assembly, and expression of Ig genes in a variety of organisms and had also provided the tools to investigate the general mechanisms by which Ig gene assem-
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genes. Much like the impact that the discovery of RAG had on elucidating the VDJ recombination process, the discovery of AID has now allowed the generation of new insights into the mechanisms that work to effect SHM and CSR at the mechanistic level. In another five years, it is likely that our knowledge of these processes will advance to the level that we now understand VDJ recombination. As for VDJ recombination, we have learned that the RAGs likely evolved from elements of a transposase, a finding that along with other aspects Ig Gene organization has provided some notion about how the Ig gene assembly system may have evolved. Studies of AID and its relatives are beginning to provide similar insights into the evolution of the CSR and SHM processes. As anticipated, gene targeted mutational studies have continued to provide great insights into the function of Ig genes and the mechanisms that regulate their expression, and have also helped to elucidate the function of many other molecules involved in the differentiation of B cells and in their activation and effector functions. Exciting developments have also take place in the area of the cellular dynamics that regulate migration of B cells for participation in effector functions at specific locations of the body. Application of immunoglobulins to clinical fields, not only diagnosis but also therapy, and understanding of molecular basis of human B cell defects and malignancy has also witnessed remarkable advances. Still, there are major questions remaining to be solved at many levels and with respect to many processes. One that
has been particularly enigmatic is the molecular details of how accessibility regulation occurs at the level of the chromosome and chromatin structure. This clearly is a problem that is not restricted to B cells but bears on generally relevant control mechanisms. There are also many aspects of B cell physiology (such as the mucosal antibody response and the biology of memory B cells) that remain to be fully dissected by molecular approaches. However, the new technologies now available will hopefully allow substantial advances to be made in the study of all these processes before the next edition appears. The chapters of Molecular Biology of B Cells, as in previous editions, are written by authors who have very actively participated in the accumulation of knowledge in the area that they cover. Moreover, in this edition, we have tried something new; now most chapters are authored by two separate authorities on the subject covered. In this way, we have hoped to achieve the most balanced view of each individual field and, in some cases, to generate novel points of view from the cooperative efforts of authors with somewhat different viewpoints. As before, we continue to look forward to many more exciting developments in research on Ig genes and B lymphocyte development and function over the next five years. Tasuku Honjo Frederick Alt Michael Neuberger
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1 Human Immunoglobulin Heavy Chain Locus FUMIHIKO MATSUDA Centre National De Genotypage 2 Rue Gaston Cremieux, 91057 Evry Cedex France and Department of Genome Epidemiology, Kyoto University Graduate School of Medicine, Yoshida Sakyo-Ku, Kyoto 606 Japan
provides evidence to estimate the relative contribution of germline VH repertoire, VDJ recombination, somatic hypermutation, and subsequent selection of B lymphocytes to the Ig repertoire. Accumulating evidence indicates that the human VH locus is highly polymorphic. It is interesting to investigate the polymorphic variation of the number and repertoire of germline VH segments and CH genes and its association with disease susceptibility. It is known that some VH segments are overrepresented in the antibody repertoire, suggesting that the utilization of each VH segment may not be random. It is important to know whether the germline organization or structure of VH segments predicts such a preferential usage. Isolation of the total human VH segments has played important roles in the generation of human Ig in J segment–disrupted mice carrying human Ig mini loci (Taylor et al., 1994; Green et al., 1994; Mendez et al., 1997). Finally, from an evolutionary viewpoint, multigene families are considered to have evolved through repeated duplication and recombination of DNA. Diversification of newly generated VH members by such events contributes to the germline VH repertoire. Evolutionary studies of VH organization and structure and, in particular, comparison of VH loci between related species, will provide insight to the molecular mechanisms that govern the evolution of multigene families. Needless to say, studies on the complete organization of the CH locus are the basis for understanding the molecular mechanism for class switching and regulation of IgH expression (see the chapter by Honjo). Significant progress was made in the structural analysis of the human IgH locus between the first (1988) and second (1995) editions of this book by completion of the physical mapping of the human VH segments using YAC clones. Since then, with a rapid evolution of genome sequencing technology, the complete nucleotide sequence of the entire
The immunoglobulin (Ig) molecule is composed of heavy (H) and light (L) chains, both of which consist of variable (V) and constant (C) regions. The V region is responsible for antigen binding, whereas the CH region specifies the isotype of Ig. Genes encoding IgH V regions are split into VH, diversity (DH), and joining (JH) segments. One each of the three segments is generally assembled into a functional VH gene by a somatic genetic event called VDJ recombination. In Homo sapiens, VH region genes are mapped to chromosome 14 q32.33 (Croce et al., 1979; Kirsch et al., 1982). Recent completion of the nucleotide sequence of the 957kilobase (kb) DNA covering the entire human VH locus demonstrated that it consists of 123 copies of VH segments, 26 DH segments, and six JH segments. Conversely, the human CH locus comprises 11 CH genes, of which 2 are pseudogenes (Matsuda et al., 1998). The VH and CH loci are physically linked on the chromosome in the order of 5¢-VHDH-JH-CH-3¢. The distance between the 3¢-most VH locus segment (JH) and the 5¢-most CH gene (Cm) is approximately 8 kb in man (Ravetch et al., 1981). The 298-kb DNA of the entire human CH locus was also sequenced (Heilig et al., 2003; Nicodeme et al., submitted). Thus, the IgH locus, which combines the VH and CH loci, constitutes a huge multigene family, encompassing the 1.3-Mb DNA of the distal end of chromosome 14. The complete knowledge of the organization and structure of the IgH locus will provide clear answers to a number of questions essential to Ig repertoire formation and Ig expression. Obviously, the total number of VH segments determines the upper limit of the germline Ig repertoire, although somatic genetic events, including VDJ recombination, hypermutation, and gene conversion further tremendously amplify the expressed repertoire. Comparisons between the total germline and expressed VH sequences
Molecular Biology of B Cells
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Copyright 2004, Elsevier Science (USA). All rights reserved.
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human IgH locus is now available. This achievement provides an enormously beneficial reference to map expressed VH genes and their polymorphisms, as well as a detailed structure of the human CH locus.
ORGANIZATION OF THE HUMAN VH LOCUS Studies on the physical mapping of the human VH locus were initiated by cosmid cloning (Kodaira et al., 1986). The distribution of VH families on 23 cosmid clones has shown that members of different VH families are interspersed, in contrast to the finding that the same family members tend to cluster in the mouse VH locus (Kemp et al., 1981; Rechavi et al., 1982). Another important conclusion from early studies is the presence of abundant pseudogenes (about 40%), many of which are highly conserved, with only a few point mutations (Givol et al., 1981; Kodaira et al., 1986). The complete physical map was constructed for the 80-kb of DNA encompassing the 3¢-most V6-1 segment, DH cluster, and JH cluster (Matsuda et al., 1988; Buluwela et al., 1988; Buluwela and Rabbits, 1988; Sato et al., 1988; Schroeder et al., 1988). A more general overview of the whole human VH locus has been provided by studies using pulse field gel electrophoresis (PFG), which allowed examination of the VH content on a few hundred kb to 1,000 kb DNA fragments. The total size of the human VH locus was estimated to be about 2.5 to 3.0 Mb (Berman et al., 1988; Matsuda et al., 1988), including the D5-hybridizing fragments that later mapped to chromosome 15. PFG analysis using twodimensional electrophoresis has provided a more precise organization of the entire VH locus of about 1.2 Mb, on which 76 human VH segments were mapped (Walter et al., 1990). The same group further refined the mapping using the deletion profile of VH segments associated with VDJ recombination in human B cell lines (Walter et al., 1991a). Introduction of the yeast artificial chromosome (YAC) vector has been key to completing the physical mapping of the entire human VH locus (Figure 1.1). The first report using YAC cloning has identified and located five VH segments proximal to the DH and JH segments (Shin et al., 1991). These authors proposed to rename all the VH segments by the family number and the order from the 3¢ end of the VH locus. The nomenclature of VH segments was controversial not only because investigators named VH segments in their own way, but also because many expressed VH sequences containing somatic mutations could not be easily assigned as different VH segments. The newly proposed nomenclature defined VH segments only when they were mapped on the chromosome, which has been well accepted by the scientific community. The same group in Kyoto has completed the mapping of 64 VH segments in 0.8 Mb of the human VH locus
by analyzing more than seven overlapping YAC clones. The nucleotide sequences of all these VH segments were determined (Matsuda et al., 1993). The transcriptional orientation of 43 VH segments that are located either 5¢-most, middle, or 3¢-most part of the contig was determined. All had the same polarity as the JH segments, unlike in the human Vk locus, where the gross inversion of the distal duplicated copy of 360-kb DNA containing 59 Vk segments and relics is observed (Kawasaki et al., 2001, see Chapter 3). The physical mapping of the human VH locus was completed by identification of a YAC clone in human subtelomeric region–specific YAC libraries (Cook et al., 1994). A 200-kb clone that physically links to the upstream portion of the 0.8Mb contig was isolated and 17 novel VH segments were identified, suggesting that the total number of the VH segment is 81. The same region was cloned independently by the Kyoto group, confirming the telomeric end of the physical map by the Cambridge group. The last and biggest effort to complete the physical map was made by the Kyoto group. The complete nucleotide sequence of the 957,090-bp DNA upstream of the human JH cluster was determined (Matsuda et al., 1998). The 5¢-most part of the locus contains a diverged human telomere repeat and subtelomeric region, confirming the proximity of the VH locus to chromosome 14q telomere. A total of 123 VH segments and pseudogenes was identified in the 883-kb DNA between 73 and 956 kb upstream of the JH cluster. The highly interspersed organization of the VH segments belonging to seven different families was confirmed. Sixteen of the 17 distal VH segments were identified at the position proposed. However, the VH sequence corresponding to the V777 segment was not identified at the suggested position even though the physical maps of the corresponding portions are exactly identical. V3-82P, the 5¢-most VH segment, is located 1,480 bp downstream of the 5¢ terminus of the locus. The distances between neighboring VH segments are quite variable, with the largest being 41.4 kb (between V1-2 and V41.1P) and the smallest as little as 418 bp (between V3-67.2P and V4-67.1P). However, clustering of VH segments, as shown in the human Vl locus, was not evident (Frippiat et al., 1995; Kawasaki et al., 1997). The transcriptional polarities of all the VH segments are the same as that of the JH segments. Southern hybridization detects many DNA fragments in YACs and cosmids that weakly hybridize with VH probes, although such hybridization is not detectable against human genomic DNA, thus suggesting the presence of additional VH-related sequences including those of novel VH families. Nucleotide sequencing newly identified 43 such sequences. However, all these VH segments were classified as a member of seven known VH families, excluding the existence of novel VH families in humans. Interestingly, they all carry defects in their structure and are categorized as pseudogenes.
1. Human Immunoglobulin Heavy Chain Locus
3
FIGURE 1.1 Organization of the entire human IgH locus. Five thick horizontal lines show the 1.3-MB DNA with the 3¢ end at the bottom right corner. VH segments are indicated by vertical lines with their names (newly identified VH segments are shown with asterisk). VH segments containing truncations are shown by shorter vertical lines. DH segments and CH genes are indicated by diamonds and open boxes, respectively. Thirteen locus-specific repeats are indicated below by boxes of different pattern. Enhancers, predicted matrix attachment region, and nonimmunoglobulin genes are also shown with their names. Modified from Matsuda et al. (1998).
The Total Number of VH Segments One of the most important goals in the study of the human VH locus is to determine the total number of functional VH segments that can participate in functional heavychain formation. Some discrepancy was noted regarding the classification of VH segments into functional and pseudo-
genes, in part due to the incomplete nucleotide sequence of some VH segments. Given the complete nucleotide sequencing of the human VH locus, 123 VH segments were classified into four different categories based on the following criteria (Matsuda et al., 1998). The 79 VH segments without open reading frames (ORF) (due to various defects including frame shift and truncation) were classified as pseudogenes.
4
Matsuda
The other 44 VH segments with a complete ORF were further subdivided into “functional,” “transcribed,” or “ORF” group as follows:
that ancient truncation events were followed by gene duplication.
• The “functional” VH segments have an intact exonintron structure, a complete ORF, and no fatal defects in recombination signal sequences (RSS). In addition, their expression was confirmed by identification of the corresponding full-length VH mRNAs. • The “transcribed” VH segments correspond to those whose sequence identity with partial VH mRNA sequences have been identified. • The “ORF” segments consist of the VH segments with a complete ORF, yet not demonstrated to be transcribed (Table 1.1). Obviously, the direct proof of the functional VH segment is to identify its sequence in the IgH amino acid sequence.
Polymorphism of the Human VH Segments
Among 44 VH segments that have a complete ORF, 39 VH segments are translated and classified as functional. V428 is >97% identical to partial VH mRNA sequences in the database. However, its translation product remains to be identified and, hence, it was classified as “transcribed.” The remaining four genes, namely, V3-16, V3-35, V3-38, and V7-81, were categorized as “ORF” due to the absence of their transcripts in the database. Indeed, V3-38 has a truncation at the 5¢ untranslated region that results in the complete loss of its 5¢ regulatory region. Moreover, these four VH segments carry a diverged RSS heptamer sequence. They might not be employed for VDJ rearrangement. The 79 VH segments classified as pseudogenes were subdivided into 29 VH segments with point mutations and 50 with truncation(s) (Table 1). Indeed, none of these 79 VH segments corresponded to any VH mRNAs. Interestingly, 12 VH3 pseudogenes have the 5¢ truncation at the same position in their introns. Similarly, 13 VH4 pseudogenes contain the common 5¢ truncation in the second exon, suggesting
RFLP and DNA sequencing have shown a number of polymorphic VH alleles. Among a variety of polymorphisms in the locus, insertion/deletion of VH segments and single nucleotide polymorphisms (SNPs) within the coding region are more likely to have functional significance. One obvious possibility is the expansion of repertoire. Polymorphic VH may affect the affinity of the antibody for its ligand, as even mutations in framework residues of the Ig have been shown to influence the binding affinity (Foote and Winter, 1992). Furthermore, expression of particular allelic variants could influence the efficiency of H-L chain pairing or interaction with B cell super-antigens. Variation of the germline VH segment copy number may associate with the preferential utilization and expression level of specific VH segments (Sasso et al., 1996). To date, three insertion/deletion polymorphisms have been mapped along the human VH locus. An insertional V74.1 segment is present between V2-5 and V4-4 in 65% and 72% of alleles among the Caucasian and Japanese populations, respectively. Another frequent insertion polymorphism of 50-kb DNA containing five functional VH segments was identified in the region between V3-31 and V3-30 by “HAPPY mapping” (Walter et al., 1993); this polymorphism is present in 73% of the Caucasian population. Interestingly, the insertion is located between a tandem homology pair of three VH segments, namely, V3-33/V3-32P/V4-31 and V330/V3-29P/V2-28, suggesting the highly recombinogenic nature of the region. A large insertion polymorphism of 80kb DNA, containing at least one each of VH2- and VH3family segments, was localized in the region between V2-70 and V1-67P by 2D-PFGE analysis (Walter et al., 1990). One
TABLE 1.1 Summary of the human VH segments VH family Chromosome
Classification
1
2
3
4
5
6
7
Total
14q32.33
Functional Transcribed ORF
9 0 0
3 0 0
19 0 3
6 1 0
1 0 0
1 0 0
0 0 1
39 1 4
Pseudogene Point mutator Truncation
3 2
1 0
21 22
2 23
0 1
0 0
2 2
29 50
14
4
65
32
2
1
5
123
15q11
Total
6
0
1
1
0
0
0
8
16p11
4
1
11
0
0
0
0
16
The number of VH segments on chromosome 14 is calculated by the results from Matsuda et al. (1998). VH segments with polymorphic insertion are not included. Information on VH segments on chrmonsome 15 and 16 is taken from Nagaoka et al. (1994) and Tomlinson et al. (1994).
1. Human Immunoglobulin Heavy Chain Locus
of the most polymorphic VH segment is V1-69, having 13 known alleles including duplication and deletion (Sasso et al., 1993); this segment is located very close to the region of the large polymorphic deletion. In addition, several VH segments mapped to chromosome 14 have not been located in the current map, suggesting the possibility of other deleterious polymorphisms. It is important to test whether VH polymorphisms are associated with disease susceptibility. However, this is a controversial area; some reports suggested the association of VH polymorphisms with autoimmune diseases such as rheumatoid arthritis, systemic lupus erythematosus, and multiple sclerosis (Yang et al., 1990; Walter et al., 1991b), whereas others failed to find a clear association (Hashimoto et al., 1993; Shin et al., 1993a). This might be due to the usage of only a limited number of VH segments or polymorphic markers in the analysis. To address this question, it would be essential to perform a large-scale genetic analysis of the entire VH locus. An SNP genotyping program is under way to determine the functional VH segments and the pseudogenes among a large number of DNA samples of multiple ethnic backgrounds (F. Matsuda unpublished). Preliminary results show that the content and frequency of SNPs differ largely between individual VH segments. The V1-69 segment has as many as eighteen alleles in ninety-six Japanese DNAs, a rather homogenous population, with different copy numbers among individuals, whereas no SNPs are detected in another VH segment. Other VH segments that have copy-number variation in the Caucasian population are V1-2, V3-23, V2-26, and V2-70. The systematic screening of VH locus haplotypes, generated by the combination of identified VH alleles against a large cohort of autoimmune and immune deficiency patients, will provide some ideas on possible associations.
Organization of the Human DH Segments The 70-kb region of the 3¢-most part of the human VH locus is occupied by DH and JH gene segments (Figure 1.1). The human JH cluster contains three pseudo JH segments interspersed among six functional JH segments (Ravetch et al., 1981). These are clustered approximately 8 kb upstream of Cm, the 5¢-most constant region gene. A human counterpart to the murine DQ52 segment is located about 100 bp upstream of the JH1 segment. Initially, a family of DH segments (D1-D4 or DLR1-DLR4) was identified, using as a probe an aberrantly rearranged DH-JH segment in a CLL cell clone (Siebenlist et al., 1981). Physical mapping studies showed that these are ordered at regular 9-kb intervals along the chromosome, suggesting the generation of the human DH cluster by gene duplication. However, the fact that these four segments corresponded to a smaller part of DH sequences in functional VDJ rearrangement raised a possibility of novel DH families in the genome. Later, a number of additional
5
human DH segments were identified, including ones homologous to the murine DFL16 segments as well as a number of those that are markedly dissimilar in size and sequence (Schroeder et al., 1987; Zong et al., 1988; Ichihara et al., 1988a, b; Buluwela et al., 1988; Sonntag et al., 1989; Shin et al., 1993b). Five novel DH families (DM, DXP, DA, DK, and DN) were identified in the order of 5¢-DM-D(LR)DXP-DA-DK-DN-3¢ by the nucleotide sequencing of a 15-kb DNA fragment covering the D(LR)1 segment and flanking regions (Ichihara et al., 1988b). Southern blot analysis strongly suggested the existence of a set of six DH segments in each copy of duplicated 9-kb DNA. An additional DH family (DIR) with unusual structure was identified in the 5¢ adjacent portion of DM family segments. The possible involvement of the DIR family in D-D rearrangement was pointed out because of its irregular spacer length (23 bp) of RSS. The definitive answer to the content of DH segments was given by the nucleotide sequencing analyses of the entire human DH locus (Corbett et al., 1997). A total of 26 DH segments was identified in four tandemly arrayed copies of the 9-kb DNA (Figure 1.1). These consist of five DM and DXP segments and four each of D(LR), DA, DK, and DN family gene segments. However, the number of DH segments shows allelic variation. One example is the polymorphic deletion of the 9-kb DNA containing the D(LR)1 segment, which occurs at a high frequency among the Japanese population (48% of alleles are D(LR)1 negative) (Zong et al., 1988). An extensive analysis of DH segment usage in rearranged heavychain sequences classified the 27 DH segments—including the unique DQ52 segment—into 25 functional and two pseudogenes (DM2 and DN3) (Corbett et al., 1997). The authors demonstrated the highly biased utilization of different DH segments and reading frames. In contrast, no evidence was obtained for the usage of DIR segments, inverted DH segments, or DD recombination in functional VDJ rearrangements.
VH and DH Segments on Chromosomes 15 and 16 Although the VH locus is located at the telomere end of chromosome 14q32, several VH and DH clusters remained unmapped for some time. The first evidence that a DH segment is located on chromosome 15 was obtained by in situ hybridization (Chung et al., 1984). Subsequently, studies using in situ hybridization, as well as human/rodent somatic hybrid cells (Cherif and Berger, 1990; Matsuda et al., 1990; Nagaoka et al., 1994; Tomlinson et al., 1994), identified two VH orphon loci on chromosome 15q11 and chromosome 16p11. Studies of cosmid and YAC clones derived from these orphon loci revealed that approximately 40% of VH segments in both loci (three out of seven VH on chromosome
6
Matsuda
16 and one out of three VH on chromosome 15) are apparently functional, without any structural defects, in the coding region as well as in RSS (Nagaoka et al., 1994) (Figure 1.2). A totally different approach, based on PCR and using somatic cell hybrid DNAs as templates, specifically amplified 24 VH segments, including 10 apparently functional ones, on chromosomes 15 and 16 (Tomlinson et al., 1994). Among them, 16 VH segments (including the above 7 VH segments) were mapped on chromosome 16, of which 14 make seven pairs of closely related VH sequences. The authors pointed out the possibility of intrachromosomal duplication of the DNA containing the seven VH segments. Taken together, the total number of VH segments on chromosome 16 would be sixteen, although the critical test for the duplication depends on their mapping along the chromosome. Florescence in situ hybridization mapped two independent contigs containing human DH segments (D5-a and D5b) to chromosome 15q11–12 (Nagaoka et al., 1994). Each consists of five DH segments in the order 5¢-DM5-D(LR)5DXP5-DA5-DK5–3¢, whereas the DN segment, the 3¢-most DH segment in the DH clusters on chromosome 14 is absent (Matsuda et al., 1990). Nucleotide sequence homology of the corresponding DH segments is much higher between D5a and D5-b clusters than between the D5 and any of the
D1–D4 clusters. One of the DH clusters (D5-b) is flanked by three VH segments. Interestingly, these three VH segments are located 3¢ to the D5-b cluster, and one of them (V13C) is apparently functional, having a complete ORF. The polarity of one of them (V3) had the same transcriptional orientation relative to DH (Matsuda et al., 1990). Quantitative hybridization estimated the copy number of D5 clusters to be at least four (Nagaoka et al., 1994). Chromosomespecific PCR amplification identified eight VH segments on chromosome 15 (Tomlinson et al., 1994). Nucleotide sequencing of both of these translocated VH loci is under way as a part of the human genome project. The future completion of the sequencing will provide us with definitive information regarding the number and organization of orphon VH and DH segments. Moreover, the evolutionary origin and mechanisms of translocation will be elucidated through comparative structural analysis between these loci and those on chromosome 14.
Nonimmunoglobulin Genes in the Human VH Locus Computer-assisted homology searches using the 957-kb DNA identified eight DNA sequences that are highly
D5-a
D5-b
M XP K LR A
M XP K LR A
V3 V54 V13C
Chr.15 82% 83%
95%
2-26 1-24P 3-22P 3-21 3-25P 3-23
77%
3-16P 1-14P 1-12P 3-15 3-13 3-11 1-17P
Chr.14 3-20
1-18 3-19P
95%
93%
96% 95%
93%
93%
95%
Chr.16 (VH-F) F2-26
0
F3-16P F3-15
50
100
F1-14P F3-13 F3-11 F1-12P 150
200(kb)
FIGURE 1.2 Comparison of VH segments on chromosomes 15 and 16 with their counterparts on chromosome 14. Corresponding VH segments are indicated with the percentage of homologies of coding and intron sequences. Neighboring VH segments of V1-18 were compared with V3 or V13C on the D5-b region (shown by dashed lines). Modified from Nagaoka et al. (1994).
7
1. Human Immunoglobulin Heavy Chain Locus
homologous to known DNA sequences in the databases. The 7883-bp cDNA of the KIAA0125 gene (Nagase et al., 1995) showed 99.8% identity to the DNA sequence between the V6-1 segment and the D gene cluster (Figure 1.1). This gene is encoded by a single exon and its relative transcriptional orientation is opposite to the VH segments. KIAA0125 has several interesting features that are often found in imprinted genes, including an extremely short putative protein coding region (77 amino acids) and very long 5¢- and 3¢-untranslated regions (1,289 and 6,087 nucleotides, respectively) and the presence of tandem repeats of 68 and 48 bp units in the 3¢-untranslated region (Neumann et al., 1995). Interestingly, its expression is limited to lymphoid organs such as spleen, thymus, and peripheral blood leukocyte (Nagase et al., 1995). The other seven DNA sequences are homologous to the human ribosomal protein S8, the metalloprotease-like, disintegrin-like, cystein-rich protein (MDC) family of Macaca, the human leukemia virus receptor 1 (GLVR1) (2 copies), and the human golgin-245 (three copies). All are processed pseudogenes.
a
ANALYSIS OF HUMAN VH SEGMENTS VH Subgroups and Families Human VH regions were divided into three subgroups based on amino acid sequences (reviewed in Kabat et al., 1991). These protein subgroups have been further subdivided into seven distinct VH families defined by the nucleotide sequence homology; VH segments that show 80% or greater identity are considered to be in the same family whereas VH segments that have less than 70% identity to one another form different VH families (Kodaira et al., 1986; Lee et al., 1987; Shen et al., 1987; Berman et al., 1988). Such criteria have been supported by construction of the phylogenetic tree of 114 VH segments (Figure 1.3) (Matsuda et al., 1998). It clearly shows three VH subgroups, namely VHI, VHII, and VHIII, subdivided into the VH1/VH5, VH2/VH4/VH6, and VH3 families, respectively. It is interesting to note that the VH4 (Lee et al., 1987), VH5 (Shen et al.,
b Truncated VH4
VH4 VH6
Human/Mouse Segregation
V3-52P/V4-51.2P
V4-44.1P
V3-22.2P/V4-22.1P
VH2
V3-50P/V4-49.1P
73 VH1
V3-32P/V4-31.1P V3-29P/V4-28.1P
132
13
VH3 VH7
35
44
100
V3-63P/V4-62.1P V3-79P/V4-78.1P V3-54P/V4-53.1P
VH5 54
75(Myr)
V3-33.2P/V4-33.1P 39
V3-30.2P/V4-30.1P 10
Truncated VH3
FIGURE 1.3 (a) A phylogenetic tree of the human VH segments based on their nucleotide sequence alignment. Three distinct sets of the VH segments, which correspond to VHI, VHII, and VHIII subgroups, are separated with boxes and indicated by Roman numerals. (b) Estimation of divergence time between 10 homologous units containing a pair of the VH3 and VH4 segments. The divergence time is indicated in million years ago (Myr) and the human/mouse divergence is shown by a vertical line. Modified from Matsuda et al. (1998).
8
Matsuda
1987), and VH6 (Berman et al., 1988) families have been identified by the comparison of nucleotide sequences of VH segments. The VH4 family members are most strongly conserved, suggesting that VH4 may have evolved most recently (Lee et al., 1987; Haino et al., 1994). However, frequent recombination between VH segments makes it difficult to estimate the precise time of divergence among VH segments. The VH5 and VH6 families contain only two and one members, respectively. Subgroup I contains a unique set of VH segments that share about 80% overall homology with the VH1 family but much less similarity to VH1 at a clustered region between framework 2 (FR2) and FR3. This group was also identified from nucleotide sequence homology and has been proposed to be classified as VH7 family (Schroeder et al., 1990). According to the above definition of the VH family, VH7 should be a subfamily of VH1 or a family captured in transition from VH1 to independence (Kirkham and Schroeder, 1994). However, Southern blot and sequencing analysis revealed that the VH7 family is a small but discrete VH family consisting of five to eight members that are dispersed within the VH locus (van Dijk et al., 1993), indicating that the classification of VH7 is practically useful. Of interest, a set of 12 VH3 pseudogenes that have the 5¢truncation at the same position constitutes an independent cluster of the VH3 family in the phylogenetic tree (Figure 1.3). Another group of 13 VH4 pseudogenes, sharing the common 5¢-truncation in the second exon, again branched off from the common ancestor. Since they are scattered across the locus, the initial truncation probably took place in an ancestral VH segment, followed by interspersion of duplicated copies throughout the locus. The V4-44.1P segment, which shares <40.6% amino acid similarity to the other human VH segments, constitutes an independent branch in the tree (Figure 1.3). Interestingly, a similar level of amino acid similarity was obtained from those of a variety of vertebrates including mouse (38.8%), rat (30.0%), rabbit (38.6%), dog (34.4%), caiman (36.4%), Xenopus (33.7%), teleost fish (36.7%), and horned shark
(28.6%). This fact suggests that V4-44.1P might be a putative ancestral VH segment or a very old pseudogene having an accumulation of the point mutations. Several VH family-specific conserved regions occur in human germline VH segments (Kabat et al., 1991; Tomlinson et al., 1992; Matsuda et al., 1993; Haino et al., 1994). Family-specific sequences were found in the codons 9–30 in FR1 and the codons 60–85 of FR3. It is important to note that the codons 60–65 in the 3¢ portion of complementarity-determining region 2 (CDR2) were conserved in a family-specific way. Generally conserved universally were codons 1–8, FR2 (codons 38–47), and codons 86–92, in which the embedded heptamer recombination signal is located. A more extensive structural comparison of VH subregions is found elsewhere (Tomlinson et al., 1992; Kirkham and Schroeder, 1994).
5¢ Regulatory Regions The 5¢ flanking region of the VH segments contains two cis-acting elements, namely the octamer motif that regulates tissue-specific expression of IgH genes and the TATA box essential for the general transcription machinery. Extensive comparison of 500-bp of 5¢-flanking sequences of 79 VH segments without 5¢ truncation revealed striking familyspecific conservation (Haino et al., 1994; Matsuda et al., 1998) (Table 1.2). Locations of the octamer motif and TATA box are conserved within the same family but are different between different families. Forty out of 44 VH segments having a complete ORF contain an octamer sequence identical to the consensus (ATGCAAAT). The V3-20, V353, and V6-1 segments carry slightly less conserved octamer sequence but are known to be translated. The V3-38 segment in the ORF group has completely lost octamer (and TATA) due to 5¢-truncation. Of note, the octamer sequence is less conserved in pseudogenes and as many as 15 of 33 pseudogenes without 5¢-truncation have diverged octamer motif.
TABLE 1.2 Summary of the 5¢ regulatory sequences and RSS of the human VH segments 5¢ regulatory region
RSS
VH family
Heptamcer
(bp)*
Octamer
(bp)*
TATA
(bp)*
7mer
(bp)
9mer
VH1 VH2 VH3 VH4 VH5 VH6 VH7
CTCATGA — — — — — TTCATGA
2 — — — — — 2
ATGCAAAT ATGCAAAT ATGCAAAT ATGCAAAT ATGCAAAT AGGCAAAT ATGCAAAT
19 26 18 39 18 19 8
TAAATAT TT(G/C)AAAA ATGAAAA TTAAATT ACTTAAA TTTAAAT GGAATAT
81 41 101 59 79 78 79
CACAGTG CACAGAG CACAGTG CACAGTG CACAGTG CACAGTG CACAGTG
23 23 23 23 23 23 23
TCAGAAACC ACAA(A/G)AACC ACACAAACC ACA(C/A)AAACC CTAAAACCC ACACAAACC TCAGAAACC
* Most common distance between the motifs is shown.
1. Human Immunoglobulin Heavy Chain Locus
In contrast, the sequence of the TATA box is well conserved within the same VH family, but very different between different families (Table 1.2). In addition, like the octamer motif, pseudogenes have a less conserved TATA motif. A heptamer sequence (CTCATGA), which is reported to be essential for full VH promoter activity in mouse lymphoid cells (Ballard and Bothwell, 1986; Eaton and Calame, 1987; Siu et al., 1987), is found in the human VH1 and VH7 families and, as in mice, similarly located (2- to 22-bp upstream of octamer). However, no heptamer element is found or similarly placed in the other VH families. Hence, this provided no further supportive evidence for the hypothesis that the heptamer element is involved in the activation of the Hchain promoters by the oct protein before the activation of the L-chain promoters that do not contain the heptamer motif (Kemler et al., 1989). Another interesting finding is that the 5¢-flanking regions of three VH3 segments, namely V3-9, V3-20, and V3-43 have a common 65-bp deletion in the region at 251- to 315bp upstream of the initiation codon (Haino et al., 1994). Since all are found in the full-length VH mRNAs, the deletion would not drastically reduce the promoter activity. No other conserved nucleotide sequences or potential candidates for a novel cis-acting element of VH transcription regulation are identified, in spite of an extensive investigation of nucleotide sequence alignment. However, future studies to correlate VH promoter activity and nucleotide sequence variation in the 5¢ regulatory region may identify such elements.
Recombination Signal Sequence The RSS of VH segments, which is located immediately 3¢ to the coding region, is composed of highly conserved heptamer (CACAGTG) and nonamer (ACAAAAACC) sequences that are separated by a 23-bp spacer. In vitro analysis of the RSS clearly showed that the first three nucleotides (CAC) in the heptamer and the fifth and sixth nucleotides (AA) in the nonamer are critical for the efficient V(D)J recombination (Ramsden et al., 1996; Couno et al., 1996). All 40 VH segments classified as either “functional” or “transcribed” maintain the first four and the last nucleotides (CACANNG) in the heptamer, and 35 of them have intact heptamers (Matsuda et al., 1998). The other five VH segments have either AA, GA, or GC at their fifth and sixth positions in the heptamer sequence. Conservation of the nonamer sequence is weaker and relatively family specific (Table 1.2). Again, the critical two nucleotides are well conserved in the more than 40 VH segments, except V2-26 and V2-70, which carry GA instead of AA. The nonamer sequence having G nucleotide at its fifth position is shown to be still active in V-J recombination in the human Vl genes (Kawasaki et al., 1997). The VH1 and VH7 families have a
9
family-specific nonamer TCAGAAACC. In the ORF group, the heptamer signal of the V3-16 (TCCTGTG) and V3-38 (TACACAG) segments are highly diverged without conservation of the first three nucleotides, suggesting their incapability for functional VDJ rearrange-ment. Likewise, V3-35 and V7-81 contain diverged heptamers (CACTGAG and CACCATG, respectively) but with the first three nucleotides intact. However, the effect of these mutations on the efficiency of VDJ recombination is not yet known. Obviously, they all maintain 23-bp spacer length. In contrast, RSS of VH pseudogenes are much less diverged. The 64 pseudogenes that have not lost RSS by 3¢truncation contain 26 pseudogenes with at least one mutation in the five critical positions. In addition, one, two, six, and seven pseudogenes have an irregular spacer length of 17, 20, 22, or 24 bp, respectively, although V segments with 22- and 24-bp spacers are acceptable for the V(D)J rearrangement in the human Vl and TCRb loci (Kawasaki et al., 1997; Rowen et al., 1996). However, roughly 35% or 28 of 79 of the pseudogenes retain the flawless RSS with a 23-bp spacer. Assuming that the VDJ recombination possibility is the same for any of the 70 VH segments (39 functional, 1 transcribed, 2 ORF, and 28 pseudogenes), the probability of productive VH to DH-JH rearrangement per allele is 1/3 (frame) ¥ {40 (functional/transcribed VH segments)/70} = 0.19 or 19%.
Primary Repertoire of the Human VH Region The number of germline VH, DH, and JH segments is largely different between different species. This corresponds to the genetic mechanism through which VH region diversity is created. One extreme example is the chicken VH locus, in which a multiple number of VH pseudogenes function as a donor of somatic gene conversion to a single functional VH segment in VDJ rearrangement (Raynaud et al., 1987). In humans, VH diversity is obtained in a “classical” way; the number of germline VH, DH, and JH segments generates the genetic basis of the VH region repertoire. The immune response of Ig-transgenic mice (xenomice) gives important hints to the minimum number of functional VH segments necessary to provide a full antibody repertoire. The xenomouse II strains that carry 35 and 18 (respectively) functional human VH and Vk segments develop a human adultlike antibody repertoire with high-affinity human antibodies against diverse antigens produced by a sufficient number of mature B-lymphocytes (Mendez et al., 1997). In contrast, in the xenomouse I strains bearing five functional VH and three functional Vk segments, maturation of Blymphocytes is severely affected and only a modest immune response is observed. This strongly suggests the importance of the primary V-region repertoire in the highly diverse human antibody response. The combinatorial diversity of
10
Matsuda
the human VH genes can be estimated as 40 (functional/transcribed VH segments) ¥ 25 (functional DH segments) ¥ 6 (functional JH segments) = 6,000. This is certainly an approximate value since the number of functional VH segments varies between alleles due to deleterious polymorphisms.
VH Segment Usage and Repertoire Formation The biased usage of particular VH, DH, and JH segments during early phases of ontogeny was first reported in mouse (Yancopoulos et al., 1984; Reth et al., 1986). In humans, VH5 and VH6 families are selectively expressed at 7 weeks of gestation, when B-lineage development is initiated (Cuisinier et al., 1989). A rapid expansion of the VH repertoire, including biased usage of specific VH segments, takes place between 8 and 15 weeks of gestation. The utilization of VH segments may be influenced by two groups of factors: (a) those affecting the recombination frequency and (b) those affecting selection of B cells expressing particular VH segments. Group (a) includes the distance between DH (or JH) and VH segments, variation in the recombination signal sequence, and locations that favor the recombinase accessibility. Group (b) includes self-antigens and bacterial super-antigens. The initial observation of the preferential usage of JHproximal VH segments in mouse led to the hypothesis that the proximity of VH segments to JH favors biased expression of VH segments in early stages of ontogeny. According to the studies investigating the utilization of VH segments in early development, those VH segments often used preferentially in early stages of ontogeny were V6-1, V1-2, V2-5, V3-13, V3-15, V3-23, V3-30, V5-51, V3-53, V4-59, and V1-69 segments (Schroeder et al., 1987; Schroeder and Wang, 1990; Cuisinier et al., 1993). In particular, the V3-30 segment is utilized at the highest rate in these studies. The V1-69 segment is also used frequently in peripheral B cells (Schwartz and Stoller, 1994), B-cell leukemia, and autoantibodies (Zouali, 1992). On the physical map, however, they are scattered across the region between 70-kb (V6-1) and 900-kb (V1-69) upstream of the JH cluster. Furthermore, none of the JH-proximal functional VH segments (including V1-2, V1-3, V2-5, V3-11) were found repeatedly in either examination. The results indicate that VH segments preferentially used in early stages of ontogeny do not necessarily cluster in the JH-proximal region. Surprisingly, three overexpressed VH segments in early ontogeny (V3-23, V3-30, and V1-69) have polymorphic variations of germline copy number. Moreover, Sasso et al. (1996) reported that the expression of V1-69 is proportional to its germline copy number. It is interesting and feasible to test the correlation between polymorphisms of specific VH segments and their utilization.
EVOLUTION OF THE HUMAN VH LOCI Evolution of the VH Locus on Chromosome 14 The evolution of multigene families, including Ig, has been driven by dynamic reorganization of the gene locus including duplication, deletion, and translocation. Comparative analysis of VH loci among different vertebrates from shark to man revealed dramatic differences of H-chain gene organization among species, yet all of them carry multiple VH segments, thus indicating that duplication of VH segments must have started quite a long time ago. It is also important to realize that reorganization of the VH locus is still ongoing on a variety of scales, as evidenced by dramatic difference in the VH locus organization between mouse and man. Recent translocation of VH and DH segments to chromosome 15 and 16 is more evidence of the dynamic reshuffling of the VH locus. Homology matrix analysis of the 957-kb sequence against itself showed that 67% of the entire VH locus is occupied by the thirteen DNA sequences of variable length (4 kb to 24 kb) that appear at least twice across the locus (Figure 1.1). These homologous units contain the DNA fragments previously shown to cross-hybridize with fourteen nonrepetitive intergenic probes using Southern blot analysis (Matsumura et al., 1994). One of such sequence, which appears 11 times across the locus, contains a pair consisting of a VH3 segment and a VH4 segment having the 5¢ truncation. Molecular evolutionary analysis by comparison of the highly conserved spacer sequence between different VH3 and VH4 units showed that the reorganization took place at least eight times between 132 and 10 million years ago (Matsuda et al., 1998). Of these, six took place after the mammalian radiation at 75 million years ago (Figure 1.3b). A similar calculation between DNA regions containing the truncated VH segments, namely, V367.3P/V3-67.2P and V3-5.2P/V3-5.1P, deduced the divergence time to be 61 million years ago, again after the divergence between mouse and human (Matsuda et al., 1998). These results strongly suggest that recent DNA reorganizations play a key role in the generation of the germline VH-region repertoire in human.
Evolution of Orphon VH and DH Loci It is striking that orphon VH and DH loci contain a remarkably high percentage (about 40%) of apparently functional VH segments. Several possible explanations for such conservation might be: • recent translocation; • functional constraint through their usage by transchromosomal rearrangement, as shown in the
1. Human Immunoglobulin Heavy Chain Locus
human g and d T cell receptor loci (Tycko et al., 1991); or • correction mechanisms, such as gene conversion (discussed below). Putative origins for the orphon VH segments on chromosome 15 and 16 were found in the 0.43- to 0.25-Mb JH proximal VH region on chromosome 14 (Nagaoka et al., 1994) (Figure 1.2). The orphon VH locus on chromosome 16 (designated as VH-F) is structurally similar to the region between the V3-11 segment and the V2-26 segment on chromosome 14. In addition, nucleotide sequence homologies of seven corresponding VH segments between the two loci total more than 93% (Figure 1.2). Most remarkable homology was found between two truncated pseudogenes VF1-12P and V1-12P, in which the homology extends into the region 3¢ to the truncation site. The time of the translocation of the VH-F locus from chromosome 14 was estimated to be, at the earliest, 20 million years ago, using the synonymous site substitution in the coding region between corresponding VH segments. In contrast, no obvious counterparts of the D5 clusters could be identified on chromosome 14. However, the V54 segment in the D5-b locus showed a significant homology of 94.7% to the V1-18 segment, which is located in the putative origin of VH-F locus, although the other two segments are less homologous to the corresponding VH segments (Figure 1.2). The segregation time of V54 and V1-18 was estimated to be approximately 13 million years ago. These findings suggest that a DNA fragment of greater than 100 kb might have been translocated simultaneously to chromosome 15 and 16 approximately 20 million years ago.
Pseudogenes and Gene Conversion The completion of the nucleotide sequence of the entire human VH locus revealed that as much as 65% (79 out of 123) of the VH segments are pseudogenes. The presence of abundant VH pseudogenes and orphon VH segments raises the question of whether they have any functional significance. Since 40% of orphon VH segments are apparently functional, they can theoretically recombine with DH-JH rearrangements on chromosome 14 through interchromosomal recombination. Recombination between a VH–DH rearrangement on chromosome 15 and a JH segment on chromosome 14 is also possible since a VH–DH fusion product was isolated from a human B-cell line (Shin et al., 1993b) and from B lymphocytes of transgenic mice carrying an IgH mini locus (Tuaillon et al., 1994). In addition, germline transcripts of orphon VH have been identified in human fetal liver (Cuisinier et al., 1993), suggesting that orphon VH loci might be targets of recombinase. Unfortunately, however, no direct evidence for the expression or recombination of translocated VH segments has been reported to date. More-
11
over, an extensive analysis of the human D segment usage ruled out the possible involvement of D5 segments on chromosome 15 in functional VDJ rearrangement (Corbett et al., 1997). Conserved pseudogenes have been already shown to serve as sequence donors for gene conversion in other species. Somatic gene conversion (or double unequal crossing-over) has been shown to take place to amplify the V region repertoire in chicken (Raynoud et al., 1987; Tompson and Neiman, 1987) and rabbit (Becker and Knight, 1990) but not in mouse and man. One attempt to identify evolutionary gene conversion is based on the theory of molecular evolution (Haino et al., 1994). In the course of evolution, the introns and the synonymous positions of the coding region evolve at high and remarkably similar rates in different genes, and base substitutions are accumulated at approximately constant rates with respect to geological time (Miyata et al., 1980; Hayashida and Miyata, 1983). Hence, the recipients of gene conversion would be found by comparing the substitution rates in the intron and the synonymous position of pairs of VH segments; clear differences in substitution rates of the two portions in a given pair of VH segments suggests some recombination events. One of the recently duplicated VH segment pairs, V3-62P and V3-60P, which show 94% nucleotide homology and share the same mutation in RSS (Kodaira et al., 1986), w ere chosen for the analysis. These two VH segments displayed a significant difference in the substitution rates in the intron (Kci = 0.1637 ± 0.0461) and the synonymous position of the coding region (Kcs = 0.0821 ± 0.0339), thus indicating possible segmental change in the intron in either V3-60P or V3-62P (Figure 1.4). Sequences of these two VH segments were compared with those of the other VH3 members, to look for the putative donor of the segmental transfer. As a result, V3-43 and V3-62P were found to have significantly smaller Kci (0.0820 ± 0.0280) than Kcs (0.4165 ± 0.0761). Such unusual homology of the 5¢ half region between V3-43 and V3-62P is most likely explained by the unidirectional segmental transfer of the V3-43 sequence to V3-62P. This example supports the hypothesis that gene conversion contributes to the maintenance of the pseudogene structure. The same method was applied to VH segments belonging to the VH4 family, which is most conserved (>90%) and richest in functional VH segments among the seven human VH families (Lee et al., 1987). Rather frequent unidirectional correction was observed between VH4 segments, thus demonstrating that the VH4 family members evolved by recent duplication, followed by gene conversion (Figure 1.4). It is to be noted that V4-55P served as a donor of two functional VH segments, V4-4b and V4-28. This might indicate that the high percentage of pseudogenes should also contribute to the generation of the germline VH repertoire.
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FIGURE 1.4 Schematic demonstration of gene conversion. The values of Kci and Kcs between VH segments are calculated for introns and synonymous positions of codons -4/92, respectively. Vertical arrows indicate direction of sequence transfer. Modified from Haino et al. (1994).
HUMAN CH LOCUS The human CH gene family is mapped to the q32 band of chromosome 14 (Kirsch et al., 1982) and consists of nine functional genes and two pseudogenes. Between mouse and human, the characteristic difference in the organization of the CH gene cluster is the presence of the duplication of 70kb DNA, consisting of two Cg genes and one each Ce and Ca genes in human (Figure 1.1) (Flanagan and Rabbitts, 1982). In addition, a pseudo Cg gene has been genetically mapped between the duplication unit (Bech-Hansen et al., 1983). The 5¢ Ce or Ce2 gene is a pseudogene with a 5¢-
truncation, resulting in the absence CH1 and CH2 exons. The other pseudogene Ce3 is processed and translocated to chromosome 9 (Battey et al., 1982). The complete nucleotide sequence of the entire CH locus permitted the precise organization of the human CH locus as follows: 5¢-JH-(8 kb)-Cm(5 kb)-Cd-(65 kb)-Cg3-(26 kb)-Cg1-(19 kb)-Ce2-(13 kb)-Ca1-(34 kb)-yCg-(20 kb)-Cg2-(18 kb)-Cg4-(23 kb)Ce1-(10 kb)-Ca2–3¢ (Ravetch et al., 1981; Flanagan and Rabbitts, 1982; Word et al., 1989; Heilig et al., 2003; Nicodeme et al., submitted), which is in agreement to that of previous studies based on hybridization (Bottaro et al., 1989b; Hofker et al., 1989).
1. Human Immunoglobulin Heavy Chain Locus
Three heavy chain transcription enhancers are known in the human CH locus, all of which are located at a place similar to those of mouse (Figure 1.1). The 5¢-lost enhancer, or Em enhancer, is located in the intron between JH and Cm (Rabbitts et al., 1983). Two nearly identical copies of enhancer arrays homologous to the mouse Ca 3¢-enhancer (3¢aE) were identified at the 3¢ flanking region of each of the two human Ca genes, namely at the 3¢ end of the CgCg-Ce-Ca duplication (Mills et al., 1997). A novel regulatory motif cluster with a potential B lymphocyte-specific enhancer function (Ed-g3) was recently identified (Mundt et al., 2001) in the 55-kb DNA between the Cd and Cg3 genes, where the existence of a strong candidate region for matrix attachment was predicted by computer programs (Nicodeme et al., submitted). This region has an exceptionally low G + C nucleotide content (average 42%) in the G + C predominant human CH locus (average 58%). Three non-immunoglobulin DNA sequences were identified in the CH locus (Figure 1.1). The 1.7-kb mRNA sequence of AK056731 showed 99.7% homology to a DNA sequence between yCg and Cg2 genes. This single exon
13
gene, having a 545-bp ORF, is expressed in placenta, but the function of the protein is unknown. A processed pseudogene of ELK2 is located upstream of the yCg gene (Harindranath et al., 1997). Another processed pseudogene of ATP6V1G1, a vacuolar ATPase, was identified in the region between the Cd and Cg3 genes.
Structure of CH Genes All the human CH genes have been isolated and sequenced completely. References for complete CH gene sequences with detailed information are available from Ig databases (for example, IGMT database; http:// imgt.cines.fr). The human CH genes for secretory forms are composed of three (d, g, and a) or four (m and e) exons, each encoding a functional and structural unit of the H chain, namely a domain (Edelman et al., 1969) (Figure 1.5). Cd has an additional exon that encodes a C-terminal tail for the secretory-form IgD 2-kb downstream of its CH3 exon. Exons corresponding to hinge regions are located between the CH1 and CH2 exons in the Cd and Cg genes, and their number
FIGURE 1.5 Exon/intron structure of human CH genes and pseudogenes. Coding exons, sterile (I) exons, hinge exons, and membrane exons are shown by open box, hatched box, vertical line, and striped box, respectively. Switch regions are indicated with vertical stripes. The exon of the Cd gene for soluble form (CH-S) is indicated. Note that I exons of Cg2, Cg4, and yCe2 and membrane exons of Cg4 and yCe2 are predicted by homology search.
14
Matsuda
and length vary between different CH genes and subclasses. The hinge region of Ca genes is exceptionally encoded by the CH2 exon, and there is no obvious hinge region in the Cm and Ce genes. In addition, one (a) or two (others) separate exons encode the hydrophobic transmembrane and short intracytoplasmic segments that are used for a membrane-form Ig. The size of each CH exon is similar to that of the CL exon, suggesting that the CH gene evolved through the duplication of a primordial single exon gene, like the CL gene. Such exon–intron organization of the CH gene is consistent with the domain hypothesis that states that the Hchain protein consists of a tandem array of three or four functional units (Edelman et al., 1969). The total length of each CH gene is therefore variable, ranging from 4 to 9 kb (sterile exons are not taken account). All functional CH genes except Cd have the switch (S) region at the 5¢ flanking region of the CH1 exon. S regions consist of tandem repeats of pentameric nucleotides and are responsible for class switch recombination. The presence of germline transcripts prior to class switching arising from untranslated exons (I exons) was reported previously for most of the CH subclasses (Sideras et al., 1989; Nilsson et al., 1991; Kuzin et al., 2000; Mage et al., 1989; Bachl et al., 1996). Expression of such transcripts is driven by a promoter located upstream of the S region. Missing information for the I exons of the Cg2, Cg4, and yCe genes was recently obtained by nucleotide sequence alignment of the 5¢ flanking region between different Cg and Ce genes (Nicodeme et al., submitted). Most of the CH genes carry a single I exon, except two Cg genes; Cg1 has three I exons as does its duplicated copy, Cg4. No putative I exons were found in the Cd gene, which is transcribed together with Cm as a single transcript. The absence of the pseudo Cg gene product, despite a complete set of coding exons without defects, is explained by the deletion of S region and I exons.
Expression of the membrane exons is controlled by differential splicing. Transcripts of the membrane exons are spliced to the 3¢-most domain exons by removing the last few residues of the secreted Ig tail. Membrane segments, except those of the Ca genes, are encoded by two exons. The hydrophobic transmembrane segment of 26 residues is relatively conserved among all the H chains, suggesting the possibility that membrane-form Ig is anchored by a common membrane protein (Yamawaki-Kataoka et al., 1982). Since the intracytoplasmic segments of the membrane-form Ig are too short (27 residues for Cg and Ce chains, 13 residues for Ca, and 2 residues for Cm and Cd) to catalyze any enzymatic activity such as phosphorylation, transduction of the triggering signal of the antigen–antibody interactions may require involvement of at least one other protein. This hypothesis has been verified by subsequent identification of Iga and b proteins (see Chapter 11).
Polymorphisms of the Human CH Locus The human CH locus is highly polymorphic, with different alleles carrying deletion and duplication of CH genes. Eleven types of deletions and eight duplications involving one or more CH genes have been identified to date (summarized in Table 1.3). One of the most common polymorphism is the Cg4 gene duplication, which is present in 44% of Caucasian chromosomes (Brusco et al., 1995a). Large differences in the frequency of the CH haplotype were observed between different ethnic groups in an inter-racial genetic study (Rabbani et al., 1996). Of interest, seven of these polymorphisms appear as both deletion and duplication, suggesting that unequal crossing-over between highly homologous regions played a major role for the CH locus polymorphisms. Looping-out excision is also conceivable as another genetic mechanism (Bottaro et al., 1989a). It is
TABLE 1.3 Summary of the human CH locus polymorphisms Reference Polymorphism Cg1 Cg1-Ca1 Cg1-Cg2 Cg1-Cg4 yCe2-yCg yCe2-Ce1 Ca1-Ce1 yCg yCg-Ca2 Cg2 Cg2-Cg4 Cg4
Approximate size (in
Deletion
Duplication
— 50 110 130 70 120 120 — 110 — 35 —
Smith et al. (1989) Rabbani et al. (1995) Smith et al. (1989) Lefranc et al. (1982) Lefranc et al. (1983) Migone et al. (1984) Migone et al. (1984) N.I. Bottaro et al. (1989a); Hendriks et al. (1989) Bottaro et al. (1989a); Hendriks et al. (1989) Olsson et al. (1991) Bottaro et al (1990)
N.I. N.I. N.I. Rabbani et al. (1996) N.I. Bottaro et al. (1991) Bottaro et al. (1991) Rabbani et al. (1996) Brusco et al. (1995a) Bech-Hansen and Cox (1986) Brusco et al. (1995a) Brusco et al. (1995a)
* The size of deletion/duplication for those comprising multiple CH genes was estimated from the physical map. N.I.; not identified to date.
1. Human Immunoglobulin Heavy Chain Locus
rather surprising that individuals with deletions of multiple CH genes have not shown any severe clinical symptoms, thus suggesting that Cg and Ca subclass genes are capable of substituting each other and that the Ce genes might not be obligatory but might facilitate efficient protection from parasite infection. Ig allotype typing is usually performed with serological methods based on hemagglutination inhibition. Allotypes of Ig are mostly explained by specific amino acid substitutions in CH regions. In humans, Ig allotypes have been identified for five human CH genes, the Cg1, Cg2, Cg3, Ca2, and Ce1 genes, and are designated as G1m, G2m, G3m, A2m, and Em, respectively. Molecular typing of Ig allotypes has been done for different CH genes. SNPs specific to G2m and G3m allotypes (Brusco et al., 1995b; Dard et al., 2001) were identified by nucleotide sequencing and were confirmed by population-based tests.
Acknowledgments We thank all our colleagues for their contribution to accomplish this study, and Dr. David H. Gelfand (Roch Molecular Systems, Inc.) for the critical reading of the manuscript. Most of the work was done in the Center for Molecular Biology and Genetics and the Department of Medical Chemistry, Kyoto University Graduate school of Medicine (on the VH locus) and Centre National de Genotypage (on the CH locus). The work was supported in part by grants from the Ministry of Education, Science, Sports, and Culture in Japan and from the Science and Technology Agency of Japan. The CNG is supported by the Ministere de la Recherche et des Nouvelles Technologies.
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2 Immunoglobulin Heavy Chain Genes of Mouse ROY RIBLET Torrey Pines Institute for Molecular Studies San Diego, California, USA
using a large number of landmarks derived from prior assembly of a C57BL yeast artificial chromosome (YAC) contig (Chevillard et al., 2002). The sequence-ready tiling path was selected by a team at Washington University, led by John MacPherson. BAC sequencing was performed by the Genome Therapeutics Corporation sequencing group directed by Douglas Smith. This summary of our findings was written in advance of primary publication.
The immunoglobulin heavy chain locus, Igh in mouse, is an unusual genetic locus that must undergo molecular recombination to yield an active expressible gene for its antibody heavy chain product. Before this genetic rearrangement occurs, the locus is comprised of an array of clusters of gene segments of four types, Variable (Vh), Diversity (Dh), Joining (Jh), and Constant (Ch) gene segments. A rearranged active heavy chain V gene is constructed by fusing together a V, D, and J segment (Sakano et al., 1980). In mouse, this array of gene segments is near the telomere of chromosome 12 and comprises about 3 Mb (million basepairs) of DNA. DNA sequence variation occurs between mouse strains across the entire locus, so that it is helpful to analyze a single allelic state of the array, such as is found in an inbred mouse strain. This allelic form of the entire length is termed a haplotype. Prior to the initiation of the mouse genome project, many years of characterization of the mouse Igh locus focused primarily on the Igha haplotype of BALB/c due to the extensive collection of mineral oil–induced plasmacytomas and their monoclonal antibody products that were available in this strain (Potter, 1977). This work was well reviewed in the last edition of this book (Honjo and Matsuda, 1995) and is covered in a comparative manner here. The DNA sequence of the C57BL/6 mouse genome is nearly completed, and this includes the Igh locus. Mouse Igh was expected to be difficult to analyze since it is two to three times longer than human IGH (Chevillard et al., 2002) and is known to contain an unusually high level of Line1 repetitive elements (Herring et al., 1998). Because Igh was accepted as a locus of high biological interest, it was sequenced from a bacterial artificial chromosome (BAC) contig rather than assembled from shotgun reads. A deeply redundant BAC contig was assembled in my laboratory
Molecular Biology of B Cells
Igh-V OR VH GENES OF THE Ighb HAPLOTYPE A search for Vh gene segments in the current assembly of the Ighb sequence identified 170 full length coding sequences (plus additional truncated gene fragments). An additional 20 to 30 sequences are expected in the unfinished 5¢ end of the locus. Of the 170 sequences, 69 have obvious defects in the coding sequence that preclude their expression as functional Vh genes. A search of GenBank, including the EST database, indicates that many of the 101 apparently functional sequences are present, at least as recovered mRNAs. Many others have not been observed and may have defects in their promoter or RSS sequences that render them unable to be expressed. The 101 potentially functional Vh coding sequences were aligned and are displayed as a neighbor joining tree in Figure 2.1. The gene relationships radiate from a central trifurcation that reflects the three Vh gene and protein subgroups originally noted by Kabat (1991). The subgroups further divide into the 15 Vh gene families described by Brodeur and other groups (summarized in Mainville et al., 1996) for the Vh sequences of BALB/c and the Igha haplotype. Strain surveys by Southern blot hybridization indicated
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Copyright 2004, Elsevier Science (USA). All rights reserved.
20
Riblet
FIGURE 2.1 Neighbor-joining tree of the C57BL/6 Vh gene segments. The 101 candidate functional Vh genes group into three major clades (the subgroups of Kabat) and 15 Vh gene families. The Vh gene coding sequences corresponding to the mature heavy chain peptide were extracted from the mouse genome assembly. Apparent pseudogenes containing termination codons were omitted. Vh sequences were aligned and a neighbor-joining tree calculated using ClustalX (Thompson et al., 1997); the tree was plotted with DrawGram in the PHYLIP package (Felsenstein, 1993).
similar gene family organization and content in C57BL/6 and many different Igh haplotypes in lab strains and wild mice (Tutter and Riblet, 1988; Tutter and Riblet, 1989b). The complexity or content of the Vh gene families in the Ighb haplotype of C57BL/6 is listed in Table 2.1. Evident pseudogenes are tabulated separately from sequences that appear functional. The numbers of genes in each Vh gene
family in the assembled C57BL/6 sequence are in general agreement with, but tend higher than, the restriction fragment counts from a C57BL YAC contig of Ighb (Chevillard et al., 2002), and the estimates from BALB/c made by cloning individual genes and counting bands on blots (Brodeur and Riblet, 1984). Most families are small, with one to six members; three of the four Group I families are
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2. Immunoglobulin Heavy Chain Genes of Mouse
TABLE 2.1 Mouse Vh repertoire in the Ighb haplotype Mouse Vh family VhQ52 Vh36–60 Vh3609P Vh12 VhJ558 VhGam3–8 VhSm7 VhX24 Vh7183 VhJ606 VhS107 Vh10 Vh11 Vh15 Vh3609N Totals
Intact genes
Pseudogenes
Total
8 6 8 1 43 4 4 1 10 5 3 2 2 1 3
3 2 4 2 31 0 0 1 16 1 2 4 1 1 1
11 8 12* 3 74* 4 4 2 26 6 5 6 3 2 4
101
69
170
* Gene numbers in these two families represent sequencing from 1.2 Mb. An additional 0.2 to 0.3 Mb remains to be sequenced.
somewhat larger with 8 to more than 12 members. The 7183 family contains 26 sequences, and the J558 family is largest, with 74 sequences currently identified. When finished, the 5¢ portion of the locus should contain an additional 10 to 20 J558 and 3609P sequences. Overall, 40% of the Vh sequences have obvious defects in the coding sequence that preclude their expression, and most families contain such pseudogenes. A majority of sequences in the Vh7183, Vh10, and Vh12 families is defective. A physical map of the Vh gene array is shown in Figure 2.2. It begins at D12Mit263, arbitrarily taken as a boundary between the Dh and Vh gene segment regions, and ends more than two million base pairs later at the 5¢ end of the current assembly. Comparison to the YAC contig and other data indicates that 200 to 300 kb remain to be sequenced. The placement of Vh families in the locus is in agreement with the deletion map of Brodeur (Mainville et al., 1996). The first megabase of the Vh array contains all of 13 of the families. Each family is localized in a subregion, interspersed with several other families. The 5¢ 1100 kb (plus 200 to 300 kb) of the locus contains exclusively VhJ558 and Vh3609P genes. Most of the Vh region is densely occupied by Vh genes, with roughly 10 kb spacing, but genes are more sparse in the distal, 5¢ 600 kb, where spacing averages 20 kb.
POLYMORPHISM IN VH GENES The array of Vh, Dh, and Jh gene segments in Igh defines the germ line, or inherited, antibody heavy chain repertoire, the spectrum of antibody structures that the B cell popula-
tion will make initially and throughout life, although its diversity will be much enhanced by somatic mutation, N region addition, and other junctional mechanisms. This is a basic measure of the universe of bindable antigens. The inherited Vh gene array, the starting library of antibody specificities, can vary between inbred mouse strains, and can affect specific antibody responses. Understanding the extent of the variation between mouse strains, and then between species, will teach us about the acceptable limits for the inherited repertoire and what antibody diversity an animal needs to start with in order to build a successful humoral immune system. We can begin to compare the repertoires of two mouse strains, C57BL/6 and BALB/c. Extensive random cDNA and focused genomic cloning and sequencing efforts have characterized all members of several Vh gene families in BALB/c, and we can compare these to the genomic C57BL/6 repertoire. Figure 2.3 shows a tree of BALB and C57BL/6 Vh genes of the Vh10, VhS107, and Vh7183 families. Both strain sequences were previously known for Vh10 (Whitcomb et al., 1999) and VhS107 (Perlmutter et al., 1985). The Igha haplotype sequences (from strain 129) for Vh7183 were recently completed (Williams et al., 2001). Figure 2.3 shows only the subset of Vh genes in each strain that are clear alleles, and it tabulates the nucleotide divergence between alleles. These differences range from zero (identity throughout the mature protein coding sequence) to 6%. Perhaps more significantly, not shown in this figure are those members of each gene family that do not have alleles in both strains. These have resulted from gene duplications and deletions that occurred independently in the history of the two haplotypes. In the Vh10 family, one additional functional member in Igha was previously known (Whitcomb et al., 1999), and we see a total of five members in C57BL/6, although the three not shown are all pseudogenes. In VhS107, an additional pseudogene occurs in C57BL/6. In Vh7183, 9 allelic pairs are shown in Figure 2.3, but there are also 11 BALB/c and 17 C57BL/6 Vh7183 genes that clearly have no allelic relationship. With the development of such extensive discordance (not sequence divergence as such) over a relatively short evolutionary span (1 to 3 million years), it is apparent that, at least in mice, the inherited library can vary quite extensively with respect to sequence. Whether this is reflected in comparable variation in functional binding specificities cannot yet be addressed.
Dh Figure 2.4 displays the physical map of the D-J-C region of Ighb. Dh sequences in BALB/c have been extensively analyzed (Kurosawa and Tonegawa, 1982; Wood and Tonegawa, 1983; Feeney and Riblet, 1993). One Dh segment, DhQ52, is only 700 bp from the first Jh segment. Additional Dh segments are scattered in a region of at least
si
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FIGURE 2.2 Physical map of the C57BL/6 Vh gene cluster. The positions of 170 full-length Vh gene segments of the Ighb haplotype are shown to scale. The genes are named according to their family, “b” haplotype, and position in the array starting at the 3¢ end. For example, Vh7183.b1Psi and Vh7183.b2 are the b alleles of E4Psi and Vh81X, respectively. Apparent pseudogenes are shown in gray. The scale is in kb, and 2.1 Mb of the Vh cluster is shown. An estimated 200 to 300 kb at the 5¢ end of the Vh array remain to be sequenced. 0p si
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2. Immunoglobulin Heavy Chain Genes of Mouse
apart, but DhFL16b and Vh7183.b1Psi (the C57BL/6 allele of E4Psi) are separated by 90 kb.
Jh The four Jh gene segments were isolated as 1,340 bp Jh locus PCR products from mouse strains of ten Igh haplotypes and sequenced by Solin and Kaartinen (1992). The J locus from C57BL/10 reported by Solin is identical to the genomic sequence of C57BL/6 and differs from BALB/c at eight nucleotides. The Jh coding segments are identical, except for one nucleotide difference resulting in an amino acid replacement in Jh1 between BALB and the C57BL strains.
Ch
FIGURE 2.3 Tree of alleles of 3 Vh families. For the Vh10, VhS107, and Vh7183 gene families the alleles from the Igha and Ighb haplotypes were aligned and a neighbor-joining tree calculated. For each allele pair, the percentage nonidentity was calculated. Vh gene segments from the Ighb haplotype are shown in black, the Igha haplotype in gray.
80 kb between DhQ52 and the first Vh gene, E4Psi. These include two DFL16 segments, nine DSP2 segments, a DST4 segment, and an undetermined number of defective D-like pseudogenes. These 13 listed Dh segments of the Igha haplotype are found in productive VDJ rearrangements (Feeney and Riblet, 1993). Comparable detailed analysis of Dh usage in the Ighb haplotype is lacking. On the basis of genomic sequence, nine Dh segments have an intact RSS on each side. These include evident homologs for 3¢ DhQ52 and DhST4 segments and the 5¢ DhFL16.1; these enclose a series of six DhSP2 segments. Additionally, eight homologs of the BALB/c D1Psi pseudogene alternate with the DhSP2 and DhFL16 segments. These apparent pseudo-Dh sequences lack one or both consensus RSS segments. A dotplot of this region reveals a pattern of 5 kb duplications yielding the alternating Dh–DhPsi pattern. The spacing between the separate clusters of Jh and Dh, and Dh and Vh gene segments is interestingly different. Jh1 and DhQ52 are only 700 bp
The heavy chain constant region gene segments (Ch) encode the C-terminal major portion of the heavy chain protein. The eight classes or isotypes of constant regions and the respective isotypes of serum immunoglobulin in mouse are m and IgM, d and IgD, g1 and IgG1, g2a and IgG2a, g2b and IgG2b, g3 and IgG3, e and IgE, and a and IgA. The physical map of the 200 kb long Ch gene region in BALB/c was determined by Shimizu et al. (1982). The sequencebased map of Ighb shown here is in agreement with BALB/c, with minor variations in intergenic spacing. The significant difference in the Ch genes of the two haplotypes is the replacement of IgG2a in BALB/c by IgG2c in C57BL/6, as previously described (Fukui et al., 1984; Morgado et al., 1989). This is apparently the result of the duplication of an ancestral g2 gene to create g2a and g2c isotypes, subsequent sequence divergence, and finally the loss of alternate genes in the two haplotypes. This explanation is confirmed by the presence of both isotypes in wild Asian and European mouse haplotypes (Fukui et al., 1984; JouvinMarche et al., 1989).
3¢ Regulatory Region (Enhancer) Downstream (3¢) of the Ca gene segment is the 3¢ regulatory region (3¢RR), which has enhancer activity for the transcription of Igh. This region is important in the regulation of isotype switching as well (Manis et al., 2003; Pinaud et al., 2001). It is characterized by a complex series of direct and inverted repeats that produce nested palindromes (Chauveau and Cogne, 1996). Recently, the complete sequence of this region was obtained from a 125 kb BAC from 129, an Igha haplotype mouse strain (Zhou et al., 2002a). This work also defined the positions of the nearest genes flanking Igh on the centromeric side: Crip and Mta1. The genomic sequence of C57BL/6 matches the 129 sequence with small variations in nucleotide sequence and intergenic distances.
24
Ce
t41 D1 2M i
Ca
D1 2M i
D1 2M i
t18
t19
Riblet
Cg2a
Cg2b
Cg1
Cm
200 Dh Dh02b Dh03b 04 b Dh 05 Dh b 06 b Dh Dh07b 08 b Dh Dh09b 10 b Dh 11 Dh b 12 b Dh Dh13b 14 b Dh 1 Dh 5b 16 b
Dh 1 D1 7b 2M it2 63
100
299
200 0
JH
Dh 01 b
Cd
Cm D1 mem 2M it8
Cg3
D1 2M i
t20
0
20
40
60
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100
FIGURE 2.4 Physical map of the Ch, Jh, and Dh clusters. 300 kb of the Ighb locus is diagrammed, starting at the 3¢ end of Igh, which is centromeric on chromosome 12. The eight Ch gene segments and the small cluster of four Jh segments occupy 200 kb. The Dh segments occupy the next 100 kb, ending at the simple sequence marker D12Mit263. The Dh segments are identified as Dh01b–Dh17b. Dh01b is the Ighb allele of the BALB DhQ52, Dh02b the allele of DhST4, and Dh16b the allele of DhFL16.1. Dh04b, Dh06b, Dh08b, Dh10b, Dh12b, and Dh14b are DhSP2 segments, and the alternating Dh03b, Dh05b, Dh07b, Dh09b, Dh11b, Dh13b, Dh15b, and Dh17b are Dh1Psi pseudogenes.
EVOLUTION Comparison of the human IGH and mouse Igh loci reveals some basic similarities: The constant region classes or isotypes, IgM, IgD, IgG, IgE, and IgA, are present in both and probably in all mammals. Each species may have different numbers of duplicated members of some isotypes, particularly IgG and IgA genes. Similarly, the Jh and Dh gene segments vary in number; the human content is higher than the mouse in both cases. In contrast, the mouse Vh gene array is larger than the human; the mouse has about 200 Vh gene segments spaced over more than 2 Mb, compared with about 100 Vh genes in 1 Mb in man. Analysis of the V sequences in both species reveals the same trifurcation into three subgroups, noted by Kabat (1991) as shown for mouse in Figure 2.1. This evolutionary trifurcation is ancient, existing in most mammals (Tutter and Riblet, 1989a). In each species, there is division of subgroups into gene families, seven in man and fifteen in mouse as shown. Similarities of certain families exist across species (Tutter and Riblet 1989a), but with no evident correspondence of individual Vh genes between mouse and man. Evolutionary analysis of individual Vh gene sequence and specificity will require comparison of different mouse haplotypes and comparisons of mouse and rat. Such studies are in progress.
GENOMIC CONSIDERATIONS The genomic sequence facilitates consideration of the Igh locus as a whole, as a functional entity, and enables us to ask what distinguishes it from surrounding genes, what
boundaries mark the edges of the locus-specific regulation of Igh? What characteristics of the locus relate to the mechanisms that activate and regulate the intricate series of genomic alterations involved in VDJ recombination, class switch recombination, somatic mutation, and allelic exclusion during the development of a B cell?
3¢ Border The genes that flank Igh on its 3¢ side, Crip and Mta1, are not B cell–specific; rather, they are expressed in many cell types and are regulated independently of Igh. The boundary of transcriptional regulation between the Igh region (the Igh structural genes and 3¢RR) and these flanking loci coincides with a remarkable developmentally regulated origin of DNA replication (Zhou et al., 2002a). In non-B cells and in mature B cells, a replication fork initiates at this origin at the beginning of S phase in the cell cycle and travels 5¢ though the 3¢ RR, Ch, Jh, and Dh regions and into the Vh genes. This progression occurs over nearly the entire span of the S phase and covers a distance of over 400 kb. Late in S phase, the remaining portion of Igh, more than 2 Mb containing most Vh gene segments, is replicated by forks moving in both directions. In a contrasting pattern in pro- and pre-B cells, where Igh becomes activated to undergo VDJ recombination, the entire locus is replicated early in S by forks moving in both directions. The significance of this unusual replication pattern is not clear, but its two transitions, first as the locus is activated for rearrangement, and second as the locus completes rearrangement and is ready for stable high-efficiency transcription, are correlated with other changes in chromatin structure and accessibility.
25
2. Immunoglobulin Heavy Chain Genes of Mouse
5¢ Border The 5¢ end of Igh is distal (telomeric) in both mouse and man. In man, IGH is immediately adjacent to the chromosome 14 telomere (Cook et al., 1994). In mouse, this is not the case; although Igh is far distal on chromosome 12 there are several million base pairs of genome before the telomere. The genes flanking the 5¢ end of Igh are Zfp386, a Kruppel-like zinc finger protein identified in EST sequencing, and Vipr2. In man, homologous sequences are located on 7q36. 1 to 2 Mb farther distal in mouse are the genes Sp4 and Dnahc11 (iv, situs inversus). In man, these are on 7p15–21. This mouse genomic information is based partially on genomic sequencing and clone assembly, and it extends previous published and unpublished genetic mapping studies (Brueckner et al., 1989; de Meeus et al., 1992).
Nuclear Location and Chromatin Structure The C57BL/6 and 129/Sv BAC contigs that were assembled for the genomic sequencing of Igh have provided the reagents for other studies of the locus as a whole. Fluorescent in situ hybridization (FISH) studies using BAC probes showed that the Igh locus undergoes a cyclical change in location in the nucleus in differentiating B cells (Kosak et al., 2002; Zhou et al., 2002b). In non-B cells and hematopoietic progenitors, the Igh locus is positioned at the nuclear periphery, associated with the nuclear lamina. In pro- and pre-B cells, Igh repositions towards the nuclear center, and in B- and plasma cells it moves back to the edge. In addition, the locus undergoes a compaction when it leaves the periphery. The ends of this 3 Mb locus are brought closer together, presumably to facilitate VDJ recombination (Kosak et al., 2002). Increased knowledge of the sequence and structure of Igh has also facilitated detailed studies of transcriptional regulation and chromatin changes across the locus during B cell development (Chowdhury and Sen, 2001; Johnson et al., 2003). These have shown a strong correlation of histone acetylation with accessibility and activation of the locus for rearrangement. It is straightforward to hypothesize that these alterations in chromatin structure and locus accessibility are mechanistically correlated with developmental alterations in replication patterns, nuclear location, and compactness of Igh. However, which, if any, of these parameters is the initiator or first link in the intricate chain of developmental events, and how these different steps are linked together, are important questions yet to be addressed.
CONCLUSION Several decades of structural studies of Igh have culminated in the nearly complete DNA sequence of this 3 Mb
locus. This has yielded a complete definition of all the gene segments in the mouse locus that can now be manipulated to answer questions about the immune repertoire and can be compared to human and other species. It has also led to novel findings of global changes in replication patterns, nuclear location, and chromatin structure that offer new avenues to study antibody gene actions and B cell development.
References Brodeur, P. H., and Riblet, R. (1984). The immunoglobulin heavy chain variable region (Igh-V) locus in the mouse. I. One hundred Igh-V genes comprise seven families of homologous genes. Eur J Immunol 14, 922–930. Brueckner, M., D’Eustachio, P., and Horwich, A. L. (1989). Linkage mapping of a mouse gene, iv, that controls left-right asymmetry of the heart and viscera. Proc Natl Acad Sci U S A 86, 5035–5038. Chauveau, C., and Cogne, M. (1996). Palindromic structure of the IgH 3¢locus control region. Nat Genet 14, 15–16. Chevillard, C., Ozaki, J., Herring, C. D., and Riblet, R. (2002). A threemegabase yeast artificial chromosome contig spanning the C57BL mouse Igh locus. J Immunol 168, 5659–5666. Chowdhury, D., and Sen, R. (2001). Stepwise activation of the immunoglobulin mu heavy chain gene locus. EMBO J 20, 6394–6403. Cook, G. P., Tomlinson, I. M., Walter, G., Riethman, H., Carter, N. P., Buluwela, L., Winter, G., and Rabbitts, T. H. (1994). A map of the human immunoglobulin VH locus completed by analysis of the telomeric region of chromosome 14q. Nat Genet 7, 162–168. de Meeus, A., Alonso, S., Demaille, J., and Bouvagnet, P. (1992). A detailed linkage map of subtelomeric murine chromosome 12 region including the situs inversus mutation locus IV. Mamm Genome 3, 637–643. Feeney, A. J., and Riblet, R. (1993). Dst4: A new, and probably the last, functional Dh gene in the BALB/c mouse. Immunogenetics 37, 217–221. Felsenstein, J. (1993). PHYLIP (Phylogeny Inference Package). Fukui, K., Hamaguchi, Y., Shimizu, A., Nakai, S., Moriwaki, K., Wang, C. H., and Honjo, T. (1984). Duplicated immunoglobulin gamma 2a genes in wild mice. J Mol Cell Immunol 1, 321–330. Herring, C. D., Chevillard, C., Johnston, S. L., Wettstein, P. J., and Riblet, R. (1998). Vector-hexamer PCR isolation of all insert ends from a YAC contig of the mouse Igh locus. Genome Res 8, 673–681. Honjo, T., and Matsuda, F. (1995). Immunoglobulin heavy chain loci of mouse and human. In Immunoglobulin genes, T. Honjo and F. W. Alt, eds. (London, Academic Press), pp. 145–171. Johnson, K., Angelin-Duclos, C., Park, S., and Calame, K. L. (2003). Changes in histone acetylation are associated with differences in accessibility of V(H) gene segments to V-DJ recombination during B-cell ontogeny and development. Mol Cell Biol 23, 2438–2450. Jouvin-Marche, E., Morgado, M. G., Leguern, C., Voegtle, D., Bonhomme, F., and Cazenave, P. A. (1989). The mouse Igh-1a and Igh-1b H chain constant regions are derived from two distinct isotypic genes. Immunogenetics 29, 92–97. Kabat, E. A., Wu, T. T., Perry, H. M., Gottesman, K. S., and Foeller, C. (1991). Sequences of proteins of immunological interest. U.S. Dept of Health and Human Services, Washington, D.C. Kosak, S. T., Skok, J. A., Medina, K. L., Riblet, R., Le Beau, M. M., Fisher, A. G., and Singh, H. (2002). Subnuclear compartmentalization of immunoglobulin loci during lymphocyte development. Science 296, 158–162. Kurosawa, Y., and Tonegawa, S. (1982). Organization, structure, and assembly of immunoglobulin heavy chain diversity segments. J Exp Med 155, 201–218.
26 Mainville, C., Sheehan, K., Klaman, L. D., Giorgetti, C. A., Press, J. L., and Brodeur, P. H. (1996). Deletional mapping of fifteen mouse Vh gene families reveals a common organization for three Igh haplotypes. J Immunol 156, 1038–1046. Manis, J. P., Michaelson, J. S., Birshtein, B. K., and Alt, F. W. (2003). Elucidation of a downstream boundary of the 3¢ IgH regulatory region. Mol Immunol 39, 753–760. Morgado, M. G., Cam, P., Gris-Liebe, C., Cazenave, P. A., and Jouvin-Marche, E. (1989). Further evidence that BALB/c and C57BL/6 gamma 2a genes originate from two distinct isotypes. EMBO J 8, 3245–3251. Perlmutter, R. M., Berson, B., Griffin, J. A., and Hood, L. (1985). Diversity in the germline antibody repertoire. Molecular evolution of the T15 VH gene family. J Exp Med 162, 1998–2016. Pinaud, E., Khamlichi, A. A., Le Morvan, C., Drouet, M., Nalesso, V., Le Bert, M., and Cogne, M. (2001). Localization of the 3¢ IgH locus elements that effect long-distance regulation of class switch recombination. Immunity 15, 187–199. Potter, M. (1977). Antigen-binding myeloma proteins of mice. Adv Immunol 25, 141–211. Sakano, H., Maki, R., Kurosawa, Y., Roeder, W., and Tonegawa, S. (1980). Two types of somatic recombination are necessary for the generation of complete immunoglobulin heavy-chain genes. Nature 286, 676–683. Shimizu, A., Takahashi, N., Yaoita, Y., and Honjo, T. (1982). Organization of the constant region gene family of the mouse immunoglobulin heavy chain. Cell 28, 499–506. Solin, M. L., and Kaartinen, M. (1992). Allelic polymorphism of mouse Igh-J locus, which encodes immunoglobulin heavy chain joining (JH) segments. Immunogenetics 36, 306–313. Thompson, J. D., Gibson, T. J., Plewniak, F., Jeanmougin, F., and Higgins, D. G. (1997). The CLUSTAL_X windows interface: Flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Res 25, 4876–4882.
Riblet Tutter, A., and Riblet, R. (1988). Duplications and deletions of Vh genes in inbred strains of mice. Immunogenetics 28, 125–135. Tutter, A., and Riblet, R. (1989a). Conservation of an immunoglobulin variable-region gene family indicates a specific, noncoding function. Proc Natl Acad Sci U S A 86, 7460–7464. Tutter, A., and Riblet, R. (1989b). Evolution of the immunoglobulin heavy chain variable region (Igh-V) locus in the genus Mus. Immunogenetics 30, 315–329. Whitcomb, E. A., Haines, B. B., Parmelee, A. P., Pearlman, A. M., and Brodeur, P. H. (1999). Germline structure and differential utilization of Igha and Ighb VH10 genes. J Immunol 162, 1541–1550. Williams, G. S., Martinez, A., Montalbano, A., Tang, A., Mauhar, A., Ogwaro, K. M., Merz, D., Chevillard, C., Riblet, R., and Feeney, A. J. (2001). Unequal v(h) gene rearrangement frequency within the large v(h)7183 gene family is not due to recombination signal sequence variation, and mapping of the genes shows a bias of rearrangement based on chromosomal location. J Immunol 167, 257–263. Wood, C., and Tonegawa, S. (1983). Diversity and joining segments of mouse immunoglobulin heavy chain genes are closely linked and in the same orientation: implications for the joining mechanism. Proc Natl Acad Sci U S A 80, 3030–3034. Zhou, J., Ashouian, N., Delepine, M., Matsuda, F., Chevillard, C., Riblet, R., Schildkraut, C. L., and Birshtein, B. K. (2002a). The origin of a developmentally regulated Igh replicon is located near the border of regulatory domains for Igh replication and expression. Proc Natl Acad Sci U S A 99, 13693–13698. Zhou, J., Ermakova, O. V., Riblet, R., Birshtein, B. K., and Schildkraut, C. L. (2002b). Replication and subnuclear location dynamics of the immunoglobulin heavy-chain locus in B-lineage cells. Mol Cell Biol 22, 4876–4889.
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3 Immunoglobulin k Genes of Human and Mouse HANS G. ZACHAU Adolf Butenandt Institut, Molekularbiologie, Universität München, Germany
that show a lower or higher degree of sequence variation, respectively. The recombination signal sequences at the 3¢ side of the Vk genes consist of conserved 7mer and 9mer sequences at a distance of 12 bp from each other. The early research on antibodies and antibody genes of mouse, human, and other species is recounted in the book by Kindt and Capra (1984). The molecular genetics of immunoglobulins is presented not only in textbooks, but also in reviews (e.g., Max, 1999).
The immunoglobulin k gene chapter in the first edition of this book covered all that was known at the time about the genes of humans and mouse (Zachau, 1989), and the chapter in the second edition concentrated on the human k genes (Zachau, 1995). Although an enormous amount of data on the structure, function, and evolution of the k genes has accumulated in the meantime, the present chapter again deals with the genes of both species. This is possible only since several aspects of the k genes are covered in other chapters of this book. With respect to the early work on k genes, the reader is referred to the previous reviews. In the present chapter, some basic facts on the k genes are recounted, but the emphasis is on the recent work. If a topic is dealt with in several publications, only the latest one is quoted here.
2. HUMAN IMMUNOGLOBULIN k GENES The work on human k genes has been reviewed by Zachau (1989, 1995, 1996, 2000), by Lefranc and Lefranc (2001), and in the database of Lefranc (2002). The results of our group are summarized on the Internet (Zachau, 2001).
1. GENERAL FEATURES OF HUMAN AND MOUSE k GENES
2.1 Elucidation of the Human k Locus
The human and the mouse k loci contain extended Vk gene regions and one Jk–Ck gene region. A typical Vk gene consists of upstream regulatory sequences, a leader sequence, the region coding for the k protein, and, at the 3¢ side, the recombination signal sequences. The upstream regulatory elements comprise, in addition to a TATA box, more or less conserved 10mer and 15mer sequences (Falkner and Zachau, 1984; Schäble and Zachau, 1993; Bemark et al., 1998). The leader sequence is interrupted by an intron, which results in an L and L¢ sequence, the latter being contiguous with the sequence coding for the k protein. Comparison of all known k protein sequences in the database of Kabat (2002; Johnson and Wu, 2001) led to the definition of three framework and three complementarity-determining regions
Molecular Biology of B Cells
A prominent feature of the human k locus is the duplication of most of its Vk gene region. The first pairs of very similar but not identical Vk genes were detected by Bentley and Rabbitts (1983) and by Pech et al. (1985). Numerous cosmid and phage l clones were mapped in our laboratory by restriction nuclease cleavage and assembled in large contigs (review Zachau, 1995). When yeast artificial chromosomes (YAC) and bacterial artificial chromosomes (BAC) became available, we also used those (BrensingKüppers et al., 1997; Kawasaki et al., 2001). A so-called Ck proximal (p) contig of 600 kb comprises, in addition to the Jk–Ck gene region, 40 Vk genes; a distal (d) contig of 440 kb contains 36 Vk genes. The p and d copies of the k locus are arranged in opposite 5¢–3¢ polarities. These
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Copyright 2004, Elsevier Science (USA). All rights reserved.
28
Zachau
are inverted repeats with a still uncloned region of 800 kb in between, which does not seem to contain any Vk genes (Weichhold et al., 1993a). The structure is largely symmetrical starting from a center in the uncloned region. The data on the region of the k locus between the first Vk gene 23 kb upstream of Jk1, which we called B3, and the k deleting element (kde) 24 kb downstream of Ck (Klobeck and Zachau, 1986) were reviewed (Zachau, 1995). No nonVk gene sequences were detected within the k locus, but a transcribed region was found 46 kb downstream of Ck; this was termed BENE (Lautner-Rieske et al., 1995) because of its homology to the membrane protein MAL (de Marco et al., 2001 and earlier literature). A Vk orphon sequence (2.5) was found 1.5 Mb downstream of Ck (Huber et al., 1994),
and another 0.5 Mb further downstream the CD8a locus was localized (Weichhold et al., 1993b).
2.2 Vk Genes, Pseudogenes, Relics, and Repetitive Elements Within the Human k Locus In our laboratory in Munich, we sequenced only the Vk genes and regions of special interest, but more recently we gave our clones to N. Shimizu’s group in Tokyo, who sequenced with them the whole locus (Kawasaki et al., 2001). The results of the mapping and sequencing work on the k locus are summarized in Figure 3.1.
FIGURE 3.1 Comprehensive map of the human immunoglobulin k locus, taken from Kawasaki et al. (2001). (A) Locations of the clones used in sequencing. Sequenced and unsequenced regions are depicted as red and green lines, respectively. (B) Locations of the genes. Vk (red), Jk1-5 (sky blue), and Ck (blue) genes with the same transcriptional polarity are indicated as small vertical lines on the same side of the horizontal line. Lines with full height, 2/3 height, and 1/3 height represent Vk genes with ORFs, pseudogenes with >200 bp, and relics with <200 bp in length, respectively. Relics consisting only of exon I are not included. The names of the Vk genes with ORFs are shown. Thick horizontal lines within the k locus represent duplicated regions; thin horizontal lines indicate regions, which exist in either the proximal or the distal unit. Wedge-shaped shadows indicate deletion events that happened after the inverted duplication. The 6-kb regions between the two wedges show sequence homology but do not seem to be inverted duplication counterparts; rather they correspond to adjacent biock duplicates generated prior to the inverted duplication. (C) Six categories of interspersed repeats are indicated; Alu (green), MIR (blue), LINE L1 and L2 (red), LTR (yellow), DNA transposons (sky blue), and others (purple). (D) The GC content was plotted with a window size of 4,000 nt and with a sliding size of 2,000 nt. (E) Sequence identity without indels between the proximal unit and the distal unit was plotted with a window size of 10,000 nt and with a sliding size of 500 nt. Thirteen homology blocks (A–M) and the average sequence identities (red dashed lines) are indicated. The scale at the bottom of the figure shows the proximal unit; it does not take into account the gaps in the distal unit. See color insert.
3. Immunoglobulin k Genes of Human and Mouse
Seventy-six Vk genes and pseudogenes were identified in the k locus by restriction mapping and sequencing. Eight solitary Vk genes and 34 gene pairs occur with 95 to 100% sequence identity between the p and d copy genes. Thirtytwo of the 76 Vk genes are potentially functional, and 25 are pseudogenes. Minor defects were found in 16 genes, and three genes have potentially functional alleles and alleles with minor defects. A minor defect was defined as a one or two 1-bp alteration in a gene; for example, the occurrence of a stop codon and/or a deviation from the canonical sequences of a regulatory element, a splice site, or a recognition sequence. The genes with minor defects are taken as a separate class of genes, since functional alleles may exist in the human population for more than the three aforementioned genes. This is not to be expected for pseudogenes, which usually carry several defects each. The Vk genes and pseudogenes, including all alleles known to us at the time, and the conserved sequence elements, were compared in a review by Schäble and Zachau (1993). The pseudogenes, the unique sequences, and the repetitive sequences of the k locus were dealt with by Schäble et al. (1994). The sequencing of the whole locus revealed (in addition to the 76 Vk genes) 55 truncated pseudogenes and relics located in the stretches between the Vk genes (Kawasaki et al., 2001). In this publication, the total number of genes and relics is given as 132, since the orphon Z0 (see 2.5) is included in the number; this is not done in this review. The truncated pseudogenes (>200 bp) and the relics (<200 bp) had not been detected by the hybridization techniques employed in our mapping work. The structural features of all sequence elements of the locus and the presence or absence of their rearrangement, transcription, and translation products are compiled in Table 1 of Kawasaki et al. (2001). The classification of k proteins into four subgroups, as used in the database of Kabat (2002), was fully confirmed when the gene sequences became known (Schäble and Zachau, 1993). For subgroups V–VII, no proteins have been found; they are defined on the basis of the nucleotide sequences only. The same is true for the truncated pseudogenes and relics, which have been grouped into five further subgroups VIII–XII. In a phylogenetic tree of the genes and relics, all subgroups can be distinguished (Kawasaki et al., 2001). The average GC content of the k locus is 40.7%, indicating that it belongs to an AT-rich L isochore (Bernardi, 2000). More than 34.9% of the locus consists of interspersed repeats. These were assigned to seven different classes (Kawasaki et al., 2001) and are depicted as a colorful bar in Figure 3.1. The nomenclature of Vk genes used in our publications (O1–O18, A1–A30, L1–L25, B1–B3) evolved as the elucidation of the locus proceeded cluster by cluster. However, in retrospect it became clear that at least some of the clus-
29
ters have a structural, probably evolution-derived significance, since their gene regions share certain sequence features. A systematic nomenclature of the 76 Vk genes and pseudogenes was proposed by Lefranc (2001). All 131 Vk genes, pseudogenes, and relics were designated by a systematic nomenclature stating the gene family and the location within the locus; sequences in homologous positions in the two copies of the locus are distinguished by the suffixes “p” and “d” (Kawasaki et al., 2001); various nomenclatures are compared of this publication. Our old nomenclature is used by many groups today. However, now that the k locus has been sequenced, anybody starting new systematic work on the k genes or gene products also may employ one of the systematic nomenclatures and correlate it to the historic one.
2.3 Rearranged and Expressed Human Vk Genes How many germline Vk genes are actually rearranged, transcribed, and translated? This question was studied by sequencing 70 clones from a cDNA library prepared from spleen mRNA, as well as by comparing numerous rearranged genomic Vk genes, cDNAs, and k proteins from the literature (Klein et al., 1993). Not all potentially functional Vk genes were found to be rearranged and expressed, whereas the alleles of some genes with minor defects are rearranged and expressed. The assignment of the products to the germline genes was summarized in a figure (Klein et al., 1993; Figure 2 in Zachau, 1995) and in a recent survey (Table 1 in Kawasaki et al., 2001). Due to somatic mutations and processes at the V–J junction, several rearrangement and expression products cannot be clearly assigned to one or the other gene of a homologous gene pair. Such products must be attributed to the gene pairs, and both genes of the pair were assumed to be active. In this way, the observed rearrangement products are derived from 42, the cDNAs from 35, and the proteins from 33 germline genes. Considering the aforementioned uncertainties of some assignments, it is reasonable to suppose that about half the 76 germline Vk genes contribute to the light chain repertoire. Possible reasons for the absence of functional products from certain potentially functional genes have been discussed (Klein and Zachau, 1995; Kawasaki et al., 2001). Tomlinson et al. (1995) defined 40 germline Vk genes as “functional” on the basis of structural considerations of the main chain conformations, or canonical structures, they encode. The authors state that all or most of those genes are rearranged. The expression of the genes was not studied. The systematic search for rearrangement and expression products also permitted two conclusions: In mapping the k locus by hybridization, we had not missed any Vk genes, and the Vk genes outside the locus (the orphons [2.5]) are not rearranged or expressed.
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The alignment of the sequences of the germline genes and their products, which were, of course, derived from many different individuals, indicated that the extent of allelic polymorphism in the k locus is low (2.4). The alignments were also useful in defining the preferred sites of somatic mutation. The 5¢–3¢ polarity of the Vk genes within the locus determines the type of Vk–Jk rearrangement. The two Jk proximal Vk genes and all Vk genes of the d copy, whose polarities are opposite to that of Jk–Ck, are arranged by an inversion mechanism (3.2), whereas the other genes rearrange by deletion of the DNA between the Vk and Jk genes (Weichhold et al. 1990). In Figure 3.1 the polarities are indicated by the gene bars pointing either up or down. The work on the germline and expressed Vk genes in apparently healthy individuals was also the basis of studies on the k repertoires in different developmental stages (e.g., Girschick and Lipsky, 2001), in particular disease states (e.g., Pyon et al., 2001), or in population studies (e.g., Padyukov et al., 2001). In a search for allelic variations in the k locus, an additional Vk gene was found in 12 of 57 individuals (Juul et al., 1998). This gene, termed La, is rearranged and transcribed, but it was not yet mapped; it was found to be only 94% similar to the closest known gene of the k locus. Transgenic mice with human immunoglobulin genes produced human immunoglobulins (e.g., Gallo et al., 2000). A Ck sequence altered in one codon was found to give rise to amyloid deposits in a patient (Solomon et al., 1998). Many aspects of immunoglobulin gene expression were reviewed in a symposium (Casali and Silberstein, 1995).
2.4 Polymorphisms in the Human k Locus Since the polymorphisms were extensively discussed earlier (Zachau, 1995; 1996), only three facts should be mentioned here. 1. The extent of allelic variation in the k locus seems to be fairly low (2.3). Therefore, no problems were encountered when we used DNA samples from different human individuals in our structural work 2. In the elucidation of the duplicated parts of the locus, the duplication-differentiating probes played an important role (Pargent et al., 1991) 3. The only startling polymorphism detected is one in which the whole d copy, with its 36 Vk genes, is missing. This so-called haplotype 11 was found in an apparently healthy individual, who is homozygous for it (Pargent et al., 1991). The d-copy gene A2 codes for the most common light chain in the Haemophilus influenzae response. Vaccination of individual 11 with the appropriate carbohydrate vaccine gave rise to antibodies whose light chains were derived, of course, from p-copy genes but carried more mutations than the usual A2-derived light chains (Scott et al., 1992). Haplotype 11 is rather rare (Schäble et al., 1993). Indirect evidence indicates that it is due to a deletion
rather than to the persistence of an evolutionary early nonduplicated structure (Weichhold et al., 1993a).
2.5 Vk Genes Outside the Human k Locus The Vk genes dispersed to positions outside the k locus were first found by Lötscher et al. (1986). They are called orphons in analogy to the histone and ribosomal RNA genes located outside the respective loci. All Vk orphons contain introns and, therefore, certainly have been dispersed at the DNA level and not by mRNA retrotranscription. A group of VkI orphons, the so-called Z family, has closely similar sequences. To date, 24 Vk orphons have been cloned and sequenced. About a dozen further orphons, some of them belonging to the Z family, were detected but not further studied; they may represent new loci or may be alleles of known orphons. Two Vk orphons have no defects in their sequences, but they are included in this group because of their location outside of the locus. The other orphons are pseudogenes, also according to their sequence characteristics. Vk orphons were localized on chromosome 1, 2, 22, and others (review Zachau, 1995). The 12 Vk orphons on the long arm of chromosome 2 (2cen-q13) are of particular interest. These were probably translocated by a pericentric inversion from 2cen-p13, where also the k locus is located. Such inversions seem to be ongoing processes, since 0.1% of the human population carries inverted chromosomes 2 with the k locus on the long arm and the group of orphons on the short arm (LautnerRieske et al., 1993). An orphon on the short arm of chromosome 2, called Z0, was found at a distance of about 140 kb beyond the last Vk gene of the distal copy of the locus (Brensing-Küppers et al., 1997). Z0 may have been the first orphon that left the k locus and became the parent of the other Z-family orphons (Kawasaki et al., 2001). The sequence of ZO is 99.3% and 97.7% identical with the sequences of the Z-family orphons on chromosomes 1 and 22, respectively, but at best 92.6% to the Vk genes within the k locus. The cluster of five orphons on chromosome 22 has no direct counterpart within the k locus either, suggesting that its precursor, as the one of the Z orphons, left an early k locus by a nonduplicative transposition (Kawasaki et al., 2001). A practical note: In searching through hybridization or polymerase chain reactions (PCR) for the germline origin of a rearranged Vk gene or a cDNA, one has a good chance of finding not only genes in the locus, but also orphon sequences.
3. MOUSE IMMUNOGLOBULIN k GENES The earliest work on immunoglobulin genes was on the k genes of the mouse. This research took place in 1974–1975
3. Immunoglobulin k Genes of Human and Mouse
in the laboratory of S. Tonegawa in Basel. In our laboratory, the work on mouse immunoglobulin k genes started in the late 1970s with studies on the chromatin structure of germline and rearranged k genes, on Vk–Jk rearrangements, on nonfunctional Vk–Jk joining, on somatic hypermutation, and on the expression of k genes (review Zachau, 1989). Human k genes were included in some of those experiments for comparison and, from the early 1980s on, we concentrated on the elucidation of the structure of the human k locus (2.1); this research took about 12 years. Only in 1992 did we return to the mouse k genes.
3.1 Elucidation of the Mouse k Locus The strategy of work on the mouse k locus was similar to that used on the human locus: identification of germline Vk genes by hybridization with subgroup-specific probes, cloning of the gene regions, contructing contigs from overlapping clones, and, finally, chromosomal walking to link the contigs. But the work proceeded faster than that on the human k locus, since YAC- and BAC-cloning and longrange PCR had been introduced into genome work in the meantime. Numerous Vk germline and rearranged genes, as well as cDNAs, had been characterized in various mouse strains (review Kofler et al., 1992), but only one contig of two Vk genes was known at the time (Lawler et al., 1992). We prepared a cosmid library of C57BL/6J mouse DNA and screened it with 18 probes, which were more or less specific for the different Vk gene families. The contigs and the still unlinked cosmid clones covered 1.6 Mb, with 85 strong and 11 weak Vk hybridization signals (Zocher et al., 1995). First experiments with Vk gene containing YACs were reported by George et al. (1995). In a systematic study, 43 YACs with C57BL/6J and C3H mouse DNA were analyzed and the first evidence for about 140 Vk genes was obtained (Kirschbaum et al., 1996). A size of 3.5 Mb of the k locus was estimated on the basis of YAC contigs (Schupp et al., 1997). To obtain a detailed restriction map of the locus and exactly localize the Vk genes, cosmid sublibraries were prepared from the YACs. Since the YACs tended to undergo recombinations and deletions, BAC clones were included into our analysis. All maps were eventually established on the level of cosmid clones and subclones thereof. Contigs were designated Z1 at the Ck end to Z9 at the 5¢ end of the locus, and this is how the maps and the gene localizations were reported in the literature up to 1999. The results on the 3¢ and the central parts of the locus were published by Kirschbaum et al. (1998; 1999) and those on the 5¢ part were published by Röschenthaler et al. (1999; 2000). Special attention was paid to the Vk genes and gene families (Thiebe et al., 1999), as well as to the relics and orphons (Schäble et al., 1999). A panel of BAC clones allowed Röschenthaler et al. (2000) to close the gaps in the 5¢ part of the k locus and to show that one of the previously defined contigs (Z7) is
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located near, but not within, the locus. The three pseudogenes of this contig are therefore orphons (3.3). Conversely, three additional Vk genes were found on the BAC clones. The picture of the k locus, as it stood in 1999–2000, at the time of my retirement when our laboratory closed, is that there is a 5¢ contig of 1.88 Mb with 82 Vk genes and a 3¢ contig of 1.04 Mb with 51 Vk genes. In between are two contigs of 65 and 105 kb with 2 and 5 genes, respectively. The detailed restriction maps of the k locus on the Internet (Zachau, 2001) represent this structure. Three gaps of 10 to 40 kb each, comprising together about 90 kb, in the central part of the locus were not bridged in our work because of internal duplications and difficulties with rearranged YACs. A new sublibrary, which would have been required to close the gaps, was not established, since the missing data should become available soon from the mouse genome project. The size of the locus is taken to be about 3.2 Mb, in fair agreement between results of pulsed field gel electrophoresis (PFGE) experiments and the addition of the sizes of the numerous clones. An a-tubulin genelike sequence and an S-adenosyl methionin decarboxylase genelike sequence were detected in the 5¢ and 3¢ contigs, respectively (Röschenthaler et al., 1999; Kirschbaum et al., 1999). The regulatory sequences upstream and downstream of Ck were studied in detail (e.g., Liu et al., 2002). A gene coding for ribose 5-phosphate isomerase was localized 40 to 50 kb downstream of Ck (Apel et al., 1995).
3.2 Vk Genes and Pseudogenes Within the Mouse k Locus One hundred and forty Vk genes and pseudogenes were localized within the k locus, cloned, and sequenced. There are indications that two to five additional Vk genes or pseudogenes, which we were not able to identify, exist in the locus. Less than a third of the Vk genes are oriented in the same 5¢–3¢ polarity as Jk–Ck; the rest are in the opposite polarity. The map positions of the genes and their 5¢ to 3¢ polarities can be seen in the above-mentioned publications and in our final form on the Internet (Zachau, 2001). A so-called signal joint remains in the locus when the rearrangement occurs by inversion. This joint was first cloned from the DNA of a mouse myeloma and sequenced by Steinmetz et al. (1980). Seventy-five of the 140 genes are functional: that is, transcription and/or translation products are known; 21 potentially functional genes have no structural defects, but no expression products have been found yet; 44 Vk genes are pseudogenes (Thiebe et al., 1999; Röschenthaler et al., 2000). In the mouse k locus, we defined no “genes with minor defects” in the human k locus. The characteristics of the genes, their accession numbers, and the expression products are compiled in tables (Kirschbaum et al., 1998; Schäble et al., 1999). A combination of the tables is
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included, together with additional information, in our Internet site (Zachau, 2001). Instead of presenting here detailed maps and tables, an illustration of the mouse k locus, as it was envisaged in 1999, is shown (Figure 3.2). The Vk genes and pseudogenes were assigned to 18 families. The heterogeneous Vk9- and Vk10-gene families of previous classifications were combined into one family, and Vkdv was established as a new one-member gene family, leaving the total number of families unchanged. In general, the members of a family yield clear cross-hybridization signals under stringent conditions and have at least 80% homology in their exon II sequences. As can be seen in Figure 3.2, some clustering of gene families and also of the 5¢ to 3¢ polarities occurs, but notable cases of interspersion also occur. Since the characteristics of the gene families have been described in detail (Thiebe et al., 1999), only a few families should be mentioned here. The Vk4/5 family, which was called “young, dynamic, successful,” is remarkable for its size: 27 functional and potentially functional genes plus 6 pseudogenes were cloned. Since the gaps in our map are in the Vk4/5-gene region, the missing genes also may belong to this family. On the other side, there are the four onemember gene families: Vk22, Vk38C, VkRF, and Vkdv. The
Vk8, Vk19/28, and Vk22 gene families are similar and may be considered a clan. As with the human Vk genes, our nomenclature of the mouse Vk genes developed cluster by cluster, as the work proceeded. An abbreviated designation of the gene family was always included in the name, and once a contig was linked to Jk–Ck, its genes were numbered consecutively: the 22 Jk–Ck proximal genes were named Vk21-1 to Vk8-22. The genes of the yet-unlinked contigs are named by a combination of two letters and the designation of the gene family. For example, ar4 is a Vk4/5 gene in the contig Z2. The attempt to develop a systematic nomenclature was published by Martinez-Jean et al. (2001), but a final nomenclature must await the sequencing of the whole locus. A huge amount of work exists in the literature on mouse Vk genes, and a few recent publications should be mentioned. The germline diversity of the Vk9 and Vk10 gene families was studied by Ulrich et al. (1997) and Fitzsimmons et al. (2002), respectively. The Vk1 and Vk22 genes were the objects of Whitcomb and Brodeur (1998). Vk genes in autoantibodies and in antibacterial antibodies were investigated by Ye et al. (1996) and Emara et al. (1995), respectively. Rapid cloning and PCR techniques applied to Vk genes were frequently studied (e.g., Wang et al., 2000).
FIGURE 3.2 A researcher’s dream of the mouse immunoglobulin k locus, taken from Thiebe et al. (1999); the data of Röschenthaler et al. (2000) are not incorporated. The Jk–Ck and Vk genes are depicted as mice. Different gene families have different colors. Mice in full color designate potentially functional Vk genes, mice sketched only in outline are pseudogenes. Relics are not included. The 5¢, 3¢ direction of the Vk genes is indicated by the direction of the mice. See color insert.
3. Immunoglobulin k Genes of Human and Mouse
3.3 Mouse Vk Relics and Orphons Vk relics are genes with substantial deletions. Those in the human k locus were systematically studied only after the whole locus was sequenced (2.2), whereas in the mouse locus they were detected during the mapping work as weak hybridization signals and immediately sequenced. This was done because such signals sometimes turn out to be caused by cross-hybridization with functional genes of another family. The 18 relics found in the mouse k locus and their characteristics, accession numbers, and locations were compiled by Schäble et al. (1999). A fair number of additional relics also will probably be found in the mouse k locus once its sequence becomes known. Two mouse Vk orphons were localized to chromosomes 16 and 19 (Schupp et al., 1997). They were members of orphon clusters (Schäble et al., 1999). A map of the cluster on chromosome 16, which contains a Vk2, a Vk9, and a Vk20 orphon, was established at a time when it was not yet known that the genes were orphons (Figure 3.2 in Zocher et al., 1995). A Vk2 orphon was found near a Vk20 orphon on chromosome 19. The Vk2 orphon on chromosome 16 contains an internal inversion, but the two Vk2 orphon sequences still are more similar to each other than to the sequences of the Vk2 genes of the k locus. In the two Vk20 orphons, the leader and the exon II segments are separated by several kb of intracisternal A particle sequences. The Vk20 orphon sequences are 99% identical but less similar to those of the Vk20 genes of the locus. Apparently, first a Vk2 and a Vk20 orphon left the locus and both of them were later duplicated. When in the BAC analysis of the 5¢ part of the k locus the contig Z7 could not be linked to the other contigs (3.1), it was shown by fluorescence in situ hybridization (FISH) to be located near the k locus on chromosome 6 but not within the locus (Röschenthaler et al., 2000). Its cytogenetic distance from the locus exceeds 20 Mb. The single Vk1 and the two Vk9 pseudogenes of the contig are therefore classified as orphons.
4. ASPECTS OF EVOLUTION OF THE k GENES When in evolution was the human k locus duplicated? The locus is not yet duplicated in chimpanzees (see below), and the human and chimpanzee clades are thought to have diverged 5 to 7 million years ago. On the basis of 1% sequence divergence between the gene regions of the p and the d copies of the locus, we postulated that the duplication might have taken place 1 to 2 million years ago (Schäble and Zachau, 1993). However, the consideration of the total sequence of the locus led to the conclusion that the duplication had occurred at an earlier time, possibly shortly after the separation of the human clades and the chimpanzee
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clades. Thirteen blocks of homology between the sequences of the p and d copies were detected (A–M in Figure 3.1E), which may have arisen by inversion-mediated recombination. If one takes as the basis for the age of the duplication the sequence difference in the most highly diverged homology blocks, one arrives at the earlier date (Kawasaki et al., 2001). Several evolutionary events must have taken place before the duplication of the human k locus and others afterwards (Zachau, 1995; 2000). The amplification and interdigitation of the Vk genes of different subgroups, and most changes that converted functional genes to pseudogenes, occurred before the duplication. The insertion of an Alu element into the d and not into the p copy of the locus (Lautner-Rieske et al., 1992), the small deletions in one but not the other copy, and at least some of the gene-conversion–like events (Huber et al., 1993) took place after the duplication. The k loci of several nonhuman primate species were studied by Ermert et al. (1995). The Ck gene sequences of human and chimpanzee were found to be 99.6% identical, and the sequences of their Ck proximal Vk genes are remarkably similar. The restriction nuclease digestion patterns of the k loci of different primate species also are quite comparable. However, on the basis of the hybridization patterns obtained with 11 duplication-differentiating probes (2.4), it became clear that the chimpanzee and gorilla have only the part of the k locus in their genomes that corresponds to the p copy of humans. Since cosmid clones from the orphon regions of the human chromosomes 1 and 22 (2.5) hybridized in situ to the homologous chromosome bands in all great apes, the translocation may have happened early in primate evolution (Arnold et al., 1995). The pericentric inversion seems to have occurred after the gorilla and before the chimpanzee clades diverged from the human evolutionary tree. The role of recent duplications and duplicative transpositions in the evolution of the genomes of closely related primates was discussed by Eichler (2001). The mouse k locus is three times the size of the human locus and contains about twice as many Vk genes. Whereas the human locus has one large duplication, the mouse locus contains several smaller duplications. In the human locus, the genes of different families are intermingled, whereas in the mouse some gene families occur clustered. Serial amplifications of gene regions in the rodent clade were postulated as the reason for this clustering (Kirschbaum et al., 1999). Distinct homologies exist between the human gene families I–VII and several mouse gene families, generally ranging from 74 to 84% (for details see Thiebe et al., 1999). Apparently, the formation of gene families predated the divergence of the human and the mouse clades. Only the “young” Vk 4/5 gene family of mouse, with its 70% or less homology to the human genes, probably arose after the point of divergence (Thiebe et al., 1999).
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The evolution of the Vk gene loci was described as a continuous process of duplications and deletions or “birth and death” of the genes (Sitnikova and Nei, 1998). Whereas in humans the k and l proteins are found in comparable percentages, 95% k chains and only 5% l chains exist in the mouse. The ratios are related to the sizes of the light chain loci in the two species (Almagro et al., 1998). A general discussion of V gene evolution is presented by Rothenfluh et al. (1995).
5. CONCLUDING REMARKS The structures of the immunoglobulin k loci of human and mouse were elucidated by classical molecular biology techniques, in parallel to functional and mechanistic studies. Some colleagues involved in the large-scale genome projects called this the “cottage industry approach” (Zachau, 2000). However, most questions concerning the assembly processes and the maturation and functioning of the k genes could be answered in principle on the basis of data obtained through the classical approach, as is obvious from various chapters of this book. Because the human and mouse genomes have been largely sequenced now, the k loci of the two species will be looked at in detail and be annotated soon. Thus, the 800-kb Vk-gene free gap between the p and the d copies of the human k locus and the three small gaps in the mouse k locus will be closed. Also, the sequences adjacent to the k loci and to the orphons may be interesting and contribute to our insight into the processes of evolution and genome dynamics.
Acknowledgments I thank the members of our group for their contributions. The work of our laboratory was supported by Bundesministerium für Forschung und Technologie and by Fonds der Chemischen Industrie.
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3. Immunoglobulin k Genes of Human and Mouse and Zachau, H. G. (1999). The central part of the mouse immunoglobulin k locus. Eur J Immunol 29, 2057–2064 Klein, R., Jaenichen, R., and Zachau, H. G. (1993). Expressed human immunoglobulin k genes and their hypermutation. Eur J Immunol 23, 3248–3262. Klein, R., and Zachau, H. G. (1993). Comparison of human germ-line Vk gene sequences to sequence data from the literature. Eur J Immunol 23, 3263–3271. Klein, R., and Zachau, H. G. (1995). Expression and hypermutation of human immunoglobulin k genes. Ann N Y Acad Sci 764, 74–83 (see Casali and Silberstein, eds.). Klobeck, H.-G., and Zachau, H. G. (1986). The human Ck gene segment and the kappa deleting element are closely linked. Nucleic Acids Res 14, 4591–4603. Kofler, R., Geley, S., Kofler, H., and Helmberg, A. (1992). Mouse variableregion gene families: Complexity, polymorphism and use in nonautoimmune responses. Immunol Rev 128, 5–21. Lautner-Rieske, A., Huber, C., Meindl, A., Pargent, W., Schäble, K. F., Thiebe, R., Zocher, I., and Zachau, H. G. (1992). The human immunoglobulin k locus. Characterization of the duplicated A regions. Eur J Immunol 22, 1023–1029. Lautner-Rieske, A., Hameister, H., Barbi, G., and Zachau, H. G. (1993). Mapping immunoglobulin gene related DNA probes to the central region of normal and pericentrically inverted human chromosome 2. Genomics 16, 497–502. Lautner-Rieske, A., Thiebe, R., and Zachau, H. G. (1995). Searching for non-Vk transcripts from the human immunoglobulin k locus. Gene 159, 199–202. Lawler, A. M., Umar, A., and Gearhart, P. J. (1992). Linkage of two pseudogenes from the Vk1 and Vk9 murine immunoglobulin families. Mol Immunol 29, 295–301. Lefranc, M.-P. (2001). Nomenclature of the human immunoglobulin kappa (IGK) genes. Exp Clin Immunogenet 18, 161–174. Lefranc, M.-P., and Lefranc, G. (2001). The immunoglobulin facts book (London, UK: Academic Press). Lefranc, M.-P. (2002). ImMunoGeneTics database, continuously updated. http://imgt.cnusc.fr:8104 Liu, Z.-M., George-Raizen, J. B., Li, S., Meyers, K. C., Chang, M. Y., and Garrard, W. T. (2002). Chromatin structural analyses of the mouse Igk gene locus reveal new hypersensitive sites specifying a transcriptional silencer and enhancer. J Biol Chem 277, 32640–32649. Lötscher, E., Grzeschik, K.-H., Bauer, H. G., Pohlenz, H.-D., Straubinger, B., and Zachau, H. G. (1986). Dispersed human immunoglobulin k light chain genes. Nature 320, 456–458. de Marco, M. D. C., Kremer, L., Albar, J. P., Martinez-Menárguez, J. A., Ballesta, J., Garcia-López, M. A., Marazuela, M., Puertollano, R., and Alonso, M. A. (2001). BENE, a novel raft-associated protein of the MAL proteolipid family, interacts with caveolin-1 in human endothelial-like ECV304 cells. J Biol Chem 276, 23009–23017. Martinez-Jean, C., Folch, G., and Lefranc, M.-P. (2001). Nomenclature and overview of the mouse (mus musculus and mus sp.) immunoglobulin kappa (IGK) genes. Exp Clin Immunogenet 18, 255–279. Max, E. E. (1999). Immunoglobulins: Molecular Genetics. In Fundamental immunology, 4th ed., W. E. Paul, ed. (Philadelphia: LippincottRaven), pp. 111–182. Padyukov, L., Hahn-Zoric, M., Blomqvist, S. R., Ulanova, M., Welch, S. G., Feeney, A. J., Lau, Y. L., and Hanson, L. A. (2001). Distribution of human kappa locus IGKV2-29 and IGKV2D-29 alleles in Swedish Caucasians and Hong Kong Chinese. Immunogenetics 53, 22–30. Pargent, W., Schäble, K. F., and Zachau, H. G. (1991). Polymorphisms and haplotypes in the human immunoglobulin k locus. Eur J Immunol 21, 1829–1835. Pech, M., Smola, H., Pohlenz, H.-D., Straubinger, B., Gerl, R., and Zachau, H. G. (1985). A large section of the gene locus encoding human
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immunoglobulin variable regions of the k type is duplicated. J Mol Biol 183, 291–299. Pyon, H. S., Ha-Lee, Y. M., Song, G. G., and Sohn, J. (2001). Analysis of the Igk light chain gene variable regions expressed in the rheumatoid synovial B cells. Scand J Immunol 53, 503–509. Röschenthaler, F., Kirschbaum, T., Heim, V., Kirschbaum, V., Schäble, K. F., Schwendinger, J., Zocher, I., and Zachau, H. G. (1999). The 5¢ part of the mouse immunoglobulin k locus. Eur J Immunol 29, 2065–2071. Röschenthaler, F., Hameister, H., and Zachau, H. G. (2000). The 5¢ part of the immunoglobulin k locus as a continuously cloned structure. Eur J Immunol 30, 3349–3354. Rothenfluh, H. S., Blanden, R. V., and Steele, E. J. (1995). Evolution of V genes: DNA sequence structure of functional germline genes and pseudogenes. Immunogenetics 42, 159–171. Schäble, K. F., and Zachau, H. G. (1993). The variable genes of the human immunoglobulin k locus. A review. Biol Chem Hoppe-Seyler 374, 1001–1022. Schäble, K. F., Thiebe, R., Flügel, A., Meindl, A., and Zachau, H. G. (1994). The human immunoglobulin k locus: pseudogenes, unique and repetitive sequences. Biol Chem Hoppe-Seyler 375, 189–199. Schäble, K. F., Thiebe, R., Bensch, A., Brensing-Küppers, J., Heim, V., Kirschbaum, T., Lamm, R., Ohnrich, M., Pourrajabi, S., Röschenthaler, F., Schwendinger, J., Wichelhaus, D., Zocher, I., and Zachau, H. G. (1999). Characteristics of the immunoglobulin Vk genes, Pseudogenes, relics and orphons in the mouse genome. Eur J Immunol 29, 2082– 2086. Schäble, G., Rappold, G. A., Pargent, W., and Zachau, H. G. (1993). The immunoglobulin k locus. Polymorphism and haplotypes of Caucasoid and non-Caucasoid individuals. Hum Genet 91, 261–267; Erratum 92, 105. Schupp, I. W., Schlake, T., Kirschbaum, T., Zachau, H. G., and Boehm, T. (1997). A yeast artificial chromosome contig spanning the mouse immunoglobulin kappa light chain locus. Immunogenetics 45, 180–187. Scott, M. G., Zachau, H. G., and Nahm, M. H. (1992). The human antibody V region repertoire to the type b capsular polysaccharide of Haemophilus influenzae. Int Rev Immunol 9, 45–55. Sitnikova, T., and Nei, M. (1998). Evolution of immunoglobulin kappa variable region genes in vertebrates. Mol Biol Evol 15, 50–60. Solomon, A., Weiss, D. T., Murphy, C. L., Hrncic, R., Wall, J. S., and Schell, M. (1998). Light chain-associated amyloid deposits comprised of a novel k constant domain. Proc Natl Acad Sci U S A 95, 9547–9551. Steinmetz, M., Altenburger, W., and Zachau, H. G. (1980). A rearranged DNA sequence possibly related to the translocation of immunoglobulin gene segments. Nucleic Acids Res 8, 1709–1720. Thiebe, R., Schäble, K. F., Bensch, A., Brensing-Küppers, J., Heim, V., Kirschbaum, T., Mitlöhner, H., Ohnrich, M., Pourrajabi, S., Röschenthaler, F., Schwendinger, J., Wichelhaus, D., Zocher, I., and Zachau, H. G. (1999). The variable genes and gene families of the mouse immunoglobulin k locus. Eur J Immunol 29, 2072–2081. Tomlinson, I. M., Cox, J. P. L., Gherardi, E., Lesk, A. M., and Chothia, C. (1995). The structural repertoire of the human Vk domain. EMBO J 14, 4628–4638. Ulrich, H. D., Moore, F. L., and Schultz, P. G. (1997). Germline diversity within the mouse Igk-V9 gene family. Immunogenetics 47, 91–95. Wang, Z., Raifu, M., Howard, M., Smith, L., Hansen, D., Goldsby, R., and Ratner, D. (2000). Universal PCR amplification of mouse immunoglobulin gene variable regions: The design of degenerate primers and an assessment of the effect of DNA polymerase 3¢ to 5¢ exonuclease activity. J Immunol Methods 233, 167–177. Weichhold, G. M., Klobeck, H.-G., Ohnheiser, R., Combriato, G., and Zachau, H. G. (1990). Megabase inversions in the human genome as physiological events. Nature 347, 90–92. Weichhold, G. M., Ohnheiser, R., and Zachau, H. G. (1993a). The human immunoglobulin k locus consists of two copies that are organized in opposite polarity. Genomics 16, 503–511.
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Weichhold, G. M., Huber, C., Parnes, J. R., and Zachau, H. G. (1993b). The CD8a locus is located on the telomere side of the immunoglobulin k locus at a distance of 2 Mb. Genomics 16, 512–514. Whitcomb, E. A., and Brodeur, P. H. (1998). Rearrangement and selection in the developing Vk repertoire of the mouse: An analysis of the usage of two Vk gene segments. J Immunol 160, 4904–4913. Ye, X. J., Marion, T. N., Terato, K., Aelion, J. A., Cremer, M. A., Tillman, D. M., Krug, M. S., Jackson, B., and Yo, T. J. (1996). Variable-region gene family usage for type II collagen autoantibodies in arthritis-susceptible DBA/1 mice. Clin Immunol Immunopathol 78, 263–275. Zachau, H. G. (1989). Immunoglobulin light-chain genes of the k type in man and mouse. In Immunoglobulin genes, T. Honjo, F. W. Alt, and T. H. Rabbitts, eds. (London, UK: Academic Press), pp. 91–109. Zachau, H. G. (1995). The human immunoglobulin k genes. In Immunoglobulin genes, T. Honjo, F. W. Alt, and T. H. Rabbitts, eds. (London, UK: Academic Press), pp. 173–191. Zachau, H. G. (1996). The human immunoglobulin k genes. Immunologist 4, 49–54. Zachau, H. G. (2000). The immunoglobulin k gene families of human and mouse: A cottage industry approach. Biol Chem 381, 951–954. Zachau, H. G. (2001). http://www.med.uni-muenchen.de/biochemie/ zachau/kappa.htm, homepage updated in 2001, containing the mouse k gene data; human k genes in the section . . . zachau/human_kappa.htm
Zocher, I., Röschenthaler, F., Kirschbaum, T., Schäble, K. F., Hörlein, R., Fleischmann, B., Kofler, R., Geley, S., Hameister, H., and Zachau, H. G. (1995). Clustered and interspersed gene families in the mouse immunoglobulin k locus. Eur J Immunol 25, 3326–3331.
Note Added in Proof This review was concluded at the time of submission in early October 2002. Before receiving the proofs a year later, several relevant publications appeared; for reasons of space only a few of them can be quoted here: Li, S., and Garrard, W. T. (2003). The kinetics of V-J joining throughout 3,5 megabases of the mouse Ig kappa locus fit a constrained diffusion model of nuclear organization. FEBS Lett 536, 125–129. Krangel, M. S. (2003). Gene segment selection in V(D)J recombination: accessibility and beyond. Nature Immunol 4, 624–630. Ohlin, M., and Zouali, M. (2003). The human antibody repertoire to infectious agents: implications for disease pathogenesis. Molec Immunol 40, 1–11. Brekke, K. M., and Garrard, W. T. (personal communication, 2003) assembled the nucleotide sequence of the mouse Igk locus using data from the recent genome sequencing effort. Their emphasis was on analyzing regulatory elements. They also counted 140 Vk genes, of which 95 were called potentially functional.
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4 Immunoglobulin Lambda (IGL) Genes of Human and Mouse MARIE-PAULE LEFRANC AND GÉRARD LEFRANC Université Montpellier II, CNRS, Montpellier, France
This chapter provides a description of the human and mouse germline immunoglobulin lambda (IGL) genes that are used to create the antibody lambda chain repertoire in human and mouse. In order to ensure accuracy, consistency and coherence in the immunoglobulin repertoire description between chain types and species, IMGT, the international ImMunoGeneTics information system® (http://imgt.cines.fr) (Lefranc, 2001a, 2003a), created in 1989 at the Université Montpellier II, CNRS, Montpellier, France, has developed a formal specification of the terms to be used in the domain of immunogenetics and immunoinformatics. This has been the basis of IMGT-ONTOLOGY (Giudicelli and Lefranc, 1999), the first ontology in the domain.
• Gene type—Three types of genes are involved in immunoglobulin lambda synthesis, the variable (V) and joining (J) genes, which encode the antigen binding sites, and the constant (C) genes. • Configuration—The configuration defines the status of the genes: “germline” or “rearranged” for the IGL V and J genes. Note that the C genes do not rearrange directly and therefore their configuration is not defined. • Chain type—The chain type identifies the nature of the peptidic chain potentially encoded by the immunoglobulin IGL genes. The chain type instances are defined by the C gene sequence characteristics. They are refered as Ig-Light-Lambda in IMGT/LIGM-DB (Lefranc, 2000c). • Functionality—The definition of functionality is based on the sequence analysis (Lefranc, 1998). As examples, the instances functional (F) (for germline IGLV and IGLJ, and for IGLC genes), and productive (for rearranged IGL V-J sequences) mean that the coding regions have an open reading frame without a stop codon, and that there is no described defect in the splicing sites, recombination signals, and/or regulatory elements. According to the gravity of the identified defects, the functionality can be defined as open reading frame (ORF) or pseudogene (P) (for germline IGLV and IGLJ, and for IGLC genes), or unproductive (for rearranged IGL V-J sequences).
IGL GENES AND IMGT-ONTOLOGY The human and mouse IGL gene description in this chapter follows the IDENTIFICATION, CLASSIFICATION, and DESCRIPTION concepts of IMGTONTOLOGY (Giudicelli and Lefranc, 1999). The first part of the chapter summarizes the rules of the IMGT Scientific chart (Lefranc et al., 1999a), based on these concepts, for the IGL genes. The second and third part of the chapter provide, for the first time, a description of the genes of a given immunoglobulin locus—the IGL locus—in two different species, human and mouse, with the same IMGT standardized rules for nomenclature and numbering.
IGL Genes and the IDENTIFICATION Concept
IGL Genes and the CLASSIFICATION Concept
The IDENTIFICATION concept allows scientists to identify immunoglobulin lambda sequences according to fundamental biological and immunogenetics characteristics (Giudicelli and Lefranc, 1999).
The CLASSIFICATION concept (Figure 4.1) organizes that immunogenetics knowledge useful to name and classify the immunoglobulin IGL genes (Giudicelli and Lefranc, 1999).
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Copyright 2004, Elsevier Science (USA). All rights reserved.
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the sequence level (Lefranc et al., 1998; Lefranc, 1998). Their sequences are compared to the reference sequence designated as *01 (see IMGT Scientific chart at http://imgt.cines.fr for IMGT description of mutations and IMGT allele nomenclature for sequence polymorphisms). Nomenclature of the Human and Mouse IGL Genes and Alleles
FIGURE 4.1 The CLASSIFICATION concept in IMGT-ONTOLOGY.
• Locus—A locus is a group of immunoglobulin genes that are ordered and localized in the same chromosomal location in a given species. The human and mouse genomes include one main IGL locus on chromosome 22 at 22q11.2 and on chromosome 16 at 13 cM, respectively. Immunoglobulin lambda genes have also been identified in the human genome in other chromosomal locations outside the main locus, thus representing new instances of the locus concept. However, the genes they contain, designated as orphons, are not functional. • Group—A group is a set of genes that share the same gene type (V, J, or C) and participate potentially in the synthesis of a polypeptide of the same chain type. By extension, a group includes the related pseudogenes and orphons. There are three groups—IGLV, IGLJ, and IGLC—for the immunoglobulin lambda genes. • Subgroup—A subgroup is a set of genes that belong to the same group, in a given species, and that share at least 75% identity at the nucleotide level (in the germline configuration for V and J genes). • Gene—A gene is defined as a DNA sequence that can be potentially transcribed and/or translated (this definition includes the regulatory elements in 5¢ and 3¢, and the introns, if present). Instances of the gene concept are gene names. By extension, orphons and pseudogenes are also instances of the gene concept. For each gene, IMGT has defined a reference sequence (Lefranc et al., 1999a). For the V and J genes, the reference sequence corresponds to a germline entity. The rules for the choice of the reference sequences are described at http://imgt.cines.fr, in the IMGT Scientific chart. • Allele—An allele is a polymorphic variant of a gene. Alleles are described, exhaustively and in a standardized way, for the “core” coding regions; that is, the germline V-REGIONs and J-REGIONs, and the C-REGIONs, from immunoglobulin lambda genes. These alleles refer to sequence polymorphisms, with mutations described at
The CLASSIFICATION concept (Figure 4.1) has been used to set up a unique nomenclature of the immunoglobulin genes (Barbié and Lefranc, 1998; Pallarès et al., 1998, 1999; Ruiz et al., 1999; Scaviner et al., 1999; Lefranc, 2000a,b, 2001b; Lefranc and Lefranc, 2001a). A four-letter root designates the group: IGLV, IGLJ, and IGLC for the immunoglobulin lambda genes. Gene names are derived from the four-letter root by adding, if necessary, number(s) and/or letter(s) to allow unambiguous identification of the gene; a single number or letter is used whenever possible. IMGT nomenclature was approved by the HUGO (HUman Genome Organization) Nomenclature Committee, HGNC (http://www.gene.ucl.ac.uk/nomenclature) in 1999 (Wain et al., 2002). All IMGT human immunoglobulin genes have been entered into GDB, Genome Database, Toronto, Canada (http://www.gdb.org); into LocusLink at NCBI (National Center for Biotechnology Information), Bethesda, USA (http://www.ncbi.nlm.nih.gov/LocusLink); and into GeneCards, Weizmann Institute, Israel (http://bioinformatics. weizmann.ac.il/cards/). Links to the IMGT gene cards are provided from GDB, LocusLink, and GeneCards. Links to the accession numbers of the IMGT reference sequences are provided from GDB (Seq@IMGT) and LinkOut at NCBI (http://www.ncbi.nlm.nih.gov/entrez/linkout/). Allele names of the IGL V-REGIONs, J-REGIONs, and C-REGIONs comprise the IMGT gene name followed by an asterisk and a two-figure number. The V-REGIONs, JREGIONs, and C-REGIONs selected as references for the allele polymorphism description have the number *01; other alleles are designated by increasing numbers (*02, *03, . . .) based, if possible, on the chronological order of their publication or confirmation of data by different authors. Note that the number *01 does not mean necessarily that other alleles are already known, but it signifies that any new polymorphic sequence will be described by comparison to that allele *01. IMGT accession numbers are assigned to each allele. Although the IMGT accession numbers are the same as those from the EMBL/GenBank/DDBJ generalist databases, the content of the IMGT/LIGM-DB flat files differs in the expert annotations added by IMGT. IMGT data are available from IMGT/LIGM-DB, IMGT Repertoire, and from Sequence Retrieval System (SRS) sites (available from the IMGT Home page, http://imgt.cines.fr).
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
IGL Genes and the DESCRIPTION Concept The DESCRIPTION concept provides a standardized description of the organization and components of the immunoglobulin sequences, and a characterization of their specific and conserved motifs. Prototypes have been set up to graphically represent the description and configuration of an immunoglobulin gene (Giudicelli et al., 1997). These prototypes give information on the order and expected length (in number of nucleotides) of the labels (Giudicelli et al., 1997; Lefranc et al., 1999a). For example, the prototype V-
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GENE represents a genomic IGLV gene in the germline configuration, whereas V-J-GENE represents genomic IGL V and J genes in the rearranged configuration (Figure 4.2).
The IMGT Unique Numbering A uniform numbering system for the immunoglobulin of all species has been established to facilitate sequence comparison and cross-referencing between experiments from different laboratories, whatever the antigen receptor
FIGURE 4.2 Prototype V-GENE of genomic IGL V gene in the germline configuration and prototype V-J-GENE of genomic V and J genes in the rearranged configuration. Labels (in capital letters) are those used for the sequence description in IMGT (http://imgt.cines.fr). One hundred seventy-seven labels are necessary to describe all structural and functional subregions that compose immunoglobulin sequences (Giudicelli et al., 1997), whereas only seven are available in EMBL, GenBank, or DDBJ.
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(immunoglobulin or T cell receptor), the chain type, or the species. The IMGT unique numbering (Lefranc, 1997, 1999; Lefranc et al., 2003) relies on the high conservation of the structure of the variable regions (V-REGIONs) and domains (V-DOMAINs). This numbering results from the analysis of more than 5,000 sequences, from fish to human. It takes into account and combines the definition of the framework (FR) and complementarity determining regions (CDR) (Kabat et al., 1987, 1991), structural data from X-ray diffraction studies (Satow et al., 1986), and the characterization of the hypervariable loops (Chothia and Lesk, 1987). The delimitations of the FR-IMGT and CDR-IMGT regions have been defined, and correspondence between the IMGT numbering and other numberings has been established (Lefranc, 1999; Lefranc and Lefranc, 2001a,b). The IMGT unique numbering has many advantages: • It allows an easy comparison between sequences coding the variable regions, whatever the antigen receptor (immunoglobulins or T cell receptors), the chain type (heavy or light chains for immunoglobulins), or the species. • In the IMGT unique numbering, the conserved amino acids always have the same position: for example, Cystein 23, Tryptophane 41, Leucine 89, Cystein 104. The hydrophobic amino acids of the framework regions are also found in conserved positions. • This unique numbering has allowed the redefinition of the limits of the FR and CDR. The FR-IMGT and CDRIMGT lengths themselves become crucial information characterizing the variable regions belonging to a group, subgroup, or gene. • Framework amino acids (and codons) located at the same position in different sequences can be compared without requiring sequence alignments. This also holds for amino acids belonging to CDR-IMGT of the same length. • The IMGT unique numbering has allowed a standardized IMGT description of mutations for the IMGT description of allele polymorphisms and somatic hypermutations of the variable regions (Lefranc et al., 1998; Lefranc, 1998). • The unique numbering is used as the output of the IMGT/V-QUEST alignment tool (Lefranc, 2003b,c), which analyses variable (germline or rearranged) sequences according to IMGT criteria (Lefranc et al., 1999a). In IMGT/V-QUEST, for example, a variable rearranged lambda sequence is compared to the appropriate sets of V-REGION and J-REGION alleles from the IMGT reference directory. The results show, aligned with the input sequence, the sequences of the most homologous IGL V-REGION and J-REGION alleles. The aligned V-REGION sequences are displayed according to the IMGT unique numbering and with the FR-IMGT and CDR-IMGT delimitations.
The IMGT/JunctionAnalysis tool displays results from position 104 (2nd-CYS) to position 118 (J-PHE of the IGL J-REGION) (Lefranc, 2003b,c). The IMGT unique numbering has been extended to the C-DOMAINs of the immunoglobulins and T cell receptors, and to all domains belonging to the V-set and C-set of the immunoglobulin superfamily (Lefranc et al., 2003). IMGT Collier de Perles The IMGT Colliers de Perles (Lefranc, 1999; Lefranc et al., 1998, 1999a; Ruiz et al., 2000) are two-dimensional graphical representations of the V-REGIONs and VDOMAINs (Figure 4.3) and C-DOMAINs (Figure 4.4), with strands and loops delimitations, according to the IMGT unique numbering. Colliers de Perles 2D representations provide information on the amino acid positions in the betastrands and loops of the lambda variable and constant domains, and in the FR-IMGT and CDR-IMGT of the VREGIONs and V-DOMAINs. They allow a quick visualization of those amino acids important for the structural configuration of the V-REGION, V-DOMAIN and CDOMAIN. For a given V-REGION or V-DOMAIN, the lengths of the three CDR-IMGT are shown in brackets and separated by dots. For example, [8.3.9] means that in the germline IGLV1–36 gene (Figure 4.3), the CDR1-IMGT, CDR2-IMGT, and CDR3-IMGT are 8, 3, and 9 amino acids long, respectively (Figure 4.3); [9.3.10] means that in the rearranged Mcg IGLV2–8*01-IGLJ1*01 gene (Figure 4.4), the CDR1-IMGT, CDR2-IMGT, and CDR3IMGT are 9, 3, and 10 amino acids long, respectively (Figure 4.4).
THE HUMAN IGL GENES Chromosomal Localization of the Human IGL Locus The human IGL locus is located on chromosome 22 (Erikson et al., 1981), on the long arm, at band 22q11.2 (Emanuel et al., 1985) (Figure 4.5). The orientation of the locus has been determined by the analysis of translocations, involving the IGL locus, in leukemia and lymphoma. Sequencing of the long arm of chromosome 22 showed that it encompasses about 35 megabases of DNA and that the IGL locus is localized at six megabases from the centromere (Dunham et al., 1999). Although the correlation between DNA sequences and chromosomal bands has not yet been made, the localization of the IGL locus can be refined to 22q11.2.
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
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FIGURE 4.3 IMGT Collier de Perles of the V-REGION and V-DOMAIN. (a) IMGT Collier de Perles for the human germline IGLV1–36 (Z73653) according to the IMGT unique numbering for V-REGION. The CDR-IMGT of the germline IGLV1–36 have a length of 8, 3, and 9 amino acids, respectively [8.3.9]. Amino acids are shown in the oneletter abbreviation. Hydrophobic amino acids (hydropathy index with positive value) and tryptophan (W) are found at a given position in more than 50% of analyzed immunoglobulin, and T cell receptor sequences are shown in gray. The CDR-IMGT are limited by amino acids shown in squares, which belong to the neighboring FR-IMGT. Hatched circles correspond to missing positions according to the IMGT unique numbering. Arrows indicate the direction of the beta sheets and their different designations in 3D structure. (b) IMGT Collier de Perles of the human Mcg IGLV2–8*01IGLJ1*01 domain (la8j) (from IMGT/3Dstructure-DB) according to the IMGT unique numbering for V-DOMAIN. The IMGT Collier de Perles is displayed on two sheets, with the hydrogen bonds indicated between amino acids of the inner sheet (amino acids in grayed circles with broad contours).
Organization of the Human IGL Locus The human IGL locus at 22q11.2 spans 1,050 kb (Figure 4.6). The human IGL locus consists of 73 to 74 IGLV genes (Frippiat et al., 1995; Kawasaki et al., 1995, 1997; Williams et al., 1996; for review Pallarès et al., 1998; Scaviner et al., 1999; Lefranc and Lefranc, 2001a), localized on 900 kb, and 7 to 11 IGLJ and 7 to 11 IGLC genes depending on the haplotypes, each IGLC gene being preceded by one IGLJ gene (Hieter et al., 1981; Taub et al., 1983; Dariavach et al., 1987; Vasicek and Leder, 1990; Bauer and Blomberg, 1991) (Table 4.1). One enhancer has been identified 8 kb downstream of the IGLC7 gene (Blomberg et al., 1991; Asenbauer et al., 1999; Combriato and Klobeck, 2002).
The Human IGLV Genes Fifty-six to 57 IGLV genes belong to 11 subgroups, whereas 17 pseudogenes that are too divergent to be assigned to subgroups have been assigned to three clans (Table 4.1) (Frippiat et al., 1995; Williams et al., 1996; Kawasaki et al., 1997). (See Pallarès et al., 1998, for an exhaustive list of the human germline IGLV genes with accession numbers, reference sequences, and sequences from the literature; Lefranc and Lefranc, 2001a, for alignments of alleles of the functional and ORF genes.) The most 5¢ IGLV genes occupy the more centromeric position, whereas the IGLC genes, in 3¢ of the locus, are the most telomeric genes in the IGL locus. All human IGLV genes have the same transcriptional orientation and rearrange by a
FIGURE 4.4 IMGT Collier de Perles of the C-DOMAIN. (a) IMGT Collier de Perles of the human IGLC1 domain (X51755) according to the IMGT unique numbering for C-DOMAIN. Amino acids are shown in the one-letter abbreviation. Hydrophobic amino acids (hydropathy index with positive value) and tryptophan (W) are found at a given position in more than 50% of analyzed immunoglobulin, and T cell receptor sequences are shown in gray. The positions 26, 39, and 104 are shown in squares, by homology with the corresponding positions in the V-REGIONs. Positions 45 and 77, which delimit the characteristic CD strand of the C-DOMAINs, and position 118, which corresponds structurally to J-PHE or J-TRP of the J-REGION, are also shown in squares. Hatched circles or squares correspond to missing positions according to the IMGT unique numbering for C-DOMAINs. (b) IMGT Collier de Perles of the human IGLC1 domain according to the IMGT unique numbering for C-DOMAIN. The IMGT Collier de Perles is displayed on two sheets. Amino acids at positions 1 and 3, 45, and 100 are involved in the Mcg, Ke, and Oz serological markers, respectively (see text and Table 4.2).
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FIGURE 4.4 (Continued)
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TABLE 4.1 Complete list of the human IGL genes on chromosome 22 at 22q11.2 Other nomenclatures
IMGT groups IGLC
IGLJ
IGLV
IMGT gene names (Pallarès et al., 1998; Lefranc, 2001)
IMGT functionality
IMGT reference sequence accession numbers
IMGT number of alleles
J00252 J00253
2 4 1 4 2 2 5 2
IGLC1 IGLC2 (*)IGLC2D1 IGLC3 IGLC4 IGLC5 IGLC6 IGLC7
F F F F P P F, P F
IGLJ1 IGLJ2 (*)IGLJ2D1 IGLJ3 IGLJ4 IGLJ5 IGLJ6 IGLJ7
F F F F ORF ORF F F
X04457 M15641 M15642 X51755 X51755 M18338 X51755
1 1 1 2 1 2 1 2
IGLV1-36 IGLV1-40 IGLV1-41 IGLV1-44 IGLV1-47 IGLV1-50 IGLV1-51 IGLV1-62 IGLV2-5 IGLV2-8 IGLV2-11 IGLV2-14 IGLV2-18 IGLV2-23 IGLV2-28 IGLV2-33 IGLV2-34 IGLV3-1 IGLV3-2 IGLV3-4 IGLV3-6 IGLV3-7 IGLV3-9 IGLV3-10 IGLV3-12 IGLV3-13 IGLV3-15 IGLV3-16 IGLV3-17 IGLV3-19 IGLV3-21 IGLV3-22 IGLV3-24 IGLV3-25 IGLV3-26 IGLV3-27 IGLV3-29 IGLV3-30 IGLV3-31 IGLV3-32 IGLV4-3 IGLV4-60
F F ORF, P F F ORF F P P F F F F F P ORF P F P P P P F, P F F P P F P F F F, P P F P F P P P ORF F F
Z73653 M94116 M94118 Z73654 Z73663 M94112 Z73661 D87022 Z73641 X97462 Z73657 Z73664 Z73642 X14616 X97466 Z73643 D87013 X57826 X97468 D87024 X97465 X97470 X97473 X97464 Z73658 X97463 D87015 X97471 X97472 X56178 X71966 Z73666 X71968 X97474 X97467 D86994 Z73644 Z73646 X97469 Z73645 X57828 Z73667
1 3 2 1 2 1 2 1 2 3 3 4 4 3 1 3 1 1 2 1 2 1 3 2 2 1 1 1 1 1 3 2 2 3 1 1 1 2 2 1 1 3
J00254 J03009 J03010 J03011 X51755
Frippiat et al., 1995; Williams et al., 1996
1a 1e 1d 1c 1g 1f 1b 2a1 2c 2e 2a2 2d 2b2 2b1 2f 3r 3q 3a2 3n 3j 3p 3i 3f 3a 3g 3l 3h 3e 3d 3m 3b 3c 3o 3k 3i1 4c 4a
Kawasaki et al., 1997
1-11 1-13 1-14P 1-16 1-17 1-18 1-19 1-23P 1-1P 1-2 1-3 1-4 1-5 1-7 1-8P 1-9 1-10P 2-1 2-2P 2-3P 2-4P 2-5P 2-6 2-7 2-8 2-9P 2-10P 2-11 2-12P 2-13 2-14 2-15 2-16P 2-17 2-18P 2-19 2-20P 2-21P 2-22P 2-23P 5-1 5-4 (continues)
44
TABLE 4.1 (continued) Other nomenclatures
IMGT groups
IMGT gene names (Pallarès et al., 1998; Lefranc, 2001) IGLV4-69 IGLV5-37 (**)IGLV5-39 IGLV5-45 IGLV5-48 IGLV5-52 IGLV6-57 IGLV7-35 IGLV7-43 IGLV7-46 IGLV8-61 IGLV9-49 IGLV10-54 IGLV10-67 IGLV11-55 IGLV(I)-20 IGLV(I)-38 IGLV(I)-42 IGLV(I)-56 IGLV(I)-63 IGLV(I)-68 IGLV(I)-70 IGLV(IV)-53 IGLV(IV)-59 IGLV(IV)-64 IGLV(IV)-65 IGLV(IV)-66-1 IGLV(V)-58 IGLV(V)-66 IGLV(VI)-22-1 IGLV(V)-25-1 IGLV(VII)-41-1
IMGT functionality F F F F ORF F F P F F, P F F F P ORF P P P P P P P P P P P P P P P P P
IMGT reference sequence accession numbers
IMGT number of alleles
Z73648 Z73672 Z73668 Z73670 Z73649 Z73669 Z73673 Z73660 X14614 Z73674 Z73650 Z73675 Z73676 Z73651 D86996 D87007 D87009 X14613 D86996 D87022 D86993 D86993 D86996 D87000 D87022 D87022 D87004 D87000 D87004 X71351 D86994 X99568
2 1 2 3 1 1 1 1 1 3 3 3 3 2 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1
Frippiat et al., 1995; Williams et al., 1996 4b 5e 5a 5c 5d 5b 6a 7c 7a 7b 8a 9a 10a 10b
V lambda A
Kawasaki et al., 1997 5-6 4-1 4-2 4-3 4-4 1-22 3-1P 3-2 3-3 3-4 5-2 1-20 1-25P 4-6 1-6P 1-12P 1-15P 1-21P 1-24P 1-26P 1-27P 4-5P 4-7P 4-8P 4-9P 5-3P 5-5P
lambdavg2 lambdavg3 lambdavg1
(*) Allelic polymorphism by gene amplification that concerns the IGLJ and IGLC genes: The additional IGLJ2D1 and IGLC2D1 genes belong to the 8-IGLC gene haplotype (see text). (**) Allelic polymorphism by insertion/deletion that concerns the IGLV5-39 gene. The IGLV genes are designated by a number for the subgroup (Chuchana et al., 1990; Frippiat et al., 1995; Williams et al., 1996) followed by a hyphen and a number for the localization from 3¢ to 5¢ in the locus (for review, see Pallarès et al., 1998; Lefranc, 2001; Lefranc and Lefranc, 2001a). In the IGLV gene name column, the IGLV genes are listed, for each subgroup, according to their position from 3¢ to 5¢ in the locus. (I), (IV), (V) (VI), (VII) refer to the clans for those pseudogenes that could not be assigned to subgroups with functional genes. Clans comprise, respectively: — clan I: IGLV1, IGLV2, IGLV6, and IGLV10 subgroup genes and pseudogenes IGLV(I)-20, -38, -42, -56, -63, -68, and -70; — clan II: IGLV3 subgroup genes; — clan III: IGLV7 and IGLV8 subgroup genes; — clan IV: IGLV5 and IGLV11 subgroup genes and pseudogenes IGLV(IV)-53, -59, -64, -65, and -66-1; — clan V: IGLV4 and IGLV9 subgroup genes and pseudogenes IGLV(V)-58 and -66; — clan VI: pseudogenes IGLV(VI)-22-1 and -25-1; — clan VII: pseudogene IGLV(VII)-41-1. A pseudogene belonging to the IGLV2 subgroup has not been locatized (IGLV2-NL1, Z22209) and is not shown in the table. For IMGT list with links to GDB and LocusLink: http://imgt.cines.fr/cgi-bin/IMGTlect.jv?query=203. Information on individual IGL genes is provided in IMGT/GENE-DB: http://imgt.cines.fr
45
46
Lefranc and Lefranc
Protein display of the allele *01 of each functional and ORF IGL J-REGION of the 7-IGLC gene haplotype is shown in Figure 4.7b. In the 8-IGLC gene haplotype, the additional IGLJ2D1 gene is functional (van der Burg et al., 2002). The potential additional IGLJ genes from the 9- to 11-IGLC gene haplotypes have not yet been characterized and sequenced. The Human IGLC Genes
FIGURE 4.5 Chromosomal localization of the human IGL locus at 22q11.2. A vertical line indicates the localization of the IGL locus at 22q11.2. The arrow indicates the orientation 5¢ Æ 3¢ of the locus and the gene group order in the locus. The arrow is proportional to the size of the locus, indicated in kilobases (kb). The total number of genes in the locus in shown between parentheses. Depending on the haplotype, there are 7 to 11 IGLC genes. In the 7- and 8-IGLC gene haplotype, each IGLC gene is preceded, in 5¢, by one IGLJ gene. Although the additional IGLC genes, in the 9-, 10-, and 11-IGLC gene haplotypes have not yet been sequenced, they are also probably preceded by one IGLJ gene. The number of functional genes define the potential IGL repertoire that comprises, in the 7IGLC gene haplotype, 37 to 43 genes (29–33 IGLV, 4–5 IGLJ, and 4–5 IGLC) per haploid genome.
deletion mechanism. IGLV allelic polymorphisms and RFLP were reported, some in association with diseases (for review, Lefranc et al., 1999b). The IMGT Protein display of the functional and ORF IGLV genes is shown in Figure 4.7a. Lengths of the IGLV CDR-IMGT are as follows: CDR1IMGT: 6 to 9; CDR2-IMGT: 3, 7, 8; and germline CDR3IMGT: 7 to 9, 11 (Scaviner et al., 1999). The expressed IGLV repertoire is mainly due to five IGLV genes: IGLV2–14, IGLV1–40, IGLV2–8, IGLV1–44, and IGLV3–21, which encode 60% of the lambda repertoire (Ignatovich et al., 1997, 1999). The Human IGLJ Genes The human IGLJ group comprises seven mapped genes, in the 7-IGLC gene haplotype (Chang et al., 1986; Dariavach et al., 1987; Udey and Blomberg, 1987, 1988; Vasicek and Leder, 1990; Bauer and Blomberg, 1991; Poul et al., 1991). Five IGLJ genes are functional (IGLJ1, IGLJ2, IGLJ3, IGLJ6, and IGLJ7) and two are ORF (IGLJ4 and IGLJ5) (Table 4.1). The IGLJ4 and IGLJ5 have a noncanonical DONOR-SPLICE and a nonconserved J-HEPTAMER. Moreover, they precede the IGLC4 and IGLC5 pseudogenes (Dariavach et al., 1987; Vasicek and Leder, 1990). The IGLJ6 is functional and can be used in a productive lambda chain when it precedes the rare IGLC6 functional allele (Dariavach et al., 1987; Poul et al., 1991), or in a truncated lambda chain when it precedes the IGLC6 pseudogene alleles (Stiernholm et al., 1995). The IMGT
In the human IGL locus, the IGLC group comprises 7 to 11 genes, depending on the haplotypes (Taub et al., 1983; Ghanem et al., 1988; Kay et al., 1992; Lefranc et al., 1999b). In the 7-IGLC gene haplotype, four to five IGLC genes are functional (Table 4.1). The additional IGLC2D1 gene of the 8-gene haplotype is functional (van der Burg et al., 2002). The additional IGLC genes in the 9-, 10-, and 11-gene haplotypes have not yet been sequenced. The IGL C-REGIONs belong to four isotypic forms that differ in limited amino acid substitutions to produce the serological markers Kern (Ke) (Ponstingl et al., 1968; Hess et al., 1971), Oz (Appella and Ein, 1967; Ein and Fahey, 1967; Ein, 1968), and Mcg (Fett and Deutsch, 1974; Fett and Deutsch, 1975). The isotypes expressed by the different human IGLC genes and the amino acid differences between the four isotypes (Dariavach et al., 1987) are shown in Table 4.2. Mcg+ proteins have Asparagine (Asn) 1 and Threonine (Thr) 3 (according to the IMGT unique numbering for C-DOMAIN) (Figure 4.4), whereas Mcg- proteins have Alanine (Ala) 1 and Serine (Ser) 3. Position 82, initially described as characteristic of the Mcg marker, is not involved. Indeed, the protein MOR is different from the other Mcg- proteins by having Ala 82 (Frangione et al., 1985), and the protein MCP (Mcg-) encoded by the IGLC7 gene (Niewold et al., 1996) has Lys 82. Ke+ proteins have Glycine (Gly) 45, whereas Ke- proteins have Ser 45. Oz+ proteins have Lys 100, whereas Oz- proteins have Arginine (Arg) 100. The IGLC1 gene encodes Mcg+ Ke+ Oz- lambda chains (Table 4.2). The IGLC2 gene encodes Mcg- Ke- Oz- lambda chains. The IGLC3 gene encodes Mcg- Ke- Oz+ (alleles IGLC3*01, *02 and *03) or Mcg- Ke- Oz- (allele IGLC3*04) lambda chains. The IGLC3 Ke- Oz- isotype has not been assigned serologically, but instead on the presence of the characteristic amino acids (Kawasaki et al., 1997). The IGLC6 gene is either functional (allele *01) in a rare haplotype and potentially encodes Mcg- Ke+ Oz- lambda chains (Dariavach et al., 1987) or a pseudogene (alleles *02 to *05) in more frequent haplotypes (Vasicek and Leder, 1990). The pseudogene alleles result from a recent insertion (duplication) of four nucleotides (agct), which leads to a frameshift and premature stop codon and, if expressed, to truncated lambda chains (Stiernholm et al., 1995). The IGLC7 gene encodes the four characteristic amino acids of the Mcg- Ke+ Oz- isotype (Ala 1, Ser 3, Gly 45, and Arg 100). The IMGT Protein display of the allele*01 of each functional and ORF IGL C-
FIGURE 4.6 Representation of the human IGL locus at 22q11.2. The boxes representing the genes are not to scale. Exons are not shown. A, B, and C refer to three distinct V-CLUSTERs based on the IGLV gene subgroup content (Williams et al., 1996). Pseudogenes that could not be assigned to subgroups with functional genes are designated by a roman number between parentheses, corresponding to the clans, followed by a hyphen and a number for the localization from 3¢ to 5¢ in the locus. IGLV(IV)-66-1 has been identified in D87004 by IMGT curators (G. Folch, V. Giudicelli, M.-P. Lefranc
[email protected]). The vestigial sequences have been attributed to the clans VI and VII as IGLV(VI)-22-1 (lvg2) and IGLV(VII)-41-1 (lvg1). IGLV(VI)-25-1 (lvg3) has been identified in D86994 by IMGT curators (N. Bosc, M.-P. Lefranc) between IGLV3–25 and IGLV3–26 (Lefranc and Lefranc, 2001a).
47
A. IGLV gene
FR1-IMGT (1-26) 1 10 20 .........|.........|...... Z73653,IGLV1-36 QSVLTQPPS.VSEAPRQRVTISCSGS M94116,IGLV1-40 QSVLTQPPS.VSGAPGQRVTISCTGS M94118,IGLV1-41 QSVLTQPPS.VSAAPGQKVTISCSGS Z73654,IGLV1-44 QSVLTQPPS.ASGTPGQRVTISCSGS Z73663,IGLV1-47 QSVLTQPPS.ASGTPGQRVTISCSGS M94112,IGLV1-50 QSVLTQPPS.VSGAPGQRVTISCTGS Z73661,IGLV1-51 QSVLTQPPS.VSAAPGQKVTISCSGS X97462,IGLV2-8 QSALTQPPS.ASGSPGQSVTISCTGT Z73657,IGLV2-11 QSALTQPRS.VSGSPGQSVTISCTGT Z73664,IGLV2-14 QSALTQPAS.VSGSPGQSITISCTGT Z73642,IGLV2-18 QSALTQPPS.VSGSPGQSVTISCTGT X14616,IGLV2-23 QSALTQPAS.VSGSPGQSITISCTGT Z73643,IGLV2-33 QSALTQPPF.VSGAPGQSVTISCTGT X57826,IGLV3-1 SYELTQPPS.VSVSPGQTASITCSGD X97473,IGLV3-9 SYELTQPLS.VSVALGQTARITCGGN X97464,IGLV3-10 SYELTQPPS.VSVSPGQTARITCSGD Z73658,IGLV3-12 SYELTQPHS.VSVATAQMARITCGGN X97471,IGLV3-16 SYELTQPPS.VSVSLGQMARITCSGE X56178,IGLV3-19 SSELTQDPA.VSVALGQTVRITCQGD X71966,IGLV3-21 SYVLTQPPS.VSVAPGKTARITCGGN Z73666,IGLV3-22 SYELTQLPS.VSVSPGQTARITCSGD X97474,IGLV3-25 SYELMQPPS.VSVSPGQTARITCSGD D86994,IGLV3-27 SYELTQPSS.VSVSPGQTARITCSGD Z73645,IGLV3-32 SSGPTQVPA.VSVALGQMARITCQGD X57828,IGLV4-3 LPVLTQPPS.ASALLGASIKLTCTLS Z73667,IGLV4-60 QPVLTQSSS.ASASLGSSVKLTCTLS Z73648,IGLV4-69 QLVLTQSPS.ASASLGASVKLTCTLS Z73672,IGLV5-37 QPVLTQPPS.SSASPGESARLTCTLP Z73668,IGLV5-39 QPVLTQPTS.LSASPGASARFTCTLR Z73670,IGLV5-45 QAVLTQPAS.LSASPGASASLTCTLR Z73649,IGLV5-48 QPVLTQPTS.LSASPGASARLTCTLR Z73669,IGLV5-52 QPVLTQPSS.HSASSGASVRLTCMLS Z73673,IGLV6-57 NFMLTQPHS.VSESPGKTVTISCTRS X14614,IGLV7-43 QTVVTQEPS.LTVSPGGTVTLTCASS Z73674,IGLV7-46 QAVVTQEPS.LTVSPGGTVTLTCGSS Z73650,IGLV8-61 QTVVTQEPS.FSVSPGGTVTLTCGLS Z73675,IGLV9-49 QPVLTQPPS.ASASLGASVTLTCTLS Z73676,IGLV10-54 QAGLTQPPS.VSKGLRQTATLTCTGN D86996,IGLV11-55 RPVLTQPPS LSASPGATARLPCTLS M34927,V-PREB
CDR1-IMGT (27-38) 30 ...|........ SSNIGNNA.... SSNIGAGYD... SSDMGNYA.... SSNIGSNT.... SSNIGSNY.... SSNIGAGYV... SSNIGNNY.... SSDVGGYNY... SSDVGGYNY... SSDVGGYNY... SSDVGSYNR... SSDVGSYNL... SSDVGDYDH... KLGDKY...... NIGSKN...... ALPKKY...... NIGSKA...... ALPKKY...... SLRSYY...... NIGSKS...... VLGENY...... ALPKQY...... VLAKKY...... SMEGSY...... SEHSTYT..... SGHSSYI..... SGHSSYA..... SDINVGSYN... SGINVGTYR... SGINVGTYR... SGINLGSYR... SGFSVGDFW... SGSIASNY.... TGAVTSGYY... TGAVTSGHY... SGSVSTSYY... SGYSNYK..... SNNVGNQG.... SDLSVGGKN...
FR2-IMGT (39-55) 40 50 .|.........|..... VNWYQQLPGKAPKLLIY VHWYQQLPGTAPKLLIY VSWYQQLPGTAPKLLIY VNWYQQLPGTAPKLLIY VYWYQQLPGTAPKLLIY VHWYQQLPGTAPKLLIY VSWYQQLPGTAPKLLIY VSWYQQHPGKAPKLMIY VSWYQQHPGKAPKLMIY VSWYQQHPGKAPKLMIY VSWYQQPPGTAPKLMIY VSWYQQHPGKAPKLMIY VFWYQKRLSTTSRLLIY ACWYQQKPGQSPVLVIY VHWYQQKPGQAPVLVIY AYWYQQKSGQAPVLVIY VHWYQQKPGQDPVLVIY AYWYQQKPGQFPVLVIY ASWYQQKPGQAPVLVIY VHWYQQKPGQAPVLVIY ADWYQQKPGQAPELVIY AYWYQQKPGQAPVLVIY ARWFQQKPGQAPVLVIY EHWYQQKPGQAPVLVIY IEWYQQRPGRSPQYIMK IAWHQQQPGKAPRYLMK IAWHQQQPEKGPRYLMK IYWYQQKPGSPPRYLLY IYWYQQKPGSLPRYLLR IYWYQQKPGSPPQYLLR IFWYQQKPESPPRYLLS IRWYQQKPGNPPRYLLY VQWYQQRPGSSPTTVIY PNWFQQKPGQAPRALIY PYWFQQKPGQAPRTLIY PSWYQQTPGQAPRTLIY VDWYQQRPGKGPRFVMR AAWLQQHQGHPPKLLSY MFWYQQKPGSSPRLFLY
CDR2-IMGT (56-65) 60 ....|..... YDD....... GNS....... ENN....... SNN....... RNN....... GNS....... DNN....... EVS....... DVS....... EVS....... EVS....... EGS....... NVN....... QDS....... RDS....... EDS....... SDS....... KDS....... GKN....... YDS....... EDS....... KDS....... KDS....... DSS....... VKSDGSH... LEGSGSY... LNSDGSH... YYSDSDK... YKSDSDK... YKSDSDK... YYSDSSK... YHSDSNK... EDN....... STS....... DTS....... STN....... VGTGGIVG.. RNN....... HYSDSDK...
FR3-IMGT (66-104) 70 80 90 100 ....|.........|.........|.........|.... LLPSGVS.DRFSGSK..SGTSASLAISGLQSEDEADYYC NRPSGVP.DRFSGSK..SGTSASLAITGLQAEDEADYYC KRPSGIP.DRFSGSK..SGTSATLGITGLWPEDEADYYC QRPSGVP.DRFSGSK..SGTSASLAISGLQSEDEADYYC QRPSGVP.DRFSGSK..SGTSASLAISGLRSEDEADYYC NRPSGVP.DQFSGSK..SGTSASLAITGLQSEDEADYYC KRPSGIP.DRFSGSK..SGTSATLGITGLQTGDEADYYC KRPSGVP.DRFSGSK..SGNTASLTVSGLQAEDEADYYC KRPSGVP.DRFSGSK..SGNTASLTISGLQAEDEADYYC NRPSGVS.NRFSGSK..SGNTASLTISGLQAEDEADYYC NRPSGVP.DRFSGSK..SGNTASLTISGLQAEDEADYYC KRPSGVS.NRFSGSK..SGNTASLTISGLQAEDEADYYC TRPSGIS.DLFSGSK..SGNMASLTISGLKSEVEANYHC KRPSGIP.ERFSGSN..SGNTATLTISGTQAMDEADYYC NRPSGIP.ERFSGSN..SGNTATLTISRAQAGDEADYYC KRPSGIP.ERFSGSS..SGTMATLTISGAQVEDEADYYC NRPSGIP.ERFSGSN..PGNTTTLTISRIEAGDEADYYC ERPSGIP.ERFSGSS..SGTIVTLTISGVQAEDEADYYC NRPSGIP.DRFSGSS..SGNTASLTITGAQAEDEADYYC DRPSGIP.ERFSGSN..SGNTATLTISRVEAGDEADYYC ERYPGIP.ERFSGST..SGNTTTLTISRVLTEDEADYYC ERPSGIP.ERFSGSS..SGTTVTLTISGVQAEDEADYYC ERPSGIP.ERFSGSS..SGTTVTLTISGAQVEDEADYYC DRPSRIP.ERFSGSK..SGNTTTLTITGAQAEDEADYYY SKGDGIP.DRFMGSS..SGADRYLTFSNLQSDDEAEYHC NKGSGVP.DRFSGSS..SGADRYLTISNLQLEDEADYYC SKGDGIP.DRFSGSS..SGAERYLTISSLQSEDEADYYC GQGSGVP.SRFSGSKDASANTGILLISGLQSEDEADYYC QQGSGVP.SRFSGSKDASTNAGLLLISGLQSEDEADYYC QQGSGVP.SRFSGSKDASANAGILLISGLQSEDEADYYC HQGSGVP.SRFSGSKDASSNAGILVISGLQSEDEADYYC GQGSGVP.SRFSGSNDASANAGILRISGLQPEDEADYYC QRPSGVP.DRFSGSIDSSSNSASLTISGLKTEDEADYYC NKHSWTP.ARFSGSL..LGGKAALTLSGVQPEDEAEYYC NKHSWTP.ARFSGSL..LGGKAALTLSGAQPEDEAEYYC TRSSGVP.DRFSGSI..LGNKAALTITGAQADDESDYYC SKGDGIP.DRFSVLG..SGLNRYLTIKNIQEEDESDYHC NRPSGIS.ERLSASR..SGNTASLTITGLQPEDEADYYC QLGPGVP.SRVSGSKETSSNTAFLLISGLQPEDEADYYC
CDR3-IMGT (105-115) 110 .....|...... AAWDDSLNG... QSYDSSLSG... LAWDTSPRA... AAWDDSLNG... AAWDDSLSG... KAWDNSLNA... GTWDSSLSA... SSYAGSNNF... CSYAGSYTF... SSYTSSSTL... SLYTSSSTF... CSYAGSSTL... SLYSSSYTF... QAWDSSTA.... QVWDSSTA.... YSTDSSGNH... QVWDSSSDH... LSADSSGTY... NSRDSSGNH... QVWDSSSDH... LSGDEDN..... QSADSSGTY... YSAADNN..... QLIDNHA..... GESHTIDGQVG* ETWDSNT..... QTWGTGI..... MIWPSNAS.... AIWYSSTS.... MIWHSSAS.... MIWHSSAS.... GTWHSNSKT... QSYDSSN..... LLYYGGAQ.... LLSYSGAR.... VLYMGSGI.... GADHGSGSNFV* SAWDSSLSA... QVYESSAN....
QPVLHQPPA.MSSALGTTIRLTCTLR NDHDIGVYS... VYWYQQRPGHPPRFLLR YFSQSDQ... SQGPQVP.PRFSGSQDVARNRGYLSISELQPEDEAMYYC AMGARSSEKEER
B. IGLJ genes 1
...... X04457 M15641 M15642 X51755 X51755 M18338 X51755
,IGLJ1 ,IGLJ2 ,IGLJ3 ,IGLJ4 ,IGLJ5 ,IGLJ6 ,IGLJ7
10 .. .|. YVFGTGTKVTVL VVFGGGTKLTVL VVFGGGTKLTVL FVFGGGTQLIIL WVFGEGTELTVL NVFGSGTKVTVL AVFGGGTQLTVL
C. IGLC genes A AB B BC C CD D DE E EF F FG G --------------> ----------> ------> -------> -----------> -------> -----------> 104 1 10 15 16 20 23 30 36 39 41 45 77 84 85 89 96 97 110 118 121 130 7654321|........|....|123|...|...... ...|.....| |.....|1234567|......|12345677654321|..........|12|....... .....|....... ..|.........| X51755 J00253 K01326 J03011 X51755
,IGLC1 ,IGLC2 ,IGLC3 ,IGLC6 ,IGLC7
(G)QPKANPTVTLFPPSSEELQ...ANKATLVCLIS (G)QPKAAPSVTLFPPSSEELQ...ANKATLVCLIS (G)QPKAAPSVTLFPPSSEELQ...ANKATLVCLIS (G)QPKAAPSVTLFPPSSEELQ...ANKATLVCLIS (G)QPKAAPSVTLFPPSSEELQ...ANKATLVCLVS
DFYP..GAVT DFYP..GAVT DFYP..GPVT DFYP..GAVK DFYP..GAVT
VAWKADGSPVKA..GVETTKPSKQSN......NKYAASSYLSLTPEQW..KSHRSYSC VAWKADSSPVKA..GVETTTPSKQSN......NKYAASSYLSLTPEQW..KSHRSYSC VAWKADSSPVKA..GVETTTPSKQSN......NKYAASSYLSLTPEQW..KSHKSYSC VAWKADGSPVNT..GVETTTPSKQSN......NKYAASSYLSLTPEQW..KSHRSYSC VAWKADGSPVKV..GVETTKPSKQSN......NKYAASSYLSLTPEQW..KSHRSYSC
QVTHE....GSTV QVTHE....GSTV QVTHE....GSTV QVTHE....GSTV RVTHE....GSTV
FIGURE 4.7 IMGT Protein displays of the human IGL genes. (a) Human IGL V-REGIONs. Only the allele *01 of each functional or ORF V-REGION is shown. The FR-IMGT and CDR-IMGT are according to the IMGT unique numbering for V-REGION. Human IGLV genes are listed, for each subgroup, according to their position from 3¢ to 5¢ in the locus. For comparison, the human V-PREB region is displayed at the bottom of the figure (only a part of the nonIg-like segment is shown) (Scaviner et al., 1999). (b) Human IGL J-REGIONs. Only the allele *01 of each functional or ORF J-REGION is shown. (c) Human IGL C-REGIONs. The IGL C-REGIONs correspond to a single C-DOMAIN. The strands and loops are according to the IMGT unique numbering for C-DOMAIN. Amino acids at positions 1 and 3, 45, and 100 are involved in the Mcg, Ke, and Oz serological markers, respectively (see text and Table 4.2).
48
EKTVAPTECS EKTVAPTECS EKTVAPTECS EKTVAPAECS EKTVAPAECS
49
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
REGION of the 7-IGLC gene haplotype is shown in Figure 4.7c. Polymorphisms by duplication of the IGLC2 and/or IGLC3 genes have been described in different populations (Taub et al., 1983; Ghanem et al., 1988) with a total number of 7 to 11 IGLC genes. The restriction fragment length polymorphism (RFLP) alleles correspond to polymorphic 8-, 13-, 18-, 23-, 28-kb EcoRI fragments (Taub et al., 1983; Dariavach et al., 1987; Ghanem et al., 1988; Kay et al., 1992; Lefranc et al., 1999b), which contain two, three, four, five, and six IGLC genes, respectively. So far, only the IGLC2 and IGLC3 genes of the 8-kb EcoRI fragment (7IGLC gene haplotype) (Hieter et al., 1981) and the addi-
tional IGLC2D1 gene of the 13-kb EcoRI fragment (8-IGLC gene haplotype) (van der Burg et al., 2002) have been sequenced.
Human IGL Orphons Six IGL orphons have been identified (Table 4.3) (Frippiat et al., 1997; Lefranc, 2001b). Two IGLV orphons are on chromosome 8 at 8q11.2 and one (belonging to subgroup 8) has been sequenced. Two IGLC orphons and two IGLV orphons have also been characterized on 22q, outside the major IGL locus (Dunham et al., 1999; Lefranc, 2001b) (see also IMGT Repertoire, http://imgt.cines.fr).
TABLE 4.2 Correspondence between serological lambda isotypes and IGLC gene and allele names Amino acid positions (1)
Serological isotype
IGLC gene and allele name
1 (6) 112
3 (8) 114
45 (46) 152
82 (57) 163
100 (83) 190
Mcg+ Ke+ Oz-
IGLC1*01, *02
Asn
Thr
Gly
Lys
Arg
Mcg- Ke- Oz-
IGLC2*01, *02, *03 IGLC3*04 IGLC2D1*01
Ala
Ser
Ser
Thr
Arg
Mcg- Ke- Oz+
IGLC3*01, *02, *03
Ala
Ser
Ser
Thr
Lys
IGLC2*04 IGLC6*01 IGLC7*01, *02
Ala
Ser
Gly
Thr
Arg
Ala
Ser
Gly
Lys
Arg
-
+
-
Mcg Ke Oz
(1) Amino acid positions according to the IMGT unique numbering for C-DOMAIN (in bold) (Figure 4.4), to the IMGT exon numbering (between parentheses) and to the Kabat numbering (in italics). Whereas the serological markers Mcg, Ke, and Oz were assigned to Bence-Jones and myeloma proteins using specific antibodies (Walker et al., 1988), their assignment to the IGLC gene and allele names is based on the presence or absence of characteristic amino acids (Dariavach et al., 1987).
TABLE 4.3 List of the human IGL orphons IMGT gene groups
IMGT gene names
IMGT functionality
IMGT reference sequences
Accession numbers
IMGT number of alleles
Chromosomal localization
IGLC
IGLC/OR22-1 IGLC/OR22-2
P P
dJ90G24.3 dJ149A16.1
AL008723 AL021937
1 1
22 (16.1 Mb from the centromere) 22 (16.26 Mb from the centromere)
IGLV
IGLV8/OR8-1 IGLV8/OR8-2 IGLV(IV)/OR22-1 IGLV(IV)/OR22-2
P, ORF (1) P P
Orphée1, TL6 Orphée2 bK390C10.1 DJ149A16.4
Y08831, U03636
2 1 1 1
8q11.2 8q11.2 22 (9.4 Mb from the centromere) 22 (16.28 Mb from the centromere)
AL008721 AL021937
Reference sequences in bold have been mapped: “mapped” refers to sequences that have been obtained from clones (phages, cosmids, YACs) either by subcloning or PCR, and does not apply to sequences obtained directly by PCR from genomic DNA. Orphon genes are designated by a subgroup number (if known) followed by a slash, OR (for Orphon), the chromosome number, a dash, and a specific gene number. References and detailed information on the orphons are available in the IMGT Repertoire, http://imgt.cines.fr. (1) Not sequenced.
50
Lefranc and Lefranc
Total Number of Human IGL Genes and Potential Genomic Repertoire The total number of human IGL genes per haploid genome is 87 to 96 (93 to 102 genes, if the orphons are included) (Table 4.4). The potential genomic human IGL repertoire comprises 37 to 43 functional genes: 29 to 33 functional IGLV genes belonging to 10 subgroups, four to five IGLJ, and four to five IGLC functional genes in the 7-IGLC gene haplotype (Table 4.5).
THE MOUSE IGL GENES Chromosomal Localization and Organization of the Mouse IGL Locus The mouse IGL locus is located on chromosome 16 at 13 cM. A complete map of the IGL locus from Mus musculus domesticus and derived laboratory mice was constructed from clone analysis and pulse field gel electrophoresis (PFGE) of large DNA fragments (Figure 4.8) (Storb et al., 1989). The mouse IGL locus spans 240 kb and consists of
TABLE 4.4 Repertoire of the human germline IGLV genes at 22q11.2 Seventy-three–74 IGLV genes on 900 kilobases: 56 to 57 genes belonging to 11 subgroups and 17 pseudogenes assigned to the clans. Twenty-nine to 30 FUNCTIONAL Five ORF Thirty-five PSEUDOGENE Three FUNCTIONAL or PSEUDOGENE One ORF or PSEUDOGENE Potential repertoire: 29 to 33 FUNCTIONAL IGLV genes belonging to 10 subgroups. Subgroup
Functional
ORF
Pseudogene
IGLV1 (B) (C) IGLV2 (A) IGLV3 (A) IGLV4 (A) (C) IGLV5 (B) IGLV6 (C) IGLV7 (B) IGLV8 (C) IGLV9 (B) IGLV10 (C) IGLV11 (C) IGLV(I) (A) (B) (C) IGLV(IV) (C) IGLV(V) (C) IGLV(VI) (A) IGLV(VII) (B) Total
Total
5 — 5 8(+2)* 1 2 3–4** 1 1(+1)* 1 1 1 — — — — — — — —
1(+1)* — 1 1 — — 1 — — — — — 1 — — — — — — —
(+1)* 1 3 12(+2)* — — — — 1(+1)* — — 1 — 1 2 4 5 2 2 1
7 1 9 23 1 2 4–5** 1 3 1 1 2 1 1 2 4 5 2 2 1
29–30(+3)*
5(+1)*
35(+4)*
73–74**
* The following genes have alleles with different functionality: ORF or PSEUDOGENE (IGLV1-41), FUNCTIONAL or PSEUDOGENE (IGLV3-9, IGLV3-22, IGLV7-46). ** An allelic polymorphism by insertion/deletion, which concerns IGLV5-39 (Frippiat et al., 1995). (A), (B), (C) refer to three distinct V-CLUSTERs based on the IGLV gene subgroup content (Williams et al., 1996). (I), (IV), (V), (VI), (VII) refer to the clans for those pseudogenes that cannot be assigned to subgroups with functional genes.
51
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
FIGURE 4.8 Representation of the mouse (Mus musculus) IGL locus on chromosome 16 at 13 cM. The boxes representing the genes are not to scale. Exons are not shown.
TABLE 4.5 Total number of human immunoglobulin lambda (IGL) genes per haploid genome, compared to the total number of kappa (IGK) and heavy (IGH) genes Major loci
Locus IGL IGK IGH
Chromosomal localization
V
D
J
22q11.2 2p11.2 14q32.33
73–74 (40a or) 76 123–129
0 0 27
7–11 5 9
C
Total number of genes in the major locus
Number of orphons
Total number of genes (including orphons)
7–11 1 11b
87–96 (46a or) 82 170–176c
6 25 36d
93–102 (71a or)–107 206–212c,d
a
Number of genes in the rare IGKV haplotype without the distal V-CLUSTER. Allelic IGHC multigene deletions, duplications, and triplications have been described in healthy individuals. The number of IGHC genes may vary from five (deletion I) to probably nineteen (triplication III) per haploid genome (Lefranc and Lefranc, 2001a). c Not included, the seven nonmapped IGHV genes. d Included, the IGHC processed gene, IGHEP2, localized on chromosome 9 (9p24.2–p24.1). b
three IGLV genes, five IGLJ genes, and four IGLC genes organized in two V-J-C-J-C clusters (Figure 4.8). The 5¢ and 3¢ clusters contain two and one IGLV gene(s), respectively. Each IGLC is preceded by one (or two) IGLJ gene(s). Enhancers have been characterized downstream of each cluster (Hagman et al., 1990; Eccles et al., 1990) (Figure 4.8). A search of the Celera database confirmed the order and transcriptional orientation of the IGL genes (IGLV2, the most 5¢ IGLV gene in the mouse locus, is at 15.6 Mb in the Celera contig, whereas IGLC1, the most 3¢ gene in the locus, is at 15.4 Mb) (Gerdes and Wabl, 2002). The Celera mouse
chromosome 16 (Mmu16) draft sequence (Mural et al., 2002) is derived from four mouse strains (A/J, DBA/2J, 129X1/SvJ, 129S1/SvImJ), whereas the public Mouse Genome Sequencing Consortium (MGSC) draft is generated from the C57BL/6J strain (http://mouse.ensembl.org).
The Mouse IGLV Genes In Mus musculus domesticus and derived laboratory mice (BALB/c), the IGL locus only comprises three IGLV genes
52
Lefranc and Lefranc
belonging to two subgroups (Bernard et al., 1978; Tonegawa et al., 1978a,b; Arp et al., 1982; Weiss and Wu, 1987; Sanchez et al., 1990) (Table 4.7). The IGLV1 and IGLV2 genes that belong to subgroup 1 are localized in the 3¢ and 5¢ cluster, respectively. The IGLV3 gene, which is the unique representative of subgroup 2, is localized downstream of IGLV2 in the 5¢ cluster. It shows a stop codon at its end, which considerably reduces the possibility of a productive rearrangement. In derived strains from wild Mus musculus musculus (MBK, PWK, MAI) and Mus spretus (SMZ, STF) mice, IGLV genes that belong to a third IGLV subgroup have been characterized (Table 4.7). This subgroup is absent in Mus musculus domesticus and in other laboratory mice (BALB/c). The first cDNAs from the IGLV3 subgroup were sequenced from two Mus musculus musculus mice from Skive (Denmark) (clone SD26) and Sladeckovce (Czech Republic) (clone CZ81) (Reidl et al., 1992) (Table 4.7). IGLV3 subgroup genes were then identified in some strains from Mus musculus musculus and North African Mus spretus species (Amrani et al., 2002). The number of IGLV3 subgroup genes varies from zero to at least five in Mus musculus musculus strains, and from zero to three in Mus spretus strains, as deduced from RFLP analysis (Table 4.8). The lost Mus spretus SPE strain probably does not have, as the B6.lambdaSEG strain, any gene belonging to the IGLV3 subgroup. Moreover, these strains lack the IGLV1 gene and only have the two IGLV genes (IGLV2 and IGLV3) of the 5¢ cluster (Amrani et al., 2002) (Table 4.7). The IMGT Protein display of the functional and ORF IGLV genes is shown in Figure 4.9A. Lengths of the IGLV CDR-IMGT are as follows: CDR1-IMGT: 7 to 9; CDR2-IMGT: 3, 7; and
germline CDR3-IMGT: http://imgt.cines.fr).
8,
11
(IMGT
Repertoire,
The Mouse IGLJ and IGLC Genes The IGL locus from Mus musculus domesticus and derived laboratory mice contains four IGLC genes belonging to two subgroups: the IGLC1 and IGLC4 genes belong to subgroup 1, whereas the IGLC2 and IGLC3 genes belong to subgroup 2. Each of these genes is preceded by one (or two) IGLJ gene(s). The IGLJ genes (Table 4.9) and IGLC genes (Table 4.10) are arranged in two clusters: J2-C2-J4C4 and J3-J3P-C3-J1-C1 (Bernard et al., 1978; Blomberg et al., 1981; Blomberg and Tonegawa, 1982; Miller et al., 1981, 1982; Selsing et al., 1982; Weiss and Wu, 1987). Probes specific for the IGLC subgroups allowed researchers to estimate the number of IGLC genes per subgroup in different wild Mus musculus musculus and Mus spretus mice and derived strains (Table 4.9). In the B6.lambda SEG strain, the 3¢ cluster V1-J3-J3P-C3-J1-C1 is deleted and the remaining single cluster displays an additional J–C duplication: V2-V3-J2-C2-J4-C4-J5-C5. However, only lambda2 chains can be expressed (with either a rearranged IGLV2-J2 or IGLV3-J2 gene) since IGLC4 is a pseudogene (Table 4.10) and IGLJ5 is an ORF (Table 4.9). This organization is also probably that of the lost strain SPE (Mami and Kindt, 1987) and of the other Mus spretus strains that do not have lambda1 and lambda3 chains (Amrani et al., 2002). The IMGT Protein displays of the allele *01 of each functional and ORF J-REGION and C-REGION are shown in Figure 4.9B and Figure 4.9C, respectively.
TABLE 4.6 Number of functional human immunoglobulin lambda (IGL) genes per haploid genome compared to the number of functional kappa (IGK) and heavy (IGH) genes Chromosomal localization
Locus size in kb (kilobases)
V
D
J
C
Number of functional genes
IGL
22q11.2
1050
29–33
0
4–5
4–5
37–43
29 ¥ 4 = 116 (m) 33 ¥ 5 = 165 (M)
IGK
2p11.2
1820
30–35
0
5
1
36–41
500a
17–19a
0
5
1
23–25a
30 ¥ 5 = 150 (m) 35 ¥ 5 = 175 (M) 17 ¥ 5 = 85 (m)a 19 ¥ 5 = 95 (M)a
1250
38–46
23
6
9b
76–84
Locus
IGH
a
14q32.33
Combinatorial diversity (range per locus)
38 ¥ 23 ¥ 6 = 5244 (m) 46 ¥ 23 ¥ 6 = 6348 (M)
In the rare IGKV haplotype without the distal V-CLUSTER. In haplotypes with multigene deletion, the number of functional IGHC genes is five (deletions I, III, and V), six (deletions IV and VI), or eight (deletion II) per haploid genome (Lefranc and Lefranc, 2001a). In haplotypes with multigene duplication or triplication, the exact number of functional IGHC genes per haploid genome is not known. The range of the theoretical combinatorial diversity indicated takes into account the minimum (m) and the maximum (M) number of functional V, D, and J genes in each of the major IGL, IGK, and IGH loci. b
TABLE 4.7 Mouse (Mus musculus, Mus spretus) IGLV germline genes Mouse (Mus musculus) IGLV IGLV subgroup
IGLV gene name
IGLV allele name
Fct
IGLV1
IGLV1
IGLV1*01
F
BALB/c
V1
J00590
IGLV1*02 IGLV2*01 IGLV2*02
F F F
BALB/c
IGLV2
BALB/c
M315/eVl1 V2 J558/eVl2
X58417 J00599 X58412
Strain
Reference sequences
Accession numbers
Sequences from the literature BALB/c, A1-13/eVl1[X58409], BALB/c [V00811] [V00815] BALB/c [X58418], BALB/c, M315e/Vl2[X58423], BALB/c [X58424]
IGLV2
IGLV3
IGLV3*01
F
BALB/c
Lg1
M34597
BALB/c, VLx (Vlambdax)[D38129]
IGLV3
IGLV4 IGLV5 IGLV6
IGLV4*01 IGLV5*01 IGLV6*01 IGLV6*02 IGLV6*03 IGLV7*01 IGLV7*02 IGLV8*01 IGLV8*02
[F] [F] [F] [F] [F] [F] [F] [F] [F]
MBK PWK PWK MAI MBK PWK MBK MAI MBK
MBK2 PWK1 PWK3 MAI2 MBK4 PWK2 MBK1 MAI1 MBK3
AF357985° AF357981° AF357979° AF357983° AF357987° AF357980° AF357984° AF357982° AF357986°
SD26[M94349]#c CZ81[M94351]#c
IGLV7 IGLV8
#c: rearranged cDNA. °: genomic DNA, but not known as being germline or rearranged.
Mouse (Mus spretus) IGLV IGLV gene name
IGLV allele name
Fct
IGLV1 IGLV2
(1) IGLV2*01
F
SPE
IGLV2SPE (Vlambda2SPE)
M17529
IGLV2
IGLV3
IGLV3*01
F
B6.lambdaSEG
VlambdaxSEG
AF357988
IGLV3
IGLV4
IGLV4*01 IGLV4*02 IGLV8*01
[F] [F] [F]
SMZ STF SMZ
SMZ1 STF2 SMZ2
AF357978° AF357975° AF357977°
IGLV subgroup IGLV1
IGLV8
Strain
Reference sequences
Accession numbers
Sequences from the literature
B6.lambdaSEG, Vlambda2SEG [AF357989]
STF, STF1[AF357976]°
Functionality (Fct) is shown between brackets when the accession number refers to genomic DNA, but not known as being germline or rearranged. Reference sequences in bold have been mapped: “mapped” refers to sequences that have been obtained from clones (phages, cosmids, YACs) either by subcloning or PCR, and does not apply to sequences obtained directly from genomic DNA. For a given gene name, each horizontal line corresponds to a different allele. (1) The cluster IGLV1-IGLJ3_IGLC3_IGLJ1_IGLC1 is deleted in B6. lambdaSEG and probably also in the lost strain SPE (Amrani et al., 2002). See Tables 4.9 and 4.10 for IGLJ and IGLC genes, respectively.
53
A. Mouse (Mus musculus) IGLV IGLV gene
° ° ° ° °
FR1-IMGT CDR1-IMGT FR2-IMGT CDR2-IMGT FR3-IMGT CDR3-IMGT (1-26) (27-38) (39-55) (56-65) (66-104) (105-115) 1 10 20 30 40 50 60 70 80 90 100 110 .........|.........|...... ...|........ .|.........|..... ....|..... ....|.........|.........|.........|.... .....|......
J00590 ,IGLV1 J00599 ,IGLV2 M34597 ,IGLV3(1) AF357985, IGLV4 AF357981, IGLV5 AF357979, IGLV6 AF357980, IGLV7 AF357982, IGLV8
QAVVTQESA.LTTSPGETVTLTCRSS QAVVTQESA.LTTSPGGTVILTCRSS QLVLTQSSS.ASFSLGASAKLTCTLS TQPSS.VSTSLGSTVKLSCKRS TQPSS.VSTSLGSTVKLSCKRS TQPSS.VSTSLGSTVKLPCKCS TQPSS.VSTSLGSTVKLSCKPS TQPSS.VSTSLGSTVKLPCKRS
TGAVTTSNY... TGAVTTSNY... SQHSTYT..... TGNIGNNY.... TGNIGNNY.... TGNIGSYY.... TGKIGNYF.... TGNIGNDY....
ANWVQEKPDHLFTGLIG ANWVQEKPDHLFTGLIG IEWYQQQPLKPPKYVME VHWYQQYMGRSPTNMIY VNWYQQYMGRSPTNMIY VHWYQQHMGRSPTNMIH MSWYQQHMGRSPTNMIY VHWYQQHMGRSPTNMIY
GTN....... GTS....... LKKDGSH... DDN....... GDD....... SDD....... RDD....... RDD.......
NRAPGVP.ARFSGSL..IGDKAALTITGAQTEDEAIYFC ALWYSNHF.... NRAPGVP.VRFSGSL..IGDKAALTITGAQTEDDAMYFC ALWYSTHF.... STGDGIP.DRFSGSS..SGADRYLSISNIQPEDEAIYIC GVDTIKEQFV*. KRPSGVS.DRFSGSIDSSSNSAFLTINNVQAED QRPTGVS.DRFSGSIDSSSNSAFLTINNVQAED QRPSGVS.DRFSGSIDSSSNSAFLTINNVQAED LRPSGVS.DRFSGSIDSSSNSAFLTINNVQAED QRPSGVS.DRFSGSIDSSSNSAFLTINNVQAED
Mouse (Mus spretus) IGLV IGLV gene
FR1-IMGT CDR1-IMGT FR2-IMGT CDR2-IMGT FR3-IMGT CDR3-IMGT (1-26) (27-38) (39-55) (56-65) (66-104) (105-115) 1 10 20 30 40 50 60 70 80 90 100 110 .........|.........|...... ...|........ .|.........|..... ....|..... ....|.........|.........|.........|.... .....|......
M17529 ,IGLV2 AF357988,IGLV3 ° AF357978, IGLV4 ° AF357977, IGLV8
QAVVTQESA.LTTSPGGTVILTCRSS QPVLTQSSS.ASFSLGASAKLTCTLS TQPSS.VSTSLGSTVKLSCKRS TQPSS.VSTSLGSTVKLPCKRS
TGAVTTSNY... SEHSTYI..... TGNIGNN..... TGNIGNN.....
AIWVQEKTDHLFAGVIG IEWYQQQPLKPPKYVMQ YVHWYQQYMGRSPTNMI YVHWYQQHMGRPPTNMI
DTS....... LKKDGSH... YDD....... YRD.......
NRAPGVP.ARFSGSL..IGDKAALTITGAQTEDDAMYFC ALWYSNHF.... SKGDGIP.DRFSGSS..SGADRYLSISNIQPEDEAIYIC GVDDNIRGQFV. NKRPSGVSDRFSGSIDSSSNSAFLTINNVQAED DQRPSGVSDRFSGSIDSSSNSAFLTINNVQAED
B. Mouse (Mus musculus) IGLJ __________________________________________ IGLJ segments __________________________________________ 1 10 .........|... V00813 J00593 J00583 J00584 J00596
,IGLJ1 ,IGLJ2 ,IGLJ3 ,IGLJ3P ,IGLJ4(1)
WVFGGGTKLTVL. YVFGGGTKVTVL. FIFGSGTKVTVL. GSFSSNGLLYAG. WVFGGGTRLTVL.
Mouse (Mus spretus) IGLJ __________________________________________ IGLJ segments __________________________________________ 1 10 .........|... M16555 ,IGLJ4(1) AF357974,IGLJ5
WVFGGGTRLTVL. WVFGGGTRLTVL.
FIGURE 4.9 IMGT Protein displays of the mouse (Mus musculus, Mus spretus) IGL genes. (a) Mouse IGL VREGIONs. Only the allele *01 of each functional or ORF V-REGION is shown. (1) The last codon of the CDR3-IMGT of the Mus musculus IGLV3 gene is a STOP-CODON, which can disappear during rearrangements. °: genomic DNA, but not known as being germline or rearranged. Partial sequences at both ends. (b) Mouse IGL J-REGIONs. Only the allele *01 of each functional or ORF J-REGION is shown. The sequences of Mus musculus and Mus spretus IGLJ4*01 are identical. (c) Mouse IGL C-REGIONs. The IGLC Protein display is according to the IMGT unique numbering for C-DOMAIN. Only the allele *01 of each functional or ORF J-REGION is shown. N-glycosylation sites (NXS/T, where X is different from P) are underlined. The IGL C-REGIONs correspond to a single C-DOMAIN. The strands and loops are according to the IMGT unique numbering for C-DOMAIN.
54
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
55
C. Mouse (Mus musculus) IGLC ____________________________________________________________________________________________________________________________________________________________ IGLC genes ____________________________________________________________________________________________________________________________________________________________ A AB B BC C CD D DE E EF F FG G --------------> ----------> ------> -------> -----------> -------> ----------> 1 10 15 16 20 23 30 36 39 41 45 77 84 85 89 96 97 104 110 118 121 130 7654321|........|....|123|...|...... ...|.....| |.....|1234567|......|12345677654321|..........|12|....... .....|....... ..|.........| J00587 J00595 J00585
,IGLC1 ,IGLC2 ,IGLC3
(G)QPKSSPSVTLFPPSSEELE...TNKATLVCTIT DFYP..GVVT (G)QPKSTPTLTVFPPSSEELK...ENKATLVCLIS NFSP..SGVT (G)QPKSTPTLTMFPPSPEELQ...ENKATLVCLIS NFSP..SGVT
VDWKVDGTPVTQ..GMETTQPSKQSN......NKYMASSYLTLTARAW..ERHSSYSC QVTHE....GHTV EKSLSRADCS VAWKANGTPITQ..GVDTSNPTKEGN.......KFMASSFLHLTSDQW..RSHNSFTC QVTHE....GDTV EKSLSPAECL VAWKANGTPITQ..GVDTSNPTKEDN.......KYMASSFLHLTSDQW..RSHNSFTC QVTHE....GDTV EKSLSPAECL
Mouse (Mus spretus) IGLC ____________________________________________________________________________________________________________________________________________________________ IGLC genes ____________________________________________________________________________________________________________________________________________________________ A AB B BC C CD D DE E EF F FG G --------------> ----------> ------> -------> -----------> -------> ----------> 104 1 10 15 16 20 23 30 36 39 41 45 77 84 85 89 96 97 110 118 121 130 7654321|........|....|123|...|...... ...|.....| |.....|1234567|......|12345677654321|..........|12|....... .....|....... ..|.........| M16554 ,IGLC2 AF357974,IGLC5
(G)QPKSTPTLTVFPPSSEELK...ENKATLVCLIS NFSP..SGVT VAWKANGTPITQ..GVDTSNPTKEGN.......KFMASSFLHLTSDQW..RSHNSFTC QVTHE....GDTV EKSLSPAECL (G)QPKATPSVNLFPPSSEELK...TKKATLVCMIT EFYA..TAVR MAWKADGTPITQ..DVETTQPPKQS........DNMASSYLLFTAEAW..ESHSSYSC HVTHE....GNTV EKNLSRAECS
FIGURE 4.9 (Continued) TABLE 4.8 Mouse (Mus musculus, Mus spretus) IGLV RFLP Mouse (Mus musculus) Number of IGLV genes IGLV subgroup IGLV1
IGLV gene namea
BALB/c
IGLV1 IGLV2
1 1
MAI
MBK
PWK
2
3
3
2
IGLV2
IGLV3
1
2
2
2
IGLV3
IGLV4–
0
5
5
5
IGLV8
Mouse (Mus spretus) IGLV subgroup IGLV1
IGLV gene namea IGLV1 IGLV2
Number of IGLV genes B6.lambdaSEG 0 1
STF
SMZ
1
3
2
IGLV2
IGLV3
1
1
1
IGLV3
IGLV4,
0
2
3
IGLV8M a
For sequenced genes (see Table 4.7).
Total Number of Mouse IGL Genes and Potential Genomic Repertoire In Mus musculus domesticus and laboratory mice, a total of 12 IGL genes exist per haploid genome. The potential genomic repertoire comprises nine functional genes: three functional IGLV genes that belong to two subgroups, three
IGLJ, and three IGLC functional genes (Tables 4.7, 4.9, and 4.10). In contrast, wild Mus musculus musculus and Mus spretus mice have a more diverse and polymorphic repertoire. Although the organization of the IGL locus in these mice is not known, preliminary data suggest that the total number of IGLV genes may vary from two to at least ten (Table 4.8), and the total number of IGLC may vary from three to at least 10, depending on the strains (Table 4.11). Sequencing will be required to evaluate the functionality of these genes. The phylogenetic tree obtained with IMGT/PhyloGene (Figure 4.10) shows that the mouse IGLV1 subgroup (IGLV1 and IGLV2 genes) is related to the human IGLV7 subgroup, the mouse IGLV2 subgroup (IGLV3 gene) to the human IGLV4 subgroup, and the mouse IGLV3 subgroup (IGLV4 to IGLV8 genes found in wild mice) to the human single IGLV6 subgroup.
CONCLUSION The IMGT classification and description of the human and mouse IGL genes and alleles allow, for the first time, an easy and standardized comparison of the genome and genetics data between species. IMGT/V-QUEST and IMGT/JunctionAnalysis online tools, based on IMGT reference sequence data sets, facilitate the analysis of the human and mouse lambda repertoire in normal and pathological situations at the allele level. Moreover, the IMGT unique numbering for V-DOMAIN and C-DOMAIN provides interesting insights into the 3D structure and function of the immunoglobulin lambda chains between mouse and human. In mouse, it was demonstrated that a selective 50-fold
TABLE 4.9 Mouse (Mus musculus, Mus spretus) IGLJ germline genes Mouse (Mus musculus) IGLJ IGLJ gene name
IGLJ allele name
IGLJ1
IGLJ1*01
F
BALB/c
J1
V00813
BALB/c, M315J1 [X58419], J1 [Js00586], BALB/c [X58411]
IGLJ2
IGLJ2*01
F
BALB/c
J2
J00593
BALB/c, M315J2 [X58420], BALB/c [X58414]
IGLJ3
IGLJ3*01
F
BALB/c
J3
J00583
BALB/c M315J3 [X58421], BALB/c [X58411]
IGLJ3P
IGLJ3P*01
ORF
BALB/c
Pseudo J3
J00584
IGLJ4
IGLJ4*01
ORF
BALB/c
PseudoJL4
J00596
BALB/c, M315J4 [X58422], BALB/c [X58414]
IGLJ allele name
Strain
Reference sequences
Accession numbers
Sequences from the literature
Fct
IGLJ4
IGLJ4*01
ORF
SPE
J4SPE
M16555
IGLJ5
IGLJ5*01
ORF
B6.lambda SEG
J4SEG2
AF357974
Fct
Reference sequences
Strain
Accession numbers
Sequences from the literature
Mouse (Mus spretus) IGLJ IGLJ gene name IGLJ2
Fct: IMGT functionality. Reference sequences in bold have been mapped: “mapped” refers to sequences that have been obtained from clones (phages, cosmids, YACs) either by subcloning or PCR, and does not apply to sequences obtained directly by PCR from genomic DNA.
TABLE 4.10 Mouse (Mus musculus, Mus spretus) IGLC genes and alleles Mouse (Mus musculus) IGLC IGLC subgroup
IGLC gene name
IGLC allele name
Fct
Strain
IGLC1
IGLC1 IGLC4
IGLC1*01 IGLC1*02 IGLC4*01
F P P
BALB/c BALB/c BALB/c
IGLC5
—
IGLC2 IGLC3
IGLC2*01 IGLC3*01
F F
IGLC gene name
IGLC allele name
Fct
Strain
IGLC1 IGLC4
— IGLC4*01
P
B6.lambdaSEG SPE
IGLC5
IGLC4*02 IGLC5*01
P F
B6.lambdaSEG B6.lambdaSEG
IGLC2*01 IGLC2*02 —
F F
SPE B6.lambdaSEG B6.lambdaSEG
IGLC2
Reference sequences
Accession numbers
Clambda1
BALB/c [X58411]
J558/aCl4
J00587 V00814 X58416
Clambda2 Clambda3
J00595 J00585
[J00592], BALB/c [X58414] BALB/c [X58415], BALB/c [X58411]
Sequences from the literature
BALB/c, M315/eCL4 [X58410], BALB/c, Clambda4 [J00598], BALB/c [X58414]
BALB/c BALB/c BALB/c
Mouse (Mus spretus) IGLC IGLC subgroup IGLC1
IGLC2
IGLC2 IGLC3
Reference sequences
Accession numbers
Clambda4S (Clambda4SPE) Clambda4SEG1 Clambda4SEG2
M16628
Clambda2SPE Clambda2SEG
M16554 AF357973
Sequences from the literature
AF357972 AF357974
Fct: IMGT functionality. Reference sequences in bold have been mapped: “mapped” refers to sequences that have been obtained from clones (phages, cosmids, YACs) either by subcloning or PCR, and does not apply to sequences obtained directly by PCR from genomic DNA. A dash indicates the absence of a gene.
56
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse
57
TABLE 4.11 Mouse (Mus musculus, Mus spretus) IGLC RFLP Mouse (Mus musculus) IGLC subgroup
IGLC gene namea
Number of IGLC genes BALB/c
MAI
MBK
PWK
IGLC1
IGLC1 IGLC4 IGLC5
1 1 0
2
3
3
2
IGLC2
IGLC2 IGLC3
1 1
2
7
6
6
Mouse (Mus spretus) IGLC subgroup
IGLC gene namea
Number of IGLC genes B6.lambdaSEG
STF
SMZ
IGLC1
IGLC1 IGLC4 IGLC5
0 1 1
2
2
3
IGLC2
IGLC2 IGLC3
1 0
1
4
5
a
For sequenced genes (see Table 4.10).
decrease in lambda1, observed in the SJL and related BSVS and FVB strains, is due to a single point mutation that changes a glycine to a valine at position 45 in the IGLC1 gene (Sun et al., 2002). Interestingly, that position is at the beginning of the CD transversal strand of the C-DOMAIN and corresponds to the position that, in the human IGLC genes, is involved in the Kern serological marker. This correlation is of particular interest since the mutation in the mouse lambda chain leads to a defect in B cell receptor signaling.
Acknowledgments We are grateful to the IMGT team members for their helpful contribution. We thank Valérie Thouvenin-Contet for her assistance in the preparation of the manuscript. IMGT is funded by the European Union’s 5th PCRDT (QLG2-2000-01287) program, the Centre National de la Recherche Scientifique (CNRS), and the Ministère de la Recherche et de l’Education Nationale.
References Amrani, Y. M., Voegtlé, D., Montagutelli, X., Cazenave, P. A., and Six, A. (2002). The Ig light chain restricted B6.k-lSEG mouse strain suggests that the IGL locus genomic organization is subject to constant evolution. Immunogenetics 54, 106–119. Appella, E., and Ein, D. (1967). Two types of lambda polypeptide chains in human immunoglobulins based on an amino acid substitution at position 190. Proc Natl Acad Sci U S A 57, 1449–1454.
FIGURE 4.10 Phylogenetic tree of human and mouse IGLV genes, using IMGT/PhyloGene (Elemento and Lefranc, 2003). The tree, constructed using a distance matrix and the neighbor-joining algorithm, is displayed with branch lengths and rooted using the midpoint procedure.
Arp, B., McMullen, M. D., and Storb, U. (1982). Sequences of immunoglobulin lambda 1 genes in a lambda 1 defective mouse strain. Nature 298, 184–187. Asenbauer, H., Combriato, G., and Klobeck, H. G. (1999). The immunoglobulin lambda light chain enhancer consists of three modules which synergize in activation of transcription. Eur J Immunol 29, 713–724. Barbié, V., and Lefranc, M.-P. (1998). The human immunoglobulin kappa variable (IGKV) genes and joining (IGKJ) segments. Exp Clin Immunogenet 15, 171–183. Bauer, T. R. Jr., and Blomberg, B. (1991). The human lambda L chain Ig locus. Recharacterization of JC lambda 6 and identification of a functional JC lambda 7. J Immunol 146, 2813–2820. Bernard, O., Hozumi, N., and Tonegawa, S. (1978). Sequences of mouse immunoglobulin light chain genes before and after somatic changes. Cell 15, 1133–1144. Blomberg, B., and Tonegawa, S. (1982). DNA sequences of the joining regions of mouse lambda light chain immunoglobulin genes. Proc Natl Acad Sci U S A 79, 530–533. Blomberg, B., Rudin, C. M., and Storb, U. (1991). Identification and localization of an enhancer for the human lambda L chain Ig gene complex. J Immunol 147, 2354–2358. Blomberg, B., Traunecker, A., Eisen, H., and Tonegawa, S. (1981). Organisation of four mouse l light chain immunoglobulin genes. Proc Natl Acad Sci U S A 78, 3765–3769.
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the international ImMunoGeneTics database. Nucleic Acids Res 25, 206–211. Hagman, J., Rudin, C. M., Haasch, D., Chaplin, D., and Storb, U. (1990). A novel enhancer in the immunoglobulin lambda locus is duplicated and functionally independent of NF kappa B. Genes Dev 4, 978–992. Hess, M., Hilschmann, N., Rivat, L., Rivat, C., and Ropartz, C. (1971). Isotypes in human immunoglobulin lambda-chains. Nat New Biol 234, 58–61. Hieter, P. A., Hollis, G. F., Korsmeyer, S. J., Waldmann, T. A., and Leder, P. (1981). Clustered arrangement of immunoglobulin lambda constant region genes in man. Nature 294, 536–540. Ignatovich, O., Tomlinson, I. M., Jones, P. T., and Winter, G. (1997). The creation of diversity in the human immunoglobulin V(lambda) repertoire. J Mol Biol 268, 69–77. Ignatovich, O., Tomlinson, I. M., Popov, A. V., Bruggemann, M., and Winter, G. (1999). Dominance of intrinsic genetic factors in shaping the human immunoglobulin Vlambda repertoire. J Mol Biol 294, 457–465. Kabat, E. A., Wu, T. T., Reid-Miller, M., Perry, H. M., and Gottesman, K. S. (1987). Sequences of proteins of immunological interest, 4th ed. Washington, D.C.: Public Health Service. Kabat, E. A., Wu, T. T., Perry, H. M., Gottesman, K. S., and Foeller, C. (1991). Sequences of proteins of immunological interest. Washington, D.C.: Public Health Service. NIH Publication 91–3242. Kawasaki, K., Minoshima, S., Schooler, K., Kudoh, J., Asakaw, S., de Jong, P. J., and Shimizu, N. (1995). The organization of the human immunoglobulin lambda gene locus. Genome Res 5, 125–135. Kawasaki, K., Minoshima, S., Nakato, E., Shibuya, K., Shintani, A., Schmeits, J. L., Wang, J., and Shimizu, N. (1997) One-megabase sequence analysis of the human immunoglobulin lambda gene locus. Genome Res 7, 250–261. Kay, P. H., Moriuchi, J., Ma, P. J., and Saueracher, E. (1992). An unusual allelic form of the immunoglobulin lambda constant region genes in the Japanese. Immunogenetics 35, 341–343. Lefranc, M.-P. (1997). Unique database numbering system for immunogenetic analysis. Immunol Today 8, 509. Lefranc, M.-P. (1998). IMGT (ImMunoGeneTics) locus on focus. A new section of Experimental and Clinical Immunogenetics. Exp Clin Immunogenet 15, 1–7. Lefranc, M.-P. (1999). The IMGT unique numbering for immunoglobulins, T cell receptors and Ig-like domains. Immunologist 7, 132–136. Lefranc, M.-P. (2000a). Locus maps and genomic repertoire of the human immunoglobulin genes. Immunologist 8/3, 80–88. Lefranc, M.-P. (2000b). Nomenclature of the human immunoglobulin genes. Curr Protocols Immunol A.1P.1–A.1P.37. Lefranc, M.-P. (2000c). IMGT ImMunoGeneTics database. Int BIOforum 4, 98–100. Lefranc, M.-P. (2001a). IMGT, the international ImMunoGeneTics database. Nucleic Acids Res 29, 207–209. Lefranc, M.-P. (2001b). Nomenclature of the human immunoglobulin lambda (IGL) genes. Exp Clin Immunogenet 18, 242–254. Lefranc, M.-P. (2003a). IMGT, the international ImMunoGeneTics database®. Nucleic Acids Res 31, 307–310. Lefranc, M.-P. (2003b). IMGT, the international ImMunoGeneTics information system®. In Methods in molecular biology. Antibody engineering: Methods and protocols, Benny K. C. Lo, ed. Humana Press, Totowa, N.J., USA. Lefranc, M.-P. (2003c). IMGT® databases, web resources and tools for immunoglobulin and T cell receptor sequence analysis, http://imgt.cines.fr. Leukemia. 17, 260–266. Lefranc, M.-P., and Lefranc, G. (2001a). The immunoglobulin FactsBook (London: Academic Press), pp. 1–458. ISBN: 012441351X. Lefranc, M.-P., and Lefranc, G. (2001b). The T cell receptor FactsBook. London: Academic Press), pp. 1–398. ISBN: 012441351X.
4. Immunoglobulin Lambda (IGL) Genes of Human and Mouse Lefranc, M.-P., Pallarès, N., and Frippiat, J.-P. (1999b). Allelic polymorphisms and RFLP in the human immunoglobulin lambda light chain locus. Hum Genet 104, 361–369. Lefranc, M.-P., Pommié, C., Ruiz, M., Giudicelli, V., Foulquier, E., Truong, L., Contet, V., and Lefranc, G. (2003). IMGT unique numbering for immunoglobulin and T cell receptor variable domains and Ig superfamily V-like domains. Dev Comp Immunol 27, 55–77. Lefranc, M.-P., Giudicelli, V., Busin, C., Bodmer, J., Muller, W., Bontrop, R., Lemaitre, M., Malik, A., and Chaume, D. (1998). IMGT, the international ImMunoGeneTics database. Nucleic Acids Res 26, 297–303. Lefranc, M.-P., Giudicelli, V., Ginestoux, C., Bodmer, J., Müller, W., Bontrop, R., Lemaitre, M., Malik, A., Barbié, V., and Chaume, D. (1999a). IMGT, the international ImMunoGeneTics database. Nucleic Acids Res 27, 209–212. Mami, F., and Kindt, T. J. (1987). C lambda 2 and C lambda 4 immunoglobulin light chain genes in a wild-derived inbred mouse strain. J Immunol 138, 3980–3985. Miller, J., Bothwell, A., and Storb, U. (1981). Physical linkage of the constant region genes for immunoglobulin light chain lI and lII. Proc Natl Acad Sci U S A. 78, 3829–3833. Miller, J., Selsing, E., and Storb, U. (1982). Structural alterations in J regions of mouse immunoglobulin lambda genes are associated with differential gene expression. Nature 295, 428–430. Mural, R. J., Adams, M. D., Myers, E. W., Smith, H. O., Gabor Miklos, G. L., Wides, R., et al. (2002). A comparison of whole-genome shotgun derived mouse chromosome 16 and the human genome. Science 296, 1661–1671. Niewold, T. A., Murphy, C. L., Weiss, D. T. and Solomon, A. (1996). Characterization of a light chain product of the human JC lambda 7 gene complex. J Immunol 157, 4474–4477. Pallarès, N., Frippiat, J.-P., Giudicelli, V., and Lefranc, M.-P. (1998). The human immunoglobulin lambda variable (IGLV) genes and joining (IGLJ) segments. Exp Clin Immunogenet 15, 8–18. Pallarès, N., Lefebvre, S., Contet, V., Matsuda, F., and Lefranc, M.-P. (1999). The human immunoglobulin heavy variable (IGHV) genes. Exp Clin Immunogenet 16, 36–60. Ponstingl, H., Hess, M., and Hilschmann, N. (1968). Complete aminco acid sequence of Bence Jones protein Kern. A new subgroup of the immunoglobulin L-chains of lambda-type. Hoppe Seylers Z Physiol Chem 349, 867–871. Poul, M.-A., Zhang, X. M., Ducret, F., and Lefranc, M.-P. (1991). The IGLJ6 joining segment as a STS in the human immunoglobulin lambda light chain constant region gene locus (located at 22q11). Nucleic Acids Res 19, 4785. Reidl, L. S., Kinoshita, C. M., and Steiner, L. A. (1992). Wild mice express an Ig V lambda gene that differs from any V lambda in BALB/c but resembles a human V lambda subgroup. J Immunol 149, 471–480. Ruiz, M., Pallarès, N., Contet, V., Barbié, V., and Lefranc, M.-P. (1999). The human immunoglobulin heavy diversity (IGHD) and joining (IGHJ) segments. Exp Clin Immunogenet 16, 173–184. Ruiz, M., Giudicelli, V., Ginestoux, C., Stoehr, P., Robinson, J., Bödmer, J., Marsh, S., Bontrop, R., Lemaître, M., Lefranc, G., Chaume, D., and Lefranc, M.-P. (2000). IMGT, the international ImMunoGeneTics database. Nucleic Acids Res 28, 219–221. Sanchez, P., Marche, P. N., Rueff-Juy, D., and Cazenave, P. A. (1990). Mouse V lambda x gene sequence generates no junctional diversity and is conserved in mammalian species. J Immunol 144, 2816–2820.
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Satow, Y., Cohen, G. H., Padlan, E. A. and Davies, D. R. (1986). Phosphocholine binding immunoglobulin Fab McPC603. An X-ray diffraction study at 2.7 A. J Mol Biol 190, 593–604. Scaviner, D., Barbié, V., Ruiz, M., and Lefranc, M.-P. (1999). Protein displays of the human immunoglobulin heavy, kappa and lambda variable and joining regions. Exp Clin Immunogenet 16, 234–240. Selsing, E., Miller, J., Wilson, R., and Storb, U. (1982). Evolution of mouse immunoglobulin lambda genes. Proc Natl Acad Sci U S A 79, 4681–4685. Stiernholm, N. B., Verkoczy, L. K., and Berinstein, N. L. (1995). Rearrangement and expression of the human psi C lambda 6 gene segment results in a surface Ig receptor with a truncated light chain constant region. J Immunol 154, 4583–4591. Storb, U., Haasch, D., Arp, B., Sanchez, P., Cazenave, P. A., and Miller, J. (1989). Physical linkage of the mouse l genes by pulse field gel electrophoresis suggests that the rearrangement process favours proximate target sequences. Mol Cell Biol 9, 711–718. Sun, T., Clark, M. R., and Storb, U. (2002). A point mutation in the constant region of Ig lambda 1 prevents normal B Cell development due to defective BCR signalling. Immunity 16, 245–255. Taub, R. A., Hollis, G. F., Hieter, P. A., Korsmeyer, S. J., Waldmann, T. A., and Leder, P. (1983). Variable amplification of immunoglobulin lambda light-chain genes in human populations. Nature 304, 172– 174. Tonegawa, S., Brack, C., Hozumi, N., and Pirrotta, V. (1978a). Organization of immunoglobulin genes. Cold Spring Harb Symp Quant Biol 42, 921–931. Tonegawa, S., Maxam, A. M., Tizard, R., Bernard, O., and Gilbert, W. (1978b). Sequence of a mouse germ-line gene for a variable region of an immunoglobulin light chain. Proc Natl Acad Sci U S A 75, 1485–1489. Udey, J. A., and Blomberg, B. B. (1987). Human lambda light chain locus: organization and DNA sequences of three genomic J regions. Immunogenetics 25, 63–70. Udey, J. A., and Blomberg, B. B. (1988). Intergenic exchange maintains identity between two human lambda light chain immunoglobulin gene intron sequences. Nucleic Acids Res 16, 2959–2969. van der Burg, M., Barendregt, B. H., van Gastel-Mol, E. J., Tumkaya, T., Langerak, A. W., and van Dongen, J. J. (2002). Unraveling of the polymorphic C lambda 2-C lambda 3 amplification and the Ke+Ozpolymorphism in the human Ig lambda locus. J Immunol 169, 271– 276. Vasicek, T. J., and Leder, P. (1990). Structure and expression of the human immunoglobulin lambda genes. J Exp Med 172, 609–620. Wain, H. M., Bruford, E. A., Lovering, R. C., Lush, M. J., Wright, M. W., and Povey, S. (2002). Guidelines for human gene momenclature. Genomics 79, 464–470. Walker, M. R., Solomon, A., Weiss, D. T., Deutsch, H. F., and Jefferis, R. (1988). Immunogenic and antigenic epitopes of Ig. XXV. Monoclonal antibodies that differentiate the Mcg+/Mcg- and Oz+/Oz-C region isotypes of human lambda L chains. J Immunol 140, 1600– 1604. Weiss, S., and Wu, G. E. (1987). Somatic point mutations in unrearranged immunoglobulin gene segments encoding the variable region of lambda light chains. EMBO J 6, 927–932. Williams, S. C., Frippiat, J.-P., Tomlinson, I. M., Ignatovich, O., Lefranc, M.-P., and Winter, G. (1996). Sequence and evolution of the human germline V lambda repertoire. J Mol Biol 264, 220–232.
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5 The Mechanism of V(D)J Recombination JOANN SEKIGUCHI
FREDERICK W. ALT
MARJORIE OETTINGER
Department of Internal Medicine, Divison of Molecular Medicine and Genetics, University of Michigan Medical School, Ann Arbor, Michigan, USA
Howard Hughes Medical Institute, The Children’s Hospital, The Center for Blood Research, Boston, Massachusetts, USA
Department of Molecular Biology, Massachusetts General Hospital, Boston, Massachusetts, USA
A quarter-century ago, the revolutionary discovery was made that DNA in lymphoid cells encoding the antigen receptors is altered from that of other somatic tissues and germline cells (Hozumi and Tonegawa, 1976). This DNA rearrangement is at the heart of the ability of B and T cells to generate a highly diverse array of antigen receptor molecules, thus allowing a virtually unlimited set of antigen molecules to be recognized with a high degree of specificity. A series of site-specific recombination events, collectively termed V(D)J recombination, serves to assemble antigen receptor genes from arrays of gene segments (for additional reviews on this topic, see Bassing et al., 2002; Fugmann et al., 2000; Gellert, 2002; Lewis, 1994). In addition to the combinatorial diversity that results from this assembly process, the V(D)J reaction itself contributes to the diversity by joining the gene segments imprecisely. This junctional diversity is achieved precisely at the complementaritydetermining region 3 (CDR3) of the antigen receptor, a major determinant of the specific interaction between antigen receptor and antigen. In broad terms, rearrangement is initiated by the lymphoid-specific V(D)J recombinase composed of the recombination activating gene 1 and 2 (RAG1 and RAG2) proteins (Oettinger et al., 1990; Schatz et al., 1989). Together, RAG1 and RAG2 bind to the recombination signal sequences (RSS) that flank each gene segment and introduce a double-strand break (DSB) between the RSS and the flanking coding DNA (Figure 5.1) (reviewed by Gellert, 2002). The DNA ends generated by cleavage are asymmetrical, with the coding end covalently sealed into a hairpin and the signal end present as a 5¢ phosphorylated blunt DNA end (Roth et al., 1992; Roth et al., 1993; Schlissel et al., 1993). This first stage of V(D)J rearrangement is the point at which
Molecular Biology of B Cells
most if not all of the regulation of the recombination reaction is imposed. The broken DNA generated by RAG cleavage can be resolved through several different pathways (Figure 5.1). The first is standard V(D)J joining, in which the hairpins are opened and joined imprecisely to each other to form a coding joint (CJ) and the two signal ends ligated heptamer to heptamer, to generate a signal joint (SJ). Joining is mediated by components of the nonhomologous end joining (NHEJ) DNA repair pathway (reviewed by Bassing et al., 2002). Alternatively, two nonstandard products of V(D)J recombination can be generated by the joining of signal ends to coding ends (Lewis et al., 1988). Rejoining of a signal to its original coding flank yields an open and shut joint, whereas joining of a signal to its reaction partner’s coding flank generates a hybrid joint (HJ). Finally, RAG1/2 bound to the signal ends can catalyze the transpositional attack of the signal ends on unrelated target DNA (Agrawal et al., 1998; Hiom et al., 1998). The mechanisms and factors involved in each of these reactions are considered separately later in this chapter. The past several years have seen an explosion in the understanding of the cleavage mechanism and the functional properties of the RAG proteins, in large part due to the ability to carry out V(D)J cleavage with purified proteins. Great strides in understanding the repair stage of the reaction have also been made. Six key factors required for repairing the broken molecules to generate signal and coding joints have been identified and some of their biochemical properties defined. This chapter considers in detail what is known about the V(D)J recombination reaction, including the biochemistry of the cleavage reactions, the activities of the RAG proteins, and the components of the NHEJ DNA repair pathway.
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FIGURE 5.1 RAG-mediated DNA rearrangements. (A) RAG1/2 initiate V(D)J recombination by nicking the RSS sequences adjacent to the coding segments, leaving a 3¢-OH on the coding flanks. RAG1/2 activate the hydroxyl group to attack the opposite DNA strand to form a hairpin coding end and blunt, 5¢ phosphorylated signal end. (B) Standard inversional V(D)J recombination is catalyzed by the NHEJ proteins to form modified coding joints (CJ) and precise signal joints (SJ). (C) Nonstandard V(D)J recombination is catalyzed by the NHEJ proteins to form open-and-shut and hybrid joints (HJ). Nucleotide loss or addition can be observed within the junctions. (D) RAG-mediated transeseterification reactions occur in the absence of the NHEJ proteins. The reactions catalyzed by the RAGs include transposition and formation of incomplete open-and-shut joints and HJs. RSSs, triangles; coding segments, rectangles.
ANTIGEN RECEPTOR GENE ASSEMBLY Immunoglobulin (Ig) genes and T cell receptor (TCR) genes exist in the germline as linear arrays of clustered gene segments. Seven antigen receptor loci exist: TCR a, b, g, and d, and IgH, k, and l. All loci contain V (variable) and J (joining) segments, and three (TCR b and d and IgH) also contain D (diversity) segments between the V and J clusters. The heterodimeric immune receptors are always composed of one polypeptide derived from a locus containing V, D, and J elements and one from a locus with just V and J elements. At each locus, the variable region exon consisting of VJ or VDJ elements is then fused to a C (constant) region through RNA splicing (Figure 5.2). Each recombinationally active gene segment is flanked by an RSS that consists of a dyad symmetric heptamer, an A/T rich nonamer, and a spacer region of conserved length (12 or 23 bp +/-1) but generally nonconserved sequence
(Figure 5.3). The consensus sequences for the heptamer (CACAGTG) and nonamer (ACAAAAACC) are also optimal for rearrangement, but considerable deviation from the consensus is tolerated, with few segments flanked by an RSS that fits the consensus sequence precisely (Lewis, 1994). The length of the spacer plays a crucial role in the reaction: efficient V(D)J recombination requires a pair of signals, one with a 12- and the other with a 23-bp spacer (Tonegawa, 1983). This relatively simple restriction, the 12/23 rule, has important biological outcomes. First, because all segments of a particular type (e.g., Vk segments) are flanked by one type of signal, and all the segments to which they could be joined (Jk) are flanked by the opposite type, this arrangement ensures that joining is restricted to events that could be biologically productive. Second, because a signal pair is required to induce the catalytic activity of the RAG proteins, the chance of introducing a double-strand break in the absence of a partner DNA to join to is greatly reduced. This restriction
5. The Mechanism of V(D)J Recombination
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FIGURE 5.3 The 12/23 rule. Two V region coding segments (Vk and Jk) are depicted as rectangles and are flanked by a 12-bp and a 23-bp spacer RSS. The consensus sequences of the conserved heptamer and nonamer are shown. Coupled cleavage of the 12/23 RSSs by RAG1/RAG2 occurs at the 5¢ end of the RSS heptamers (arrows). The coding ends are joined after further modification, and the heptamers at signal ends are joined precisely.
FIGURE 5.2 Immunoglobulin heavy (IgH) chain gene assembly and expression. The events involved in the assembly of the IgH chain into a complete immunoglobulin molecule are depicted. H chain gene assembly begins with the rearrangement of a DH segment to a JH segment, followed by rearrangement of a VH segment to the pre-assembled D-JH segment. Transcripts originating from the VH promoter that encode the V-D-JH and Cm gene segments are differentially spliced and give rise to both the membrane and secreted forms of IgM. An IgH chain protein combines with an IgL chain protein to form a typical monomeric subunit of an Ig molecule.
therefore decreases the potential for RAG-induced genomic instability. The RSS sequences are all that is required to render a piece of DNA a substrate for V(D)J recombination. As shown in Figure 5.4, the orientation of the signal sequences with respect to each other determines the outcome of the reaction. Rearrangement can result in retention of the coding joint in the chromosome and deletion of a circular molecule containing the signal joint. Alternatively, recombination can lead to inversion of the DNA between the RSSs with retention of both the SJ and CJ in the chromosome. Both of these arrangements are found in vivo (Fujimoto and Yamagishi, 1987; Okazaki et al., 1987; Zachau, 1993). As an experi-
FIGURE 5.4 Deletional and inversional V(D)J recombination. (A) The intervening sequences between the recombining coding segments can be deleted when the 12/23 RSSs are oriented, as depicted, to form a CJ on the chromosome and a SJ on an extrachromosomal circle. (B) RSSs oriented in the same direction along the chromosome, as depicted, lead to inversional V(D)J recombination in which both CJ and SJ remain on the chromosome.
mental convenience, plasmid-based synthetic recombination substrates can be generated that retain either the signal or the coding joint on the plasmid, allowing for recovery of the joined molecule and a detailed analysis of the junction (Hesse et al., 1987; Lewis et al., 1985). Such substrates also permit a detailed analysis of the sequence requirements for a functional RSS and flanking coding DNA. An outline of a standard assay for V(D)J recombination in tissue culture cells is shown in Figure 5.5.
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FIGURE 5.5 In vitro V(D)J recombination assay. Extrachromosomal plasmid recombination substrates contain two RSSs (triangles) that can undergo site-specific recombination (Hesse et al., 1987). The substrates harbor a prokaryotic promoter and a drug resistance or color marker gene (i.e., chloramphenicol resistance [CAT] or LacZ genes). These elements are separated by a prokaryotic transcription terminator flanked by the RSS sequences. The plasmid recombination substrates are introduced into cells along with constructs expressing RAG1 and RAG2. Recombination between the RSSs deletes the transcription terminator sequence, thus allowing transcription through the selectable marker genes. The efficiency of recombination can be measured via transformation into bacteria and selection on media containing the appropriate antibiotics. Coding and RSS joining can be measured based on the orientation of the RSSs within the substrate.
MUTATIONAL ANALYSIS OF RECOMBINATION SIGNAL SEQUENCES The RSSs are remarkably well conserved among vertebrates, with the same motifs found from sharks to humans. Using the assay outlined in Figure 5.5, the RSS requirements for V(D)J recombination in vitro have been extensively explored (Akamatsu et al., 1994; Akira et al., 1987; Connor et al., 1995; Hesse et al., 1987; Hesse et al., 1989; Nadel et al., 1998; Ramsden and Wu, 1991). The first three
nucleotides of the heptamer sequence are the most crucial, with considerable variation tolerated at the remaining four positions. The CAC sequence also shows the greatest degree of conservation among naturally occurring RSSs. Within the nonamer, alterations in positions 5, 6, and 7 cause the greatest decrease in recombination, but generally the sequence of the nonamer is less critical than the heptamer. In fact, recombination between a signal pair in which one signal contains only a heptamer, while the other has the consensus sequence, is still observable (down 20- to 50-fold) (Hesse et al., 1989). Nucleotide substitutions have similar effects when incorporated into a 12-RSS or a 23-RSS, suggesting that the recognition of these two signals by the recombinase is similar. Early thoughts that the dyad symmetry of the RSS would allow for pairing between signals as part of the joining process have not proved to be correct, as such homology can be disrupted without affecting recombination of a signal pair (Hesse et al., 1989). Although the spacer sequence is not well-conserved, there is mounting evidence that its sequence can considerably influence RSS usage (for example, see Jung et al., 2003; Nadel et al., 1998). Although the initial description of the sequence requirements for an RSS was determined in cell culture experiments, these same requirements are seen in vitro. The actual RSS sequence used at an antigen receptor locus is rarely the consensus sequence. This variation may influence segment usage, with segments flanked by RSSs closer to the consensus favored over others. For example, the RSSs at Vk are generally closer to consensus than those at Vl, perhaps providing one level of explanation for the favored usage of Vk segments. This preference (up to 100-fold) can be reproduced with kappa and lambda RSSs in synthetic substrates (Feeney et al., 2000; Ramsden and Wu, 1991). However, as discussed later, restrictions on RSS usage go beyond this 12/23 regulation.
“BEYOND 12/23” RESTRICTION OF V(D)J REARRANGEMENTS As indicated above, the organization of 12- and 23-RSSs, which flank V, D, and J segments within the Ig and TCR loci, facilitates proper rearrangement. However, additional restrictions on RSS usage must exist. For example, the simple 12/23 rule cannot fully account for the rearrangement patterns observed at the TCRb locus. At this locus, the Vs are flanked with 23-RSSs and Js with 12-RSSs, whereas the D has a 5¢ 12-RSS and a 3¢ 23-RSS. Direct Vb to Jb joining is rarely observed in vivo, although it would be in accordance with the 12/23 rule. The vast majority of TCRb variable region genes are generated via D to Jb and then Vb to DJb rearrangements (Born et al., 1985; Ferrier et al., 1990; Sleckman et al., 2000).
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These observations have been explained by further studies showing that some pairs of functional RSSs are restricted from recombining with each other. This restriction “beyond the 12/23 rule” (B12/23) was first suggested by placing transgenic recombination substrates in the TCRb locus, and more definitively shown by gene-targeted mutation (Bassing et al., 2000; Sleckman et al., 2000; Wu et al., 2003). Targeted replacement of the 5¢ Db1 12-RSS with the Jb 12-RSS prevented Vb rearrangement. However, replacement of the Jb 12-RSS with the 5¢ Db1 12-RSS in a simplified locus (lacking the Db2Jb2 cluster) in which Db1 had been deleted allowed direct Vb–Jb joining but only to the Jb that contained the replaced RSS (Bassing et al., 2000). Thus, the 5¢ Db1 12-RSS, but not the Jb 12-RSS, is an acceptable target for the rearrangement of the Vb segments. The notion that particular RSSs restrict specific rearrangements in a B12/23 manner was further illustrated by studies in which the Vb14 23-RSS was replaced with the 3¢ Db1 23-RSS (Wu et al., 2003). The Vb14/3¢DbRSS replacement dramatically increased the usage of Vb14, predominantly due to an increase in the relative level of primary Vb14 to DJb rearrangement. However, the 3¢Db1 23-RSS replacement also broke the TCRb locus B12/23 restriction and allowed direct Vb14 to Jb1 rearrangement. Thus, the Vb14 23-RSSs contributes to B12/23 restriction by preventing Vb14 to Jb1 rearrangement; furthermore, the high recombination potential of the 3¢ Db1 23-RSS may have evolved to ensure that complete Db to Jb rearrangements occur prior to Vb rearrangement. The B12/23 restrictions observed in vivo at the TCRb locus were recapitulated in nonlymphoid cells using extrachromosomal V(D)J recombination substrates containing various combinations of Vb 23-RSSs, 5¢ Db 12-RSSs, 3¢ Db 23-RSSs, and Jb 12-RSSs, as well as with in vitro cleavage assays using purified RAG proteins (Jung et al., 2003; Tillman et al., 2003). All Vb 23-RSSs analyzed in these studies preferred the 5¢ Db1 12-RSS over the Jb1 12 RSSs. Thus, consistent utilization of the Db gene segment is largely ensured by the constraints imposed on the formation of functional cleavage complexes (discussed later) containing an RSS pair, the RAG proteins, and HMG1.
INFLUENCE OF CODING FLANKS In addition to the RSS, the first two or three nucleotides of the coding flank immediately abutting the heptamer of the RSS can have considerable effects on the efficiency of the recombination reaction. Although most sequences are essentially neutral, certain nucleotide combinations can be favorable or unfavorable to the reaction. A run of T’s (5¢ to 3¢ toward the heptamer) substantially reduces recombination (Boubnov et al., 1995; Ezekiel et al., 1995; Ramsden and Wu, 1991). Some dinucleotides such as 5¢ TG 3¢ favor
recombination and are termed “good flanks”), while others such as 5¢ AC 3¢ and 5¢ GG 3¢ are unfavorable (“bad flanks”) (Sadofsky et al., 1995). The effect of the coding flank sequence appears to be primarily at the cleavage phase of the reaction (Cuomo et al., 1996; Ramsden et al., 1996), although there may be some effect on end-processing and the later joining stage of the reaction.
THE BIOCHEMISTRY OF V(D)J CLEAVAGE V(D)J cleavage requires only RAG1 and RAG2, a divalent metal ion (Mn2+ or Mg2+), and a DNA substrate containing the RSS (McBlane et al., 1995). In addition, the nonspecific DNA bending protein HMG1 (or HMG2) can serve to augment the reaction, as discussed here. No external source of energy is needed (McBlane et al., 1995; van Gent et al., 1995). With these simple components, the cleavage reaction can be reproduced, including the RSS selectivity (Cuomo et al., 1996; Ramsden et al., 1996) and the requirement for a 12/23 signal pair (Eastman et al., 1996; Kim and Oettinger, 1998; van Gent et al., 1997; van Gent et al., 1996). In general, the reactions have been studied using truncated “core” portions of the RAG proteins (mouse RAG1 amino acids 384 to 1,008 of 1,040 and RAG2 amino acids 1 to 382 of 527). These endonucleolytically active core portions have been more readily purifiable than their fulllength counterparts (McBlane et al., 1995; Sawchuk et al., 1997). When expressed in tissue culture cells, the core proteins permit V(D)J recombination to occur (Cuomo and Oettinger, 1994; Kirch et al., 1996; Sadofsky et al., 1994; Sadofsky et al., 1993; Silver et al., 1993) though some differences from the full-length proteins have been seen. All enzymatic activities of the RAG proteins require the cooperation of RAG1 and RAG2; individual activities of either protein have not been described. Cleavage itself occurs in two separable steps (McBlane et al., 1995). In the first step, a nick is introduced on the top strand adjacent to the recombination signal, leaving a 3¢ hydroxyl on the coding side and a 5¢ phosphoryl group on the signal end (see Figure 5.1). In the second step, the 3¢ hydroxyl from the top strand attacks the phosphodiester bond at the same position on the opposing strand, resulting in the formation of the covalently sealed hairpinned coding end, and the blunt signal end. The energy required for the formation of the new bond is derived from the breakage of the old one. Stereochemical studies have shown that this conservative reaction occurs with the inversion of chirality, indicating that a covalent bond between the RAG proteins and DNA is not formed (van Gent et al., 1996). This distinguishes the RAG cleavage reaction from that of a number of other site-specific recombinases, such as Cre, Flp, and Lambda-Int, which rely on a covalent intermediate. Instead, the direct transesterifi-
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cation mechanism used by RAG1/2 is similar to that of bacterial transposases and HIV integrase (Engelman et al., 1991; Mizuuchi and Adzuma, 1991; van Gent et al., 1996). The in vivo recombination reaction is largely a coupled process. That is, two signal sequences are required not just for a complete recombination event, but also for the initiating cleavage events. In addition, a 12/23 signal pair is preferred by approximately 50-fold over a 12–12 pair (Steen et al., 1996). The in vitro reaction can reproduce both this requirement for a signal pair and approximately the same extent of 12/23 preference as observed in vivo. Both these properties are intrinsic features of the RAG proteins, and the full extent of 12/23 preference can be observed upon the addition of HMG1 and nonspecific competitor DNA (Eastman et al., 1996; Kim and Oettinger, 1998; van Gent et al., 1997; van Gent et al., 1996). However, by altering the reaction conditions (substituting Mn2+ for Mg2+) cleavage can be uncoupled, thus allowing a break at a single RSS to be made. Studying the reaction under these conditions allows an examination of requirements for DNA recognition and cleavage as distinct from the requirements for synaptic complex assembly.
RSS Requirements for Cleavage A simple oligonucleotide containing an RSS can serve as a substrate for V(D)J cleavage (McBlane et al., 1995). By modifying this substrate, the precise DNA sequence requirements for binding and cleavage have been determined and compared with the rules for V(D)J recombination derived from in vivo experiments. Both a 12- and a 23-RSS can be recognized and individually cleaved by the RAG proteins. However, a 12-RSS is cleaved more efficiently than a 23-RSS. As with V(D)J recombination in vivo, a consensus RSS serves at the optimal for cleavage (Cuomo et al., 1996; Ramsden et al., 1996). The first three nucleotides of the heptamer are again the most sensitive to mutation. However, substitutions at these nucleotides primarily affect hairpin formation, influencing the initial nicking step to a much lesser extent. In the complete absence of a heptamer, the nonamer alone can still direct some nicking (but no hairpinning). This somewhat imprecise and very inefficient nicking occurs where the boundary of the heptamer would have been in a 12- or 23-RSS (that is, 19 or 30 nucleotides from the end of the nonamer), as if the proteins reach out a defined distance from the bound nonamer. Substrates containing a heptamer alone also work, with cleavage reduced by only a few-fold. The length of the spacer sequence is also important (Cuomo et al., 1996; Ramsden et al., 1996). The length difference between the 12- and 23-RSS is almost precisely one helical turn. This disposition suggests that having the two recognition elements, the heptamer and nonamer, in the same rotational phase is important for binding and cleavage.
Alteration of the length of the spacer supports this view. Adding an extra one-half helical turn (18- or 29-bp) substantially inhibits cleavage, below that seen with an isolated heptamer, whereas adding one full turn (33- or 34-bp) permits a substantial level of cleavage. Taken together these results suggest that each element can function on its own and act together synergistically with the proper spacing, but they conflict when the spacing is wrong. Alteration of the structure of the substrate DNA suggests that DNA unpairing and structural distortion might play a role in V(D)J cleavage (Cuomo et al., 1996; Ramsden et al., 1996). Unpairing of the first few nucleotides of coding sequence, when those bases are unfavorable for cleavage, can significantly enhance hairpin formation, suggesting that unpairing of the DNA sequence may be an important part of the cleavage reaction. More dramatically, with the coding flank remaining as duplex DNA, the RSS of a nucleotide substrate can be made single-stranded and still serve to direct site-specific binding and hairpin formation. In this reaction, only the heptamer appears important. This reaction is very efficient, again suggesting that DNA unwinding, perhaps due to RAG binding, may play an important role in the cleavage reaction. The specific sequence of the heptamer may have evolved not only to serve as a specific binding site for the RAG proteins, but also to be readily unpaired. The CACA/GTGT sequence of the heptamer is considerably distorted both in free solution and in crystals (Cheung et al., 1984; Patel et al., 1987; Timsit et al., 1991). Good flanks extend this unusual structure (purine/pyrimidine alternation) (Sadofsky et al., 1995), perhaps explaining their effect.
RAG1/2-RSS BINDING RAG1 and RAG2 together are required for highly specific binding to DNA; RAG1/2 prefers an RSS sequence over nonspecific DNA by ~50-fold in competition experiments (Hiom and Gellert, 1997). Two distinct species of RAG1/2 bound to a single RSS can be resolved by gel retardation experiments (Mundy et al., 2002; Swanson, 2002). These two complexes differ in protein content, but not in other properties. The RAG1/2-DNA complex, once formed, is highly stable. It remains bound (and active for cleavage) up to 8 hours after assembly, and resists very high levels of competitor DNA (Akamatsu and Oettinger, 1998; Hiom and Gellert, 1997; Li et al., 1997; Mundy et al., 2002). Both the heptamer and nonamer (with proper spacing) are required for maximal binding (Akamatsu and Oettinger, 1998; Hiom and Gellert, 1997; Nagawa et al., 1998; Swanson and Desiderio, 1998). Footprinting of the RAG1/2 complex on a single RSS reveals contacts in both the heptamer and nonamer (Akamatsu and Oettinger, 1998; Nagawa et al., 1998; Swanson and Desiderio, 1998), and photocrosslinking experiments indicate that the heptamer is
5. The Mechanism of V(D)J Recombination
touched by both RAG proteins (Eastman et al., 1999; Mo et al., 1999; Swanson and Desiderio, 1998). RAG1 binds to DNA on its own, but with considerably lower affinity and specificity, and appears only to contact the nonamer (Akamatsu and Oettinger, 1998; Mo et al., 1999; Swanson and Desiderio, 1998). Thus, in the presence of RAG2, the DNA contacts of RAG1 appear to change. Moreover, the enhancements of chemical cleavage observed with dimethyl sulfate (DMS) protection and phenanthroline-copper (OP-Cu) DNA footprinting support the idea that some DNA unwinding occurs near the heptamer–coding DNA border and that this is a result of the binding of RAG1 together with RAG2 (Akamatsu and Oettinger, 1998; Mo et al., 1999; Swanson and Desiderio, 1998). Although RAG1/2 together can bind to a single RSS, it is the synaptic “paired complex” (PC) containing a 12/23 signal pair that is competent to generate double-strand breaks under restrictive coupled-cleavage (Mg2+) conditions (Hiom and Gellert, 1998). The synaptic PC is a very stable species, resistant to high levels of nonspecific competitor DNA (Hiom and Gellert, 1998). Footprint analysis of this complex shows greatly enhanced protection of the heptamer sequence over that seen in a single-site complex (Nagawa et al., 2002). PC contains a dimer of RAG2 and either a dimer or tetramer of RAG1 (Landree et al., 2001; Mundy et al., 2002; Swanson, 2002); the ambiguity in RAG1 content arises from similar experiments that yield differing results (Mundy et al., 2002; Swanson, 2002), though additional experiments support the conclusion that RAG1 binds as a tetramer (Godderz et al., 2003). At this step of synaptic complex assembly the 12/23 rule is at least largely enforced with a 12/23 pair greatly preferred over a 12/12 or 23/23 pair (Hiom and Gellert, 1998; Mundy et al., 2002). It has also been suggested that the cleavage step itself may also contribute to 12/23 restriction (West and Lieber, 1998; Yu and Lieber, 2000). Although the 12/23 rule is enforced at the binding step, it is only hairpin formation that is subject to this control, because nicking can occur without synapsis. Although it was generally expected that each RSS would serve as a half-site, binding half the content of RAG1/2 that would later be present in the PC, this turns out not to be the case (Jones and Gellert, 2002). The RAG protein content of the slower mobility SC2 complex does not differ from PC (Mundy et al., 2002). Instead, SC2 and PC differ only by the addition of the second RSS containing DNA (Mundy et al., 2002). In other words, the RAG proteins bind to one signal first (with a strong preference for the 12RSS, as shown in competition experiments), then recognize and bind the second signal (Jones and Gellert, 2002). Because SC2 is not competent to form hairpins under restrictive Mg2+ conditions, even though all the necessary RAG1/2 proteins are present (Mundy et al., 2002), it is highly likely that binding to the second signal induces some conformational change in
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the RAG1/2 complex to render it catalytically active. Such a process would help to regulate cleavage in vivo, thus reducing the chance of the inappropriate introduction of double-strand breaks into the genome.
RAG1/2 POST-CLEAVAGE COMPLEX After cleavage, the RAG1/2 complex remains bound to the DNA. Stable post-cleavage complexes of RAG1/2 bound to a pair of signal ends (Agrawal and Schatz, 1997; Jones and Gellert, 2001) or to all four DNA ends (two hairpin and two coding ends) (Hiom and Gellert, 1998) have been observed in vitro. The finding that cleaved products can be resolved to form SJ, CJ, and hybrid or open and shut joints (a joining of coding end to signal end) suggests that all four ends are held together in a complex in vivo as well (Lewis et al., 1988). Several lines of evidence suggest that the bound RAG proteins participate in the later resolution stages of the reaction. First, deproteinization of the signal ends is required prior to joining by NHEJ factors in vitro (Leu et al., 1997; Ramsden et al., 1997). Second, there are mutants of both RAG1 and RAG2 that cleave the RSS but fail to support complete V(D)J recombination, thus suggesting the RAGs are involved in joining (Huye et al., 2002; Qiu et al., 2001; Tsai et al., 2002; Yarnell Schultz et al., 2001). Third, bluntend joining in yeast is normally imprecise, but following V(D)J cleavage, blunt signals are rejoined precisely, suggesting that the RAGs play a role in this process (Clatworthy et al., 2003). Fourth, the nonstandard resolution products of RAG cleavage observed in vitro indicate a role for the RAG proteins post-cleavage. RAG1/2 complexed with cleaved signal ends can bind to unrelated target DNA in the target capture step of transposition (discussed later), and the formation of a hybrid joint (in vitro but not necessarily in vivo) can be formed by a RAG-mediated attack of a signal end on a hairpin coding end (discussed later). It has been suggested that the RAG proteins bound to the cleaved ends may serve as a scaffold and may recruit the NHEJ factors to facilitate end-processing and joining (Huye et al., 2002; Tsai et al., 2002). In this regard, mutations that affect the ability of the RAGs to maintain the broken ends in stable postcleavage complexes may lead to misrepair of the DSBs, and thereby may have the potential to cause oncogenic chromosomal aberrations (Huye et al., 2002; Tsai et al., 2002).
A ROLE FOR HMG1 (OR HMG2) IN V(D)J RECOMBINATION Although RAG1 and RAG2 are the only lymphoidspecific proteins required for cleavage, the high mobility group protein 1 (HMG1) or HMG2 may be a generally
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important co-factor for RAG-mediated activities. HMG1 and -2 are ubiquitously expressed, abundant nuclear proteins that bind to DNA without sequence specificity and can bend linear DNA. HMG1 and -2 are also associated with chromatin, and appear to play important roles in the assembly of nucleoprotein complexes involved in DNA repair and transcription (reviewed by Thomas and Travers, 2001). The addition of HMG1 to the V(D)J cleavage reaction generally has little effect on the efficiency of cleavage of a 12RSS but substantially increases cleavage at a 23RSS (Sawchuk et al., 1997; van Gent et al., 1997). The nonspecific binding or bending activity of HMG1 suggests its role may be to facilitate the distortion of the DNA to allow for the same components of an active RAG complex to bind the two different signal sequences. Formation of the synaptic complex is greatly enhanced by the presence of HMG1 (or HMG2) (Hiom and Gellert, 1998; West and Lieber, 1998; Swanson, 2002), and the addition of HMG1 increases the preference for a 12/23 pair over a 12/12 pair (van Gent et al., 1997; Hiom and Gellert, 1998; Kim and Oettinger, 1998). HMG1 (or -2) is also required for RAG-mediated transposition, both for paired complex formation and for capturing target DNA (Agrawal et al., 1998; Hiom et al., 1998; Swanson, 2002) (discussed later). Finally, the addition of HMG1 augments V(D)J cleavage of RSS sequences assembled into nucleosomes, suggesting it may play an important role in facilitating RAG binding at endogenous loci (Kwon et al., 1998). However, an in vivo role for HMG1 or -2 during V(D)J recombination has not yet been established.
A CLOSER LOOK AT RAG1 AND RAG2 The RAG genes were originally identified based on their ability, when expressed in fibroblasts, to induce the V(D)J recombination of an artificial recombination substrate (Oettinger et al., 1990; Schatz et al., 1989). As such they are the only lymphoid-specific factors required to induce V(D)J recombination even in a nonlymphoid cell where this reaction would not normally occur, because the other required factors are generally expressed and can be recruited to complete the joining reaction. In the absence of RAG1 or RAG2, no V(D)J recombination can occur and thus mice with targeted disruptions of either gene lack mature B and T cells (Mombaerts et al., 1992; Shinkai et al., 1992). However, such mice do not exhibit any defects outside of the immune system, indicating that RAG function is limited to the lymphoid lineage. RAG1 and RAG2 share no sequence similarity. However, the genomic structure of the RAG genes is highly unusual. In all species examined, the two genes are adjacent and convergently transcribed. Although adjacent, their lack of
sequence similarity indicates that they did not arise via gene duplication. In most species (human, mouse, chicken, Xenopus) they also share the unusual feature of encoding the entire structural gene within a single exon. Two exceptions have been found: the zebra fish and trout RAG1 coding sequences contain introns (Hansen and Kaattari, 1995; Willett et al., 1997). This unusually compact structure of the RAG genomic locus led to the suggestion that the RAG genes might have evolved from (been co-opted from) a primordial transposon (Oettinger et al., 1990; Thompson, 1995), a presumption strengthened by biochemical demonstrations of RAG transposase activity (discussed later) (Agrawal et al., 1998; Hiom et al., 1998). Although the RAG genes do not show sequence similarity to each other, the RAG genes are highly conserved between species. Between pufferfish and human, there is 66% and 61% amino acid similarity for the RAG1 and RAG2 proteins respectively (Peixoto et al., 2000). Interestingly, whereas RAG1 is highly conserved across the entire structural gene, that conservation is biphasic, with the region between 411 and 1,036 even more highly conserved (75% amino acid identity) (Peixoto et al., 2000). This region roughly corresponds to the “core portion” of RAG1, the minimal region required for catalysis. A comparison of the two RAG proteins with known structures and structural motifs has led to the proposal that RAG2 has two distinct domains separated by a “hinge” region (Aravind and Koonin, 1999; Callebaut and Mornon, 1998), and studies with limited proteolysis confirm that two protease resistant domains exist (Arbuckle et al., 2001; Kim et al., 2003). The core portion of RAG2 (the domain required for catalysis) is proposed to fold into a structure resembling a six-bladed propeller where each blade contains a kelch motif (Aravind and Koonin, 1999; Callebaut and Mornon, 1998). Each kelch motif, originally identified in a Drosophila regulatory protein, would contain a four-stranded twisted antiparallel beta sheet. In other cases, such structures are involved in protein–protein interactions, suggesting that this domain may not only bind DNA but allow for interaction with RAG1 or an additional RAG2, a proposal consistent with studies of RAG1/RAG2 protein–protein interaction (Corneo et al., 2000; Gomez et al., 2000; Landree et al., 1999). The C-terminal region of RAG2 contains an acidic portion (amino acids 352 to 410, 42% acidic), a Cys-His rich PHD motif (aa 420 to 480), and a binding site for CDK2 (thr490) (Lee and Desiderio, 1999). Amino acids from 383 to the end of the protein (aa 527) are absent in the recombinationally active RAG2 core protein so that the study of the functions of these regions has been limited. Recent success in purifying the full-length protein has led to the observation that the presence of the C-terminal domain of RAG2 diminishes RAG1/RAG2 mediated transposition following V(D)J cleavage (Elkin et al., 2003; Tsai and Schatz, 2003). Thr490
5. The Mechanism of V(D)J Recombination
is involved in regulation and is required for the proper cellcycle control of RAG2 protein levels (Lee and Desiderio, 1999). Thr490 phosphorylation leads to translocation of RAG2 from the nucleus to the cytoplasm, where it is degraded by the ubiquitin–proteosome system during S phase (Mizuta et al., 2002). PHD motifs are found in a number of regulatory proteins, many of them thought to affect chromatin structure or bind to chromatin components. Such a role is of interest given the indications that the absence of the C-terminal domain of RAG2 leads to a decrease in assembly of particular antigen receptor loci (Akamatsu et al., 2003; Kirch et al., 1998; Liang et al., 2002). Identifiable sequence motifs within RAG1 are minimal, though it is notable for containing several putative zinc fingers. Limited proteolysis has defined three distinct domains, the N-terminal (and dispensable) domain and two in the active core (Arbuckle et al., 2001). The most Cterminal domain displays DNA binding activity on its own (Arbuckle et al., 2001), although additional sites within the core are required for formation of a functional RAG1/RAG2 DNA complex. The structure of the N-terminal part of RAG1 has been solved and contains one zinc finger of the RING family and two additional zinc finger motifs (Bellon et al., 1997; Freemont et al., 1991; Rodgers et al., 1996). Such RING motifs are often found in proteins that serve as ubiquitin ligases, so-called E3 proteins. Recently it has been shown that this domain of RAG1 does indeed have E3 ubiquitin ligase activity (Yurchenko et al., 2003). Although this domain is not required for RAG1 enzymatic activity, its deletion is associated with some alterations in V(D)J recombination, and the E3 activity implies a regulatory role in V(D)J recombination for this domain. A simple search for homologies or motifs that might identify the active site within RAG1 or RAG2 did not yield obvious candidates. However, the knowledge that the RAG proteins cleave DNA using the same chemistry as transposases suggested that the RAG active site might share similarities with these enzymes. Many transposases use a triad of Asp and Glu residues, often termed a DDE motif, to bind a divalent metal at the active site (Rice et al., 1996). Mutational analysis of acidic residues in RAG1 and RAG2 led to the identification of three residues in RAG1—D600, D708, and E962—that are absolutely required for V(D)J recombination in vivo and V(D)J cleavage in vitro, but not for DNA binding (Fugmann et al., 2000; Kim et al., 1999; Landree et al., 1999). A role in metal binding has been established for D600 and D708 (Kim et al., 1999; Landree et al., 1999). Mutations in either of these residues eliminates hydroxyl radical cleavage activity of RAG1 (Kim et al., 1999). In addition, as has been seen for other transposases (Sarnovsky et al., 1996), the substitution of Asp with Cys restores some cleavage in Mn++ but not Mg++ (Kim et al., 1999; Landree et al., 1999). These same experiments failed to show that E962 was directly involved in metal binding, and its func-
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tion remains unclear. RAG1 may not actually contain a classical DDE motif, as this third amino acid is at a greater distance from the first two than is typically seen and appears to be located in a separate domain of RAG1 (Arbuckle et al., 2001), whereas all three residues of the known DDE motifs are contained in a single domain. Despite exhaustive mutagenesis of RAG2, no acidic residues were seen to be required (Landree et al., 1999). Other RAG2 point mutations do disrupt catalysis (Qiu et al., 2001), suggesting that RAG2 helps to establish the full active site. A crystal structure would be most useful in understanding how the active site is formed. Naturally occurring mutations in the RAG proteins are responsible for some forms of human severe combined immunodeficiency (SCID) (Schwarz et al., 1996; Villa et al., 1998; Villa et al., 2001). Complete immunodeficiencies, where the patients lack both B and T cells arise when RAG activity is absent. Partial loss-of-function mutations can give rise to a partial SCID phenotype or the related immunodeficiency, Omenn syndrome, in which T cells are more severely affected than B cells (Villa et al., 1998; Villa et al., 2001). Several of the RAG2 SCID mutations cluster along one surface of the predicted kelch propeller, leading to the suggestion that this region is involved in interactions with RAG1 (Corneo et al., 2000). Additional mutations have helped to further define the regions in which these two proteins interact with each other and with DNA (Fugmann and Schatz, 2001; Gomez et al., 2000; Landree et al., 1999). As indicated here, the core domains of RAG1 and RAG2 are sufficient to mediate V(D)J recombination in vivo. However, some notable differences occur between the behavior of the full-length proteins and the core versions. First, the frequency of V(D)J recombination on exogenous or integrated substrates in fibroblast cells is lower with the core than full-length proteins (Cuomo and Oettinger, 1994; Kirch et al., 1996; Sadofsky et al., 1995; Sadofsky et al., 1994; Silver et al., 1993). Second, recombination by the core RAG proteins in fibroblast cells leads to a greater accumulation of signal ends than is observed with full-length proteins (Steen et al., 1999). Third, the absence of the amino terminus of RAG1 results in reduced D to J rearrangement, with differential effects observed on the assembly of endogenous T cell receptor and immunoglobulin genes (Noordzij et al., 2000; Roman et al., 1997; Santagata et al., 2000). Fourth, mice that express core RAG1 in the absence of wild type RAG1 exhibit reduced frequency of both D-toJH and VH-to-DJH chromosomal rearrangements in RAG1c/c mice, which most likely reflects a decrease in overall V(D)J recombination efficiency (Dudley et al., 2003). Fifth, in both pro-B cell lines and mice that express core RAG2, Dh-to-Jh joining is mildly lowered, whereas Vh-to-DJh joining is severely reduced (Akamatsu et al., 2003; Kirch et al., 1998; Liang et al., 2002). Thus, the noncore regions of
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RAG1 and RAG2 clearly play important roles in ensuring efficient V(D)J recombination in vivo.
Hybrid Joining Although normal V(D)J recombination results in the formation of signal and coding joints, nonstandard joining products, hybrid joints (HJ), and open-and-shut joints are also observed in vivo (Lewis, 1994; Lewis et al., 1988). Such products arise when a signal end is rejoined to a coding end; rejoining to the original coding flank yields an open-and-shut joint (detectable if base loss or addition occurs), whereas rejoining to the partner flank yields a hybrid joint. These products are detectable both in artificial recombination substrates, where they can account for up to 10% of the recombination products, and at lower frequency in the antigen receptor loci themselves (Lewis et al., 1988). A reaction that would lead to the generation of hybrid or open-and-shut joints can be carried out with purified proteins in vitro (Melek et al., 1998). In this reaction, the RAG proteins initially perform a standard coupled cleavage reaction, giving rise to hairpin coding ends and blunt signal ends. Following this cleavage, the RAG proteins then catalyze the attack of the free hydroxyl of the RSS end onto the coding hairpin at or near the tip, thereby joining the signal end to coding DNA on one strand (see Figure 5.1). Attack on the original coding DNA would yield an open and-shut joint, while an attack on the partner coding end would result in HJ formation. Such a reaction would give rise to a perfect or near-perfect HJ, with no further nucleotide loss or addition. These in vitro products are joined only on the strand where the transesterification occurred (Melek et al., 1998), and can be generated by the core and, to a lesser extent, the fulllength versions of RAG1 and RAG2 (Elkin et al., 2003; Tsai and Schatz, 2003). The core RAG proteins can mediate formation of this same type of precise but incomplete HJ in NHEJ-deficient cells (Bogue et al., 1997; Han et al., 1999; Sekiguchi et al., 2001). In contrast, the full-length RAGs form very few HJs in NHEJ-deficient cells and the majority of the joints contain large deletions and appear to be repaired by an alternative DNA repair pathway; thus, the full-length RAGs do not efficiently facilitate HJ formation in a cellular context (Sekiguchi et al., 2001). These results led to the proposal that the noncore regions suppress the ability of the RAGs to catalyze HJ in vivo (Sekiguchi et al., 2001). Consistent with this notion, in vitro experiments subsequently demonstrated that the full-length RAGs exhibit decreased HJ activity in comparison to core RAGs (Elkin et al., 2003; Tsai and Schatz, 2003). However, because the full-length RAGs can form detectable levels of precise HJs in vitro, additional cellular factors may be responsible for further influencing the pathways of HJ formation in vivo.
Transposition Mediated by RAG1/2 Whereas RAG1/2 serves as an endonuclease in V(D)J cleavage, RAG1/2 can also perform transposition, inserting the cleaved RSSs into unrelated DNA (reviewed by Fugmann, 2001). Several observations led to the experiments that demonstrated RAG1/2 transposase activity. First were the stereochemical studies of hairpin formation which, as discussed earlier, demonstrated that these were formed by direct transesterification rather than by a reaction requiring a covalent intermediate (van Gent et al., 1996). This type of conservative DNA strand transfer (the generation of the hairpin bond requires the breakage of the opposing DNA strand) is typical of transposases (Engelman et al., 1991; Mizuuchi and Adzuma, 1991). Second was the demonstration that purified core RAG1/2 could form hybrid joints in vitro (Melek et al., 1998). This type of hybrid joint formation uses the same chemistry as transposition. Efficient transposition can be achieved with the purified core RAG proteins (Agrawal et al., 1998; Hiom et al., 1998). Transposition requires a 12/23 RSS pair, presumably to activate the RAG proteins, and relies on the same active site as is used for RSS cleavage. Transposition need not be coupled to RSS cleavage, as precut RSS ends can also be used. Although an RSS pair is required for transposition, both double-ended and single-ended insertions can be observed (Figure 5.6). As with other transposases that attack the two DNA strands at staggered positions, the DNA insertion sites for the two RSS ends in a coupled attack on the opposite strands of DNA are offset, in this case by 3 to 5 bp. The target DNA sites are generally GC-rich and a preference for insertion into DNA that can form a hairpin loop has been reported. Both intra- and intermolecular transposition can occur. Despite the efficiency of transposition in vitro, attempts to detect RAG-mediated transposition by expressing RAG proteins in cultured mammalian cells have been unsuccessful. However, two examples of RAG-mediated transposition of TCR a signal ends into the HPRT gene in a T cell isolate from a normal individual have recently been described, indicating that transposition in mammalian cells is not totally excluded (Messier et al., 2003). Regulatory mechanisms likely exist in lymphocytes to suppress the propagation of transposable elements, because frequent transposition events involving the rearranging antigen receptor loci would be highly detrimental to the host genome. Indeed, the lack of efficient full-length RAG-mediated HJ formation (which is mechanistically similar to transposition) in NHEJdeficient cells suggests that this RAG activity is downregulated in a cellular context (Sekiguchi et al., 2001). Furthermore, in vitro experiments using full-length RAG1 and RAG2 have clearly demonstrated that the noncore regions significantly inhibit RAG-mediated transposition (Elkin et al., 2003; Tsai and Schatz, 2003). In addi-
5. The Mechanism of V(D)J Recombination
FIGURE 5.6 RAG-mediated transposition. (A) Two-ended transposition. Upon cleavage of the 12/23 RSSs by RAG1/2, the RSS ends can be used in an attack on another, nonspecific DNA duplex (dashed lines). This coupled attack leads to the integration of the RSS flanked DNA fragment at positions staggered by 3 to 5 bp, resulting in target site duplication at the integration site. (B) One-ended transposition. RAG1/2 can also mediate attack of a single RSS end on a DNA molecule (depicted as a duplex circle).
tion, RAG-mediated induction of transposition has been demonstrated in yeast, indicating that the proteins are fully capable of carrying out transposition in vivo (Clatworthy et al., 2003). Therefore, active mechanisms must be in place within mammalian cells to channel RSS ends toward signal joint formation and to inhibit transposition and, in vivo, the full-length RAGs have evolved regulatory mechanisms to significantly downregulate this activity. It is not surprising that RAG-mediated transposition is prevented in developing lymphoid cells, because active transposition could lead to harmful genomic alterations, such as the generation of potentially oncogenic chromosomal translocations or inactivation of essential or tumor suppressor genes. Although many lymphoid tumors are associated with translocations initiated by V(D)J cleavage, the vast majority of these tumors appear to result from intrachromosomal V(D)J recombination or by misrepair of RAG-generated DSBs at antigen receptor loci. As discussed later, lymphomas resulting from the latter class of translocations can be eliminated by removal of the RAG genes. However, other than the one example mentioned, there is still no evidence for RAG-mediated transposition as a major pathway leading to translocations. However, generation of mice harboring
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FIGURE 5.7 The NHEJ pathway joins RAG-liberated coding and signal ends. The Ku heterodimer, XRCC4, and Lig4 are required for both coding and signal joining, whereas DNA-PKcs and Artemis are more important for coding joining. The RAGs also play an important role during the joining phase of V(D)J recombination in the context of post-cleavage synaptic complexes.
appropriate RAG mutations may help to elucidate the potential existence of such a pathway in vivo.
CODING AND SIGNAL JOINT FORMATION REQUIRES THE NHEJ PATHWAY The DNA ends generated by the RAG1/2 endonuclease cleavage reaction are joined by generally expressed cellular DNA repair machinery. The coding and RSS ends produced by RAG cleavage form different substrates for the joining phase of the V(D)J recombination; however, both types of end structures are fused by the ubiquitously expressed nonhomologous end-joining (NHEJ) pathway of DNA double strand break (DSB) repair (Figure 5.7) (reviewed by Bassing et al., 2002). In this regard, hairpinned coding ends must be opened and further processed before joining, whereas blunt RSS ends can be directly fused. Extensive nucleotide sequence analyses of endogenous joints have shown that hairpin coding ends normally are opened at or near the apex. Cleavage of a hairpin away from the apex leaves an over-
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hanging flap, which if incorporated into the joint results in a P (palindromic) nucleotide addition (Figure 5.8) (Lafaille et al., 1989; McCormack et al., 1989). These additions are one source of “junctional diversity” (reviewed by Lewis, 1994). The opened hairpin ends can be further modified by nuclease action, which can remove a self-complementary overhang or cut further into the original coding sequence. Finally, the lymphoid-specific terminal deoxynucleotidyl transferase (TdT) enzyme can add nontemplated (N) nucleotides to the ends (Alt and Baltimore, 1982; Gilfillan et al., 1993; Komori et al., 1993). N regions are believed to play a major role in the somatic diversification of the repertoire of antigen receptor variable regions (Davis et al., 1997). Finally, additional junctional diversity comes from the nucleolytic activities that remove potential coding end nucleotides. Thus, the joining phase of the V(D)J recombination provides a major source of diverse junctional sequences for V(D)J coding joins (Figure 5.8). In this regard, the region of the Ig sequence encodes CDR3 and also
encodes an analogous region of the TCR chains; thus, such junctional diversification mechanisms provide a major source of antigen receptor diversity (Davis et al., 1997).
Double Strand Break Repair by Nonhomologous DNA End-Joining DNA double strand breaks (DSBs) can be introduced by external agents such as ionizing radiation (IR) or radiomimetic drugs, by normal cellular metabolism, and in the context of specific developmental programs such as V(D)J recombination. DSBs are one of the most dangerous lesions that a cell can suffer, potentially leading to adverse consequences such as cell death or chromosomal translocations that can contribute to cancer. In this context, mammalian cells employ two different pathways to repair DNA double strand breaks (DSBs). Homologous recombination leads to accurate repair of DSBs by copying intact information from a homologous DNA template and is generally
FIGURE 5.8 Processing of coding ends prior to joining. Subsequent to RAG1/2 cleavage and concomitant formation of blunt, 5¢ phosphorylated signal and hairpin coding ends, several different events can modify the coding ends prior to ligation. P elements may be added if the hairpins are opened at sites away from the apex, the TdT enzyme can add nontemplated N nucleotides to the open coding ends, and the coding ends can undergo deletion. The events are depicted here as independent, but can occur concurrently during V(D)J rearrangements.
5. The Mechanism of V(D)J Recombination
thought to be most prominently used in the S and G2 phases of the cell cycle, when such templates are most readily available (reviewed by Thompson and Schild, 2001). On the other hand, NHEJ rejoins broken ends irrespective of sequence, can result in deletions or insertions at the junctions, and appears most prominent in the G1 phase of the cell cycle, the phase during which RAG activity is also predominant (Lin and Desiderio, 1995; Takata et al., 1998; reviewed by Jackson, 2002). The NHEJ pathway is known to involve at least six proteins, including Ku70, Ku80, DNA-dependent protein kinase catalytic subunit (DNA-PKcs), XRCC4, DNA Ligase IV (Lig4), and Artemis. The first five proteins were linked, directly or indirectly, to the NHEJ pathway by studies of mutant cells that were both sensitive to IR and defective in V(D)J recombination (reviewed by Bassing et al., 2002; Taccioli and Alt, 1995). Artemis was identified as the gene mutated in one form of human SCID (Moshous et al., 2001); see below). Notably, after finding their role in NHEJ in mammalian cells, Ku70, Ku80, XRCC4, and Lig4 homologs were found to participate in a NHEJ pathway conserved in yeast (Boulton and Jackson, 1996; Boulton and Jackson, 1996; Feldmann et al., 1996; Herrmann et al., 1998; Mages et al., 1996; Milne et al., 1996; Schar et al., 1997; Siede et al., 1996; Teo and Jackson, 1997; Wilson et al., 1997). However, DNA-PKcs and Artemis appear to have evolved more recently in vertebrates and, as described later, appear to play a more restricted role in the NHEJ reaction (Jeggo and O’Neill, 2002). The recent identification of a human SCID cell line not defective in any of these genes indicates that additional factors may also be required (Dai et al., 2003).
Identification of Mammalian NHEJ Proteins The importance of the NHEJ pathway during V(D)J joining was established by discoveries that certain IR sensitive mutant rodent cells also exhibit a severe impairment in ability to join RAG-induced DSBs (Bosma and Carroll, 1991; Taccioli et al., 1993). Through the analysis of different complementation groups of radiosensitive Chinese hamster ovary (CHO) cell lines (Taccioli et al., 1993), two known proteins, Ku80 and DNA-PKcs, were identified as NHEJ factors (Blunt et al., 1995; Kirchgessner et al., 1995; Smider et al., 1994; Taccioli et al., 1994; Taccioli et al., 1994). A complementation cloning approach utilizing an additional IR-sensitive CHO line led to the identification of XRCC4, a previously unknown gene, as another NHEJ factor (Li et al., 1995). Subsequently, the roles for these proteins in V(D)J recombination in vivo were confirmed by gene-targeted mutation studies (Gao et al., 1998; Gao et al., 1998; Kurimasa et al., 1999; Nussenzweig et al., 1996; Taccioli et al., 1998; Zhu et al., 1996), as well as by the fact that the spontaneously arising scid mutation in mice (Bosma
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and Carroll, 1991) actually involved a mutation in the carboyl terminus of the large DNA-PKcs gene (Araki et al., 1997; Blunt et al., 1996). The roles for two additional NHEJ proteins, Lig4 and Ku70, in NHEJ and V(D)J joining were implicated based on their interaction with XRCC4 and Ku80, respectively (Critchlow et al., 1997; Grawunder et al., 1997; Mimori et al., 1986). Subsequent gene-targeted mutation studies definitively showed that Lig4 and Ku70 were required both for normal DNA DSB repair and V(D)J recombination (Frank et al., 1998; Grawunder et al., 1998; Gu et al., 1997; Gu et al., 1997; Ouyang et al., 1997). Artemis deficiency in humans leads to radiosensitivity and a V(D)J recombination defect (Moshous et al., 2001) and its similar role in mice was confirmed by gene-targeted mutation analyses (Rooney et al., 2003; Rooney et al., 2002). To date, Artemis is the only NHEJ factor identified that has been implicated in human SCID, possibly because other known NHEJ factors may be more necessary for cellular proliferation and survival in humans than in mice (Li et al., 2002).
Functions of NHEJ Proteins The functions of the various NHEJ proteins are beginning to emerge, both from biochemical characterization of their activities as well as by analyses of the steps in the V(D)J reaction that are impaired in cells carrying homozygous inactivating mutations of genes encoding individual factors. Ku70 and Ku80 Ku70 and Ku80 form a heterodimer, Ku, which possesses DNA end-binding activity (Mimori and Hardin, 1986). Purified Ku protein was found to promote the association of two DNA molecules in vitro; thus, it was proposed to possess end bridging or alignment activity (Ramsden and Gellert, 1998). The crystal structure of Ku bound to DNA revealed that the Ku heterodimer forms a ring that encircles duplex DNA and positions the DNA helix in a defined path, thus providing structural evidence in support of an end alignment function (Walker et al., 2001). Upon binding to DNA ends, Ku associates with and activates the serine–threonine protein kinase activity intrinsic to DNA-PKcs, thus forming the trimeric DNA-PK holoenzyme (Khanna and Jackson, 2001); one potential role for this function may be inferred from the interaction between DNA-PKcs and Artemis. Studies in yeast have supported the notion that Ku may serve an end-protection function as well (Lee et al., 1998). Additional Ku functions during V(D)J recombination have been suggested based on in vitro studies (reviewed by Featherstone and Jackson, 1999; Tuteja and Tuteja, 2000) and may include end remodeling, or recruitment of factors
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in addition to DNA-PKcs, including the XRCC4/Lig4 complex (Chen et al., 2000; Nick McElhinny et al., 2000); however, it has not yet been proved that Ku plays such roles in vivo (Doherty and Jackson, 2001). XRCC4 and DNA Ligase IV Although several other DNA ligases are present in mammalian cells, they cannot compensate for a defect in Lig4 activity during V(D)J joining or general DNA DSB repair. Cells deficient for Lig4 (or XRCC4) are severely impaired for both coding and RSS joining and are markedly radiosensitive (Frank et al., 1998; Gao et al., 1998; Grawunder et al., 1998). XRCC4 binds to Lig4, and this interaction stimulates Lig4 in vitro (Critchlow et al., 1997; Grawunder et al., 1997) and stabilizes it in vivo (Bryans et al., 1999). However, it is possible that XRCC4 may have additional functions during NHEJ, because it can bind nonspecifically to DNA in the absence of Lig4 (Modesti et al., 1999). The crystal structure of XRCC4 indicates that it forms a stable dimer that interacts with Lig4 (Junop et al., 2000; Sibanda et al., 2001). DNA-PKcs and Artemis A number of observations indicate that DNA-Pkcs functions primarily in coding joint formation. A role for DNAPKcs in coding versus RSS joining was originally indicated by the finding that cells from SCID mice, later found to be DNA-PKcs–deficient (Blunt et al., 1995; Blunt et al., 1996; Miller et al., 1995), were much more severely impaired for coding versus RSS joining (Blackwell et al., 1989; Lieber, 1998; Malynn et al., 1988). Moreover, hairpin coding ends were found to accumulate in DNA-PKcs–deficient developing lymphocytes (Roth et al., 1992), suggesting a difficulty in processing them. Also, unusually large P nucleotide additions are found in the rare coding joints recovered from DNA-PKcs–deficient cells, suggesting that the hairpins have been improperly opened further from the apex than is normal (Lewis, 1994). More recently, these findings generally have been reproduced in DNA-PKcs–deficient cells and mice generated by gene-targeted mutation and which lack DNA-PKcs protein (Gao et al., 1998; Kurimasa et al., 1999; Taccioli et al., 1998). DNA-PKcs is a serine–threonine protein kinase containing a phosphatidylinositol 3 kinase (PI3K) catalytic domain that is activated upon interaction with Ku bound to DNA ends (Smith and Jackson, 1999). DNA-PK phosphorylates a variety of targets in vitro, including p53, transcription factors, WRN, XRCC4, Ku, and Artemis (Ma et al., 2002; Smith and Jackson, 1999) and is capable of autophosphorylation (Chan et al., 2002; Chan and Lees-Miller, 1996; Merkle et al., 2002); however, in addition to Artemis (see below), the physiological relevance of its in vitro substrates is unclear. In addition to its roles in the context of Ku and
Artemis complexes, DNA-PKcs itself may be capable of synapsing broken DNA ends (DeFazio et al., 2002), and thus may also serve a structural role during end joining. Finally, DNA-PKcs may function outside the NHEJ pathway (Gurley and Kemp, 2001; Sekiguchi et al., 2001), playing roles that may overlap with those of the ataxia telangiectasia mutated (ATM) protein. ATM, like DNA-PKcs, is a serine–threonine protein kinase with a PI3 kinase domain and is involved in controlling cellular responses to DNA DSBs (reviewed by Shiloh, 2001). Thus, the novel NHEJindependent roles for DNA-PKcs may involve damage signaling related to checkpoint control (Jackson, 2002). The discovery and characterization of Artemis provided a major insight into one potential in vivo function of DNAPKcs. Artemis is a 77.6 kDa protein that is a member of the metallo-b-lactamase superfamily (Callebaut et al., 2002; Moshous et al., 2001), of which some members appear to be involved in the repair of interstrand cross-links (ICL) in mice and yeast. The RS-SCID patients having Artemis mutations lack B and T lymphocytes and show increased radiosensitivity of bone marrow cells and skin fibroblasts (Cavazzana-Calvo et al., 1993; Moshous et al., 2001; Nicolas et al., 1996; Nicolas et al., 1998). Moreover, transient V(D)J recombination substrate studies showed that RSSCID fibroblasts are more defective for coding than RSS joins, much like DNA-PKcs deficient cells (Moshous et al., 2001; Moshous et al., 2000; Nicolas et al., 1998). In addition, a large proportion of the rare coding joints recovered from Artemis-deficient ES cells contain longer than average P nucleotide additions, reminiscent of those recovered from DNA-PKcs-deficient cells (Rooney et al., 2003). Thus, it was proposed that Artemis may function to open hairpin DNA coding ends (Moshous et al., 2001). Strong support for this notion came from in vitro studies showing that DNAPKcs forms a complex with and phosphorylates Artemis, leading to the activation of an endonuclease activity that can cleave RAG-generated hairpins (Ma et al., 2002). Thus, these results led to the hypothesis that a DNA-PKcs/Artemis complex, perhaps recruited by Ku, opens coding hairpin ends in vivo (Karanjawala et al., 2002; Ma et al., 2002). Indeed, in support of this notion, hairpin coding ends accumulate in Artemis-deficient thymocytes (Rooney et al., 2002), as they also do in Ku and DNA-PKcs–deficient thymocytes, consistent with a role for the entire Ku/DNA–PKcs/Artemis complex in this reaction (Gao et al., 1998; Roth et al., 1992; Zhu et al., 1996; Zhu and Roth, 1995). The more limited role of the DNA-PKcs/Artemis complex in V(D)J recombination, that of opening of coding end hairpins, as opposed to the four evolutionarily conserved factors that are required for both RSS and coding joins, also may give further insight into the evolution and function of these proteins. Thus, the four conserved factors may be components of a conserved complex that forms a
5. The Mechanism of V(D)J Recombination
basic end-ligation function. DNA-PKcs and Artemis, as also suggested by other lines of evidence (Gao et al., 1998; Rooney et al., 2003, see below), may have evolved more recently to function to process ends that cannot be simply ligated (e.g., blocked ends or hairpins) to a form that can be joined by the basic end-ligation apparatus. Other Activities Several additional activities are predicted to be required during V(D)J joining. One such activity is a DNA polymerase responsible for filling in short gaps at coding junctions that may be generated by end modifications. Eukaryotic DNA polymerases of the pol X family [e.g., Pol4 in S. cerevisiae (Wilson and Lieber, 1999) and Pol m in mammals (Mahajan et al., 2002)] have been implicated as potential candidates for such a V(D)J polymerase. Human pol m, which has homology to TdT, has been found to interact with Ku and requires Ku, XRCC4, and Lig4 for stable DNA binding in vitro (Mahajan et al., 2002). Pol m is upregulated and forms foci upon exposure of cells to IR, suggesting a role in general DNA DSB repair (Mahajan et al., 2002). A subset of mice deficient in pol m exhibit a significant depletion of B cells in peripheral lymphoid organs, thus indicating a function for pol m during B cell development (Bertocci et al., 2002). However, currently no compelling in vivo evidence exists to point to a specific DNA polymerase that functions during V(D)J recombination; thus, the identity of the V(D)J polymerase remains unknown. It is evident that normal V(D)J recombination involves a nucleolytic activity that deletes nucleotides at the coding ends. Several potential candidates include the Mre11/Rad50/Nbs1 (MRN), RAG1/RAG2, and Artemis/ DNA–PKcs complexes, which may fulfill the role of a V(D)J nuclease during processing of open coding hairpin ends. The MRN complex is required for DNA DSB repair in vivo and in vitro and has been demonstrated to possess endo and exonuclease activities (D’Amours and Jackson, 2002). Nbs1 has been found in foci at V(D)J induced breaks (Chen et al., 2000), and mice expressing a hypomorphic allele of Nbs1 exhibit defects in lymphocyte development (Kang et al., 2002). However, mutations in Nbs1 that result in DNA DSB repair defects do not have any obvious effects on coding junction sequences in vitro or in vivo (Harfst et al., 2000; Kang et al., 2002; Yeo et al., 2000). Thus, the Mre11/Rad50/Nbs1 complex may play a more indirect role in V(D)J recombination, such as in DSB detection and/or signaling. In vitro, truncated forms of RAG1/RAG2 have been demonstrated to open hairpin coding ends and cleave 3¢ flap structures (Besmer et al., 1998; Santagata et al., 1999; Shockett and Schatz, 1999). Although it appears that the RAGs do not play a significant role in hairpin end opening in vivo (Zhu and Roth, 1995; Zhu et al., 1996; Rooney et al., 2002), they may play a role
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in further processing the open hairpins. Artemis/DNA–PKcs possesses endonuclease activity on 5¢ and 3¢ single strand overhanging flaps and Artemis, in the absence of DNA-PKcs, has intrinsic 5¢ to 3¢ exonuclease activity on single strand DNA (Ma et al., 2002); thus, it may play roles in coding end processing in addition to nicking hairpins. In support of this notion, rare coding joints recovered from Artemis-deficient ES cells in transient transfection V(D)J recombination assays exhibit significantly less nucleotide loss at the junctions compared to those recovered from wild type ES cells (Rooney et al., 2003). A clearer picture of which of these factors, if any, are involved in coding end processing awaits additional detailed studies. As mentioned previously, a major source of junctional diversity comes from the addition of nongermline encoded nucleotides at V-D, D-J, and some V-J junctions, which are referred to as N regions (Alt and Baltimore, 1982). Although it was proposed early on that N regions were added by TdT, this was proved by gene-targeted mutations studies that clearly demonstrated the absence of N regions in TdTdeficient lymphocytes (Gilfillan et al., 1993; Komori et al., 1993). TdT is not expressed substantially during fetal development and therefore most Ig and TCR junctions formed in the fetal repertoire lack N regions (reviewed by Benedict et al., 2000; Komori et al., 1996). In addition, junctions formed in the absence of N region addition also often used short homologies to form “canonical” junctions that appear very frequently in the absence of TdT [e.g., in fetal repertories; (Benedict et al., 2000; Komori et al., 1996)]. Thus, TdT expression during B and T cell development in the adult diversifies repertoires both by N region addition and by the diminution of canonical junctions, which form much less frequently in the presence of N regions.
Mice Deficient in the NHEJ Factors Mice deficient for all known lymphoid-specific and general V(D)J recombination factors have been generated by gene-targeted mutation (reviewed by Bassing et al., 2002; Rooney et al., 2002). RAG-1 or -2 deficient mice have a severe combined immune deficiency (SCID) due to the inability to initiate V(D)J recombination. Other than a complete block in B and T cell development at the progenitor stage, RAG-deficient mice do not exhibit any other phenotypes. This very specific phenotype is consistent with the notion that RAGs evolved only for their role in effecting antigen receptor variable region gene assembly in developing lymphocytes (Shinkai et al., 1992). Mice deficient for TdT, the only other known lymphocyte-specific V(D)J recombination factor besides RAG1 and 2, exhibit relatively normal V(D)J recombination levels; however, V(D)J coding junctions lack N-region additions (Gilfillan et al., 1993; Komori et al., 1993). This phenotype is consistent with the
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nonessential role for TdT in V(D)J recombination, which involves the diversification of V(D)J junctions via the addition of nontemplated N nucleotides to coding ends. Deficiencies in the NHEJ factors also result in impaired lymphocyte development. However, the NHEJ mutant mice and cells exhibit phenotypes beyond defects in V(D)J recombination owing to the importance of the NHEJ pathway in general DSB repair. Classical SCID mice, which express a nearly full length but catalytically inactive form of DNA-PKcs, and Artemis-deficient mice exhibit a “leaky” SCID phenotype with some T and B cells appearing in older mice (e.g., Bosma et al., 1988; Rooney et al., 2002; Taccioli et al., 1998). This leaky V(D)J joining occurs at very low levels and is catalyzed by the basic NHEJ pathway (or by an alternative repair pathway) following the opening of coding end hairpins by some lower level activity. However, this interpretation is somewhat complicated by the fact that some lines of DNA-PKcs–deficient mice generated by gene-targeted mutation to completely lack DNA-PKcs protein have not been found to be leaky (Gao et al., 1998). Thus, these and certain other minor phenotypic differences between complete DNA-PKcs knock-out mice and SCID mice (e.g. Bosma et al., 2002; Manis et al., 2002) may reflect some residual activity of the latter. DNA-PKcs and Artemis deficiencies result in variable cellular IR sensitivity (Gao et al., 1998; Rooney et al., 2003). Thus, ES cells harboring targeted inactivating mutations in either DNA-PKcs or Artemis are not radiosensitive; but murine embryonic fibroblasts (MEFs) homozygous for the same mutations are significantly more IR sensitive than wildtype MEFs, suggesting potentially redundant factors in ES cells (Gao et al., 1998; Rooney et al., 2003). Also, whereas DNA-PKcs– and Artemis-deficient ES cells do not display substantial IR sensitivity, they do display more significant sensitivity to bleomycin, which is a radiomimetic drug (Rooney et al., 2003). As bleomycin and IR may lead to a different spectrum of broken ends (Povirk, 1996), these findings are consistent with the notion, outlined earlier, that DNA-PKcs and Artemis are employed in NHEJ for repairing a specific subset of DNA damage that requires processing prior to ligation. Other than variable cellular IR sensitivity, DNA-PKcs– and Artemis-deficient mice have no other obvious consistent phenotype. Ku-deficient mice also have a SCID phenotype (Gu et al., 1997; Nussenzweig et al., 1996; Ouyang et al., 1997). In this regard, the SCID phenotype of Ku70-deficient mice is leaky (Gu et al., 1997; Ouyang et al., 1997), likely due to the lowlevel joining of RAG-induced DSBs, similar to that observed in DNA-PKcs– and Artemis-deficient mice. However, unlike DNA-PKcs and Artemis deficiencies, Ku deficiency results in mice that are significantly smaller than littermates (Gu et al., 1997; Nussenzweig et al., 1996; Ouyang et al., 1997) and that show increased apoptosis of
newly generated, post-mitotic neurons in embryos (Gu et al., 2000). In addition, Ku-deficient cells exhibit growth defects, premature senescence, and IR sensitivity (Gu et al., 1997; Nussenzweig et al., 1996; Ouyang et al., 1997). The phenotypic differences between Ku- and DNA-PKcs–deficient mice reinforce the notion that Ku possesses functions separate from any it may effect in the context of the DNA-PK holoenzyme. Finally, Ku70-, but not Ku80-, deficient mice also show an increased incidence of thymic lymphomas (Gu et al., 1997; Li et al., 1998). The reason for this difference is not clear but could relate to relative leakiness of the defects, as leaky SCID mice also are more prone to T cell lymphomas on certain backgrounds (Custer et al., 1985; Jhappan et al., 1997). XRCC4- and Lig4-deficiency leads to late embryonic lethality accompanied by severe neuronal apoptosis throughout the central nervous system (Barnes et al., 1998; Frank et al., 1998; Gao et al., 1998). In addition to cellular defects analogous to those of Ku-deficient mice, XRCC4and Lig4-deficient embryos exhibit a complete block in B and T cell development in fetal lymphoid organs. Notably, the breeding of XRCC4- or Lig4-deficient mice into a p53 deficient background rescues their embryonic lethality and neuronal apopotosis defects, but not their V(D)J recombination or lymphocyte development defects (Frank et al., 2000; Gao et al., 2000). Thus, the embryonic lethality and severe neuronal apoptosis of XRCC4- or Lig4-deficient mice appears to result from a p53-dependent response to unrepaired DSBs and not from the inability to repair the breaks via NHEJ per se (Frank et al., 2000; Gao et al., 2000). Conversely, defective lymphocyte development appears to result primarily from the inability to repair the RAGinitiated DSBs to generate the functional antigen receptor genes necessary to drive further development. However, it is also clear that XRCC4- or Lig4-deficient progenitor lymphocytes pools are severely depleted due to a p53-dependent response to the unrepaired RAG-initiated DSBs (Frank et al., 2000; Gao et al., 2000). XRCC4-, Lig4-, and Ku-deficient mice appear quite similar in a p53-deficient background (Difilippantonio et al., 2002; Difilippantonio et al., 2000; Frank et al., 2000; Gao et al., 2000; Lim et al., 2000; Zhu et al., 2002), suggesting that the major differences in Ku- versus XRCC4- or Lig4deficient phenotypes are likely quantitative, as indicated by the greater “leakiness” in NHEJ and a somewhat lower level of apoptotic cell death in Ku-deficient mice which, for example, allows the generation of a functional nervous system (Gu et al., 2000; Sekiguchi et al., 1999). Thus, deficiencies in the evolutionarily conserved NHEJ factors, Ku, XRCC4, and Lig4, exhibit similar phenotypes, albeit with varying severity, whereas, DNA-PKcs and Artemis have milder phenotypes consistent with their involvement in a more limited set of NHEJ functions.
5. The Mechanism of V(D)J Recombination
Recognition of RAG-Initiated DSBs by the DNA Repair and Cell Cycle Checkpoint Machinery Initiation of V(D)J recombination by RAG1/2 is tightly coupled to the cell cycle, as evidenced by the accumulation of RAG-generated DSBs in G0/G1 cells and the periodic accumulation of the RAG2 protein during G0/G1 and its subsequent degradation at the G1–S transition (Desiderio et al., 1996; Schlissel et al., 1993). This form of regulation would be optimal to ensure joining via NHEJ. Developing lymphocytes containing unrepaired RAG-initiated DSBs normally undergo programmed cell death resulting from induction of the p53-dependent cell cycle checkpoint (reviewed by Lu and Osmond, 2000). Likewise, progenitor populations of developing lymphocytes are dramatically reduced in NHEJ-deficient animals. This decrease appears to be caused by the extensive apoptosis of progenitors harboring unrepaired RAG-induced DSBs, as p53-deficiency leads to the increased survival and proliferation of NHEJdeficient lymphocyte progenitors (Difilippantonio et al., 2000; Frank et al., 2000; Gao et al., 2000; Guidos et al., 1996), which in turn leads to the development of aggressive progenitor-B cell lymphomas. Thus, the efficient recognition of RAG-generated DNA ends by the NHEJ pathway is clearly imperative to avoid DSB detection and induction of p53-dependent apoptosis. Normally, coding ends are joined rapidly (Ramsden and Gellert, 1995; Zhu and Roth, 1995); however, in contrast, RSS ends persist throughout G1 and are joined at the G1/S transition (Ramsden and Gellert, 1995; Roth et al., 1992; Schlissel et al., 1993). However, such persistent expression does not lead to p53 induction in normal progenitor populations (Guidos et al., 1996). Thus, the prolonged presence of RSS ends appears to escape detection by the cell cycle checkpoint machinery, possibly by sequestration in a stable postsynaptic cleavage complex. In addition to p53, other proteins that monitor DNA damage and repair also appear to interplay with the V(D)J recombination reaction. The ATM protein is noteworthy in this regard. Although ATM is not directly involved in V(D)J recombination (Barlow et al., 1996; Elson et al., 1996; Hsieh et al., 1993; Xu et al., 1996), deficiency for this protein leads to lymphoid malignancies (reviewed by Khanna et al., 2001), which, in mice, are predominantly T cell lymphomas that frequently harbor translocations involving their TCRa/d locus (Barlow et al., 1996; Liyanage et al., 2000; Petiniot et al., 2000). Histone H2AX is another class of factor that has been implicated in some aspect of the V(D)J rearrangement process, which may include linkage with DNA repair and/or checkpoint pathways. H2AX is a histone H2A variant that phosphorylates upon DNA damage, such as is induced by IR, and is found in a phosphorylated form within foci of repair factors at DNA DSBs (Rogakou et al., 1998). Targeted muta-
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tional studies have shown that H2AX, which is phosphorylated by ATM and related kinases (Burma et al., 2001; Ward and Chen, 2001), is required for normal DNA DSB repair and for maintenance of genomic stability (Bassing et al., 2002; Celeste et al., 2002). Moreover, phosphorylated H2AX has also been observed to co-localize in foci with Nbs1 at DSBs induced during V(D)J recombination (Chen et al., 2000). Although mice deficient for H2AX do not exhibit severe defects in lymphocyte development (Bassing et al., 2002; Celeste et al., 2002), the localization data suggestes that H2AX may be involved in monitoring V(D)J rearrangements in the context of DNA checkpoints to suppress oncogenic translocations, possibly through modulation of chromatin structure. In this regard, very recent findings have shown that p53-deficient mice that are also deficient or haplo-insufficient for H2AX, are prone to lymphomas, including B lineage lymphomas with translocations that appear to involve RAGgenerated DSBs (Bassing et al., 2003; Celeste et al., 2003).
NHEJ Factors and Suppression of RAGInitiated Translocations In addition to their roles in V(D)J recombination and DNA DSB repair, the NHEJ factors also play important roles in maintaining genomic stability. A number of different types of chromosomal aberrations are observed in cells lacking a functional NHEJ pathway, including chromosome fragments, fusions, and translocations (reviewed by Ferguson and Alt, 2001). The importance of the NHEJ factors as genomic caretakers is highlighted by the fact that NHEJ-deficiencies, including inactivating mutations in Ku, XRCC4, Lig4, Artemis, and the classical SCID mutation in DNA-PKcs, in combination with deficiencies in the p53 cell cycle checkpoint protein in mice, predispose to lymphomagenesis (Difilippantonio et al., 2000; Frank et al., 2000; Gao et al., 2000; Gladdy et al., 2003; Lim et al., 2000; Nacht et al., 1996; Rooney, Sekiguchi, and Alt, in preparation). Equally notable is that fact that in all cases, the predominant tumor is a pro-B cell lymphoma that has translocations and amplifications involving the c-myc and IgH loci (Difilippantonio et al., 2000; Gao et al., 2000). Various lines of evidence have shown that the initiating lesions that cause the oncogenic chromosomal aberrations in these NHEJ/ p53-deficient pro-B lymphomas are RAG-induced DSBs (Difilippantonio et al., 2002; Gladdy et al., 2003; Vanasse et al., 1999; Zhu et al., 2002). Thus, the translocations involve JH region sequences, and the introduction of RAG mutation into these mutant backgrounds eliminates the occurrence of pro-B lymphomas bearing the hallmark chromosomal anomalies (Difilippantonio et al., 2002; Gladdy et al., 2003; Vanasse et al., 1999; Zhu et al., 2002). These findings implicate a pro-B cell lymphomagenesis model in which RAG-initiated DSBs in p53/NHEJ pro-B
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cells are neither repaired nor eliminated via a G1 checkpoint. Thus, these mutant pro-B cells can progress into S phase where the RAG-initiated DSBs at their JH locus are replicated. Subsequently, these replicate to generate dicentric chromosomes which, in the p53-deficient background, can rapidly generate the amplification of genes conferring a selective growth advantage via a breakage bridge fusion mechanism (Difilippantonio et al., 2002; Zhu et al., 2002). Although it is unclear if this mechanism contributes to human B cell lymphomas, it might be involved in advanced human solid tumors and potentially in some B lineage tumors, including advanced stage myelomas (Mills et al., in press). Finally, Ku, DNA-PKcs, and Artemis also may play roles in telomere maintenance, as cells deficient in these factors lead to defects in telomere capping (Bailey et al., 2001; Bailey et al., 1999; Espejel et al., 2002; Goytisolo et al., 2001; Rooney et al., 2003; Samper et al., 2000). In addition, Ku and DNA-PKcs deficiencies may also result in dysregulation of telomere length (d’Adda di Fagagna et al., 2001; de Lange, 2002; Espejel et al., 2002; Hsu et al., 2000; Samper et al., 2000).
Acknowledgments F.W.A. is an Investigator of the Howard Hughes Medical Institute. J.S. is a Special Fellow of the Leukemia and Lymphoma Society. This work was supported by NIH grants AI35714 and NCI CA92625 (FWA) and GM48025 (MAO).
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6 Transcription of Immunoglobulin Genes KATHRYN CALAME
RANJAN SEN
Departments of Microbiology and Biochemistry & Molecular Biophysics, Columbia University College of Physicians and Surgeons, New York. New York, USA
Department of Biology, Brandeis University, Waltham, Massachusetts, USA
Given the abundance of mRNA encoding immunoglobulin (Ig) light and heavy chains in murine plasmacytoma lines, immunoglobulin cDNAs and genes were among the first to be cloned in the late 1970s. The transcription of immunoglobulin genes quickly attracted the attention of many laboratories because of its strict B-cell specificity and developmental stage–specific regulation. In addition, with the understanding that functional Ig genes were created by a unique process of VDJ DNA rearrangement in B lymphocytes, the location and character of transcriptional regulatory elements was of particular interest. Ig gene transcriptional regulation became more intriguing with the discovery, in 1983, of the Ig heavy chain intronic enhancer (Em) (Banerji, Olson et al., 1983; Gillies, Morrison et al., 1983; Mercola, Wang et al., 1983). This enhancer, located between the JH and Cm gene segments, provided an explanation for transcriptional activation of rearranged VH gene promoters while unrearranged VH promoters remained inactive. Furthermore, Em was the first transcriptional enhancer identified in a mammalian gene and, like the SV40 enhancer, it could activate transcription in a distance- and orientationindependent manner. Indeed, it soon became obvious that most regulatory elements for both light and heavy chain immunoglobulin genes resided in enhancers that were either located in intervening sequences between J and C gene segments or 3¢ of C gene segments, or both. The molecular mechanism(s) by which these enhancers act has been, and continues to be, a central challenge for understanding Ig gene transcription. Both previous editions of this book contained chapters that summarized our understanding of immunoglobulin gene transcriptional regulation. At this time, most of the elements and DNA binding proteins involved are probably identified
and an updated summary of this information is presented. This chapter presents general characteristics of the Ig regulatory elements and discusses areas of current research. In addition, we discuss how, in a striking example of serendipity in science, research on Ig transcriptional regulation has provided unexpected insights into other aspect of immune system biology.
Molecular Biology of B Cells
TRANSCRIPTIONAL REGULATORY ELEMENTS IN IMMUNOGLOBULIN HEAVY AND LIGHT CHAIN GENES The regulatory elements in Ig genes have been described in previous editions of this volume and in many reviews (Leanderson and Hogbom, 1991; Li, Rothman et al., 1991; Staudt and Lenardo, 1991; Eckhardt, 1992; Kadesch, 1992; Ernst and Smale, 1995; Henderson and Calame, 1995; Henderson and Calame, 1998; Magor, Ross et al., 1999; Khamlichi, Pinaud et al., 2000). Our current understanding of these elements is summarized in Figures 6.1 and 6.2 and discussed below, followed by a discussion of the proteins that bind these elements.
Ig Promoters Each functional Ig V gene segment has a transcriptional initiation site, a TATA element, and regulatory sequences comprising a promoter extending approximately 100 to 200 bp 5¢ of the leader coding sequences. Both heavy and light chain gene promoters are remarkably simple (Figures 6.1 and 6.2). The most important regulatory element in both light and heavy chain promoters is an octamer element,
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Copyright 2004, Elsevier Science (USA). All rights reserved.
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FIGURE 6.1 Summary of transcriptional regulatory elements in the immunoglobulin heavy chain locus. Boxes indicate protein binding sites that are thought to be functional. Where known, the proteins that bind these sites are shown with activators in green and repressors or inhibitors of the activators in red. The arrow indicates the transcription initiation site. Distances are not to scale. See color insert.
which is usually located within 100 bp of the transcription initiation site. Initially, it was hypothesized that this element would confer B-cell specificity to V gene promoters; however, as discussed below, the roles of different B cellspecific and non-B cell-specific octamer proteins remain unclear, leaving the questions of oct-dependent B cell specificity unresolved. A few other regulatory elements (E, mE3) have been identified in VH and Vk promoters, but in functional assays their roles are less important than the oct sites (Avitahl and Calame, 1996). Indeed, it is interesting that C/EBP family proteins, which often bind VH and Vk pro-
moters, have recently been shown to interact with octamer proteins (Hatada, Chen-Kiang et al., 2000), underscoring the role of octamer protein for V promoters. Matrix attachment regions (MARs) have also been found 5¢ of many VH promoters (Goebel, Montalbano et al., 2002). Different VH promoters have been found to have different strengths and different degrees of enhancer dependence (Buchanan, Hodgetts et al., 1995; Love, Lugo et al., 2000). In general, however, the strong enhancer-dependence of V gene promoters in vivo renders the intrinsic activity of the promoters themselves less important for understanding Ig gene
6. Transcription of Immunoglobulin Genes
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FIGURE 6.2 Summary of transcriptional regulatory elements in the immunoglobulin light chain loci. Boxes indicate protein binding sites that are thought to be functional. Where known, the proteins that bind these sites are shown with activators in green and repressors or inhibitors of the activators in red. Parentheses indicate sites where proteins are presumed to bind but have not been shown experimentally. The arrow indicates the transcription initiation site. Distances are not to scale. See color insert.
expression, although it may be important for determining the accessibility of the V gene segments to recombinase activity (Sikes, Suarez et al., 1999).
Em The Em heavy chain intronic enhancer was the first Ig enhancer to be identified, and it has been extensively studied. It has strong, classical transcriptional enhancer activity that is promoter-, distance-, and orientation-
independent. Transgenes in which expression is dependent on Em show that the enhancer is active throughout B cell development from earliest pro B cells to plasma cells and it also has some activity in medullary thymocytes (Cook, Meyer et al., 1995). Multiple protein binding sites are present in Em, and experimental evidence indicates that those shown in Figure 6.1 are functionally important. The activity of many individual sites appears to be redundant with other sites, since mutation of individual sites usually has only a minor effect on activity whereas deletion of mul-
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tiple sites has significant impact. As detailed below, many sites in Em can be recognized by multiple proteins, some of which activate and some of which repress transcription, providing ample possibilities for subtle regulation. The “core” enhancer, which provides transcriptional activation in B cells, is flanked by two MARs that have been postulated to positively or negatively modulate its activity. As discussed below and elsewhere in this volume, a requirement for Em in VDJ recombination has been shown by gene targeting. However, there are no studies in which Em has been deleted from a rearranged heavy chain gene. This would assess its importance for transcription of a rearranged VH promoter in normal B cells, although there are cell lines that express Ig heavy chains normally from rearranged alleles that lack Em (Klein, Sablitzky et al., 1984; Wabl and Burrows, 1984).
Enhancers 3¢ of Ca Cell lines in which Em was deleted, but which continue to transcribe Ig heavy chains, provided the first indication that other enhancers might be present in the IgH locus. Indeed, it is now clear that a complex region 3¢ of Ca and more than 200 kb 3¢ of Em has enhancer activity (Figure 6.1). A recent review on this enhancer region provides a detailed summary of studies to determine its role and activity (Khamlichi, Pinaud et al., 2000). The region denoted HS1,2 is 16 kb 3¢ of Ca in the mouse IgH locus and was the first B-cell specific enhancer in this region to be identified. Subsequently, others, which are numbered based on the occurrence of DNaseI hypersensitive sites, were also identified. As indicated in Figure 6.1 the region contains several inverted repeats; HS3a and HS3b are 97% identical but in opposite orientations. Transfection studies suggested that HS1,2 had highest activity in activated B cells (Dariavach, Williams et al., 1991), and this was largely confirmed by transgene studies, although complete B-cell specificity was not observed (Arulampalam, Grant et al., 1994). HS3a and b and HS4 have weaker activity, primarily in activated B cells, and HS4 appears to be active throughout B cell development. The entire region displays locus control activity (Madisen and Groudine, 1994; Madisen, Krumm et al., 1998). Em and the 3¢ enhancers appear to synergize in a position- and distance-dependent manner (Mocikat, Kardinal et al., 1995), and the 3¢ enhancers probably function in vivo as co-enhancers. Mice with a targeted deletion of HS1,2, but retaining Em, had normal IgH transcription and selective defects in CH germline transcripts (Cogne, Lansford et al., 1994). Mice lacking the entire region 3¢ of Ca have not yet been described.
of these enhancers and has shown that they both play a role in VkJk recombination and that each has redundant and unique functions (Inlay, Alt et al., 2002). The intronic enhancer appears to be more important for secondary rearrangements that allow receptor editing (Nemazee, 2000) and for monoallelic demethylation that is required for ordered kappa rearrangement (Mostoslavsky, Singh et al., 2001). The developmental stage specificity of kappa gene rearrangement and expression was originally thought to be largely determined by the binding of NF-kB/rel family proteins in the kB site of the intronic enhancer. However, in vivo footprinting studies showed that the kB site was occupied in both pro and pre B cells; changes in occupancy of Cre, BSAP, and mB, NF-EM5 sites in the 3¢ enhancer, suggest these sites may be more important for the developmental stage-specific rearrangement and expression of kappa genes (Shaffer, Peng et al., 1997).
Lambda Enhancers Enhancers 3¢ of the constant gene segments have been found in lambda loci in mouse and human. Both the murine and human elements are illustrated in Figure 6.2 because the human element has been studied in some detail recently (Asenbauer, Combriato et al., 1999). Most protein binding sites in these enhancers are also found in other Ig enhancers, but a role for Mef proteins appears to be unique to the lambda enhancers (Satyaraj and Storb, 1998). A role for PU.1/IRF-4 in the murine lambda enhancer was evident in early studies (Pongubala, Nagulapalli et al., 1992; Eisenbeis, Singh et al., 1993) and provided a paradigm for understanding the activity of PU.1, in conjunction with other proteins, in many Ig enhancers.
PROTEINS BINDING IN IG TRANSCRIPTIONAL REGULATORY ELEMENTS Most proteins that bind to individual sites in the Ig promoters and enhancers have been identified and studied in detail. Since much of this basic information has been discussed in earlier editions of this volume and in other reviews, it has been summarized in Table 6.1 along with pertinent references. Below, we discuss some general features of the regulation and mechanism of action of proteins that bind sites in the Ig promoters and enhancers.
Kappa Intronic and 3¢ Enhancers
Ig Enhancer Activities Are Regulated by Multiple Sites and Mechanisms
In an arrangement similar to the IgH locus, the murine kappa locus has both an intronic and a 3¢ enhancer (Figure 6.2). Gene targeting has been used to compare the activities
A consistent finding has been that the Ig enhancers are complex elements and their B-cell and developmental stage specificity is not easily explained by a single B cell or devel-
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6. Transcription of Immunoglobulin Genes
TABLE 6.1 Protein
Sites
Activity
ATF/CREB
Cre
Pos.
Bright
MARs
CBF
Family bZip
Expression
Comments and references
Ubiquitous
Regulated by cAMP (De Cesare and Sassone-Corsi, 2000)
Pos.
B cells
Competes with Cux/CDP; related to HMG and Swi/Snf; may remodel chromatin (Webb, 2001)
Pos.
Ubiquitous
Associates with E2A and PU.1 (Erman, Cortes et al., 1998)
C/EBPb
E
Pos.
bZip
Wide
Induced by LPS and negatively regulated by heterodimerizing, short forms (LIP) and C/EBPg, (Lekstrom-Himes and Xanthopoulos, 1998)
Cux/CDP
MARs
Neg.
Hox
Non-B cells
Competes with Bright (Wang, Goldstein et al., 1999)
E2-A/HEB/ E2-2
mE5, mE2, kE2
Pos.
bHLH
Ubiquitous
E47 homodimera are found in B cells; negatively regulated by heterodimerizing Id proteins (Kee, Quong et al., 2000)
Ets family
mA
Pos.
Wide
Many family members; usually require association with other proteins for activity (Nikolajczyk, Sanchez et al., 1999)
Fos/Jun
AP1
Pos.
bZip
Wide
AP-1, often inducible with mitogens; negatively regulated by JunB (Shaulian and Karin, 2002)
IRF4
NF-EM5 lB
Pos.
IRF
Lymphoid, Plasma cells
Associates with PU.1 and E2-A proteins (Eisenbeis, Singh et al., 1995)
Maf/Bach2
MARE
Neg.
bZip/
Bach2 in early B cells
Heterodimer negatively regulates transcription (Muto, Hoshino et al., 1998)
Mef2
lA
Pos.
MADS
Ubiquitous
Large family, some important for myocytes (Satyaraj and Storb, 1998)
MiT
mE3, kE3
Pos.
bHLHzip
Wide
TFE3, TFEB or USF; homo- or heterodimerize; association of TFE3 with ets proteins; possible enhancer–promoter interactions (Rehli, Den Elzen et al., 1999)
NF-kB/rel
kB
Pos.
Rel
Wide
Heterodimers activate, IkB proteins regulate nuclear localization and respond to many signaling pathways (Li and Verma, 2002)
Oct1/2
Oct
Pos.
Pou/hox
Wide
Require association with the B-cell specific coactivator OCA-B (Matthias, 1998)
Pax5
BSAP
Pos/Neg.
B cells, not
Activity depends on gene context plasma cells (Nutt, Eberhard et al., 2001)
PU.1
mB, kB, lB
Pos.
Ets
B cells, Myeloid
Usually requires association with another protein such as IRF4 (Singh, Dekoter et al., 1999)
YY1
mE1, kE1
Pos./Neg
Zn finger
Ubiquitous
Recruits enzymes that modify histone acetylation; associates with many other proteins (Thomas and Seto, 1999)
ZEB
mE5
Neg.
Zn finger
Ubiquitous
Competes with E2A proteins for binding (Genetta, Ruezinsky et al., 1994)
opmental stage-specific protein. Some mechanisms contributing to the lineage and stage-specific activity of these elements are discussed below. Dimerizing Proteins with Different Partners Several families of transcriptional activators bind DNA as obligate dimers, providing the opportunity for shortened forms to act as dominant negative regulators by forming nonfunctional heterodimers. For example, the bHLH proteins encoded by E2-A, HEB, and E2–2 are negatively regulated by Id proteins, encoded by four genes (Id1–4) (Engel and Murre, 2001). The shorter HLH Id proteins lack both an activation domain and a basic region. Thus, Id/bHLH heterodimers fail to bind DNA and cannot activate transcription. Regulated expression of Id proteins is important for regulating E2-A, HEB, and E2–2 activity during B cell development (Sun, Copeland et al., 1991; Barndt and Zhuang, 1999; Becker-Herman, Lantner et al., 2002).
C/EBPb, an important activator of Ig promoters and enhancers, is a bZip protein that binds DNA as an obligate dimer. LIP, a shorter form of C/EBPb that lacks an activation domain, is generated by alternate translation initiation (Descombes and Schibler, 1991). A similar shorter form is also encoded by a separate gene, C/EBPg (Roman, Platero et al., 1990). Both short forms act as dominant negative inhibitors by forming DNA-binding heterodimers that cannot activate transcription. Both C/EBPb and the dominant negative short forms are regulated during B cell development, suggesting that activity of their binding sites is determined by both absolute and relative levels of these proteins. A similar situation has been described for the bHLHZip protein TFE3, wherein differential RNA splicing creates a truncated form that acts as a dominant negative in heterodimers with full-length proteins (Roman, Cohn et al., 1991). The Maf family of bZip proteins also contains both activating and short, nonfunctional forms. However, for this
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family an additional twist is present in B cells where a small, nonfunctional Maf protein heterodimerizes with a bZip protein called Bach2 that represses transcription (Muto, Hoshino et al., 1998). Bach2 has B-cell and neuron-specific expression; it is present in most B cells but absent in plasma cells and represses transcription by association with the corepressor SMRT. Different Proteins Binding to One Site Bright (B-cell-restricted regulator of IgH transcription) binds to the A/T rich sequences present in a subset of MARs, including those present in Em (Webb, Zong et al., 1999). Bright levels vary at different stages of B cell development and it is absent in plasma cells. Although it can activate transcription in some artificial settings, in vivo Bright may be important for chromatin remodeling early in B cell development. Interestingly, in non-B cells where Bright is absent, a negative regulator, Cux/CDP, binds to the Em MAR sites (Wang, Goldstein et al., 1999). Cux/CDP is absent in B cells, and it has been suggested that switching from Cux/CDP to Bright provides a MAR-mediated switch for Em activity. bHLH proteins encoded by the E2-A, HEB, and E2–2 genes bind sites in Ig enhancers (Figures 6.1 and 6.2). A “two-handed” zinc finger protein called ZEB has also been shown to bind the E5 site in Em in non-B cells (Genetta, Ruezinsky et al., 1994). ZEB is a transcriptional repressor that is ubiquitously expressed. The reason bHLH activator proteins overcome ZEB repression in B cells is intriguing, but incompletely understood at present. Octamer Proteins The striking conservation of oct sites in both heavy chain and light chain V gene promoters and the presence of oct sites in Em and heavy chain 3¢ enhancers, suggested that oct binding proteins might be critical for B-cell specificity of Ig transcription. However, despite extensive studies, the roles of oct sites and the proteins that bind them remains murky (Matthias 1998; Bertolino, Tiedt et al., 2000). Two octbinding proteins are present in B cells: Oct-1, which is ubiquitously expressed, and Oct-2, restricted to lymphoid and central nervous sytems cells. Both proteins interact with a B-cell specific co-activator, OCA-B, suggesting a way in which oct-dependent activation could be B-cell specific. However, B cell development and IgM secretion are normal in mice lacking OCA-B, whereas expression of secondary isotypes and the entry of B cells into peripheral pools is defective (Kim, Qin et al., 1996; Nielsen, Georgiev et al., 1996). Thus, at a minimum, OCA-B does not confer nonredundant B-cell-specific regulation on Ig transcription in vivo. It is also possible that another B-cell specific coactivator, like OCA-B, may exist. Oct sites are most important functional elements in V gene promoters and altered speci-
ficity mutants (Shah, Bertolino et al., 1997), and knock-out mice (Schubart, Massa et al., 2001) suggest that Oct-1 is more important in this context than Oct-2. Lineage and Stage Specificity of Ig Enhancers via Regulation of PU.1 and Pax5 PU.1 is an ets family protein, expressed in hematopoietic cells, which appears to be important for the B-cell specificity of Em (Nelsen, Tian et al., 1993; Shaffer, Peng et al., 1997). PU.1 also binds to kappa and lambda enhancers (Figure 6.2) and in the kappa 3¢ enhancer occupation of the PU.1 site, detected by in vivo footprinting, correlates with pre B cell, but not pro-B cell activity of the enhancer (Shaffer, Peng et al., 1997). Pax5, also called B cell lineage specific activator protein (BSAP), has a B-cell specific expression pattern but is not present in plasma cells (Nutt, Eberhard et al., 2001). Pax5, along with YY1, is a transcriptional regulator that can either activate or repress transcription, depending on the gene context of its binding site. In the enhancers HS1,2 and HS 4 3¢ of Ca, and in the 3¢ kappa enhancer, Pax5 appears to repress enhancer activity. Thus, it is likely that the decreased expression of Pax5 in plasma cells is important for the high activity of these enhancers in terminally differentiated B cells.
Cooperative Interactions Are Important for the Activity of Many DNA Binding Proteins That Regulate Ig Transcription Our current understanding of transcriptional activators that bind in promoter regions near the start of transcription is that they recruit, either directly or via co-activators and/or chromatin remodeling machines, components of the basal transcription machinery to form a stable transcription initiation complex. Ig promoters, however, are very simple and most transcriptional regulatory elements in Ig genes reside in enhancers (Figure 6.1 and 6.2). The molecular mechanism(s) by which these elements activate transcription from distances of several kilobases remains an intriguing puzzle. The question is further complicated by the complexity of most Ig enhancers. Many protein binding sites exist and complicated patterns of both functional redundancy and functional cooperativity have been observed in transfection studies. Ets Family Proteins and Their Partners PU.1 is an ets family protein that preferentially binds mB or lB sites in Em, the 3¢ kappa enhancer and the lambda enhancers (Figure 6.1 and 6.2). In the lB and 3¢ kappa enhancer sites, PU.1 associates with a lymphoid-restricted
6. Transcription of Immunoglobulin Genes
IRF family protein, IRF4, and activates transcription (Eisenbeis, Singh et al., 1995). Further study indicates that PU.1 in this context may play an architectural role in recruiting IRF4, which actively promotes transcription (Pongubala and Atchison, 1997). In Em, a tripartite region containing mA, mE3, and mB is sufficient to activate transcription in B cells (Nelsen, Tian et al., 1993; Nikolajczyk, Cortes et al., 1997), and the spacing of these three sites is important for their activity, due at least in part to the ability of PU.1 to bend DNA (Nikolajczyk, Nelsen et al., 1996). The Mi-T bHLHZip protein TFE3 cooperates with PU.1 and Ets-1 to activate transcription dependent on this region (Tian, Erman et al., 1999). Similar to the situation in light chain enhancers, the transactivation domain of PU.1 is not important for cooperative transcriptional activation (Erman and Sen, 1996). Other Enhancer-Binding Proteins and Cooperativity Many other examples of cooperative interactions among Ig transcription factors have been reported. IRF4 interacts with the E2-A proteins E12 and E47 to activate transcription from the 3¢ kappa enhancer (Nagulapalli and Atchison, 1998). C/EBPb associates with Oct1 and Oct2 in solution and forms a ternary complex on Ig heavy chain and kappa promoters, implying functional cooperativity (Hatada, Chen-Kiang et al., 2000). Functional synergy has been shown for the bHLHZip protein TFE3, binding in Em at the mE3 site with both Ets-1 and bHLH protein E47 (Nikolajczyk, Dang et al., 1999).
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transgene expression in single cells revealed that, in fact, enhancers do not increase the rate of transcription initiation, but instead increase the probability that transcription will initiate at a given promoter without affecting the rate of transcription initiation (Walters, Fiering et al., 1995; Fiering, Whitelaw et al., 2000). For enhancers in Ig genes it is important to remember that in addition to activating the transcription of V gene promoters, they are also required for VDJ recombination. Although it is not clear if the same DNA-binding proteins and the same mechanism(s) are involved in their effects on transcription and their effects on DNA recombination, it is reasonable to assume some or most may be in common. Tracking How do enhancers increase the probability of transcription initiation when they are often located significant distances from their target promoters? One model suggests some form of a tracking mechanism, by which activator proteins are recruited to enhancers and then track along the DNA until they encounter a promoter, at which point they act to facilitate transcription initiation. This idea is consistent with, but not proven by, the finding that some DNA elements (called insulators) can block enhancer activity when placed between an enhancer and a promoter (Felsenfeld, Boyes et al., 1996). However, neither the mechanism of insulators nor enhancers are known, and they may or may not involve protein tracking on DNA. Looping
AREAS OF CURRENT RESEARCH Mechanism(s) of Enhancer Action In all the Ig loci, transcriptional enhancers located several kilobases 3¢ to the V gene promoters activate the promoters and are critical for regulated gene expression. In spite of much effort, we still do not understand the molecular mechanism(s) by which enhancers in the Ig loci, or in any mammalian gene, actually work. Indeed, because there is little consensus on what an enhancer does, we discuss below some of the possibilities that have been considered. It is likely that enhancers associated with different loci will incorporate one or more of these possible mechanisms depending on the regulatory requirements of the locus. Probability of Transcription Initiation Enhancers increase the amount of transcription initiation at target promoters, and it was originally believed that they did this by increasing the rate of transcription initiation at each promoter. However, analysis of enhancer-dependent
Alternatively, looping models show that proteins bound at enhancers directly associate with proteins bound at the promoter to facilitate transcription initiation. Since in vivo enhancer-dependent transcriptional activation occurs in the context of chromatin, this “looping” of DNA may actually involve or depend on the remodeling of chromatin. Consistent with this idea, in Escherichia coli enhancer activity requires DNA supercoiling (Liu, Bondarenko et al., 2001), and in in vitro transcription reactions using mammalian genes, enhancer-dependent transcription requires a chromatinized template (Barton and Emerson, 1994). Past studies on Ig genes have addressed the issue of enhancer–promoter interactions using transfection assays, and enhancers have robust activity in these assays. However, it is important to remember the limitations of such systems, especially since chromatin structure and nuclear sublocalization may not be faithfully recapitulated in these systems. Several proteins have binding sites in both VH promoters and Em and, when isolated protein binding sites were tested for their ability to activate transcription from a distance, TFE3, but not C/EBPb or octamer proteins were able to mediate activation (Artandi, Cooper et al., 1994). This finding sug-
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gested that the self-association of bHLHZip dimers might be important for enhancer–promoter interactions. Similarly, more recent studies have shown that Bright, which binds to MARs associated with both VH promoters and Em, can selfassociate and might mediate long-range enhancer–promoter interactions in vivo (Webb, Zong et al., 1999). Enhancer–promoter interactions may also involve associations between different proteins bound at the enhancer and the promoter, although to date there is no direct evidence for this. However, distant enhancers cannot activate transcription in the absence of at least one proximal transcriptional activator and a recent study has explored this observation in the context of an Ig core promoter. The data show that a truncated octamer protein, containing only the POU domain, is sufficient to mediate activation through a distant enhancer (Bertolino, Tiedt et al., 2000). The POU domain binds DNA and recruits TBP to the TATA box. The ability of this minimal protein to mediate enhancer activity suggests that initial recruitment of TFIIB is both independent of the enhancer and required for enhancer activity. Subsequently, RNAPolII/mediator complexes recruited to the distant enhancer, may, via looping, mediate the assembly of the complete transcription initiation complex. The nature of protein–protein associations involved in the enhancer– promoter interaction are not elucidated in this paper, but the data open the possibility of associations between enhancer–bound proteins with TFIIB as well as activators bound at the promoter. Chromatin Structure Another model for enhancer activity, which is not mutually exclusive with either tracking or looping mechanisms, is that enhancers block gene silencing by preventing the localization of a gene to centromeric heterochromatin (Francastel, Walters et al., 1999). Many proteins that bind in Ig enhancers are “classical” transcription factors that often bind in the promoter elements of other genes, and few features distinguish proteins that act proximally versus those that act distally. However, two Ig enhancer binding proteins have a unique activity that may have important implications for enhancer activity. The bHLH E2-A proteins and the ets family protein PU.1, when overexpressed in T cells, are capable of inducing Em activity, evidenced by sterile mu
transcripts (Choi, Shen et al., 1996; Nikolajczyk, Sanchez et al., 1999). This has been interpreted as indicating that these proteins are capable of binding to nucleosomal DNA and activating silent chromatin, although overexpression experiments may not replicate in vivo conditions and could be misleading. In the context of models in which enhancers act by changing chromatin structure or by facilitating the localization of the gene to particular regions within the nucleus and thus affecting chromatin structure, it is important to consider the activity of the matrix attachment regions (MARs) that are associated with many Ig enhancers. In Em, the MARs appear to be important for extending chromatin activity for transcription over long distances (Jenuwein, Forrester et al., 1997), and their activity includes blocking DNA methylation and extending the domains of histone acetylation (Forrester, Fernandez et al., 1999; Fernandez, Winkler et al., 2001). Interestingly, the MARs do not appear to be required for Em to activate VDJ recombination (Sakai, Bottaro et al., 1999), suggesting differences in the mechanism of action of Em in transcription and VDJ recombination. Certainly chromatin structure and subnuclear localization are likely to be important for enhancer activity. However, the activity of enhancer elements in transient transfection assays, in which the chromatin structure of the transfected DNA only partially resembles endogenous chromatin and in which nuclear localization is unlikely to recapitulate that of endogenous loci, suggest that these features may not be entirely responsible for enhancer activity. The challenge for future experiments will be to develop assays in which enhancer activity can be systematically dissected in a context wherein the genes are in their physiological context with respect to location on the chromosome, subnuclear localization of the chromosome, and chromatin structure. Activation of the IgH Locus for Rearrangement and Transcription In its germline (unrearranged) state, the immunoglobulin heavy chain gene locus spans approximately 2.5 to 3 Mb close to the telomere of the short arm of murine chromosome 12 (Chevillard, Ozaki et al., 2002) (Figure 6.3). Approximately 1.5 to 2 Mb of this comprises multiple VH gene segments, the sixteen DH gene segments are spread
FIGURE 6.3 Schematic representation of the IgH locus including VH gene segments and all heavy chain isotypes, showing some approximate distances, not drawn to scale, on the top. Although some VHJ558 genes are interspersed with other families, the VHJ558 family is the most DH-distal and VH 7183 family is the most DH-proximal VH gene family. DFL16.1 and Dq52 are the 5¢- and 3¢- most DH gene segments, respectively.
6. Transcription of Immunoglobulin Genes
over 40 kb, and the JH/Cm/Cd region extends another 10 to 15 kb. This part of the locus is activated during antigenindependent B cell differentiation in the bone marrow. The first gamma isotypes lie approximately 50 kb 3¢ of Cd, followed by the other isotypes spread over 100 kb, which culminate in Ca. Activation of the IgH locus for rearrangement and expression has been and continues to be studied extensively, serving as a paradigm for understanding the activation of all Ig loci. Gene Rearrangement and Transcription Transcription of unrearranged (germline) Ig gene segments precedes both VDJ recombination and class switch recombination (CSR), suggesting that transcription and/or transcriptional control elements play a role in regulating these two critical DNA rearrangements. Deletion of Em inhibits VDJ recombination, with a greater effect on VDJ than DJ recombination (Serwe and Sablitzky, 1993). Surprisingly, only the core of Em is necessary, and the MARs are dispensable (Sakai, Bottaro et al., 1999). In the kappa locus, both the intronic and 3¢ enhancers are important for VJ recombination (Inlay, Alt et al., 2002). Both Em (Sakai, Bottaro et al., 1999) and the enhancers 3¢ of Ca (Cogne, Lansford et al., 1994) appear to be important for CSR. Understanding how these transcriptional control elements function to control DNA recombination is an area of intensely active study. Since the molecular mechanisms responsible for VDJ recombination and CSR are discussed in detail elsewhere in this volume, we will not detail these studies here. However, models for enhancers’ role in these DNA rearrangements include: 1) altering chromatin structure to make gene segments more accessible to recombinase machinery, 2) activating transcription which, either via the process itself or via the mRNA produced, is required for the process of recombination, or 3) recruiting proteins directly involved in recombination. Activation of the DH-Cm Region Acetylation of lysine residues at the N-termini of histones H3 and H4 has recently emerged as a marker of activated regions of the genome (Workman and Kingston, 1998). Genes that are transcriptionally active, or those that are poised to be transcribed, are associated with acetylated histones and can be assayed by immunoprecipitating DNA/protein complexes using antimodified histone antibodies and scoring for the gene of interest by the polymerase chain reaction (PCR). Analysis of the unrearranged IgH locus by this assay shows that the locus is activated in discrete, independently regulated steps during B cell differentiation. The first domain of hyperacetylation is approximately 90 to 100 kb and includes all the DH gene segments, the JH gene segments, and the Cm exons. It is
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likely that DH to JH recombination is initiated within this domain (Chowdhury and Sen, 2001). Because the VH regions are hypoacetylated at this stage, and therefore inactive by the criteria described above, these observations also provide a simple explanation for DH to JH recombination preceding VH to DJH recombination. There are several additional features of this domain. First, only two closely located DNase 1 hypersensitive sites have been identified within this region (Chowdhury and Sen, 2001). One marks the IgH m intron enhancer and the second a region, close to the 3¢ most DH gene segment, Dq52. The latter probably marks a sterile promoter. The region between the two hypersensitive sites contains the four JH gene segments and is immunoprecipitated more efficiently with antiacetylated histone antibodies than flanking sequences. These observations suggest that the JH cluster is contained within a microdomain of increased histone acetylation, which may play a role in targeting the V(D)J recombinase to this part of the locus. Interestingly, the JH gene segments are located asymmetrically within the 90 kb domain; the 3¢ end of the domain ends abruptly within 10 kb 3¢ of the JHs between the Cm and Cd exons, whereas the domain extends at least 60 kb 5¢ to include even the most distal DH gene segment, DFL16.1. It is possible that the short 3¢ extension minimizes abortive scanning of the genome 3¢ to JHs by the recombinase, where no other recombinogenic gene segments exist. The nature of the domain boundary between Cm and Cd is unclear. The few boundary elements and insulators that are known are marked by DNase1 hypersensitive sites. However, no hypersensitive site exists between Cm and Cd, suggesting that this boundary may be generated by a different mechanism. The Em has been shown to activate V(D)J recombination in engineered substrates. Yet deletion of the enhancer has no effect on DH to JH recombination (Sakai, Bottaro et al., 1999), although VH to DJH recombination is severely diminished. The identification of a second Dq52 hypersensitive site suggests that this element may provide the requisite recombinational enhancer activity, in the absence of Em, to allow DH to JH recombination. This region has also been deleted from the genome (Nitschke, Kestler et al., 2001). Unlike Em, however, this mutation permits both DH to JH as well as VH to DJH recombination. Thus, while each element may substitute for the other to activate DH to JH recombination, Em is uniquely essential for the second step of IgH gene assembly. All DH gene segments are not marked by a proximal hypersensitive site. No such sites were found in 10 kb spanning DFL16.1 and DSP2.2 gene segments. Because these gene segments recombine efficiently, it is unlikely that a closely associated hypersensitive site is required for recombination. However, it cannot be ruled out that there are other regulatory sequences within the DH region that contribute to activating the 90 kb domain.
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VH Gene Activation Histone acetylation and other studies suggest that the VH locus contains at least two independently regulated domains (Chowdhury and Sen, 2001; Johnson, Angelin-Duclos et al., 2003). The largest region comprises the telomere-proximal VHJ558 and VH3609 gene families that are activated by interleukin-7 (IL-7) in adult pro-B cells. Both H3 and H4 are acetylated; however, the acetylation is limited to about 1 kb surrounding each gene segment and does not extend into the intergenic regions (Johnson, Angelin-Duclos et al., 2003). These genes make up more than half of all VH genes. The DH/Cm-proximal VH genes, including 7183, S107, VGAM, V10, and SM7 families, are not highly acetylated in adult pro-B cells prior to DJH recombination, but become hyperacetylated in cells that contain DJH joins (Chowdhury and Sen, 2001). Although the connection is correlative, this observation raises the interesting possibility that DJH recombination itself may trigger the next step of IgH recombination. An obvious, but untested, mechanism could be that DJH joining brings the 3¢ VH genes closer to, and therefore under control of, Cm-proximal regulatory sequences such as Em (the Dq52 element being deleted by any DJH recombination other than Dq52 itself). This region of low acetylation extends approximately halfway into the VH locus and includes some DH-proximal J558 genes. A characteristic feature of IgH gene assembly, which is most obvious during fetal development, is that 3¢ VH genes, such as VH7183 family members, recombine preferentially in early B cell ontogeny (Yancopoulos, Desiderio et al., 1984; Jeong and Teale, 1989; Malynn, Yancopoulos et al., 1990; ten Boekel, Melchers et al., 1997). In the fetal liver, these genes, as well as the more DH-distal VH genes, are associated with acetylated H4 following culture in IL-7, but prior to DJH recombination (Johnson, Angelin-Duclos et al., 2003). These observations highlight a basic difference in the mechanisms that activate VH genes in the fetus versus the adult. Interestingly, the absence of Pax 5 also causes a complete loss of VH to DJH recombination in the fetal liver, but only a loss of JH-distal VH gene recombination in the adult (Nutt, 1997; Hesslein, Pflugh et al., 2003). This also makes the case for differences in VH gene regulation in fetal and adult ontogeny. However, histone acetylation of proximal and distal VH genes is similar in Pax 5-/- and normal mice, suggesting that Pax 5 may regulate VH to DJH recombination at a step other than altering the state of histone acetylation (Hesslein, Pflugh et al., 2003). Differential activation of segments of the VH locus provides insight into the basis for ordered rearrangements in adult developing B cells. Two factors may contribute to the overall outcome. First, proximal VH genes may be activated early in response to DJH recombination, as suggested above. In addition, the differential IL-7 sensitivity of developing pro-B cells may delay activation of the large cluster of VHJ558 genes. Early pro-B cells express low levels of the
IL-7 receptor a chain and are generally less responsive to IL-7 (Marshall et al., 1998). As a result of weak IL-7 signaling, the distal VHJ558 genes may not be effectively activated early to compete with the proximal genes for recombination. The net result is that proximal VH genes recombine early and the distal genes recombine later. Thus, preferential rearrangement of VH7183 family is the result of independent control of different parts of the VH locus and the complex pattern of IL-7 sensitivity of developing B cells. However, much work remains to fully understand the mechanisms and signals that differentially control histone acetylation and VH gene rearrangements. Nuclear Sublocalization Developmentally regulated changes in IgH locus chromatin structure are accompanied by alterations in the nuclear organization of the locus. Three kinds of changes have been noted. In non-B lineage cells, such as thymocytes or ES cells, both IgH alleles are located close to the nuclear periphery (Kosak, Skok et al., 2002). In pro-B cells IgH alleles were found to be more centrally located in the nucleus and away from centromeric DNA regardless of the rearrangement status. Centromeric heterochromatin has been implicated in keeping genes turned off, and these observations are consistent with both alleles being simultaneously active for transcription and recombination. The state of the locus is not permanent, however, because one allele co-localizes with centromeric DNA in mature splenic B cells activated to enter the cell cycle (Skok, Brown et al., 2001). Singh and colleagues also made the intriguing observation that in-situ hybridization signals from two ends of the VH locus were closer together in T cell nuclei (where it is peripherally located) compared to pro-B cell nuclei; they suggested that the decreased compaction in pro-B cells may reflect some aspect of recombination control or may facilitate VH to DJH rearrangements (Kosak, Skok et al., 2002). Third, a correlation has been noted between replication pattern of the IgH locus and its nuclear location (Zhou, Ermakova et al., 2002). Pre- and pro-B cell lines that replicate IgH early in the S phase localize this locus centrally in the nucleus, whereas mature B and non-B cells that follow a triphasic replication pattern localize the IgH locus to the nuclear periphery. Matrix Attachment Regions The core of the m heavy chain gene enhancer is flanked by matrix attachment regions (MARs). These A/T-rich sequences have been implicated in positive and negative regulation of Ig expression. Evidence for negative regulation stems from the observation that the tissue-range in which the m enhancer is active in transfection assays is increased if the flanking MARs are missing (Weinberger, Jat et al., 1988; Scheuermann and Chen, 1989). Conversely,
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expression of a functionally rearranged IgH transgene is significantly increased when MARs are included in the normal location flanking the enhancer (Forrester et al., 1994). Based on these studies, Grosschedl and colleagues have proposed that MARs help to propagate enhancer effects over long distances (Jenuwein et al., 1997; Fernandez et al., 2001). Recently, the IgH intron MARs have also been individually, or jointly, deleted from the genome. However, MAR deletion does not alter VDJ recombination or IgH gene expression from the altered allele (Sakai et al., 1999). The apparent discrepancy between transgenic and endogenous locus studies is best reconciled by considering that other MARs within the locus may compensate for the loss of the intronic MARs. If so, an interesting corollary is that position within the locus is not important for MAR function. Another MAR binding protein, SATB1, was identified based on its ability to bind A/T-rich sequnces that unwound easily upon torsional stress (also referred to as base unpairing regions, BURs) (Dickinson, Joh et al., 1992). SATB1 DNA binding in vitro is significantly diminished if the propensity to unwind DNA is weakened by appropriately placed mutations. Since many MARs contain BURs, the possible role of MAR binding proteins in stabilizing alternate DNA conformations should not be overlooked. In addition, both BRIGHT and SATB1 have features that underscore their importance in chromatin structure. For example, the BRIGHT DNA binding domain is similar to that found in SWI1 (a component of the chromatin remodeling complex SWI/SNF) and SATB1 is complexed to histone de-acetylases and nuclear co-repressors in cells. Genetic deletion of SATB1 inhibits T cell development and alters the structure of the interleukin-2 receptor alpha chain gene (Yasui, Miyano et al., 2002). Understanding how classical enhancers and MARs coordinately regulate gene expression remains a challenge for the future.
DISCOVERIES RESULTING FROM THE STUDY OF IG GENE TRANSCRIPTION Since studies on Ig gene transcriptional regulation were initiated early, when little was understood regarding mammalian gene regulation, and since Em was the first mammalian transcriptional enhancer to be identified, studies in this field have not only illuminated our understanding of Ig gene regulation, but have also established paradigms for understanding general mechanisms of transcriptional regulation. In addition, several regulatory proteins first identified and studied for their roles in Ig gene regulation upon further study have been found to play critical roles in the immune system unrelated to their regulation of Ig genes. Two of the most important “additional” discoveries are discussed below.
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Certain Ig Transcriptional Regulators Are Critical for Early Stages of Hematopoietic Cell Development The ets Family Protein PU.1 in Early Hematopoiesis Gene targeting studies originally revealed a requirement for PU.1 in the development of both myeloid and lymphoid lineages (Scott et al., 1994). Interestingly, in in vitro reconstitution studies, graded levels of PU.1 differentially regulate macrophage versus B cell differentiation, with higher levels being associated with macrophage development (DeKoter and Singh, 2000). PU.1 also appears important for mast cell and dendritic cell development (Singh et al., 1999; Anderson et al., 2000).
Pax5 and Commitment to the B Cell Lineage and B Cell Development Pax5 is an interesting transcription factor that can either activate or repress transcription, depending on gene context (Wallin et al., 1998). In hematopoietic cells, its expression is limited to the B lymphoid lineage. Cells lacking Pax5 are not committed to the B lineage (Nutt et al., 1999) and recently Pax5 has been shown to inhibit the Notch pathway, required for T cell commitment (Souabni et al., 2002). Thus, Pax5 has a unique role in lineage commitment. Roles for Pax5 also have been demonstrated during early B cell development and in the germinal center (Nutt et al., 2001). Like some other transcription factors, Pax5 is shut down during plasma cell differentiation, thus relieving the repression of genes such as J chain and XBP-1 that are expressed in Ig secreting cells (Schebesta et al., 2002).
E2-A Proteins in Early B Cell Development B cell development is arrested at the pro-B stage in mice lacking the E2A gene (Barndt and Zhuang, 1999; Kee et al., 2000). Furthermore, E12, encoded by E2-A, induces early B cell factor (EBF) (Kee and Murre, 1998), and together the EBF and E2-A gene products synergize in early B cell development (O’Riordan and Grosschedl, 1999).
NF-kB/rel Proteins Are Important in Many Immune and Inflammatory Processes NF-kB/rel proteins were first discovered because of their binding to the kB site in the Igk intronic enhancer. However, it soon became obvious that this family of transcriptional regulators plays an important role in many other immune cells, as well as in B cells. NF-kB/rel proteins exist in inactive forms in the cytoplasm of most cells and, in response to a wide variety of signals, they rapidly become activated and enter the nucleus to affect expression of target genes.
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These proteins are important in a wide range of diseases ranging from inflammation to cancer (Yamamoto and Gaynor, 2001; Li and Verma, 2002). They are critical for inflammatory responses and have an anti-apoptotic role in both normal and malignant cells. They are critical for normal splenic architecture, B cell survival and B cell-dependent immune responses (Caamano, Rizzo et al., 1998; Franzoso, Carlson et al., 1998), dendritic cell survival and differentiation (Ouaaz, Arron et al., 2002), and T cell survival following activation (Wang, Guttridge et al., 1999). The family of Rel homology domain (RHD)-containing proteins consists of p105/NFkB1 (the precursor to p50), p110/NFk2 (the precursor to p52), p65/RelA, c-Rel, and RelB. Most family members can homo- or heterodimerize to produce transcription factors that recognize the DNA sequence GGG(A/T)4CCC, often referred to as a kB element (for recent more comprehensive reviews see Chen and Ghosh, 1999; Li and Verma, 2002). Of the various proteins, the term NF-kB usually refers to the p50/p65 heterodimer, which is usually the most abundant form of the factor detected by electrophoretic mobility shift assays. The approximately 300-amino acid RHD is sufficient for dimerization, DNA binding, nuclear localization, and association with a family of regulatory proteins called inhibitors of NFkB (IkB) (Ghosh and Karin, 2002). X-ray crystallographic structures of several Rel proteins reveals a novel DNA binding motif utilizing loops that protrude from more defined secondary structures (Chen, Huang et al., 1998; Chen, Ghosh et al., 1998; Huxford, Huang et al., 1998; Jacobs and Harrison, 1998; Huang, Chen et al., 2001). Cocrystals of Rel/IkBa complexes show extensive contacts between the subunits and conformational alteration of the N-terminus of the RHD that contains most of the DNA binding residues. Structural similarity of the RHDs is reflected in their recognizing closely related DNA sequences, such that it is virtually impossible to ascertain the functional Rel protein from the sequence of the kB-like element in a gene. However, mice deficient in Rel genes show different phenotypes, indicating that even very similar Rel genes serve distinct functions in-vivo (Beg, Sha et al., 1995; Sha, Liou et al., 1995; Weih, Carrasco et al., 1995; Kontgen, Grumont et al., 1995). Other than p50-/p65-mice, analyses of double-deficient Rel mice are ongoing (Grumont, Rourke et al., 1998; Grossmann, Metcalf et al., 1999; Gugasyan, Grumont et al., 2000; Grumont, 1998; Pohl, Gugasyan et al., 2002). Regulation by Subcellular Localization via IkB Proteins In most cells, Rel/IkB interactions were proposed to sequester the complex in the cytoplasm by hiding the nuclear localization sequence (NLS) of the Rel protein. That the mechanism was more complex was first revealed by the
crystallographic structure of the p65/IkBa complex in which the p65 NLS was not obscured by IkBa. Furthermore, because IkB proteins did not interact with p50 or p52, heterodimers containing these subunits would be expected to have at least one available NLS for nuclear import. Recent studies show that cytoplasmic localization by IkB proteins is a dynamic process. In particular, IkBa contains a very strong nuclear export sequence (NES) located in the Nterminus of the protein (Johnson, Van Antwerp et al., 1999; Huang, Kudo et al., 2000; Tam, Lee et al., 2000). This NES interacts with the nuclear export receptor CRM1, which directs IkBa and any associated proteins out of the nucleus. That IkBa-associated proteins are in constant flux is best visualized by treating cells with the drug leptomycin B (LMB), which blocks CRM1-dependent export. In LMBtreated cells, IkBa and associated Rel proteins accumulate in the nucleus. Similar results are observed in yeast with exogenously introduced p65 and IkBa proteins in a strain with a hypomorphic mutation in the yeast crm1 gene (Tam, Lee et al., 2000). Finally, increased nuclear distribution of Rel/IkBa complexes is observed when the IkBa NES is mutated (Johnson, Van Antwerp et al., 1999; Huang, Kudo et al., 2000; Tam, Lee et al., 2000). Taken together, the combined genetic and pharmacological experiments suggest that Rel/IkBa complexes are continuously shuttling between the nucleus and the cytoplasm. The net cytosolic location observed in earlier studies is therefore the result of nuclear export dominating over nuclear import; an imbalance in the import–export equilibrium, such as that created by LMB, results in the net subcellular redistribution of the complexes. This mechanism of cytoplasmic localization also provides a ready explanation for the availability of functional NLSs in Rel/IkBa complexes. In contrast to IkBa, IkBb and IkBe do not contain strong NESs and also interact more closely with Rel proteins in the vicinity of the NLS (Malek, Chen et al., 2001; Tam and Sen, 2001). Thus, these molecules probably truly sequester Rel proteins in the cytoplasm, as envisaged earlier for all IkBs. One of the benefits of the dynamic mechanism may be that the same properties of IkBa that mediate cytosolic localization in unactivated cells can also be used to restore cells to a resting state after termination of an activating signal. NF-kB induction by diverse stimuli leads to IkBa gene transcription and new protein synthesis. The newly synthesized IkBa can enter the nucleus (Chiao, Miyamoto et al., 1994; Arenzana-Seisdedos, Thompson et al., 1995) either by passive diffusion or aided by a nonclassical NLS (Sachdev, Hoffmann et al., 1998), disrupt Rel/DNA complexes and export Rel/IkBa complexes out to the cytoplasm to await retriggering by another signal. Indeed, continued signals result in cyclical Rel protein expression in the nucleus due to the dynamics of retrieval and reinduction in the cytoplasm (Hoffmann, Levchenko et al., 2002). It is unclear whether transit of IkBa-containing complexes through the nucleus
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serves additional biological function. An untested intriguing possibility remains that nuclear kinases or phosphatases may participate in NF-kB regulation, for example in response to nuclear inducing signals such as DNA double strand breaks. In this regard, it is noteworthy that DNA-dependent protein kinase (DNA-PK) has been shown to phosphorylate IkBa in vitro (Liu, Kwak et al., 1998), and NF-kB activation is diminished in ATM-deficient mouse embryo fibroblasts (Li, Banin et al., 2001). Signaling to Activate NF-kB/rel Proteins Only B lymphocytes contain nuclear Rel proteins that bind DNA (that is, are not complexed to IkBs) prior to any activating signal. This activity consists largely of p50/c-Rel heterodimers and lower levels of p50–p65 heterodimers (Liou, Sha et al., 1994; Miyamoto, Schmitt et al., 1994). Increased IkBa turnover has been proposed as the basis for constitutive nuclear NF-kB in B cells (Miyamoto, Chiao et al., 1994). However, the mechanism of IkBa turnover remains unclear. Miyamoto and colleagues have shown that constitutive IkBa degradation is insensitive to proteasome inhibitors and may be mediated by calpainlike proteases (Fields, Seufzer et al., 2000; Shen, Channavajhala et al., 2001). Further studies are required to identify the features of IkBa that target it for increased basal turnover in B cells. The dominance of nuclear p50 and c-Rel heterodimers has been proposed to be due to inefficient export of these complexes from the nucleus (Tam, Wang et al., 2001). This model is based on two observations: that p65/RelA contains an NES in its C-terminal domain, and that c-Rel/IkBa complexes are only found in B cells. Enhanced IkBa degradation in B cells thus creates nuclear pools of both p65 and c-Rel containing homo- and heterodimers. However, the p65 NES leads to more efficient export of p65-containing complexes, with the result that c-Rel containing complexes, accumulate in the nucleus. The central feature of the NF-kB family is its inducible activation to a nuclear DNA binding form by multiple signals. This occurs by signal-induced phosphorylation of IkB proteins at two conserved serine residues within the Nterminal domain. This domain is sometimes also referred to as the signal receptor domain. Phosphorylation of IkBs marks them for proteasome-mediated degradation. Released from the inhibitory influence of IkB proteins, DNA-binding Rel dimers translocate to the nucleus to activate gene expression. IkB phosphorylation is mediated by a heterotrimeric IkB kinase (IKK) complex that consists of IKKa, b, and g (Karin and Ben-Neriah, 2000; Karin and Delhase, 2000; Ghosh and Karin, 2002). IKKa and b are catalytic subunits that homo- or heterodimerize via leucine zipper–containing dimerization domains. Either kinase can phosphorylate IkBa in vitro at the appropriate residues, though IKKb usually appears to be the more active kinase.
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Indeed, a complex consisting of catalytically inactive IKKb with an active IKKa fails to activate NF-kB in response to most pro-inflammatory stimuli; in contrast, a complex of catalytically inactive IKKa with normal IKKb is functional under most conditions. Recently it has been shown that IKKa may play an essential role in NF-kB activation mediated by the cytokines RANKL and Blys in B cells (Cao, Bonizzi et al., 2001; Schiemann, Gommerman et al., 2001; Thompson, Bixler et al., 2001). No catalytic activity has been attributed to IKKg; rather, it is believed to serve as a scaffold that targets IKKa/b to the right substrates. However, IKKg is essential for NF-kB induction (Makris, Godfrey et al., 2000; Schmidt-Supprian, Bloch et al., 2000) and small peptides that disrupt IKKg interactions with IKKa/b inhibit NF-kB activation (May, D’Acquisto et al., 2000). Mutations in IKKg have also been implicated in human immunodeficiencies (Smahi, Courtois et al., 2000; Courtois, Smahi et al., 2001; Jain, Ma et al., 2001). Although the central importance of the IKK complex is well established, it is less clear how diverse stimuli converge at IKK. Catalytic activity of IKKa and b is induced by phosphorylation of two conserved serine residues in an activation loop present in each protein. Consequently, several “upstream” kinases have been implicated in IKK activation, although the physiological relevance of many of these remains to be established. Gene knock-out studies have verified the importance of two other kinases for NF-kB activation. The kinase RIP lies in the TNFR1 signaling pathway, and protein kinase C theta is necessary for NF-kB induction via the T cell receptor (Kelliher, Grimm et al., 1998; Sun, Arendt et al., 2000). Interestingly, catalytically inactive RIP can restore NF-kB activation, suggesting that it may play the role of an adapter (Hsu, Huang et al., 1996). Similar function has been attributed to another kinase, PKR, which is required for NF-kB induction by double-stranded RNA (Bonnet, Weil et al., 2000). Recently the CARDdomain–containing proteins Bcl10 and CARD11 have been shown to be essential for NF-kB induction by B- and T-cell antigen receptors (Gaide, Favier et al., 2002; Pomerantz, Denny et al., 2002; Wang, You et al., 2002). The connection of these (nonkinase) proteins to PKC theta or the IKK complex remains to be determined. The cytoplasm is likely to be the site of IkB phosphorylation since the IKK resides here. However, phospho-IkBs must be recognized by the bTrCP/SCF complex (Yaron, Hatzubai et al., 1998), which ubiquitinates IkB at a lysine residue also located in the N-terminal signal receptor domain (Spencer, Jiang et al., 1999; Winston, Strack et al., 1999). Poly-ubiquitinated IkB then is a target for the proteasome. In an intriguing twist to the compartmentalization problem of components involved in NF-kB activation, bTrCP/SCF has been shown to be predominantly nuclear (Davis, Hatzubai et al., 2002). If IkB phosphorylation only occurs in the cytoplasm, these observations suggest that
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phosphorylated IkB must translocate to the nucleus to find bTrCP/SCF. This is feasible for IkBa complexes because they shuttle through the nucleus, but more difficult to imagine for other IkBs. Other possibilities are that there may be a small but functionally relevant amount of bTrCP/SCF in the cytoplasm, or that this protein may also shuttle through the nucleus by a presently undefined pathway. Overall, all the emerging evidence serves to underscore the dynamic state of NF-kB regulation in resting as well as activated cells.
CONCLUSION The transcriptional regulation of immunoglobulin genes has been actively studied for more than twenty years. As reviewed in this chapter, we now have a good understanding of most transcriptional regulatory elements in these genes and appreciate the key roles of various enhancers. Families of DNA-binding transcriptional regulators, binding to individual sites in the promoter and enhancer elements, have also been identified and their mechanisms of action defined at least in vitro. In addition, as described above, these studies have illuminated transcription factors that play important roles in the early development of hematopoietic and lymphoid cells and in many other aspects of immune function. However, important questions remain. The most pressing is that the mechanism(s) by which enhancers activate transcription is still unknown. In the Ig loci, these enhancers also play a role in allowing DNA rearrangements; how this occurs and how it may relate to transcriptional activation also remains unknown. Related questions involve the relationship(s) between transcription, DNA rearrangement, and DNA replication, as well as the role of subnuclear localization in determining the activity of a gene. Recent advances in studying chromatin structure and how it may be regulated by histone modification and remodeling machines, chromatin immunoprecipitations to monitor the association in vivo of particular proteins with DNA sequences, and the ability to track the subnuclear localization and replication times of particular genes may help us finally unravel the remaining secrets of immunoglobulin gene regulation.
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7 Early B Cell Development to a Mature, Antigen-Sensitive Cell FRITZ MELCHERS
PAUL KINCADE
Department of Cell Biology, Biozentrum, University of Basel, Basel, Switzerland
Oklahoma Medical Research Foundation, Oklahoma City, Oklahoma, USA
The development of immunoglobulin (Ig)-producing B lymphocytes proceeds, like the development of any other cell lineage within a multicellular organism, through a series of developmental steps. These steps can be defined by cellular stages in which a selected set of genes of the total genome is expressed. The products of these genes function to control cell proliferation, migration and location, survival and apoptosis, and cellular differentiation to changed gene expression programs and changed cellular stages and functions. For the B lymphocyte pathway of development only very few genes are selectively and only expressed in that lineage and in no other cell lineage of the organism. First, and most important, the Ig heavy (H) and light (L) chain genes are assembled from V, (D), and J segments in a stepwise fashion during development. Next, the VpreB and l5 genes, are assembled, from which the surrogate light chain is assembled at selected precursor cell states. Then, from the Iga and Igb genes the molecules are made that anchor Ig molecules composed of H and L chains as B cell receptors (BCR) for antigen in the surface membrane of B cells. These anchor the pre-B cell receptors (pre-BCR), composed of H and surrogate L chain, on the surface of precursor B cells. In addition, CD19 and CD20 are so far the only other B lineage-specific genes that are not found expressed in other cell lineages. These are expressed on the surface of B cells and appear to function in concert with BCRs to control B lymphocyte responses to stimulation. All other genes expressed in B lineage cells can also be found expressed in other cell lineages, though in other combinations. This chapter describes the cellular pathways of B lymphocyte development from the earliest identifiable progenitors, with many options for different lineage decisions to the apparently highly specialized, Ig-synthesizing B cells committed
to one B lineage. It describes the molecular-genetic programs of these different cellular stages of development, as much as they are understood at present, and the functions they play in the many decisions that cells have to make to become a B cell. The ordered development predicts that each step along the way of differentiation is in a defined state and has a high probability to develop in only one way, in one direction. However, it will become evident that this apparent unidirectionality of development can be influenced by mutations from within and by environmental influences from without, revealing a plasticity of cellular states that allows alternate options of development. This may not be too surprising in view of the experimental observation that the nucleus of a fully differentiated B lymphocyte producing one set of H and L chains (i.e., one Ig molecule) can be introduced into an enucleated embryonic stem cell from which a whole organism, a mouse, can be developed again (Gurdon et al., 1975; Hochedlinger and Jaenisch, 2002). Therefore, the descriptions of the developmental pathways to the stage of mature, antigen-reactive B lymphocytes are always reflections of experimental measurements of the most probable molecular and cellular states but never the only possible states in these processes. They are, in a way, the manifestations of a “Heisenberg uncertainty principle” of biology (Graf, 2002).
Molecular Biology of B Cells
THREE WAVES OF HEMATOPOIESIS DURING EMBRYONIC DEVELOPMENT The mouse embryo is colonized by three waves of hematopoietic cell development (Ling and Dzierzak, 2002; Godin and Cumano, 2002) (Figure 7.1). The first originates
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FIGURE 7.1 Early stages of hematopoietic development to the lymphoid progenitors.
in extra-embryonic tissue (i.e., in the yolk sac) at day 7 to 7.5 of development. This so-called “primitive” hematopoiesis (showing similarities to hematopoiesis in lower vertebrates) generates, at apparently accelerated rates, “primitive” erythrocytes (which synthesize fetal hemoglobin with globin e and bH1 chains), megakaryocytes (with lower ploidy) and platelets, and myeloid cells (such as macrophages, with a special set of enzymes) (Cumano et al., 2001). Lymphocytes are not generated (Figure 7.1). Although these “primitive” blood cell lineages are developed at the original extra-embryonic, ventral site of the embryo, they migrate into the embryonic, dorsal sites as soon as blood circulation through the development of vascular endothelium is established at day 8 to 9 of development. A comparable development occurs in human embryos between days 13 and 24 after fertilization. At day 8.5 to 9 of murine embryonic development (days 25 to 30 in the human) the second wave of hematopoiesis is initiated within the intra-embryonic part of the embryo, more specifically within the anterior portion of the aorta-gonad-mesonephros (AGM) region (Medvinsky and Dzierzak, 1996; Cumano et al., 1996). Hematopoietically
undifferentiated, apparently pluripotent stem cells (Ohmura et al., 2001), which develop from the caudal intra-embryonic mesoderm near the AGM, then migrate within the next 2 to 3 days (in humans between days 30 and 40) of development through the blood and colonize the thymic rudiment and the fetal liver. At these sites the second wave of hematopoiesis, so-called “definitive” hematopoiesis, is initiated (Figure 7.1). This wave it generates “definitive” erythrocytes (which produce adult-type hemoglobin, with a and b major globin), “definitive” megakaryocytes and platelets, and myeloid cells. Fetal liver also produces a first, apparently synchronous wave of B lymphocytes (Strasser et al., 1989). The characteristic features of fetal liver-derived B cells and their differences from other B cells will become apparent at various points of this article. Another early site of B cell development, which generates B cells with similar properties to those of fetal liver, is the omentum (Kincade, 1981; Owen et al., 1975; Melchers, 1979; Solvason and Kearney, 1992; Strasser et al., 1989; Rolink et al., 1995). The third wave of hematopoiesis (e.g., the second of “definitive” hematopoiesis) is initiated in bone marrow
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
between days 17 and 19 of development, at around birth of the mouse (days 60 to 90 in human, hence far before birth). This is, in fact, a continuous process of hematopoiesis throughout life—a flood of hematopoiesis. Bone marrowderived B cells differ from fetal liver-derived ones (Kincade et al., 2002). A number of defective mutant mice, generated by gene targeting, have provided insight into these early steps of blood cell development. Genes that affect “primitive” hematopoiesis lead to death around days 8 and 11 of development, whereas those affecting only “definitive” hematopoiesis cause death between days 12 and 19. These genes occur in three groups: 1) those affecting “primitive” and “definitive” hematopoiesis by encoding the transcription factors TAL-1 (Shivdasani et al., 1996; Robb et al., 1995, 1996; Porcher et al., 1996), LMO2 (Warren et al., 1994; Yamada et al., 1996), GATA-2 (Tsai et al., 1994) and GATA-1 (Pevny et al., 1991, 1995), and the receptor tyrosine kinases Flk-1 (Shalaby et al., 1995, 1997; Schuh et al., 1999); 2) those genes affecting “definitive” hematopoiesis only by encoding the transcription factors AML-1 (Okuda et al., 1996), CBF-b (Sasaki et al., 1996), and EKLF (Nuez et al., 1995); 3) and those genes that influence migration or homing of pHSCs b1 and a4 integrins (Hirsch et al., 1996; Potocnik et al., 2000; Yang et al., 1995, Arroyo et al., 1996, 1999) and the interactions of the chemokine SDF-1 and its receptor CXCR4, expressed on hematopoietic precursors (Nagasawa et al., 1996, Egawa et al., 2001; Ma et al., 1998, 1999; Kawabata et al., 1999; Melchers et al., 1999) (Figure 7.1). Most of these mutations have been identified with blastocyst complementation assay (Chen, 1996). This assay does not distinguish between mutations that completely or only partially shut down development, since the RAG-deficient hosts have normal erythroid, megakaryocytic, and myeloid cell development and also generate lymphoid progenitors up to the cellular stages before V to DJ rearrangements. Hence, progenitors of the mutant mice must compete with those of the RAG-deficient hosts and often may simply be outgrown. The a4 and b1 integrins, and the chemokine SDF-1 and its receptor CXCR-4, appear to control by adhesion and chemoattraction the migration of the pHSC or their progenitors through the vascular endothelium into the bloodstream. Mutations in the Flk-1, TIE-2 (Takakura et al., 1996), and SDF-1 genes generate defects in the generation and functioning of the vascular endothelium, whereas mutations in CXCR-4 affect the proper homing of hematopoietic progenitors. In summary, these mutations highlight a requirements of early embryonic hematopoiesis: Progenitors cannot reach their sites in hematopoietic differentiation when blood vessels do not form or because the hematopoietic progenitor cells cannot adhere, or be chemoattracted (i.e., cannot migrate). These cell developments and migrations set the
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stage for B lymphopoiesis from pluripotent hematopoietic stem cells (pHSC).
PLURIPOTENT HEMATOPOIETIC STEM CELLS All these waves of hematopoiesis are expected to originate from pluripotent hematopoietic stem cells (pHSCs) (Jordan et al., 1990; Osawa et al., 1996a,b; Morrison et al., 1995; Spangrude et al., 1991; Smith et al., 1991).
Self-Renewal When pHSC divide, at least one daughter cell retains the property of a pHSC, while the other daughter cell may enter further stages of differentiation. Hence, they have the capacity of self-renewal. Lines of continuously proliferating, or symmetrically self-renewing pHSC have not been established in either mouse or human tissue culture. However, a number of factors have recently been identified that might eventually help to grow such lines for extended periods (Figure 7.1). First, in germline stem cells, self-renewal has been found to be specified by JAK-STAT activation (Kiger et al., 2001). It is likely that the receptor tyrosine kinase flt-3/flk-2, expressed on pHSC is stimulated by its ligand, which is expressed on microenvironmental stromal cells, to proliferate pHSCs (Gilliland and Griffin, 2002). The transgenic expression of the transcription factor HOXB4 in pHSC enhances their engraftment (as well as progenitor cells of pHSC, including ES cells) (Kyba et al., 2002) and their selfrenewal capacity (Buske et al., 2002). Growth factors that act in organogenesis during embryogenesis may be utilized in adult life to maintain stem cells. Among such embryonic growth factors are bone morphogenic proteins (BMP), Hedgehogs (Hhs), Wnts, NOTCH ligands, and fibroblast growth factors (FGF). It was found that BMP-4 induces ectodermal cells in the frog embryo to form blood (Maeno et al., 1996). The dominant-negative form of the BMP receptor abrogated this blood development (Graff et al., 1994), and mice deficient in BMP-4 are incapable of developing the mesoderm from which blood cells are formed (Hogan, 1996). Recently, Bhardwaj et al. (2001) found that Sonic Hedgehog (Shh) induces hematopoiesis in culture, whereas antibodies against Shh block it. Shh induces the formation of BMP-4 and an inhibitor of it, called Noggin. It appears that BMP-4 maintains pHSC, whereas Shh induces their proliferation. Hence, pHSCs are expected to have receptors for BMP-4 as well as for Shh (Zon, 2001) (Figure 7.1). Hematopoietic stem cells appear to control their proliferative expansion by signals that are connected with lnk, an adaptor protein. Hence, in lnk-deficient mice the number of hematopoietic progenitors and their proliferative capacities
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are significantly increased (Takaki et al., 2000, 2002). This increased lymphopoiesis is detectable in further differentiated subpopulations of the B lymphocytic lineage (Takaki et al., 2003). The proliferative, self-renewing capacity of pHSC might be quite extensive and might depend—apart from the proper environmental stimuli—on the capacity of pHSC to resynthesize telomeres as they shorten with every division, normally by 60 to 90 base pairs. Human cells with telomeres of 6-kb length would be able to divide 60 to 80 times (Hayflick, 1965), while mouse cells with 60-kb telomeres would do so for 600 to 800 divisions, unless telomerase is induced in pHSC to resynthesize the lost telomere length (Hodes, 1999; Alsopp et al., 1992; Greider, 1996) or unless the rate of telomere loss per cell division changes. LT pHSCs (see below) appear to have increased telomerase activity (Morrison et al., 1996), whereas serial pHSC transplants that increase pHSC cell cycle activity show a shortening of telomeres (Alsopp et al., 2001).
Migration and Homing Upon transplantation into a suitable host, pHSCs migrate to the proper sites in the body in which they find a supportive microenvironment for their survival, for the retention of their stem cell properties, and for self renewal (Wright et al., 2001). Some adhesion molecules and the chemokine–chemokine receptor interactions operative in this capacity of pHSCs have been described (Alsopp et al., 2001; Peled et al., 1999).
Pluripotency pHSCs can differentiate into all the blood cell lineages— into erythrocytes, megakaryocytes, and platelets; myeloid cells, such as monocytes and macrophages; dendritic cells; osteoclasts; granulocytes including eosinophils, neutrophils, basophils, and mast cells; NK cells; and into lymphoid cells of the T and B lineage. A single pHSC can be induced to differentiate to all these different cell lineages, hence pHSC are pluripotent. A vigorous documentation of this pluripotency is described below, using clones of PAX-5–deficient precursor cells of the B lineage. The influence of the microenvironment on a pHSC—the cytokines and cell contacts provided from the stroma to the hematopoietic cells—induces differentiation along different lineages of the blood cell system. Thus, erythropoietin helps to induce erythropoiesis, whereas thrombopoietin does so for the development of megakaryocytes and platelets. In concert with multilineage cytokines such as IL-3, IL-6, and stem cell factor (SCF), pHSC can be conditioned to be inducible by IL-6 and G-CSF to granulocytes (Liu et al., 1997). In the presence of M-CSF they differentiate to mono-
cytes and macrophages, while the presence of GM-CSF induces pHSC to dendritic cell differentiation (Inaba et al., 1992; Banchereau and Steinman, 1998). The cytokine TRANCE, normally presented on osteoblasts, induces pHSC to osteoclast formation (Kong et al., 1999), and IL-15 (in vitro also IL-2) stimulates NK cell development (Ogasawara et al., 1998; Rolink et al., 1996). Lymphoid cell development, at least in the adult mouse, is dependent on IL-7, but additional factors provided by the microenvironment of the thymus, such as delta-1 (Jaleco et al., 2001), are critically required to induce development of T-lineage lymphocytes. The environment of fetal liver and bone marrow must do so similarly for B lineage lymphocyte development.
Long-Term Reconstitution Potential pHSCs transplanted into a receptive host not only home to the primary sites of hematopoiesis in the adult the bone marrow, they also reside there for long periods, retaining their original properties of self-renewal, migration and homing, and pluripotency, which they possessed in the primary organism or donor. Hence, they are capable of longterm reconstitution and are called LT-pHSC. LT-pHSCs can be transgenically marked by green fluorescent protein, expressed from its gene under the control of the promoter of the sca-1 gene (de Bruijn et al., 2002). LTpHSCs can also be identified by their capacity to express RUNX-1 (North et al., 2002). LT-pHSCs also express CD27 (Wiesman et al., 2002) and flt-2/flk-3 (Christensen and Weissman, 2001) (Figure 7.1). Two types of pHSC have been experimentally identified in transplantation experiments. These differ in the fourth capacity—their capacity to reconstitute the transplanted host. The LT pHSCs will repopulate the host for longer periods and, therefore, allow continuous hematopoiesis. The other type of pHSCs, called short-term (ST) pHSCs, allow one wave of pluripotent hematopoiesis, which ceases because the pHSCs have been lost in the host by differentiation. These have been found in mice and humans (Spangrude et al., 1991; Guenechea et al., 2001). These ST pHSCs can be distinguished phenotypically from LT pHSCs, because they downregulate the expression of flt-2/flk-3 (Christensen and Weissmann, 2001) and upregulate the expression of flt-3/flk-2 (Adolfsson et al., 2001).
Hemangioblasts as Early Progenitors of pHSC and Vascular Endothelium The progenitors of hematopoiesis found in the AGM region of the embryo do not give rise to pluripotent hematopoiesis upon transplantation (Ohmura et al., 2001).
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
They appear to need to migrate to the primary sites of hemato-lymphopoiesis (bone marrow, fetal liver, etc.) before they can do so. Conversely, they—or their close progeny—appear to be even more pluripotent than the pHSCs, since they have been seen to give rise not only to pHSCs but also to vascular endothelium (Roberts et al., 1999, 2000; Ogawa et al., 1999; Nishikawa et al., 1998). These progenitors are called hemangioblasts. The actions of the flk-1, V-CAM4, and VEGF might well control decisions to enter either the hematopoietic or the endothelial pathway of differentiation (Gerber et al., 2002). Energizing pluripotent hematopoietic cells have been found in the human embryo and fetus in the vascular walls of the embryonic aorta, yolk sac, fetal liver, and fetal bone marrow (Oberlin et al., 2002). In these experiments CD34+ CD31+ CD45- progenitor cells from the vascular endothelium of all these embryonic sources yielded myelolymphopoietic cells in culture, thus supporting the notion of a common hemangioblast.
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Plasticity Versus Stability of pHSC The pluripotency of a single pHSC to differentiate along all possible hemato-lymphopoietic pathways of blood cell differentiation [most clearly documented with cloned PAX5deficient precursor cells (Rolink et al., 2000)] is proof of the plasticity of pHSC. These changes in the differentiation capacity of pHSC appear to be transdifferentiation events caused by cells moving forward, sideways, or backwards in hematopoietic lineages. This plasticity may, at least in part, explain the heterogeneity of early progenitor and precursor phenotypes of the B lymphocytic lineage pathway(s) of differentiation documented below (see Figures 7.1 and 7.2). Redifferentiation can be initiated by external stimuli, or by changes in transcription factor gene expression programs or signal transduction programs inside the cell. This plasticity might be useful in a host response to external stress using either one or the other parts of the its innate or adap-
FIGURE 7.2 Development of B lineage cells from early lymphoid progenitors (pL) over precursor B cells (pre-B cells) to immature and mature B cells of the B1 and the conventional B lineages. The earlier stages of this development (pL1 to pL4) are oversimplified. Plasticity of cells, heterogeneity of cell populations, and age-dependent changes are likely to make this scheme more complicated in reality. (1) The PAX-5 deficiency induces a change to a more immature progenitor cell that has the capacity (2) to develop to practically all known hematopoietic cell lineages, possibly including pHSC. (3) Development of wildtype pre-B-I cells in vitro and in vivo.
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tive immune system. In this way, B cell development is subject to stress-induced changes in the rates of B cell production from pHSC and B cell progenitors (Fulop et al., 1986; Osmond et al., 1985; Rico-Vargas et al., 1995; Medina et al., 1993; Kincade et al., 1994; Kincade et al., 2002).
Transdifferentiation to and from NonHematopoietic Cell Lineages It is more difficult to judge recent reports of a much wider plasticity of pHSCs that claim these cells can generate cell lineages in the brain (Brazelton et al., 2000), in muscle (Ferrari et al., 1998), or in liver (Petersen et al., 1999; Lagasse et al., 2000; Theise et al., 2000), or into several lineages (Krause et al., 2001). Conversely, pluripotent stem cells of the brain (Björnsson et al., 1999) and of muscle (Jackson et al., 1999) have been seen to give rise to pHSCs. As long as stem cells (pHSC or of other lineages) cannot be cloned and established as clonal cell lines, the possibility remains that a mixture of stem cells committed to these different cell lineages is at the origin of all these findings.
PATHWAYS OF HEMATOPOIETIC PROGENITOR CELLS TOWARD B LYMPHOCYTE LINEAGE COMMITMENT AND DIFFERENTIATION The developmental pathways from pHSCs into the different lineages of blood cells is balanced by the proliferative expansion of progenitors and precursors, appropriate differentiation into different lineages, and cell death at different rates in different hematopoietic lineages. Two hierarchical schemes have been proposed, one from experiments that have analyzed clonal growth and differentiation in vitro and lineage development after transplantation in vivo (Kondo et al., 1997; Akashi et al., 2000), the other from experiments that have analyzed the developmental defects induced by mutations in genes encoding transcription factors, cytokines, growth factors, or signal transducing molecules (Singh, 1996; Orkin, 1992; Tsai et al., 1994; Scott et al., 1994, Georgopoulos et al., 1994; Wang et al., 1996). The two schemes differ in the earliest stages of development from pHSC. One assumes an early separation of erythropoiesis and myelopoiesis from lymphopoiesis to yield common myeloid (CMP) and common lymphoid precursors (CLP). The other depicts a linear degression of potential from a pluripotent to a myeloid/lymphoid and then to a lymphoid progenitor stage (discussed in Rolink et al., 2000; Schaniel et al., 2002). If early stages of hematopoietic cells can display a plasticity of responses to environmental stimuli such as cytokines and cell contacts, then it could be
expected that the phenotypes of those early progenitors and their numbers may well differ in different organs of hematolymphopoiesis (e.g., in bone marrow or fetal liver) at different times of development, exposed to different stimuli (Kondo et al., 2000; Montecino-Rodriguez et al., 2001; Cumano et al., 1992; Graf, 2002).
Ordering of Lymphoid Progenitors by Marker Expression The differential expression of four receptor tyrosine kinases flk-1, flt-3/flk-2, flt-2/flk-3, and c-kit (Ogawa et al., 2000; Gilliland and Griffin, 2002; Christensen and Weissman, 2001; Adolfson et al., 2001; Rolink et al., 1996; Morrison et al., 1995); of sca-1, CD27 (Wiesman et al., 2000), AA4.1, thy-1, and CD4; and of mostly B lymphoid lineage-related markers (RAG-1, RAG-2, TdT, IL-7Ra, CXCR4, CD25, VpreB, l5, Iga, Igb, B220, CD19, IgD, CD21, CD23, pTa, MHC class II, and PAX-5); and the rearrangement status of the Ig H and L chain gene loci (Ogawa et al., 2000; Igarashi et al., 2002; ten Boekel et al., 1995; Ghia et al., 1996, 1998) have allowed an ordering of hematolymphopoietic progenitors as well as B lineage precursors on their way to becoming B cells. For all these cell stages, the order has also been established by in vitro or in vivo tests of their capacity to develop to later stages in the B lineage. More recently three transgenically marked strains of mice have further clarified the earliest developmental stages. One expresses a human IL-2 receptor a chain gene (huCD25) under the control of the promoter of the l5 gene of the surrogate L chain of the pre-B cell receptor (Mårtensson et al., 1997). The other expresses the same human CD25 gene inserted in the genome under the control of the promoter of the pTa gene of the pre-T cell receptor (Gounari et al., 2002). The third expresses the gene encoding green fluorescent protein (GFP) inserted into and, hence, under the control of the RAG-1 gene active in Ig and TcR gene rearrangement (Igarashi et al., 2002).
The Earliest Lymphoid Progenitor Cells The earliest stage of progenitors from which B lymphocytes, T lymphocytes, and NK cells can be developed is a recently identified population of Lin- CD27+ ckithi sca-1+ RAG-1+ (i.e., GFP+) cells (Igarashi et al., 2002). In this article, we call these progenitors of the lymphoid T, B, and NK lineages pL cell compartments. The earliest compartment is denoted pL1 in Figure 7.2. All pL compartments develop from the pHSC and are prior to fully DJ/DJrearranged pre-T and pre-B (I) cells. All pL cells develop poorly, if at all, into erythroid and myeloid cells. These cells also express TdT, though individual cells of this population may either express both TdT and RAG, or only TdT, or only RAG. Both RAG-1 and RAG-2 are expressed. pL1 cells also
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
express E47 and low levels of EBF, two transcription factors that control B lymphocyte development positively, and Id, which controls it negatively (see below). A small number of DH-JH rearrangements are detected in this cell population, indicating that they are beginning to develop into B lymphocytes. However, it is clear from DH-JH rearrangements found in myeloid cells, T cells, and NK cells that DH-JH rearrangements do not commit cells irreversibly to the B lymphocyte lineage. The next stage toward B lineage development is the LinCD27+ ckitlo sca-1lo RAG-expressing cell population also identified by Igarashi et al. (2002) and denoted pL2 in Figure 7.2. In contrast to pL1 cells, some of these cells express Iga, the a chain of the IL-7 receptor, and the transcriptions factors aiolos and PAX-5. As described next, PAX-5 commits progenitors to the B lymphoid lineage of development. pL2 cells no longer express Id. DH-JH rearrangements in the pL2 population are more frequent than in pL1.
Myeloid Progenitor (pM) Cells Although pL1 and pL2 cells have the capacity to develop into lymphoid cells such as T, B, and NK cells, the LinCD27+ ckithi sca-1hi cells not expressing RAG (i.e., GFP-) do not develop to lymphoid cells, but are ten times as likely as their RAG-expressing counterparts to develop myeloid cells (Igarashi et al., 2002). These cells are also expected to be IL-7Ra-. Therefore, RAG and TdT and the activation of the rearrangement machinery (and subsequently IL-7Ra expression) signifies an increased potential for lymphoid (and a decreased potential for myeloid) development. These results extend a scheme of hematopoiesis proposed by Kondo et al. (1997) and Akashi et al. (2002), although an alternative scheme (Singh, 1996) cannot be totally ruled out, if for example, the RAG- early progenitors are unable to home efficiently into the proper sites of hematopoietically active organs (such as bone marrow) upon transplantation (Figure 7.2). pL2 cells have also been identified as B220+ CD27+ ckitlo flt-3/flk-2hi CD19- cells not expressing the l5 component of the surrogate L chain, as detectable by the human CD25 reporter gene under l5 promoter control (Mårtensson et al., 1997; Ogawa et al., 2000). These cells develop spontaneously in tissue culture into sIg+ B lymphocytes from their original, only partly DHJH-rearranged status of B lymphoid development.
The Earliest Lymphoid Progenitors Expressing the Surrogate Chains of the Pre-Lymphocyte Receptors The next cell population in line of lymphoid differentiation, denoted pL3 in Figure 7.2, has increased quantities of DHJH-rearranged IgH loci. These cells express the l5 com-
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ponent of the surrogate L chain as well as the pTa chain of the pre-TcR (Ogawa et al., 2000; Gounari et al., 2002). As in all preceding populations, this cell population is likely to be heterogeneous. For example, it is not yet clear whether l5 and pTa are expressed in the same, in partially overlapping, or in different cell populations. This pL3 population of B220+ CD19- cells is likely to include the progenitors of NK cells identified by Rolink et al. (1996), which are a separate population from those cells in the same population expressing CD4 or MHC class II molecules. pL3 cells may well include the last tripotent T, B, and NK progenitors that may be expected to migrate from bone marrow to the thymus to develop along the T lineage pathway, or remain in the bone marrow to continue B lymphoid development. On the way to becoming B lymphoid cells, a population with lowered expression of flt-3/flk-2 and now expressing the B lymphoid-specific CD19 (denoted pL4) has been characterized as a potential intermediate on the way to a fully DHJH/DHJH CD19+, B220+, flt-3/flk-2-, ckitlo pre-B-I cell (Ogawa et al., 2000). Pregnant or estrogen-treated mice develop a depression in T and B lymphopoiesis, whereas hypogonadal, castrated male, ovaryectomized female, and androgene receptor–deficient mice show abnormally elevated T and B cell development. The primary targets of this hormonal action appear to be the lymphoid progenitor cells in the pL1 and pL2 compartments (Medina et al., 2001).
CONTROL OF LYMPHOID CELL DEVELOPMENT BY TRANSCRIPTION FACTORS Blastocyst complementation assays with ES cells bearing mutations in the transcription factor genes Rbtn-1 (Warren et al., 1994), TAL-1 (Porcher et al., 1996), and GATA-2 (Orkin, 1992; Weiss and Orkin, 1995; Tsai et al., 1994) have revealed defects in general hematopoiesis during the generation of either LT or ST type pHSC (Figure 7.1). Mice fully defective in the PU-1 gene have a general defect in the development of myeloid and lymphoid cells, but do develop erythrocytes, megakaryocytes, and platelets (Klemsz et al., 1990; Hromas et al., 1993; Goebl, 1990; Singh, 1996). Target genes of the PU-1 transcription factors, a member of the ets domain proteins encoded by the Spi-1 proto-oncogene, include the mH chain gene (Nelsen et al., 1993), the L chain genes (Eisenbeis et al., 1993), and the gene encoding Iga (Hagman and Grossschedl, 1992; Feldhaus et al., 1992; Shin et al., 1993). Since this mutation was generated by the deletion of the exon encoding the ets-DNA binding domain, it could affect its action in a dominant-negative fashion in a complex with other transcription factors, thus replacing the wildtype PU-1 form in the complex.
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In PU-1-/- mice, in which the expression of the PU-1 gene has been deleted altogether, only the formation of macrophages and osteoclasts is inhibited. This suggests that in the complete absence of PU-1, other transcription factors may take its place to allow lymphoid development. Low PU-1 expression specifically promotes B lymphoid development, whereas high PU-1 expression suppresses it and promotes the formation of macrophages, which are members of the myeloid cell lineage (de Koter and Singh, 2000). Low levels of PU-1 expression induce the expression of the IL-7Ra chain, whereas high levels inhibit it (de Koter et al., 2001). Retroviral expression of the IL-7Ra chain in PU-1–deficient progenitors alone restores B lymphoid potential in these cells (de Koter et al., 2002). If the observed development of macrophages at high levels of PU-1 expression is a sign of general myeloid cell development in other cell types such as granulocytes, osteoclasts, and dendritic cells—and not a manifestation of a change in the balance of a later, bipotent B lymphoid/macrophage precursors (Cumano et al., 1992)—then these experiments indicate that PU-1 expression critically controls the decision between myelopoiesis and lymphopoiesis.
Transcription Factors Controlling the Decisions Toward T, B, or NK Lymphoid Development The decision to enter T and B lymphoid development appears to be controlled by the Icaros gene. The deletion of the DNA-binding domain within Icaros abolishes lymphoid development (Georgopoulos et al., 1994). Also, a mutant form of Icaros acts in a dominant-negative fashion (Molnar and Georgopoulos, 1994; Nichogiannopoulou et al., 1999). Target genes of Icaros include the RAG and TdT genes of the rearrangement machinery, the IgH and L chain genes, the Iga gene, and members of the CD3 complex (Brown et al., 1997). However, Icaros acts as a nonclassical transcription activator, possibly removing inhibitors from the vicinity of target genes and thereby remodeling chromatin (Koipally et al., 2002; Georgopoulos, 2002). Although Icaros has strong actions in lymphoid progenitors, it also influences earlier steps of hematopoiesis.
of E2A-deficient mice no DH-JH rearrangements (or VL-JL rearrangements) are detectable. Also, transcripts of sterile mH chain message, of RAG-1 and RAG-2, Iga, Igb CD19, VpreB, l5, and PAX-5 are strongly reduced or absent (Bain et al., 1994; Zhuang et al., 1994; Lin and Grossschedl, 1995; Sigvardsson et al., 1997; Kee and Murre, 1998). Transfection and the expression of E47 in fibroblasts activates the expression of TdT and of the IgH chain locus (Choi et al., 1996). When, in addition, the ectopic expression of RAG-1 and RAG-2 is provided together with either E2A or EBF in a nonlymphoid cell line, such as an embryonic kidney cell line, DH-JH rearrangements are induced at the endogenous IgH chain loci (Romanow et al., 2000). Endogenous L chain genes are more selectively rearranged in the same nonlymphoid cells: E2A, together with RAG-1 and RAG-2 allows endogenous Vk to Jk rearrangements, whereas EBF does so for Vl to Jl rearrangements (Romanow et al., 2000). This agrees with multiple E2A binding sites being found in the Ig enhancer elements; these are thought to be requisite (Serwe and Sablihky, 1993; Chen et al., 1993), although may be not sufficient (Inlay et al., 2002) for V(D)J recombination. Not all V, D, and J segments appear to be equally accessible for recombination (Goebel et al., 2001), and OcaB has recently been found to be required for a subset of Vk gene segments in their transcription and V(D)J recombination activities (Casellas et al., 2002). EBF is more restrictedly expressed in progenitors and pre-B cells (Feldhaus et al., 1992; Hagman et al., 1991). Its defect in mice generates blocks in B lymphopoiesis that are quite similar to those seen in E2A-deficient mice; that is, before the onset of DH-JH rearrangements and the development of B lineage cells that harbor them (Lin and Grossschedl, 1995).
NK Development The development of NK cells is controlled by the helixloop-helix transcription factor Id2 (Yokota et al., 1999) (Figure 7.1). Id2 is expressed in NK cells, suppressing the action of other transcription factors to induce other lymphoid lineage development (Ikawa et al., 2001). In one such inhibitory action, it might complex E2A into E2Ainactive heterodimers (Benezra et al., 1990; Engel and Murre, 2001).
B Lymphoid Development The decision to enter the B lymphoid pathway is critically controlled by two transcription factors: the basic helixloop-helix protein E2A and the early B cell factor (EBF) (Figures 7.1 and 7.2). Alternate splicing of the E2A gene generates the E12 and E47 proteins. Binding sites for these proteins are found in the IgH and IgL enhancers. Although E2A is broadly expressed in hematopoietic cells, its absence affects mainly the B lymphoid lineage. In B220+ progenitors
T Lymphoid Development Early T cell development, some of it occurring extrathymically in bone marrow, is controlled by signaling through NOTCH-1 (Figure 7.1). NOTCH-1 is activated by its ligands, members of the Jagged and Delta families of proteins. When these ligands bind to NOTCH-1 receptors on the surface of lymphoid progenitor cells, the intracellular part is cleaved from the receptors to function as a transcrip-
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
tion factor (Radtke et al., 1999; Pui et al., 1999; Andersson et al., 2000; Izon et al., 2002). The ectopic expression of NOTCH-1 (“active” NOTCH-1; see Figure 7.1) allows early thymocyte development in the absence of a thymus, and inhibits B lymphoid development (Pui et al., 1999). Also, Delta-expressing stromal cells induce human CD34+ progenitor cells to differentiate to thymocytes, but not to B lymphoid cells (Jaleco et al., 2001). Inactive NOTCH-1, on the other hand, inhibits T cell development at the earliest T lineage-related stage—that is, the DN1 CD44+ CD25- stage. At the same time, it promotes B cell development, even in the thymus (Radtke et al., 1999; Wilson et al., 2001). NOTCH-1 can also be inactivated by Lunatic Fringe (Koch et al., 2001), and by Deltex (Izon et al., 2002); such inactivations lead, again, to the arrest of T lymphopoiesis and the promotion of B lymphopoiesis (“inactive” NOTCH-1; see Figure 7.1). All available evidence suggests that a possibly heterogeneous pL3 population of lymphoid progenitors affects the stage at which these decisions between T, B, and NK lymphoid development are affected by E2A, EBF, Id2, and NOTCH-1 in active and inactive forms.
Commitment to B Cell Development Although the transcription factors E2A and EBF are required to initiate the expression of essential B lineage– specific and –related genes and V(D)J recombination, they are not sufficient to allow B cell development to the pre-BcR+ and BcR+ stages of differentiation (Figure 7.2). In the absence of PAX-5, as occurs in PAX-5–deficient mice, B cell development becomes arrested at a pre-B-I-like stage of development (Urbanek et al., 1994). These cells appear pre-B-I-like since they are DHJH-rearranged on both H chain alleles and because they proliferate for long periods of time in vitro on stromal cells in the presence of IL-7 (Rolink et al., 1999; Schaniel et al., 2002) as pre-B-I cells from wildtype mice. They express, among other genes, the VpreB and l5 genes encoding the surrogate L chain, Iga and Igb, RAG-1 and RAG-2, and the transcription factors OCT-1, OCT-2, OBF, SOX-4, PU-1, Icaros, E2A, and EBF. The fact that E2A and EBF are expressed in PAX-5–deficient pre-B cells places PAX-5 downstream of E2A and EBF (Schebasta et al., 2002). PAX-5-/- pre-B cells express ckit and surrogate L chain, but not CD25 on their surface, and wildtype pre-B cells have the same properties. However, they differ in a variety of properties from wildtype pre-B-I cells. Interestingly, such cells do not develop in fetal liver, as wildtype cells do. They develop in bone marrow, but appear to have the strong long-term proliferative capacities that wildtype pre-B-I cells from fetal liver exhibit (Nutt et al., 1997). PAX-5-/- pre-B cells do not express CD19 (which is under the direct control of PAX-5),
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and express flk-2/flt-3. Hence they have some of the phenotype of pL4 cells or even earlier pL stages. PAX-5-/- pre-B cells are blocked from entering VH to DHJH rearrangements at normal frequencies. Hence, they are blocked from generating large pre-B-II cells expressing a pre-BcR and expanding by proliferation, and they are blocked from entering VL to JL rearrangements at detectable levels and becoming immature and mature B cells (Figure 7.2). This deficiency is evident in vitro and in vivo when pre-B cells are induced to mature, by the removal of IL-7 in vitro, or by transplantation into severe combined immunodeficient (SCID) RAG-/- hosts.
PLASTICITY OF PAX-5–DEFICIENT PRE-B CELLS Conditional PAX-5 inactivation in wildtype pre-B-I cells, expanded by proliferation on stromal cells in the presence of IL-7, returns these cells back to the pre-B-I-like or even pL4 or earlier pL stage of differentiation (Figure 7.2, green 1) (Mikkola et al., 2002). Hence, PAX-5 expression is continuously required to maintain B cell differentiation to the pre-B-I cell stage. Interestingly, PAX-5 remains to be expressed in all subsequent stages of B cell differentiation, over pre-B-II, immature to mature B cells, but not to plasma cells (Urbanek et al., 1994; Busslinger and Urbanek, 1993). It will be interesting to see whether conditional PAX-5 inactivation in B lineage cells at such later stages can also induce dedifferentiation to earlier, even pre-B-I-like, pL4like, stages of development. This plasticity of B lymphoid cells deficient in PAX expression becomes even more evident when these PAX-5-/- pre-B cells are exposed to different environmental stimuli. Whereas IL-7 in tissue culture with stromal cells retains PAX-5-/- cells in their pre-B-I-like phenotype thus making IL-7 dominant over all other influences, the removal of IL-7 and the subsequent exposure to different cytokines and cell contacts induces myeloid and NK cell differentiation (Nutt et al., 1999; Schaniel et al., 2002a). In the presence of IL-2 these DHJH-rearranged cells develop into NK cells, while M-CSF induces macrophage, M-CSF plus GMCSF dendritic cell, TRANCE osteoclast, and G-CSF granulocyte development. All these differentiated blood cells carry the characteristic DHJH/DHJH rearrangements of an initial pre-B-I-like clone of pre-B-I-like PAX-5-/- cells thus indicating that the original PAX-5-/- pre-B cells are multipotent. The expression of the B lineage–related and specific markers, notably VpreB and l5, are lost in these differentiations, indicating that neither DH-JH rearrangements nor surrogate L chain expression irreversibly commit cells to the B lineage pathway (Figure 7.2, red 2). The induction of differentiation by in vivo transplantation into SCID or RAG-/- hosts reveals additional differen-
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tiation potencies of PAX-5-/- pre-B cells. In vivo, not only CD8- but also CD8+ dendritic cells develop. Furthermore, normal T cell development in the thymus and in the periphery is established (Rolink et al., 1999). At a longer term (i.e., after 2 to 3 months) the in vivo development of myeloid cells and erythrocytes becomes detectable (Schaniel et al., 2002a). Furthermore and, again, in contrast to wildtype preB-I cells, PAX-5-/- pre-B cells migrate back to the original sites in bone marrow, from where they can be reisolated, expanded again in tissue culture, and retransplanted again and again, thus showing that PAX-5-/- pre-B cells have selfrenewal and long-term reconstitution potential (Schaniel et al., 2002b). PAX-5-/- pre-B cells, therefore, are close relatives of pHSCs, lacking only the capacity to protect a lethally irradiated host from death, probably because the erythroid and myeloid cell lineages develop too slowly from the PAX-5-/- cells (Figure 7.2, red 2). In conclusion, it appears that the reactivity to IL-7 and PAX-5 expression defines three discernible states of early B cell development. The first descends from LT-pHSC over pL states to an early pre-B-I cell-like or possibly pL4 state, at which PAX-5-deficient cells are arrested. In vitro, and probably also in vivo, this state in PAX-5-/- cells is stabilized (and, hence, probably controlled) by IL-7 signaling through the IL-7 receptor. The second state is that of a pre-B-I cell, again stabilized by IL-7/IL-7 receptor signaling. Removal of IL-7 from wildtype pre-B-I cells induces the third state, a sequence of cellular developments to the sIg+ B cell (Figure 7.2, blue 3). Removal of IL-7 from the pre-B-I-like state of PAX-5–deficient cells allows dedifferentiation back to the LT-pHSC and to all erythroid, myeloid, NK, and T lineage cells (Figure 7.2, 2) (Schebasta et al., 2002; Mikkola et al., 2002; Rolink et al., 2002).
THE SURROGATE LIGHT CHAIN The surrogate L chain (SL chain) is assembled from the VpreB and l5 proteins. In humans one, and in mouse two, VpreB genes encode VpreB protein, whereas l5 is encoded by one gene in both humans and mouse. Assembly of the two proteins is spontaneous; the V region-like VpreB proteins provide b pleated sheets for a noncovalent assembly. When this seventh b pleated sheet is deleted in l5, the assembly of VpreB with l5 is abolished (Minegishi et al., 1999). The non-Ig portions at the carboxy terminal end of VpreB, and the amino terminal end of l5, protrude from the SL molecule at the site where the third complementaritydetermining region (CDR3) would form in a normal L chain. Its function still must be classified, as deletions of these nonIg portions do not abolish the assembly of VpreB and l5 to an SL. Subsequent covalent binding via an S-S bond between
the Cl5 domain and the first C1 domain of the mH chains (and other classes of H chains) functions normally (reviewed in Melchers, 1999, and Melchers et al., 2000). SL chains are expressed in pL3, pL4, and pre-B-I cells, before mH chains are expressed. In these early progenitors, these SL chains are found associated with complexes of glycoproteins, forming what has been called the pre-B cell receptor. The function of SL chains in these early cells is unknown (Melchers, 1999; Karasuyama et al., 1993; Ohnishi et al., 2000).
Structure and Assembly of the Pre-B Cell Receptor Whenever mH chains are first made from productively rearranged H chain loci in pre-B-II cells, they are probed for assembly with the SL chain. It is remarkable to note that half of all mH chains first translated from rearranged H chain loci are incapable of pairing with SL chain to form a pre-B cell receptor (pre-BcR) that can be deposited on the cell surface (ten Boekel et al., 1997; Keyna et al., 1995). This could, in part, be due to incompatible structures of the CDR3 regions of mH chains generated by N region insertions during V(D)J recombination. VH81x without N regions, made in fetal liver, bind well to SL chain, whereas most mH chains with the same VH domain, made in adult bone marrow, are unable to pair. Two regions appear to be important in allowing the association between VH with VL in normal Ig molecules. One is the VH-specific G-L-E-W hydrophobic, the P-hydrophilichydrophobic-L-hydrophobic framework 2 sequence motifs, and their accompanying b bulges (Frazer and Capra, 1999). The other is the W/F-G-X-G motif in framework 4 which, together with b bulges, is the second major contact site between VH and VL. Although these structures may be perturbed if the CDR3 region is too bulky, or otherwise structurally incompatible, it may also be possible that VpreB may interact slightly differently with VH domains, and thus may prevent the proper assembly with some germlineencoded VH segments. Once the structures of pre-BcR become known, these discussions will be more clearly defined. VpreB1 and VpreB2 proteins can pair alone with mH chains. The ability or inability of a given mH chain (with a given VH domain) to pair with an SL chain coincides with its ability or inability to pair with a VpreB protein. Hence, VpreB can be regarded as the prime probing device of the total pre-BcR for its fitness with a given mH chain (Seidl et al., 2001). The variability of VpreB–VH interactions, due to structural variations both in VH-encoded and CDR3 chance-generated sequences, predicts a spectrum of avidities for these interactions. Mass law predicts that with a given, constant
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
concentration of surrogate L chain expressed in a preB-II cell, the formation of pre-BcR molecules, and hence their numbers on the cell surface, will be determined by avidity. In contrast to VpreB protein, the l5 protein cannot associate alone with a mH chain, pairing or not, to form the classical S-S bonded heterodimer with a cm1 domain. This suggests that in cells synthesizing mH chains and l5 proteins, both proteins are “protected” from proper association. The mH chain may be bound to a chaperone, such as BIP, whereas l5 could exist in a non-Ig-like conformation (Minegishi et al., 1999). The addition of VpreB protein, synthesized in the same cells, allows the prompt assembly of SL chains with pairing mH chains, thus suggesting that VpreB, in binding to l5, induces a conformational change of l5 which, in turn, induces the mH chain to be receptive to association and S-S bonding, possibly by first replacing BIP (discussed in Melchers et al., 2000).
Ligands for the Pre-B Cell Receptor It has been proposed that pre-BcRs may have ligands that control the functions of the cells that either express SL (i.e., pre-B-I cells) or pre-BcRs (i.e., pre-B-II cells). If pre-BcR recognition were determined by the SL component, constant in all pre-BcRs, the observations that a number of different transgenic L chains can repair SL deficiencies would argue against this possibility (Pelanda et al., 1996; Rolink et al., 1996). We have argued previously that structural elements preserved in all VH domains, or the recognition of a ligand by VpreB (which could still be associated with mH L chain complexes in and on pre-B-II cells of l5-/-, L chain transgenic mice) could still serve as receptive elements of the pre-BcR. Moreover, mAbs specific for the pre-BcR—specific for VpreB, l5, or mH chains—do not perturb pre-B cell development either positively or negatively, either when injected in vivo or when added to fetal liver organ cultures in vitro. Conversely, mAbs appearing against IL-7 and its receptor inhibit pre-B-II cell expansion in the same in vitro cultures (Ceredig et al., 2000). The same mH chain–specific mAbs, on the other hand, inhibit the development of immature sIgM+ B cells in these in vitro cultures. Finally, pre-B-II cells isolated ex vivo and cultured in vitro without cytokines and stromal cells will undergo two to five divisions (Rolink et al., 2000), a result that argues for a pre-BcR occupancy-independent proliferation of large pre-B-II cells. Nevertheless, this proliferation does not occur with l5-deficient pre-B cells, arguing for the importance of the presence of pre-BcRs in these cell membranes. In view of the finding described here, it is all the more surprising that Galectin-1, an S-type lectin, anchored to
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glycosylated counterreceptors on stromal cells, interacts with the non-Ig portion of the l5 protein expressed on preB cell lymphoma lines and induces the relocalization of preBcR on the surface of the lymphoma cells and synapse formation with the stromal cells (Gauthier et al., 2002).
Expression of the Surrogate L Chain The surrogate L chain is first expressed in a part of pL3 cells (Figure 7.2). It is detectable as mRNA, protein, and as the product of a reporter gene, human CD25, expressed under control of the l5-specific promoter as a transgene (Mårtensson et al., 1997). The expression of SL chain genes on an mRNA level is turned off when pre-B-II cells begin to express a pre-BcR. Protein expression remains detectable in large pre-B-II cells, because substantial quantities of the SL protein are expressed cytoplasmically and are detectable by immunofluorescence with specific mAbs. SL chains are detectable on the surface of pre-B-I and large pre-B-II cells, although an apparently constitutive downregulation of pre-BcR expression from the surface of large pre-B-II cells makes the detection experimentally more demanding. In pre-B-I cells, SL chains appear on the surface associated with a complex of proteins, including a special E-cadherin called BILL cadherin (Ohnishi et al., 2000; Karasuyama et al., 1993). The function of this protein complex remains to be elucidated. From BILL cadherin–deficient mice it is evident that its function may be required, but is not mandatory, for pre-B-I cell development and further B lineage cell differentiation. However, its possible function in the allelic exclusion of the H chain locus has not yet been tested. Normally, SL chain mRNA and protein becomes undetectable in small pre-B-II cells and all subsequent stages of B cell development, although some B lineage tumors may be able to express both SL and conventional L chains. In the peripheral B cell compartments of humans, especially of rheumatoid arthritis patients, CD10+ CD27+ CD19+ sIgM+ B cells have been found which co-express conventional and surrogate L chains. It is not clear why these cells have not turned off SL expression, as their H chain V regions appear hypermutated; that is, capable—and a result—of antigenic, T cell-dependent stimulation (Meffre et al., 2000). A “re-expression” of pre-B cell–specific markers, such as RAG-1, RAG-2, VpreB, and l5 in germinal center cells activated by immunization with an antigen remains controversial (Han et al., 1996; Hikida et al., 1996; Papavasiliou et al., 1997). It is likely that at least part of this “re-expression” is, in fact, due to the influx of pre-B cells into the germinal center, activated in the bone marrow by the stresslike action of the immunization (Yu et al., 1999; Fulop and Osmond, 1983a, b).
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PRE-B CELLS AND THEIR DIFFERENTIATION TO MORE MATURE B LINEAGE CELLS Pre-B-I Cells During the development of B lineage cells in mice and humans, the Ig gene loci are V(D)J-rearranged in an ordered fashion (for a review see Melchers and Rolink, 1999). This, in turn, has allowed the ordering of precursor B cell phenotypes, as they have been analyzed by single cell PCR for their status of rearrangements of the H and L chain gene loci (Ehlich et al., 1994; ten Boekel et al., 1995; Ghia et al., 1996) (see also Figure 2). In the normal, nonmutant development of mouse and human B lineage cells, the first stable state is reached when both H chain alleles are DHJH-rearranged. Over 99% of all cells in these two species reach this state; immature and mature B cells with H chain alleles in germline configuration are less than 0.1% of the total populations, in contrast to some other species, such as the rabbit (Tunyaplin and Knight, 1997). Although mouse and human B cell development is strikingly similar (Ghia et al., 1998), development in the mouse is presented and discussed here. It has been proposed by Alt and colleagues (1980) that the protein products of in-frame V(D)J-rearranged Ig H and L chain genes instruct the developing B cell that the second allele can no longer be rearranged; that is, that it would be allelically excluded from expression. This would allow one B cell to produce only one H and one L chain—hence one Ig molecule—with a given specificity. With one exception, DHJH-rearranged H chain loci do not allow the translation of a DHJHCm protein. Hence, it is acceptable in Alt et al.’s hypothesis of allelic exclusion that the H chain locus is not allelically excluded at the DHJHrearranged level. The one exception is a DHJH-rearranged H chain locus in the mouse which, when rearranged in reading frame 2, allows transcription of an mRNA that potentially encodes a DHJHCm protein. The expression of this protein may contribute to the observed suppression of the representation of reading frame 2–rearranged H chain loci (in line with Alt et al.’s hypothesis). However, a DHJHCm protein has been identified only once. Since such proteins cannot be made in humans, a more general function for them in B cell development appears unlikely (reviewed in Melchers and Rolink, 1999). In the microenvironment of mouse bone marrow, or of fetal liver, DHJH/DHJH-rearranged pre-B-I cells form a pool of approximately 5 ¥ 106 cells which, in numbers, gradually decrease as the individual ages (Rolink et al., 1993; Ghia et al., 2000). Depending on the rate of influx of cells from early pL progenitors into this pool, pre-B-I cells are expected to produce pre-B-II cells by mostly asymmetrical divisions. In cells leaving this microenvironment (hence, probably the
chemoattraction of SDF-1), VH to DH-JH rearrangements are begun. Pre-B-I cells are ready to do so, because they express the rearrangement machinery RAG-1 and RAG-2. In bone marrow, but not in fetal liver, they express the enzyme TdT (reviewed in Melchers et al., 2000). Therefore, VH to DH joins, as well as the previously produced DH to JH joins, contain N regions (and consequently a higher diversity in CDR3 regions of the H chain) only in B cells made in bone marrow.
VH to DH-JH Rearrangements at the H Chain Locus at the Transition from Pre-B-I to Pre-B-II Cells It is not clear whether both DHJH-rearranged Ig H chain alleles are equally accessible in one cell for VH to DH-JH rearrangements. If only one allele were open for rearrangements, opening of the second allele could be regulated by the productive rearrangement of the first allele; that is, by a mH chain. If both alleles were open, then the mH chain would have to signal the closure of the second allele. VH to DH-JH rearrangements produce randomly in- and out-of-frame rearrangements, so that approximately half of the emerging pre-B-II cells are VDJ/DJ and the other half VDJ/VDJ-rearranged. This ratio is stable throughout development to mature B cells. The existence of VDJ/DJrearranged cells indicates that allelic exclusion—the inability to VØDHJH rearrange the second allele—is operative. Since the majority of VDJ/VDJ-rearranged cells are in-frame or productively rearranged on one allele, and outof-frame or nonproductively on the other, it suggests that a first nonproductive VDJ-rearranged allele is not recognized by the pre-B cell for either keeping closed or closing the second allele. When the gene segment encoding the transmembrane portion of the mH chain is experimentally deleted, mH chains can no longer be inserted into cell membranes. Such H chain alleles no longer function in allelic exclusion; they do not signal the other DHJH-rearranged allele to stop V(D)J recombination, even when the deleted domain is productively rearranged (Kitamura and Rajewsky, 1992). All evidence suggests that a mH chain inserted into pre-B cell membranes initiates signals that prevent VH to DH-JH rearrangements at the second H chain allele.
Responses of Pre-B-II Cells to Signaling from the SL-Containing Pre-B Cell Receptor The deposition of pre-BcR in the membranes of pre-B-II cells induces these cells to enter the cell cycle and divide two to five times (Figure 7.2). Pre-BcR–deficient cells of mMT-/-, l5-/-, VpreB1-/-, plus VpreB2-/-, and triple VpreB1-/-, plus VpreB2-/-, plus l5-/- mice do not all enter this proliferative
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
phase of B lineage cell expansion (Kitamura et al., 1992; Mundt et al., 2001; Shimizu et al., 2002). These defects in the formation of the pre-BcR, however, do not abolish, but only impede B cell development. It is important to recognize that wildtype as well as l5-/- (and probably also all other deficient) pre-B-I cells are induced in vitro (by the removal of IL-7 from the IL-7/stroma cell tissue cultures) to enter, without proliferation, V(D)J recombination at H and L chain loci. These also generate sIg+ as well as sIg- cells (in- and out-of-frame) with kinetics and in rates that are indistinguishable between wildtype and mutant cells (Rolink et al., 1993). However, whereas pre-B-II cells of wildtype mouse bone marrow expand in vivo to around 2 ¥ 107 cells, pre-B-II cells of wildtype fetal liver expand in vitro in fetal liver organ cultures (Ceredig et al., 1998), and ckit+ pre-BI cells at the transition to pre-B-II cells proliferate as single cells in medium only (Rolink et al., 2000), l5-/--deficient pre-B cells do not. Since l5-/- pre-B-I cells have an apparently unaltered capacity to differentiate, their inability to proliferate does not generate sufficient numbers of pre-B-II cells—in which subsequent L chain gene rearrangement occurs—to generate sIg+ B cells. Many more pre-B-I cells should be present to allow the same numbers of pre-B-II cells to enter L chain gene rearrangements. In this view, B cell differentiation in pre-BcR–deficient mice is not leaky, but simply inefficient. It underlines the importance of the pre-BcR for the proper maintenance of sufficient numbers of mature B cells in the antigen-reactive peripheral compartments. As soon as pre-BcRs are formed in pre-B-II cells, expression of the VpreB and l5 genes is turned off (Grawunder et al., 1995). However, the intracellular pools of mRNA, and particularly of protein, are used up more slowly by the formation of new pre-BcR molecules and by SL protein degradation. Those mH chains pairing with higher avidities in the pre-B-II cells need a lower concentration than those pairing with lower avidities. Consequently, as pre-B-II cells continue to divide, cells expressing low avidity for associating mH chains will stop dividing before those producing high avidity-pairing chains, if an effective number of pre-BcR must be inserted in newly synthesized membranes on dividing cells to keep up cell cycle and divisions. It can, therefore, be expected that the best-fitting mH chains will be expanded most in the developing pre-B-II cell repertoire before L chain gene rearrangements are initiated (Melchers, 1999).
Signaling Reactions Initiated by the Pre-B Cell Receptor The molecular details of the signaling reaction initiated by the pre-BcR still must be worked out. Partial blocks in B cell development at the transition from pre-B-I to pre-B-II cells in syk-deficient (Cheng et al., 1995; Turner et al., 1995)
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and BLNK (SLP-65)-deficient mice (Jumaa et al., 1999, Pappu et al., 1999, Hayashi et al., 2000) suggest that these molecules participate in these signaling reactions. BLNK (SLP-65)-deficient pre-B-II cells, furthermore, appear to express increased levels of pre-BcR on their surface and proliferate more extensively. These changed properties are dependent on SL expression, thus suggesting that BLNK (SLP-65), an adapter protein for the pre-BcR, acts as a downregulator of pre-BcR expression and proliferative expansion of large pre-B-II cells. Moreover, reconstitution of RAG-deficient mice with constitutive active forms of Ras (Shaw et al., 1999) or Raf (Iritani et al., 1999) restores B cell development even without Ig expression, thus suggesting that these two proteins are also involved in signaling. Finally, mice double deficient for the interferon response factor genes IRF-4 and IRF-8 have a block in pre-B cell development that closely resembles that of the BLNK (SLP-65)-deficient mice (H. Singh, personal communications), thus indicating that these two gene products cooperate in the same signaling pathway that involves the BLNK (SLP-65) gene products mediated by the SLcontaining pre-BcR.
A Role of the Pre-B Cell Receptor in Allelic Exclusion at the H Chain Locus? Among the mH chain-producing pre-B-II, immature, and mature B cells, 2 to 4% of the cells carry two productively rearranged H chain alleles (ten Bockel et al., 1998). In all these cells, however, only one mH chain has been found to be able to pair with an SL chain. This suggested that allelic exclusion could be maintained by pre-BcR expression, perhaps at the surface of pre-B-II cells. However, and in contrast to findings by Löffert et al. (1996), l5-deficient or pre-BcR-deficient, pre-B-II and mature B cells have a comparable percentage of double mH chain producers, again with only one chain capable of pairing. How could a cell sense the pairing if it were missing the component—the complete SL chain that was sensing the pairing? From these results, it had been suggested that the sensing could be done by VpreB alone, which can bind to mH chains in the absence of l5 and form a pre-BcR-like molecule. However, this possibility has now been excluded. Both the VpreB/VpreB2 double-deficient (Mundt et al., 2001), as well as the VpreB1/VpreB2 /l5 triple deficient mice (Shimizu et al., 2002) still show allelic exclusion at the Ig H locus to the same extent as wildtype littermates. Since the deletion of the transmembrane portion of the mH chain—the lack of surface deposition of mH chains—in B lineage cells allows allelic inclusion (Kitamura and Rajewsky, 1992), we are left searching for a way by which membrane-bound mH chain could signal allelic exclusion without an SL chain, either alone or in complexes with other proteins (Figure 7.3).
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FIGURE 7.3 Two pre-B cell receptors on pre-B-I cells.
A Second Pre-B Cell Receptor with mH Chains But Without SL Chains Signals Allelic Exclusion? Four other partners for mH chains have been suggested: 1) the heat shock protein 70 chaperone H chain binding protein (Hendershot, 1990); 2) the 8HS20-encoded VpreB3 (Ohnishi and Takemori, 1994); 3) the complex of proteins associated with SL chains in pre-B-I cells, including the BILL cadherin (Ohnishi et al., 2000); and 4) prematurely rearranged L chains (Ehlich et al., 1993). The last possibility is made unlikely by our finding that all L chains analyzed in small pre-B-II cells in bone marrow do not carry N region insertions, which they should, if they were rearranged during H chain gene rearrangements (Rolink et al., 1996). As soon as H chains are formed, they associate with other proteins. It is assumed that H chains alone cannot fold properly and need the association to the chaperone protein BIP (Hendershot, 1990). BIP is then displaced by L chains in pre-B-II cells by surrogate L chain, in small pre-B-II, immature and mature B cells, and all antigen-stimulated later stages of B cell development. We expect that one of the partner proteins that form heterodimers in pre-B-II cells with mH chains is involved in the signaling complex that turns off rearrangement at the second DHJH-rearranged H chain allele. One of the most rapid responses of large pre-B-II cells after mH chain expression and membrane deposition is the downregulation of expression, both on mRNA and protein levels, of the components of the rearrangement machinery— RAG-1, RAG-2, and TdT (Grawunder et al., 1995). The previously synthesized mRNA and protein molecules are rapidly degraded. This is certainly one way by which the
pre-B-II cell avoids VH to DH-JH rearrangements at the second allele, and possibly secondary VH replacements to already VHDHJH-rearranged, often nonproductive H chain alleles. It is conceivable that the proposed second pre-BcR, with mH chain but without SL chain, signals the downregulation of expression of the rearrangement machinery. Furthermore, in order to avoid any future rearrangements at DHJH-rearranged loci, or VH replacements at VHDHJHrearranged loci, these loci must be permanently closed or never be opened to access of the rearrangement machinery. This rearrangement machinery will be reactivated in small pre-B-II and immature B cells for rearrangements at the L chain gene loci. The chromatin domain containing the Ig H locus should be modeled in a way that allows differential accessibility of the V(D)J rearrangement machinery (Georgopoulos, 2002). Active loci in chromatin can be distinguished from inactive ones by a variety of changes. Inactive IgH loci in hematopoietic progenitors and pro-T cells are preferentially positioned at the nuclear periphery, but become centrally configured in B lineage cells. During this change in localization, the IgH locus undergoes large-scale compaction (Kosak et al., 2002). The ectopic expression of E2A and EBF, together with the V(D)J recombining RAG-1 and RAG-2 genes, allows V(D)J recombination in nonlymphoid cells (Romanow et al., 2000), suggesting that key regulatory factors involved in chromatin remodeling and control of transcription render Ig loci accessible for V(D)J recombination (Stanhope-Baker et al., 1996). Although transcription from the Ig loci (Blackwell et al., 1986; Schlissel and Baltimore, 1989) is important for V(D)J recombination, and the level of transcription (e.g., controlled by OcaB at the kL chain locus) appears to influence accessibility of certain subregions of the locus for V(D)J recombination (Casellas et al., 2002), it is not clear how the intensity of transcription [possibly together with DNA demethylation and histone acetylation (Kwon et al., 2000; McMurry and Krangel, 2000)] correlates with the capacity of a certain subregion of the Ig loci to be rearranged (Goebel et al., 2001). All these studies investigated the problem of how to open a locus for V(D)J rearrangement, but did not address the problem of how to close, or keep closed, the transcriptionally active DHJH-rearranged allele for rearrangement. Again, the proposed second pre-B cell receptor, with membrane-bound mH chain but without SL chain, could signal and thereby control the accessibility of the chromatin regions of the H chain locus.
REARRANGEMENTS AT THE L CHAIN LOCI AT THE TRANSITION FROM LARGE TO SMALL PRE-B-II CELLS Rearrangements at the kL and lL Chain Loci When large, proliferating pre-B-II cells cease to divide and become small, resting cells, the rearrangement machinery is reactivated. In mouse cells, TdT is not activated again,
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
whereas it is in human cells. Hence, VJ joints of mouse L chains do not contain N regions—additional variability for antigen recognition—but human L chains do. The Ig kL and lL loci are opened for rearrangements. At the mouse kL chain locus only one allele becomes accessible for V(D)J recombination (Mostoslavsky et al., 1998). The kL chain gene locus may open before the lL chain loci (Engel et al., 1999), but rearrangements at the k and l loci are independent of each other. Thus, in Ck-deficient and JCk-deficient mice small pre-B-II cells develop in normal numbers in the bone marrow, but only 15% to 25% of them carry a VlJlrearranged L chain locus. All others have all L chain loci in germline configuration (Yamagami et al., 1999a). Hence, the rate of VL to JL rearrangement appears five to ten times higher at the kL than at the lL locus, providing an explanation for the kL to lL ratio of 10 : 1 in mouse Ig molecules. This is also one of several cases, discussed in detail below, where a given cellular state of B cell differentiation, in this case the small pre-B-II cell stage, can be reached without concomitant Ig gene rearrangements and expressions.
Vk Gene Segment Usage Rearrangements at the kL chain locus occur randomly in- and out-of-frame (Yamagami et al., 1999b). There is no strong preference for usage in Vk to Jk rearrangements of any Vk segments within the locus (Kirschbaum et al., 1996, 1998, 1999; Roschenthaler et al, 1999; Schable et al., 1999; Thiebe et al., 1999; Andersson J., Yamagami T., and Melchers F., unpublished results). However, different Vk segments within the locus are differently accessible for V(D)J recombination. Thus, deficiency in the OcaB enhancer of Ig gene transcription does not allow a subset of Vk gene segments to be rearranged in small pre-B-II cells (Casellas et al., 2002). Although the absence of OcaB still allows DNA methylation and histone acetylation, a subset of Vk segments is no longer transcribed efficiently enough to allow V(D)J recombination.
Vk Jk Rearrangements at a Single Allele In a wildtype mouse, about half of the surface Ig positive (sIg+) B cells have rearranged only one kL chain allele, preferentially to Vk1, the Vk proximal J segment. The other allele remains in germline configuration (Yamagami et al., 1996). In marked contrast, small pre-B-II cells show a strongly increased frequency of multiple kL chain rearrangements. These multiple rearrangements are frequently found on one allele, seen most clearly in wildtype/JCk–deficient F1 heterozygous B lineage cells (Yamagami et al, 1999a, b). Single cell PCR analyses can track the individual rearrangements to specific sites within the Vk cluster of gene segments (Kirschbaum et al., 1996, 1998, 1999; Roschenthaler et al., 1999; Schable et al., 1999; Thiebe et al., 1999) and order them in the sequence by which they have taken place before.
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It is evident that VkJk rearrangements can start almost anywhere within the cluster, but subsequent rearrangements are most often found in closer proximity within the Vk gene cluster (Andersson J., Yamagami T., and Melchers F., unpublished results). Taken together these results suggest that Vk to Jk rearrangements start at one allele and continue, if needed, on the same allele with a much higher probability and within closer proximity of the starting point for rearrangements than on the other allele. In most cases, therefore, the first allele may be used up by rearrangement to the RS sequences before the second allele is used.
Multiple VL-JL Rearrangements in Single B Lineage Cells and L Chain Editing Within multiple Vk-Jk rearrangements at a single allele, both nonproductive and productive rearrangements have been detected. Within a tracked sequence of rearrangements (i.e., first to Vk1, then to Vk2, and so on), productive rearrangements may be followed by nonproductive ones (Yamagami et al., 1999b). This indicates that an L chain could have been produced in these cells, but did not stabilize that cell as an sIg+ B cell. Also, it was found that one fifth of all small pre-B-II cells express kL chains in their cytoplasm, but not on the surface, although half these cells carry productive Vk-Jk rearrangements. Since over 95% of the small pre-B-II cells express mH chains in their cytoplasm, we can think of at least two reasons why these cells are not sIg+ and not already in the pool of immature, sIg+ B cells (figure 7.2). One possibility is that the L chains do not pair with the particular mH chain expressed in that pre-B-II cell. The other possibility is that the L chain has paired and formed an sIg, but one that has recognized an autoantigen in the environment of the bone marrow. Binding of autoantigen would result in the downregulation of surface expression of the Ig. Such secondary VL-JL rearrangements, induced by autoantigens in immature B cells, have been demonstrated with mouse B cells (Tiegs et al., 1993; Prak et al., 1994; Gay et al., 1993; Radic et al., 1993) and are likely in human B cells (Dorner et al., 1998). Secondary rearrangements induced by autoantigen recognition at high avidity (figure 7.2) would give the B lineage cell at the interphase between an immature and small pre-B-II the chance to change the specificity of its BcR away from autoantigen recognition—to “edit” its receptor—and, avoid death by apoptosis. It is not easy to estimate how much “editing” contributes to the total frequency of secondary VL-JL rearrangements (Prak and Weigert, 1995; Retter and Nemazee, 1998). Editing is, in part, achieved also by VL replacements, rather than secondary VL-to-JL rearrangements (Casellas et al., 2001). In this context, it should also be noted that such secondary rearrangements are often found, to a similar extent and in similar frequencies, in Ck-deficient mice,
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which can rearrange Vk to Jk, but which cannot make a kL chain (Yamagami et al., 1999b). These results, in fact, argue for the alternate, nonexclusive possibility that nonproductive and nonfitting rearrangements do not turn off the rearrangement machinery, which is constitutively expressed in pre-B-II and immature B cells until an sIg+ B cell has been generated. This sIg+ B cell is not high avidity-autoreactive and can therefore either enter the B1 or the conventional B cell pathway of differentiation (figure 7.2) (Harada and Yamagishi, 1991; Hertz et al., 1998; Shimizu et al., 1991; King and Monroe, 2001; Yu et al., 1999; Monroe et al., 1999). In conclusion, secondary L chain gene rearrangements are expected to be in part BcR stimulation–dependent, and also independent of this stimulation.
Multiple Vk-Jk Rearrangements in lL Chain-Producing B Cells Secondary Vk to Jk rearrangements, down to Jk4, Jk5 and RS, are accumulated in Vl-to-Jl–rearranged, lL chain–expressing sIgM+ immature and mature B cells. In the mouse, this could be expected from the observed rate differences for rearrangements at the kL and lL chain gene loci (Yamagami et al., 1999a, b). Over 95% of the lL chain+ B cells carried such kL chain rearrangements, and these were already found in immature B cells in the bone marrow, suggesting that these secondary rearrangements occurred during primary B cell development and not during antigen-driven peripheral B cell responses. It is more surprising that in the human over 95% of the lL chain–producing B cells carry such secondary Vk-to-Jk rearrangements (again, to Jk4, Jk5, and RS), and in nonproductive as well as productive forms (Bräuninger et al., 2001). In contrast to mice, where 95% of all immature and mature B cells express kL chains and 5% express lL chains, 40% of human B cells express lL chains and 60% express kL chains. Unless the repertoires of lL chain+ B cells and of kL chain+ B cells are subjected to very different positive or negative selective pressures in mouse and human, these results cannot be explained by simple rate differences of rearrangements at the two L chain loci in the human. They suggest that pre-B-II cells at the transition from large cycling to small resting cells rapidly induce multiple kL chain gene rearrangements at one allele before they enter, perhaps more slowly, lL chain gene rearrangements (Engel et al., 1999).
IMMATURE B CELLS Immature B cells are characterized and distinguished from mature B cells by a number of properties. They express IgM, but little if any IgD on their surface; do not yet express CD21 and CD23; express the Clq-like receptor (AA4.1)
recognized by the mAb 493; turn over rapidly (with halflives of 2 to 4 days); and respond to IgM-specific mAb in vitro not by proliferation, but by apoptosis (reviewed in Melchers and Rolink, 1999). Of importance, they continue to express RAG-1 and RAG-2. Hence, they are capable of continued secondary rearrangements at the L chain gene loci and possibly also of V gene replacements at the H and L chain loci. Immature B cells are found in bone marrow and spleen. However, it might be that only the immature cells in bone marrow are capable of secondary L chain gene rearrangements or “editing” (Sandel and Monroe, 1999).
Vk-to-Jk Rearrangements in Immature B Cells Cells with a single Vk-to-Jk rearrangement are more frequent in immature and mature B cells (40 to 45%) than in small pre-B-II cells (25%) (Yamagami et al., 1999). Thus, secondary rearrangements at the kL chain loci are more frequent (over 60%) in small pre-B-II cells than in immature or mature B cells (30 to 40%). This indicates that cells with a single Vk-to-Jk rearrangement are preferentially chosen into the short-lived immature, and later into the longer-lived mature, B cell pools, whereas secondary rearrangements continue in small pre-B-II cells, the precursors to the immature and mature B cells, as long as they live and are not chosen to become sIg+ B cells. The most frequent rearrangement in the immature and mature B cells is to Jk1 (25 to 30%, but only 10% in small pre-B-II cells). However, in the kL chain expression–deficient Ck-/JCk- mice, in which the Ck- allele can still undergo Vk-Jk rearrangements without L chain expression— that is, without selection by protein—these 10% Vk to Jk1 rearrangements are found not only in pre-B-II cells, but also in (lL chain+) immature and mature B cells. These results indicate that in wildtype kL allele–containing mice, kL chain+ sIg+ B cells are preferentially selected at the transition from pre-B-II to immature (and mature) B cells by the expression of kL chain+ surface IgM. It suggests that Vk to Jk rearrangements begin with those to the Jk1 segment most proximal to the Vk cluster, and have thus an advantage to be preferentially selected into the sIg+ B cell pools.
Rapid Selection of Successful Vk-to-Jk Rearrangements and Allelic Exclusion at the L Chain Gene Loci In immature and mature B cells of wildtype kL chain allele homozygous mice, almost 70% of all cells have one kL chain allele and the lL chain alleles in germline configuration. In the same cells, more than half have one or several secondary rearrangements at the same, rearranged allele. It suggests that kL chain gene rearrangements begin at one allele while the second allele remains inaccessible even for
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
secondary rearrangements (Yamagami et al., 1999; Mehr et al., 1999; Prak and Weigert, 1995). Demethylation and histone acetylation studies of the rearranging and nonrearranging kL chain alleles suggest that only one allele is initially accessible for the rearrangement machinery (Mostoslavsky et al., 1998). If the rearrangement of Vk-to-Jk1 on the first allele is 1) the first event in L chain gene rearrangements, 2) occurs randomly in- and out-of-frame, 3) all L chains made in this way pair with mH chains in such cells, and 4) more of the sIgM formed by this process is autoreactive, one would expect 33% of all immature cells to be of this type. However, only 8 to 10% were found, indicating that many of the L chains initially formed either cannot pair or generate an autoreactive BcR, so that secondary rearrangements occur to try to correct this initial L chain expression.
SELECTIONS OF IMMATURE B CELLS Adult mice produce approximately 2 ¥ 107 immature B cells per day (Osmond, 1991). Between 10 and 20% of these immature B cells, made in the bone marrow, migrate to the spleen (Allman et al. 1993; Rolink et al., 1998). They enter through the terminal branches of central arterioles and arrive in the marginal zone blood sinusoids (MacLennan and Chan, 1993), from where some of them then also penetrate into the outer zone of the periarteriolar lymphocyte sheath (PALS). There they become part of the B cell–rich follicular areas (MacLennan and Gray, 1986; Lortan et al. 1987). Hence, the largest loss of sIg+ B cells occurs at the transit from the bone marrow to the spleen. In fact, no mutations are known so far that affect the transition from small pre-B-II to immature B cells in bone marrow, whereas several mutations block the transfer of immature B cells from the bone marrow to the spleen (Rolink et al., 1999; Schubart et al., 1996, 2000, 2001; Oka et al., 1996). The mutations, which involve either OBF together with Oct-2 or btk deficiencies, or CD40 together with btk deficiencies, or a deficiency in the Aa chain of MHC class II molecules (figure 7.2), do not yet reveal the molecular mechanisms of this inhibition. Since crosslinking of the BcR on immature B cells induces apoptosis (Rolink et al., 1998), it is not unlikely that this loss of immature cells may be due to the recognition of autoantigens causing deletion of the autoreactive repertoire within the immature B cells. If this were the cause of deletion, it would predict that the loss due to autoreactivity of immature B cells in the spleen at the transition to mature B cells should be minimal, since practically all immature B cells become mature (Rolink et al., 1998). It is interesting to note that immature B cells in bone marrow and in spleen, and mature B cells in spleen, do not again change the rearrangement status of frequencies of secondary Vk-Jk rearrangements during these stages of B cell
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development (Yamagami et al., 1996). Hence, if the editing of kL chains by secondary rearrangements occurs as a response to autoantigen recognition, it affects the small preB-II, but not the immature B cell repertoire (and all subsequent repertoires), in a detectable way. The repertoires of immature B cells that originally emerge from small pre-B-II cells all must to express Ig on their surface in order to be able to transit into the peripheral B cell compartments – sIg- B cells are normally not found in the periphery (Lam et al., 1997). For any antigen, these repertoires are expected to contain a collection of cells expressing BcRs with varying avidities affinities. Depending on these avidities affinities, immature B cells generate a range of different responses (Kouskoff et al., 1998). Three major types of immature B cell reactions can be distinguished: induction of apoptosis or anergy (Nossal and Pike, 1980; Nemazee and Buerki, 1989; Goodnow et al., 1989), induction of an antigen-excited, “tickled” state with increased survival (discussed in Pillai, 1999, and in Potter and Melchers, 2000), or retention of the original short-lived state, due to lack of recognition.
Negative Selection by Arrest of Differentiation and Induction of Anergy and Apoptosis The exposure of immature B cells to antigen at high avidities results in the downregulation of expression of sIgM and B220 (CD45R). This creates an sIgMlow B220low “transitional” cell that begins to express the CD21 and CD23 not expressed on the original immature cells (Carsetti et al., 1995). Development is the arrested at this point, as demonstrated in vitro and in vivo with immature B cells from transgenic mice expressing hen egg lysozyme (HEL) (Hartley et al., 1993). It has yet to be discovered how such autoantigens are presented in the primary lymphoid organ to the immature B cells, but it appears that membrane deposition of HEL is helpful for negative selection. A special autoantigenpresenting cell type has not yet been seen. The arrest of differentiation is most clearly seen when the developing B cells express only a transgenic, autoantigenspecific BcR, but cannot express endogenously rearranged Ig genes, as in RAG-deficient hosts. Exposure of such BcRtransgenic immature B cells to the fitting autoantigen in bone marrow results in the arrest of differentiation and apoptosis of the arrested cells, so that the peripheral B cell compartments remain empty. In the absence of the autoantigen, these peripheral B cell compartments are filled with transgenic monoclonal BcR-expressing B cells (Hartley et al., 1993). Antigens that crossreact with the deleting autoantigens, and which are administered to the primary lymphoid organs at the sites of negative selection, may be able to interfere with this deletion process. Thus, in a RAG-deficient mouse
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expressing a transgenic mAb specific for TNT (trinitrophenyl) and crossreactive with double-stranded (ds) DNA, developing B cells are normally arrested, probably by the exposure to dsDNA in the bone marrow. Injection of the T cell–independent antigens TNP-Ficoll relieves this inhibition, so that large numbers of transgenic, anti-TNP (dsDNA)–producing B cells appear in the periphery (Andersson et al., 1995).
Positive Selection of Immature B Cells into the B1 Cell Compartment In analogy to T cell development, where a low avidity interaction of T cell receptors on T cells with MHC peptide complexes on antigen-resenting cells is required for longterm survival of the T cells in the periphery (Takeda et al., 1996), autoreactive B cells can be positively selected (Hayakawa et al., 1999) into the B1 compartments of the peripheral pools of mature B cells (Hayakawa et al., 1986; Herzenberg and Kantor, 1992). Experiments by Martin and Kearney (2000) suggest that the marginal zone of spleen contains such positively selected B cells. B1 cells express BcR and are often found to crossreact with a variety of other autoantigens, as well as with foreign antigens, often of bacterial origin. B1 cells, furthermore, appear in a lowly activated, “tickled” state, in which they do not divide in response to foreign antigens, as proliferating B cells do in germinal centers, but which apparently allows them to escape from the short half-life of a previously immature B cell. The continuous presence of autoantigens in the periphery allows them to maintain this state, and transplantation into secondary hosts will let them keep this state, thus making them easily transplantable cells. Their continuous “excited” state may also make them a prime site for further transformation events leading to B cell malignancies (discussed in Potter and Melchers, 2000). Activation by crossreactive bacterial infections could induce not only the secretion of Ig molecules as a first line of defence against an infection, but may also result in autoimmune disease manifestations on the basis of an apparent antigenic mimicry between autoantigens and antigens of infectious agents (Oldstone, 1989). B1 cells have the tendency to migrate to peripheral sites outside lymphoid organizations such as the B cell follicles in spleen, lymph nodes, and gut-associated lymphoid follicles. These are often seen as single cells in the epithelia, for example, in the lamina propia in the gut. Whether the homing of these cells to such sites is influenced by their BcR specificities remains to be investigated. Although B1 cells may be initially positively selected by autoantigens of low avidities without the help of T cells, such low avidity autoreactive cells may also arise in a germinal center response of follicular B cells to T cell–dependent antigens. In this case, switched, hypermutated B cells may be generated, which
occasionally and by chance, gain low avidity specificity for an autoantigen, away from the foreign antigen that stimulated the germinal center response (discussed in Potter and Melchers, 2000). B1 B cells are most clearly distinguished from the conventional B cell populations by their apparent inability, or insufficiency, to be stimulated by interactions of the B cell–specific TNF ligand family member, BAFF, with its TNF receptor family member, BAFF-R (reviewed in Rolink and Melchers, 2002).
Selection of the Ignored Immature B Cells into Mature, Long-Lived Conventional B Cell Compartments The TNF family ligands BAFF and APRIL, and their receptors BCMA, TACI, and BAFF-R control the selection of short-lived immature B cells with no apparent positively (or negatively) selecting specificities for autoantigens to long-lived mature B cells. Experiments of the in vivo administration of soluble BAFF-R ligands and of soluble decoy receptors, and the analysis of BAFF-transgenic, BAFF-deficient and BAFF-receptor (BAFF-R)–deficient mice, as well as the in vitro responses of immature and mature B cells to BAFF (all reviewed in Rolink and Melchers, 2002) have shown that immature B cells from bone marrow and spleen (initially immature as well as “transitional” B cells) and mature B cells respond to BAFF by polyclonal maturation to long-lived B cells without proliferation. BAFF and BAFF-R deficiencies arrest B cell development at the transition from immature to transitional B cells. Although the action of BAFF in vitro is polyclonal and independent of BCR occupancy, it remains to be seen whether ligand selection through BCR occupancy plays a role in this selection of the conventional “virgin” antigenreactive mature B cells.
Pre-BcR and BcR-Independent B Cell Development B lineage cells that cannot express Ig molecules on their surface are restricted to the primary lymphoid organs and will die there. Ablation of the expression of surface-bound Ig in peripheral, mature B cells induces their rapid death (Lam and Rajewsky, 1997). Therefore, neither immature nor further differentiated B lineage cells, down to the memory and plasma cell phenotypes, are ever detectable in the peripheral immune system without expressing Ig. However, several observations, many of them made in vitro, suggest that B lineage cells from the pre-B-I cell stage (and maybe even from earlier stages of pL cells) can differentiate all the way to a mature, memory type B lineage cell without ever expressing Ig. First, L chain gene rearrangement can be induced from pre-B-I cells that have never
7. Early B Cell Development to a Mature, Antigen-Sensitive Cell
expressed mH chains (i.e., a preBcR), simply by removing IL-7. This allows their differentiation to pre-B-II and immature B cells (Grawunder et al., 1993). The transgenic expression of constitutively active forms of ras (Shaw et al., 1999) or raf (Iritani et al., 1999) induce RAG-deficient precursor B cells to develop pre-B cells with pre-B-II and immature B cell-like phenotypes. H chain rearrangement–deficient cells readily progress under such signaling to L chain gene rearrangements in small Pre-B-II-like cells. The most striking example is the development of RAG-deficient pre-B-Ilike cells in vitro, used by the removal of IL-7 and under the stimulation by CD40-specific mAb and IL-4, to sm-seswitched cells (having no V(D)J-rearranged H or L chain loci) of mature phenotype (Rolink et al., 1996) (Figure 7.2). This extreme flexibility of B lineage cells indicates that the differentiation of cells, including class switching, is controlled by cell–cell contacts and cytokines provided by cooperating cells. The action of BAFF may be one example of such in vivo action. The roles of pre-BcR and BcR also become apparent from such a scenario: These receptors control, positively and negatively, the proliferation (or anergy and apoptosis) of B lineage cells and, thereby, ascertain the provision of normal numbers of B cells in the immune system.
CONCLUSION The development of B lineage cells from early progenitors to mature, antigen-reactive cells and their controls by molecular actions must be one of the best described cellular pathways, within the body of a vertebrate. Nevertheless, it is evident that our descriptions only touch the surface and deeper probing will further clarify this development. With all genes of the human genome already known, and of the mouse genome soon to be known, the cellular stages of this development defined by gene expression at RNA (Hoffmann et al., 2002), the protein and post-translational modifications of proteins will define better all possible cellular stages and their plasticity, especially when all cells can be individually accounted and described in such a way. However, it is probable that we will not be able to predict the full capacities and reactivities of all cells in this system at any given time, especially since all of its member cells turn over at different rates and are under the influence of an uncontrollable environment that might influence their plasticity and reactivity. The better we understand this developmental cell system, the more the uncertainty of the description of the whole system becomes apparent.
Acknowledgments Fritz Melchers is supported by a research grant from the Swiss National Funds (3100-066682.01/1).
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8 Allelic Exclusion, Isotypic Exclusion, and the Developmental Regulation of V(D)J Recombination MICHAEL S. KRANGEL
MARK S. SCHLISSEL
Department of Immunology, Duke University Medical Center, Durham, North Carolina, USA
Department of Molecular and Cell Biology, Division of Immunology, University of California, Berkeley, California, USA
Antigen receptor gene assembly occurs in a highly regulated fashion that is closely coordinated with the complex developmental programs of early B and T lymphocytes (Muljo and Schlissel, 2000) (Figure 8.1). In the B cell lineage, V(D)J recombination begins at the Ig heavy chain locus, with D-to-J rearrangement occurring first, and V-toDJ rearrangement occurring subsequently. Pro-B cells that generate an “in-frame” VDJ heavy chain allele express a signaling complex known as the pre-B cell receptor (BCR), which consists of a clonotypic heavy chain, the surrogate light chains VpreB and l5, and the accessory chains Ig-a and Ig-b. Pre-BCR signaling results in clonal expansion and the temporary cessation of V(D)J recombination. Late preB cells exit the cell cycle and initiate rearrangement at the Igk and Igl light-chain loci, leading ultimately to the production and surface expression of a BCR. The early development of ab T cells is strikingly similar, with TCRb and TCRa gene segments rearranging in double negative (DN) and double positive (DP) thymocytes, respectively. Thus, V(D)J recombination events at Ig and TCR loci are regulated according to cell lineage and developmental stage. Moreover, as envisaged in the clonal selection hypothesis, each lymphocyte should be restricted to express a single antigen receptor (Burnett, 1959). B cells typically express Igk or Igl, but rarely both, a phenomenon known as isotypic exclusion (Bernier and Cebra, 1964); they productively rearrange only a single allele per locus, a phenomenon known as allelic exclusion (Weiler, 1965; Pernis et al., 1965). Here we explore current knowledge regarding the mechanisms that impart developmental control to V(D)J recombination and that yield allelically and isotypically excluded antigen receptor repertoires.
RAG EXPRESSION
Molecular Biology of B Cells
The lymphoid specificity of V(D)J recombination reflects the regulated expression of recombinase proteins RAG1 and RAG2 (Oettinger et al., 1990). Although reports exist documenting RAG expression in nonhematopoietic tissues, transcripts are either at too low a level to support recombinase activity or one RAG protein is expressed without the other. RAG gene expression likely begins at a stage of hematopoiesis just prior to lymphoid commitment, accounting for the occasional presence of DJH rearrangements in NK cells (Igarashi et al., 2002). Variation in RAG gene expression and protein stability accounts for the two waves of V(D)J recombination events in developing B and T lymphocytes (Figure 8.1). RAG gene expression is high in pro-B cells and in double negative (DN) T cells, is downregulated by pre-BCR and pre-TCR signaling, and is upregulated in late-stage pre-B cells and in double positive (DP) T cells (Wilson et al., 1994; Grawunder et al., 1995). In developing T cells, RAG gene transcription is inactivated upon positive selection (Borgulya et al., 1992; Brandle et al., 1992). In the B cell lineage, IgM+ IgD- immature B cells express RAG mRNA while IgMlo IgDhi mature B cells do not (Grawunder et al., 1995), but the signals that result in RAG inactivation are not well understood. Finally, although there were reports of RAG gene reactivation in both peripheral B and T cells (Han et al., 1997; Hikida et al., 1996; McMahan and Fink, 1998), the data have not been supported by more recent studies (Monroe et al., 1999a; Yu et al., 1999).
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ACCESSIBILITY HYPOTHESIS
FIGURE 8.1 Schematic comparing gene rearrangement events during early B and T cell development. IgH and Igk rearrangements are depicted in developing B cells; Igl rearrangement, not shown, occurs in late-stage pre-B cells and immature B cells. TCRb and TCRa rearrangements are depicted in developing T cells; TCRg and TCRd rearrangements, not shown, occur in DN thymocytes. Two periods of RAG gene expression are indicated. Pre-BCR and pre-TCR expression result in feedback inhibition of VDJH and VDJb rearrangement, respectively, as well as developmental transition and proliferative expansion.
THE 12/23 RULE As noted in previous chapters, V(D)J recombination occurs only between pairs of gene segments flanked by dissimilar RSSs, the so-called 12/23 rule. This biochemical constraint on the recombination reaction is critical for several aspects of regulation. First, and most obviously, the 12/23 rule prevents recombination between different members of the same class of gene segments. For example VH-to-VH rearrangement does not occur. Second, in the IgH locus, which undergoes two distinct recombination events, the disposition of VH, DH, and JH RSSs ensures the inclusion of the DH gene segment. Interestingly, this is not the case for TCRb, where Vb-to-Db-to-Jb and direct Vb-to-Jb rearrangement would both be permitted according to the 12/23 rule. Nevertheless Vb-to-Jb rearrangement is not observed, and recent work has shown that individual RSS sequences can regulate gene-segment utilization at a level beyond the simple 12/23 rule. Specifically, the 5¢ Db 12-RSS is a much more efficient rearrangement partner for a diverse repertoire of Vb 23-RSSs than are the Jb 12-RSSs, independent of the positions of these sequences within the TCRb locus (Bassing et al., 2000; Sleckman et al., 2000). There is uncertainty as to the precise mechanisms that enforce both the 12/23 rule and restrictions “beyond 12/23.” With respect to 12/23 regulation, although RAG1 and RAG2 alone show a preference for 12/23 restricted pairwise RSS cleavage in vitro, nonhistone chromosomal proteins HMG1 and HMG2 and perhaps other nuclear factors may play important roles as well (van Gent et al., 1996; Sawchuk et al., 1997). However, in interpreting these experiments one must keep in mind that all biochemical studies of V(D)J recombination to date utilize only the “core” domains of RAG1 and RAG2. The behavior of full-length proteins may be different.
Neither RAG gene expression nor the RSS constraints noted above can account for the locus-specific developmental control of V(D)J recombination. Rather, developmental control is thought to occur largely through regulation of RAG protein access to RSSs within chromatin. Various investigators have observed that most if not all rearranging gene segments are transcribed prior to or coincident with the activation of their rearrangement potential (Sleckman et al., 1996; Schlissel and Stanhope-Baker, 1997; Hesslein and Schatz, 2001). These so-called “germline” transcripts serve as markers of rearrangement competence. Yancopolous, Alt, and co-workers first suggested that germline transcripts might correlate with the accessibility of RSSs to the recombinase within chromatin (Yancopoulos and Alt, 1985). Transcription could be a direct cause of chromatin accessibility or could reflect another process from which transcription and accessibility follow as independent consequences. The nature of this relationship is still uncertain and will be considered in greater detail later. Nevertheless, compelling evidence in support of the accessibility hypothesis has been obtained from experiments in which purified recombinant RAG proteins were used to perform in vitro RSS cleavage assays (Stanhope-Baker et al., 1996). RAG proteins recognize and efficiently cleave RSSs in oligonucleotide or plasmid substrates (McBlane et al., 1995). Using purified total genomic DNA as substrate, Stanhope-Baker et al. detected dsDNA breaks at RSSs from each of the Ig and TCR loci. In contrast, when nuclei purified from RAGdeficient pro-B cells were used as substrate, breaks were introduced into Ig gene RSSs but not TCR gene RSSs. When RAG-deficient DN thymocyte nuclei were used as substrate, the RAGs cleaved TCR RSSs but not Ig RSSs (StanhopeBaker et al., 1996). These and similar experiments led to the conclusion that RSS accessibility to the recombinase was a developmentally regulated property of chromatin structure.
ENHANCER AND PROMOTER CONTROL OF V(D)J RECOMBINATION Transcriptional Enhancers Stimulated by the correlation between germline transcription and V(D)J recombination, much attention has been focused on the roles of transcriptional control elements as regulators of V(D)J recombination (Figure 8.2). These studies have made use of chromosomally integrated V(D)J recombination reporter substrates in transfected cell lines and transgenic mice, as well as gene targeting at endogenous Ig and TCR loci. Gene targeting experiments have been instrumental in establishing that V(D)J recombination is critically
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the redundancy issue. Finally, combined elimination of 3¢ECg1 and a second element, HsA, from the Cg1 gene cluster of the TCRg locus had only a modest effect on Vgto-Jg rearrangement (Xiong et al., 2002). Other linked enhancers may provide redundant activity. The use of transgenic reporters has been instrumental in demonstrating that transcriptional enhancers impart developmental control to V(D)J recombination. For example, Em was shown to stimulate D-to-J rearrangement within a transgenic minilocus in developing B and T lymphocytes, reflecting the lineage-nonspecific nature of endogenous DH-to-JH rearrangement (Ferrier et al., 1990). Eb and Ed were shown to direct developmentally appropriate transgenic minilocus rearrangement in DN thymocytes. In contrast, Ea directed the rearrangement of the same constructs in DP thymocytes (Capone et al., 1993; Lauzurica and Krangel, 1994; Hernandez-Munain et al., 1999). FIGURE 8.2 Schematic depicting the organization of gene segments and cis-acting regulatory elements at murine Ig and TCR loci. Gene segments are identified by filled rectangles, promoters are identified by bent arrows, and enhancers and other regulatory elements are identified by filled ovals. Only promoters discussed in the text are identified. The Igl locus and portions of the TCRg locus are not shown because detailed information about cis-acting regulators of V(D)J recombination is lacking. The diagram is not drawn to scale and does not accurately represent gene segment numbers.
dependent on transcriptional enhancers. For example, Eb was shown to be necessary for both Db-to-Jb and Vb-toDJb rearrangement at the endogenous TCRb locus (Bories et al., 1996; Bouvier et al., 1996). Similarly, Va-to-Ja rearrangement at the TCR a/d locus was shown to depend critically on Ea (Sleckman et al., 1997). In other instances, the results have been more complex, presumably due to functional redundancy among regulatory elements. Thus, at the Igk locus, Vk-to-Jk rearrangement was significantly impaired by the targeted deletion of iEk, but elimination of both iEk and 3¢Ek was required to abolish Igk rearrangement (Gorman et al., 1996; Xu et al., 1996; Inlay et al., 2002). TCRd rearrangement was found to be inhibited but not completely blocked in Ed knockout mice, suggesting redundancy with another element (Monroe et al., 1999b). Interestingly, targeted deletion of Em had minimal effect on DH-to-JH rearrangement, although VH-to-DJH rearrangement was strongly inhibited (Serwe and Sablitzky, 1993; Sakai et al., 1999). Other elements may function redundantly with Em to regulate the DH-to-JH step. One candidate is the 3¢ IgH regulatory region, which is important for class-switch recombination (Cogne et al., 1994). Another candidate is the DQ52 promoter, which flanks the most JH-proximal DH segment and displays intrinsic enhancer activity (Kottmann et al., 1994). Gene targeting revealed this promoter to mildy influence IgH rearrangement (Nitschke et al., 2001), but elimination of both DQ52 and Em will be required to clarify
Transcriptional Promoters Transcriptional enhancers appear to function, at least in part, by activating germline promoters. These promoters then appear to influence V(D)J recombination in a relatively localized fashion. For example, TEA is an Ea-dependent germline promoter situated upstream of the Ja cluster. Whereas elimination of Ea impaired Va rearrangement to the entire array of Ja segments, elimination of TEA impaired rearrangement to the most 5¢ Jas only (Villey et al., 1996). Therefore, TEA functions as a local, Ea-dependent regulator of V(D)J recombination. A similar role is played by the germline TCRb promoter PDb1. This promoter was shown to be required for all rearrangements involving the Db1-Jb1 cluster, but to be irrelevant for rearrangements involving the Db2-Jb2 cluster (Sikes et al., 1999; Whitehurst et al., 1999). The influence of PDb1 is local and depends critically on its position relative to nearby RSSs (Sikes et al., 2002). Igk rearrangement was inhibited by deletion of either the KI/KII motifs that are associated with a proximal Jk promoter, or deletion of a distal Jk promoter region; simultaneous elimination of both elements produced the most dramatic inhibition (Ferradini et al., 1996; Cocea et al., 1999). Finally, promoters can impart developmental control to V(D)J recombination: the developmental pattern of Vg rearrangement in adult thymocytes was modified by exchange of the Vg2 and Vg3 promoter regions within a transgenic reporter (Baker et al., 1998).
Accessibility and Beyond In most cases that have been examined, promoter or enhancer deletion results in an inhibition of rearrangement at the earliest step, the formation of double strand breaks between RSSs and coding segments (McMurry et al., 1997; Whitehurst et al., 2000). This is as would be expected for an
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effect of promoters and enhancers on accessibility to RAG. Interestingly, however, gene-targeted elimination of Eb resulted in a dramatic impairment in coding joint formation that could not be fully accounted for by the observed reductions in signal ends and signal joints (Hempel et al., 1998). Although not yet detected for other enhancers, this apparent effect of Eb on the joining step of V(D)J recombination opens the possibility that cis-acting elements might regulate recombination at levels beyond that of template accessibility.
TRANS-ACTING FACTORS Although it has been relatively straightforward to establish the importance of cis-regulatory elements for V(D)J recombination, it has been more difficult to critically evaluate the identities and roles of the specific factors that are recruited to these elements. The introduction of specific enhancer and promoter mutations in V(D)J recombination reporter constructs has provided some insight, but the factors that function at these sites in vivo have often not been unambiguously established. Interpreting the data from transcription factor knockout mice has been problematic due to either factor redundancy, developmental perturbations, or the potential for other indirect effects. Moreover, although the ectopic expression of transcription factors in cell lines has yielded interesting data, the direct targets of these factors typically have not been identified. For example, much evidence implicates bHLH proteins E2A and HEB as regulators of V(D)J recombination at Ig and TCR loci (Quong et al., 2002). The activation of DH-to-JH rearrangement may reflect E protein binding to critical sites in Em (Fernex et al., 1995). However, the E protein targets that influence Ig and TCR V segment rearrangement are undefined. Igk and TCRa rearrangements are activated across welldefined and experimentally accessible developmental transitions by pre-BCR and pre-TCR signaling, respectively. Despite this, it has been difficult to define the specific factors that trigger locus activation. For example, although NF-kB is an important regulator of iEk, in vivo footprinting experiments indicated that its binding site is equivalently occupied in pro- and pre-B cells (Shaffer et al., 1997). Moreover, although there are developmental changes in occupancy of binding sites for Pax-5, CREB, and PU.1 within 3¢Ek (Shaffer et al., 1997), the developmental onset of Igk rearrangement appeared normal in 3¢Ek knockout mice (van der Stoep et al., 1998). In the case of TCRa, there is a single enhancer that triggers Va-to-Ja rearrangement as thymocytes differentiate from DN to DP (Sleckman et al., 1997; Hernandez-Munain et al., 1999). Ea-binding proteins LEF1 and TCF-1 are known to function redundantly to coordinate the assembly of a multiprotein complex on Ea and to permit locus transcription and rearrangement in vivo (Giese
et al., 1995; Okamura et al., 1998). Nevertheless, occupancy of binding sites for these and other factors is virtually identical in DN thymocytes, in which Ea is inactive, and in DP thymocytes, in which it is active (Hernandez-Munain et al., 1999; Spicuglia et al., 2000). Enhancer activation might occur by post-translational modification of an enhancerbound factor, or by the association of an enhancer-bound factor with a DNA-nonbinding co-activator protein. Against this background, two recent sets of experiments are of particular interest. Several studies have shown that TCRg locus transcription, rearrangement, and accessibility are all dependent on IL-7R signaling (Maki et al., 1996; Durum et al., 1998; Schlissel et al., 2000). An elegant series of experiments using both cell lines and fetal thymus organ culture has made a very strong case for Stat5 to function downstream of the IL-7R as a direct regulator of TCRg rearrangement (Ye et al., 1999; Ye et al., 2001). Activated Stat5 was shown to bind to both the germline Jg1 promoter and Eg, and to influence promoter function and regional chromatin structure. The second set of experiments involves Oca-B, a transcriptional co-activator that associates with transcription factors Oct-1 and Oct-2, which bind to octamer motifs in Vk promoters. Oca-B-/- mice displayed relatively normal B cell development through the pre-B cell stage. However, the mice displayed reduced transcription and rearrangement of a subset of Vk segments that have relatively weak promoters (Casellas et al., 2002). It appears likely that Oca-B directly regulates k rearrangement through interactions with these promoters.
CHROMATIN DYNAMICS AND V(D)J RECOMBINATION Chromatin Structural Modifications Chromatin consists of genomic DNA noncovalently associated with a series of histone and nonhistone proteins (Workman and Kingston, 1998). The basic building block of chromatin structure is the nucleosome, an octamer consisting of two molecules each of histones H2a, H2b, H3, and H4. The histone octamer forms a disk-like structure around which 146 bp of DNA is wrapped twice. Genomic DNA is packed into long arrays of nucleosomes, which then undergo multiple higher levels of compaction, ultimately resulting in the packaging of ~1 meter of DNA into a nucleus only several microns in diameter. Nucleosome structure places a severe constraint on the accessibility of DNA sequences to certain DNA binding proteins and enzymes. For example, in vitro transcription of RNA from well-characterized promoter sequences is strongly inhibited by the assembly of the substrate into a nucleosomal structure. The inhibition of DNA reactivity by nucleosome packaging can be overcome by at least two means: post-
8. Allelic Exclusion, Isotypic Exclusion, and the Developmental Regulation of V(D)J Recombination
translational histone modification and ATP-dependent chromatin remodeling activities (Narlikar et al., 2002). Histones can be modified by acetylation, methylation, and phosphorylation. Many studies have documented striking correlations between specific post-translational modifications and gene activity, leading to the concept of the “histone code,” namely, that the precise modification of histones at specific amino acid residues is a major determinant of gene activity (Strahl and Allis, 2000; Jenuwein and Allis, 2001). Perhaps the best studied histone modification is the acetylation of lysine residues in the amino terminal tails of histones H3 and H4. This is a reversible modification that is regulated by the activities of both histone acetyltransferases (HATs) and histone deacetylases (HDACs), and one which can directly influence factor binding to nucleosomal DNA. ATPdependent chromatin remodeling complexes can also modify nucleosome structure, but do so in a noncovalent fashion (Narlikar et al., 2002). Some of these remodeling complexes can increase the availability of DNA on the surface of a nucleosome, whereas others can catalyze a physical displacement of nucleosomes. Both HATs and ATPdependent chromatin remodeling enzymes can be targeted to specific loci via protein–protein interactions with enhancer- or promoter-bound transcription factors, providing a link between enhancer and promoter activity and the structure of surrounding chromatin (Naar et al., 2001). The question of whether the RAG proteins can recognize and cleave an RSS that is stably associated with a nucleosome is controversial. One group showed that RSS cleavage is dramatically inhibited by nucleosomal association but could be increased by the addition of HMG-1, a known activator of the recombinase; by the acetylation of histones; or by the inclusion of an ATP-dependent chromatin remodeling activity (Kwon et al., 1998; Kwon et al., 2000). Another group found that nucleosomal RSSs were completely resistant to recombinase-mediated cleavage regardless of histone acetylation or the presence of HMG-1 (Golding et al., 1999). This latter result predicts that to be accessible to RAG binding and cleavage RSSs would have to be situated in the linker region that separates adjacent nucleosomes or in nucleosome-free gaps within chromatin. Nevertheless, the in vitro studies to date are compromised by the simple nature of the mononucleosomal substrate and the use of core rather than full-length RAG proteins. Future studies will need to assess more complex and more physiological components. Recent studies of a transgenic V(D)J recombination reporter and of endogenous TCR loci indicated that histone acetylation status correlates well with V(D)J recombination activity (McMurry and Krangel, 2000; Mathieu et al., 2000; Agata et al., 2001; Huang et al., 2001). Moreover, in several instances it was shown that the short-term culture of thymocytes in the presence of HDAC inhibitor trichostatin A (TSA) could increase levels of rearrangement (McBlane and
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Boyes, 2000; Mathieu et al., 2000; Agata et al., 2001; Huang et al., 2001). To the extent that the latter observations reflect direct effects of TSA on the particular loci, they suggest that histone acetylation may directly contribute to recombinase access. Nevertheless, a stably transfected V(D)J recombination reporter that contains an enhancer but no promoter was recently shown to be hyperacetylated but inaccessible to the recombinase (Sikes et al., 2002). Therefore, RSS packaging within nucleosomes containing hyperacetylated histones is not sufficient for accessibility. Additional promoterdependent remodeling events are required.
Germline Transcription Germline transcription is a consequence of enhancer and promoter function that has historically been correlated with competence for rearrangement (Sleckman et al., 1996; Schlissel and Stanhope-Baker, 1997; Hesslein and Schatz, 2001). However, whether transcription contributes directly to recombinase targeting has never been resolved. The assembly of transcription complexes at promoters and enhancers can result in local changes in chromatin structure in the absence of transcription (Kuo et al., 2000; Agalioti et al., 2000), suggesting that transcription and accessibility might be separable. On the other hand, transcriptional elongation can cause chromatin disruption at sites distal to a promoter (Brown and Kingston, 1997), and elongating RNA polymerase II complexes may contain associated HATs (Travers, 1999). Thus, transcription can directly influence chromatin structure. Despite the extensive correlations between germline transcription and competence for V(D)J recombination, there are numerous instances in which the two appear to have been dissociated. For example, Vb segments in a minilocus V(D)J recombination reporter were transcribed but did not rearrange in the developing B cells of transgenic mice (Okada et al., 1994). Certain truncated forms of Em and Eb were found to support transcription within a transgenic reporter construct, but could not support V(D)J recombination (Fernex et al., 1995; Tripathi et al., 2000). These examples suggest that recombinase accessibility may have requirements beyond those for transcriptional activation, but do not address whether transcription might play a role in accessibility. One study found that VH gene segments that appeared transcriptionally inactive in the subclones of a transformed RAG-/- pro-B cell line could still rearrange following RAG gene transfection (Angelin-Duclos and Calame, 1999). More recently, promoter inversion was shown to dramatically reduce germline transcripts in a transfected V(D)J recombination reporter, but had no effect on construct rearrangement (Sikes et al., 2002). However, neither these nor other studies can rule out that transcription occurring at low levels or in a fraction of cells could be critical for accessibility.
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DNA Methylation A substantial literature correlates the hypomethylation of CpG dinucleotides with transcriptional activity and hypermethylation of CpG dinucleotides with transcriptional inactivity (Bird, 2002). CpG methylation can inhibit gene expression by directly occluding transcription factor binding sites, but the more important influence is likely to depend on methyl-CpG binding proteins, which can associate with and recruit HDACs and other chromatin remodeling activities to hypermethylated DNA. Methylation was found to inhibit V(D)J recombination within reporter substrates in transfected cells (Hsieh and Lieber, 1992; Cherry and Baltimore, 1999). However, the most striking in vivo data has come from studies of the TCRb and Igk loci. Targeted deletion of promoter PDb1 resulted in a general increase in local DNA methylation that was associated with diminished recombinase accessibility (Whitehurst et al., 2000). Interestingly, methylation of a CpG dinucleotide within the Db RSS heptamer appeared to completely block Db-to-Jb rearrangement. This is unlikely to represent a general mechanism because CpG dinucleotides are rare in RSSs. The relationship of methylation to V(D)J recombination has been most intensively studied in the Igk locus. Bergman and colleagues found that CpG dinucleotides near the Jk segments are extensively methylated in non-B cells and become demethylated during early B cell development (Mostoslavsky et al., 1998). Demethylation is controlled by iEk and 3¢Ek (Mostoslavsky et al., 1998; Inlay et al., 2002). Interestingly, in many cells only one of the two k alleles was found to undergo demethylation, and this allele was found to be the preferred target of the recombinase (Mostoslavsky et al., 1998; Goldmit et al., 2002). As discussed below, mono-allelic demethylation may contribute to Igk allelic exclusion. However, because demethylation per se is insufficient to activate Igk rearrangement (Cherry et al., 2000), it may represent only one of several changes to chromatin associated with Igk locus activation. Moreover, demethylation is not always necessary for V(D)J recombination: Vb segments (Senoo and Shinkai, 1998; Mathieu et al., 2000) and Ja segments (Villey et al., 1997) rearrange despite being hypermethylated in vivo. Effects on V(D)J recombination and chromatin structure may vary according to the location and density of CpG dinucleotides.
Nuclear Localization Processes such as transcription occur in distinct subnuclear structures. The inspection of interphase nuclei shows that active and inactive genes tend to segregate into distinct nuclear subcompartments, with inactive genes segregated into foci associated with centromeric heterochromatin or to the nuclear periphery (Lamond and Earnshaw, 1998; Cockell and Gasser, 1999). Enhancers and other cis-acting
elements can prevent the localization of genes to heterochromatic regions, an effect that is dissociable from transcriptional activation per se (Francastel et al., 1999; Schubeler et al., 2000). Several recent studies have investigated whether changes occur in subnuclear localization of Ig loci that correlate with rearrangement or expression. Using in situ hybridization to interphase nuclei, the IgH and Igk loci were often found near the nuclear periphery in nonlymphoid and T cells, whereas such localization was rare in B cell lines and primary pro-B cell cultures (Kosak et al., 2002; Zhou et al., 2002). Additional studies showed that these peripherally localized alleles were not associated with constitutive heterochromatin, such as g-satellite DNA, but did seem to co-localize with the nuclear lamina. Movement of the Igk locus into the nuclear center occurred well before the activation of germline transcription or rearrangement, indicating that relocalization is not the proximal cause of k locus activation. Remarkably, it was also found that the two ends of the VH region, separated by about 1.5 megabases of DNA, were closer together in pro-B cells than in T cells (Kosak et al., 2002). This large scale reorganization may promote VH-to-DJH rearrangement by juxtaposing the two ends of the IgH locus. The potential relevance of this observation was enhanced by the finding that IgH locus condensation was greatly diminished in IL7Ra-/- mice. These mice have a defect in VH-to-DJH rearrangement that preferentially involves the distal VH segments (Corcoran et al., 1998). Additional work has shown that unexpressed Ig alleles are often localized near heterochromatic g-satellite sequences in activated mature splenic B cells, whereas active alleles are not (Skok et al., 2001). This observation is unlikely to contribute to the establishment of allelic exclusion, since in pro-B cell clones, neither IgH allele was associated with heterochromatin. Rather, this may represent a relatively late event associated with transcriptional silencing only.
ORDERED REARRANGEMENT WITHIN IG AND TCR LOCI A defined developmental order, D-to-J followed by V-toDJ, is a distinctive property of V(D)J recombination at both the IgH and TCRb loci. The mechanisms underlying this ordering are of particular interest because it is the second step that is tightly regulated in the context of allelic exclusion. We will review current knowledge regarding how developmental order is established before considering the allelic exclusion problem.
IgH A recent study of IgH chromatin structure has provided insight into the molecular basis for ordered IgH rearrange-
8. Allelic Exclusion, Isotypic Exclusion, and the Developmental Regulation of V(D)J Recombination
ment (Chowdhury and Sen, 2001). In pro-B cells of RAG2-/- mice, these investigators defined a 120-kb hyperacetylated chromatin domain that extends from the most 5¢ D segment, Dfl16.1, to downstream of Cm. Interestingly, VH gene segments were hypoacetylated in these cells, suggesting that ordered rearrangement is enforced by developmentally programmed accessibility that initially involves only the D and J segments. VH chromatin was found to be activated in two cirumstances. Distal VH gene segments were hyperacetylated when RAG-2-/- pro-B cells were cultured in IL-7, whereas proximal and distal VH segments were both hyperacetylated in wildtype pro-B cells (Chowdhury and Sen, 2001). The effect of IL-7 is consistent with previous work indicating the rearrangement of these VH segments to be impaired in IL-7Ra-/- mice (Corcoran et al., 1998). The basis for proximal VH activation is unclear; the authors proposed that there might be a requirement for prior DJH rearrangement (Chowdhury and Sen, 2001). The results of this study are significant because they mechanistically segregate the modification of DH and JH chromatin from the modification of VH chromatin. Although these data suggest that chromatin structure plays a primary role in ordering rearrangement, other factors may contribute. RAG proteins themselves were speculated to play a role based on the observation that full-length RAG2 efficiently stimulated both DH-to-JH and VH-to-DJH rearrangement in an AMuLV transformed pro-B cell line, whereas the truncated core RAG2 was preferentially impaired in its ability to stimulate VH-to-DJH rearrangement (Kirch et al., 1998). This result was recently confirmed in experiments that utilized core RAG2 knock-in mice (Liang et al., 2002). These observations could reflect a distinct chromatin substrate specificity conferred by the RAG2 carboxy terminus, but could also reflect a reduced potency of the core RAG2 that might be most apparent at the VH-to-DJH step.
TCRb The molecular basis for developmentally ordered TCRb rearrangement is unclear. Experimental manipulations that prevent Db-to-Jb rearrangement have demonstrated that such rearrangement is not a prerequisite for Vb-to-Db rearrangement (Sleckman et al., 2000). Developmental order could be directed by the staged activation of Db and Jb accessibility prior to Vb accessibility, but there is no direct data on this point. A necessary precondition is that Vb accessibility must be regulated distinctly from Db and Jb accessibility. This appears to be the case, since Eb was found to regulate chromatin structure across the Db, Jb, and Cb segments only; Vb chromatin is unperturbed in Eb-/- DN thymocytes and appears to be under distinct control (Mathieu et al., 2000). Interestingly, this is true not only for the main cluster of Vb segments, which is separated from Db-Jb-Cb
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by a 250-kb region containing trypsinogen genes, but also for Vb14, which lies just downstream of Eb. Importantly, studies of TCRb (Mathieu et al., 2000; Tripathi et al., 2002), IgH (Chowdhury and Sen, 2001), and TCRa/d (McMurry and Krangel, 2000) locus chromatin structure all suggest these loci to be composed of several discrete regulatory units, and further suggest that the wellcharacterized enhancers may exert their functions over particular regions rather than globally. In fact, remarkably little is known about V gene segment chromatin and whether it might be regulated by an as yet undiscovered set of longrange cis-acting elements. It will be important to better characterize the various regulatory units at Ig and TCR loci, and the mechanisms that help define them (for example, boundary elements, promoter competition), in future studies.
ALLELIC EXCLUSION AT IG AND TCR LOCI IgH Allelic exclusion at the IgH locus is highly stringent. Only about 0.01% of IgM+ splenic B cells appear to be phenotypically allelically included; that is, to co-express on their surfaces the products of both alleles (Barreto and Cumano, 2000). Alt et al. (1984) initially proposed IgH allelic exclusion to be enforced by a feedback mechanism that senses the production of a functional VDJH rearrangement and inhibits the VH-to-DJH step on the second allele. Indeed, the rearrangement of endogenous IgH alleles is inhibited, primarily at the VH-to-DJH step, by transgenes encoding membrane Igm (Weaver et al., 1985; Rusconi and Kohler, 1985; Nussenzweig et al., 1987; Manz et al., 1988). Moreover, elimination of the Igm transmembrane exon by gene targeting causes a loss of allelic exclusion in heterozygous mice (Kitamura and Rajewsky, 1992). Allelic exclusion requires the assembly of membrane Igm with surrogate light chains (Loffert et al., 1996; ten Boekel et al., 1998) and additional signaling components of the pre-BCR (Muljo and Schlissel, 2000). Nevertheless, allelic exclusion appears intact in the few B cells that traverse the developmental block in surrogate light chain mutant mice. An alternative pre-BCR/BCR, composed of prematurely expressed conventional light chains (Papavasiliou et al., 1996; Pelanda et al., 1996) probably accounts for both developmental progression and allelic exclusion in these cells. The inhibition of VH-to-DJH rearrangement that characterizes IgH allelic exclusion must be enforced in pre-B cells that express RAG proteins and actively undergo Vk-to-Jk rearrangement, suggesting a retargeting of the recombinase. Consistent with this, signal ends (SEs) at 5¢DH RSSs are not produced in pre-B cells that actively produce SEs at Jk RSSs (Constantinescu and Schlissel, 1997) and are inhibited by
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Igm transgenes (Schlissel et al., 1993; Schlissel and Morrow, 1994; Stanhope-Baker et al., 1996). Because SE formation occurs in a coupled reaction requiring two substrates in vivo, these observations do not indicate whether recombinase retargeting depends on changes in VH segments, DH segments, or both. However, several observations point to control at the level of VH segments. First, SEs at JH RSSs can be detected at later stages of B cell development and are not inhibited by Igm transgenes (Schlissel et al., 1993; Schlissel and Morrow, 1994; Chang et al., 1999). Thus, DHto-JH rearrangement is permitted and DH and JH accessibility maintained under conditions of allelic exclusion. Second, when challenged with RAG proteins in vitro under conditions permitting uncoupled cleavage, VH and 5¢DH RSSs both serve as substrates in nuclei of RAG-2-/- pro-B cells, whereas only 5¢DH RSSs are substrates in mature B cells (Stanhope-Baker et al., 1996). Do these changes in RAG protein access reflect changes in VH chromatin structure per se? Because VH and DJCm chromatin are subjects of distinct developmental programs (Chowdhury and Sen, 2001), this notion is plausible. Maes et al. (2001) compared the DNase I sensitivity of VH and JH chromatin in AMuLV transformed pro- and pre-B cell lines from RAG-2-/- and RAG-2-/- ¥ Igm transgenic mice, respectively. JH segments were highly accessible in both pro- and pre-B cells, whereas VH segment accessibility was high in pro-B cells, moderate in pre-B cells, and low in mature B cells and non-B cells. However, VH accessibility was still substantial in the pre-B cell lines, and VH chromatin histone acetylation was not significantly different between pro- and pre-B cells. Of note, AMuLv-transformed pre-B cell lines with productive VDJH rearrangements were previously shown to undergo VH-to-DJH rearrangement in apparent violation of allelic exclusion (Schlissel et al., 1991). Moreover, some VH segments that are hypoacetylated in pro-B cells of RAG-2-/mice are hyperacetylated in the AMuLV-transformants of these cells (Chowdhury and Sen, 2001). Thus, these cell lines do not faithfully reflect the properties of their in vivo counterparts. It will be crucial to analyze chromatin structure in natural cell populations in future experiments. Interestingly, VH chromatin replicates in early S phase in pro-B cells and AMuLV transformed pre-B cell lines, but reverts back to a late S phase replication pattern in immature and mature B cell lines and splenic B cells (Zhou et al., 2002). Further, upstream VH segments replicate in late S phase even on alleles in which rearranged VDJH and CH segments replicate early. Because late S phase replication is a characteristic of inactive genes (Simon and Cedar, 1996), the relationship of this change to IgH allelic exclusion warrants further attention. For a feedback mechanism to work effectively, it must be highly unlikely that VH-to-DJH rearrangement is attempted in a similar time frame on both alleles. Allelic asynchrony could be stochastic and could reflect a relatively inefficient
rearrangement process on two equally accessible alleles. Alternatively, allelic asynchrony could be regulated, in the sense that only one allele per cell is initially made accessible. Although both IgH alleles replicate relatively early in S phase and are similarly positioned in pro-B cell nuclei (Skok et al., 2001; Kosak et al., 2002; Zhou et al., 2002), the two alleles have been shown to replicate asynchronously, with the early replicating allele usually (but not always) the first to rearrange VH-to-DJH (Mostoslavsky et al., 2001). Thus, the early replicating allele appears to act as a better substrate for VH-to-DJH rearrangement, perhaps diminishing the likelihood of simultaneous rearrangement on the two alleles. The detection of low-frequency VH-to-DJH rearrangement on the late replicating allele could indicate that it is less frequently chosen as the accessible allele, or that it usually displays reduced, but functionally significant, accessibility. The latter would most easily explain how VH-to-DJH rearrangement could occur on both alleles in a substantial fraction of B cells. It is unclear how the allelic replication pattern is established and how it relates mechanistically to a bias in V(D)J recombination. The replication pattern is fixed prior to rearrangement since the alleles replicate asynchronously even in non-B cells. Early replicating alleles might represent preferred substrates for V(D)J recombination because they compete better for limiting pools of transcription factors or incorporate distinct chromatin components, thus promoting heightened accessibility (Wolffe, 1996). However, early replication could also be a consequence of a more accessible chromatin structure.
TCRb TCRb, like IgH, is subject to stringent allelic exclusion, with the product of a functional rearrangement providing a potent feedback signal that blocks further rearrangement at the V-to-DJ step (Uematsu et al., 1988). Co-expression of two functional TCRb proteins has been detected, but occurs rarely (Padovan et al., 1995; Davodeau et al., 1995). The feedback inhibition of rearrangement depends on the assembly of TCRb with additional components of the pre-TCR, which signals not only allelic exclusion, but also proliferation and developmental progression to the DP stage (Muljo and Schlissel, 2000; Khor and Sleckman, 2002). TCRb allelic exclusion must be enforced in DP thymocytes despite ongoing RAG expression and TCRa gene rearrangement (Wilson et al., 1994), thereby necessitating some form of locus-specific control. The basic mechanisms underlying IgH and TCRb allelic exclusion may be quite similar. Available evidence suggests that TCRb allelic exclusion is associated with changes in Vb but not DbJbCb chromatin. SEs indicative of Db-to-Jb rearrangement are present in both DN thymocytes (pre-allelic exclusion) and DP thymocytes (post-allelic exclusion) (Whitehurst et al., 1999).
8. Allelic Exclusion, Isotypic Exclusion, and the Developmental Regulation of V(D)J Recombination
Consistent with this, the DbJbCb region is characterized by high-level germline transcription, DNA hypomethylation, DNase I sensitivity, and histone hyperacetylation in both DN and DP thymocytes (Senoo and Shinkai, 1998; Chattopadhyay et al., 1998; Tripathi et al., 2002). In contrast, for many Vb segments, germline transcription, DNase I sensitivity, and histone acetylation are all significantly reduced in DP as compared to DN thymocytes. Although these data are consistent with accessibility control, several anomalous observations suggest additional complexity. Vb14, situated 3¢ of the DbJbCb cluster, displays an unexpected increase in germline transcription in DP thymocytes (Senoo and Shinkai, 1998; Chattopadhyay et al., 1998). Moreover, at least one Vb segment in the large upstream cluster still displays substantial DNase I sensitivity and histone acetylation in DP thymocytes (Tripathi et al., 2002). Depending on how well the experimental models reflect the natural cell populations, and how well these measures reflect accessibility to RAG proteins per se, other mechanisms may be required to fully account for allelic exclusion. As for IgH, TCRb alleles replicate asynchronously (Mostoslavsky et al., 2001). This may be associated with an allelic bias to rearrangement, although a direct linkage between replication timing and rearrangement has not been reported in this instance. A significant difference between TCRb and IgH is that TCRb contains two distinct DbJbCb clusters. An allelic exclusion signal would have to prevent Vb rearrangement not only on an allele that had yet to undergo Vb-to-DbJb rearrangement, but also on an allele that had already undergone Vb rearrangement to the Db1Jb1 cluster. If not, an initial out-of-frame Vb-to-Db1Jb1 rearrangement could be followed by in-frame Vb-toDb2Jb2 rearrangement on the same allele, even after inframe rearrangement on the other allele.
Igk The feedback inhibition of endogenous Igk rearrangement by a rearranged Igk transgene can be efficient, contingent on assembly of light chain with membrane Igm (Ritchie et al., 1984). However Igk transgenes encoding autoreactive antibodies may fail to exclude endogenous Igk or Igl rearrangements. Moreover, in-frame VJk rearrangements can be followed by the rearrangement of upstream Vks to downstream Jks or by RS rearrangement on the same allele, particularly if the initially rearranged VJk encodes an autoantibody. Such “editing” is thought to depend on BCR signals that prolong RAG expression in pre-B and immature B cells (Nemazee, 2000). However, BCR signaling is also required for the normal termination of RAG expression that would inevitably exclude further k rearrangement (Shivtiel et al., 2002); the details of these signaling events are not well understood. In the face of compromised feedback control, Igk allelic exclusion is thought to be maintained, at least
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in part, by a bias towards secondary rearrangement on the allele that had rearranged initially, before attempts on the second allele (Mehr et al., 1999). As discussed previously, single cell analysis has shown that in most developing B cells the Jk region is demethylated on a single allele. Moreover, the demethylated allele demonstrates much greater accessibility and represents the preferred substrate for Vk-to-Jk rearrangement (Mostoslavsky et al., 1998; Goldmit et al., 2002). As for IgH and TCRb, Igk alleles replicate asynchronously prior to rearrangement, and Vk-to-Jk rearrangement is biased towards the early replicating allele (Mostoslavsky et al., 2001). Thus, early replication may be associated with allelic remodeling that may provide a monoallelic bias to both initial and secondary rearrangement events. However, several interrelated observations indicate that this cannot be the entire story. First, it is well documented that about 30% of peripheral B cells display Vk-to-Jk rearrangement on both alleles (Coleclough et al., 1981). Second, it was observed that the late replicating allele rearranges first in a fraction of developing B cells (Mostoslavsky et al., 2001). Finally, demethylation was found to be biallelic nearly 30% of the time (Goldmit et al., 2002). Rather than a strict commitment to monoallelic accessibility, the two Igk alleles appear to have distinct probabilities of becoming accessible. Thus, feedback control would still be critical to enforce allelic exclusion. Feedback control of Igk rearrangement could be effected by downregulating RAG expression without any change in the k locus per se. However, Vk segments revealed much reduced sensitivity to DNase I digestion in a plasma cell line as compared to a pre-B cell line (Maes et al., 2001). This difference was not observed for Jk chromatin, suggesting an effect on chromatin that is targeted specifically to Vk segments. This must be confirmed in physiologic cell populations.
TCRa and Other TCR Loci TCRa, like Igk, rearranges in a single step, V to J, and is organized in a fashion permissive for multiple rounds of nested secondary rearrangements. However, beyond this are striking and instructive contrasts. First, the potential for secondary TCRa rearrangement is increased by the large Ja array and the low probability of generating a signal that terminates the process. Initial rearrangements are targeted to the more 5¢ Ja segments by the TEA promoter, and secondary Va-to-Ja rearrangements proceed from 5¢ to 3¢ across the array (Villey et al., 1996; Yannoutsos et al., 2001; Guo et al., 2002). Rearrangement is terminated by RAG downregulation once a TCR is produced that supports positive selection (Borgulya et al., 1992; Wang et al., 1998). Moreover, TCRa rearrangement routinely occurs on both alleles. As judged by the coordinated progression of
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secondary rearrangements along allelic Ja arrays, the two alleles seem equivalent substrates for the recombinase (Davodeau et al., 2001; Huang and Kanagawa, 2001; Mauvieux et al., 2001). The absence of an allelic bias, coupled with the low frequency of positive selection, results in allelic inclusion rather than exclusion. In fact, ample evidence exists for peripheral T cells that bear two distinct TCRa chains, although phenotypic allelic exclusion can still occur by a posttranslational mechanism (Gascoigne and Alam, 1999). Like TCRa, the TCRg and TCRd genes rearrange without evidence of allelic exclusion (Davodeau et al., 1993; Sleckman et al., 1998). Among TCR genes, only TCRb is allelically excluded.
Igl Although there are three tandemly arrayed functional Igl genes in mice, analysis of Igl+ hybridomas indicates that only one of the loci (and presumably only one allele) is typically rearranged in an individual B cell (Nadel et al., 1990). Asynchronous replication may bias initial rearrangement to a single allele, as at other loci (Mostoslavsky et al., 2001). Igl can provide feedback control, as Igl transgenes were found to suppress endogenous l and k rearrangements (Hagman et al., 1989; Neuberger et al., 1989). However, it has been speculated that Igl rearrangement may be limited to a single attempt due to inefficiency and time constraints, obviating an absolute requirement for feedback (Nadel et al., 1990). Accessibility changes at the l locus have not been analyzed.
IG LIGHT CHAIN ISOTYPIC EXCLUSION As noted previously, B cells are isotypically excluded in that they usually express k or l light chains, but not both. Early studies showed that Igk+ B cells only rarely display Igl rearrangements, whereas Igl+ B cells display either nonfunctional k rearrangements or k alleles deleted by RS rearrangement (Alt et al., 1980; Korsmeyer et al., 1981). These observations suggest that isotypic exclusion reflects a defined developmental sequence of light chain rearrangement, with k preceding l, or with a much higher probability of k rearrangement. Subsequently, in vivo pulse labeling with BrdU indicated that the developmental onset of k rearrangement precedes that of l rearrangement by about a day (Arakawa et al., 1996). Moreover, Igk RSSs were found to support V(D)J recombination at much higher frequencies than those of Igl (Ramsden and Wu, 1991). As a consequence of these factors, there is thought to be a high probability that pre-B cells will not proceed to l rearrangement until they have undergone multiple rounds of k rearrangement and have exhausted their opportunities for further
rearrangement at the k locus (Arakawa et al., 1996; Mehr et al., 1999). It is clear, however, that prior k rearrangement is not required for l rearrangement, since Igl+ B cells are generated efficiently in mice in which k rearrangement has been inactivated by gene targeting (Zou et al., 1993; Chen et al., 1993; Inlay et al., 2002). Moreover, rare l producers have Igk genes in germline configuration (Pauza et al., 1993). The detection of rare B cells expressing both k and l indicates that isotypic exclusion is not absolute (Pauza et al., 1993; Giachino et al., 1995).
FUTURE DIRECTIONS The concept of accessibility control has been a powerful one that has driven research in this area for many years. Much has been learned about the regulatory programs at Ig and TCR loci through studies of cis-acting elements and the various correlates of accessibility. However, it remains an important challenge to move from descriptive correlates to a mechanistic understanding of RAG protein access. Moreover, it is important that this problem be addressed not only at the level of local chromatin chemistry, but also within the context of a highly compartmentalized but as yet poorly understood nuclear organization. Although the accessibility problem is often visualized in terms of diffusible RAG proteins, it may be more relevant to consider how pairs of chromosomal RSSs are brought to the recombinase. Major questions remain unanswered regarding the regulation of V gene segment chromatin, including the mechanisms that establish an allelic bias and enforce feedback. Finally, it will be a challenge to understand how accessibility control integrates with other levels of regulation to maintain precise developmental programs at Ig and TCR loci.
Acknowledgments Work in the authors’ laboratories was supported by NIH grants GM41052 and AI49934 (to M.S.K.) and HL48702 and AI40227 (to M.S.S.). We thank Barry Sleckman, Annette Jackson, and Amber Meade for their helpful comments.
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9 The Development of Human B Lymphocytes PETER D. BURROWS,1 TUCKER LEBIEN,1 ZHIXIN ZHANG,1 RANDALL S. DAVIS,2 AND MAX D. COOPER1 1 Division of Developmental and Clinical Immunology, Departments of Medicine, Pediatrics, Microbiology and Pathology, University of Alabama at Birmingham, Birmingham, Alabama and the Howard Hughes Medical Institute, Birmingham, Alabama; and University of Minnesota Cancer Center, Minneapolis, Minnesota, USA 2 Divisions of Developmental and Clinical Immunology and, Hematology/Oncology, Department of Medicine, University of Alabama at Birmingham, Birmingham, AL 35294
B lineage cells in humans are progeny of the lymphoid progenitors that derive from multipotential hematopoietic stem cells (Galy et al., 1995; Rossi et al., 2002). Their orchestrated development begins in lympho-hematopoietic sites in the fetal liver and then continues in bone marrow throughout life (Gathings et al., 1977). With a few notable differences, B lineage differentiation in humans follows the same basic rules elaborated in mice and other vertebrate species. In this chapter, we describe the genotypic and phenotypic features that mark the progression of the cells along this developmental pathway, noting significant species differences in the process and indicating where human mutations have provided important clues to gene function. We outline the progression of the V(D)J gene rearrangements required for the expression of immunoglobulin (Ig), the antigen receptor and effector molecule of the B lineage, and describe the contribution of secondary V(D)J rearrangements to the human B cell repertoire. We conclude with an overview of two important types of abnormal human B cell development, the primary immunodeficiency diseases and acute lymphoblastic leukemias of B lineage.
three types of gene segments, VH (variable), DH (diversity), and JH (joining), whereas the formation of the k and l VL exons require only a single joining reaction VL Æ JL. The pro-B cells are Ig negative, but can be identified as B lineage cells by the expression of other markers and initiation of the IgH gene rearrangement process (Figure 9.1). The pre-B cells express intracellular m heavy chains, and a limited portion of these associate with surrogate light chain proteins to form a pre-B cell receptor (pre-BCR). The delivery of the pre-BCR to the cell surface and its signaling function require participation of Iga and Igb, two transmembrane proteins that are expressed within early B lineage cells even prior to Ig gene rearrangements. B cells express transmembrane Ig molecules in the B cell receptor (BCR) for antigen, whereas plasma cells preferentially synthesize the secretory form of antibodies. The pro-B cells are derived from a common lymphoid progenitor (CLP) that has the potential to differentiate into B, T, natural killer (NK), and dendritic cell (DC) lineages. Except for a macrophage default pathway, the CLP apparently have lost the capacity to differentiate along the myeloid, erythroid, or megakaryocytic pathways. CLP have been best characterized as murine bone marrow cells, which are negative for markers of mature blood cell lineages (Lin-) and have the following phenotype: interleukin 7 receptor a chain (IL-7Ra)+, Thy-1-, Sca-1lo, and c-Kitlo (Kondo et al., 1997). An earlier lymphoid progenitor has been identified recently on the basis of the activation of the recombination activating gene (RAG) locus (Igarashi et al., 2002). In humans, the CD34 marker can be used in conjunction with other markers to define multipotential hematopoietic stem cells (HSC) and their lineage-restricted progeny. CD34 is a sialo-mucin that is expressed on HSC,
STAGES OF HUMAN B CELL DIFFERENTIATION The most unambiguous marker of differentiation along the B lineage pathway is the expression of Ig heavy (H) and light (L) chains in a classification scheme that can be refined through definition of the Ig genotype (Figure 9.1). Functional Ig genes are generated somatically by a recombinatorial process to be described later. The exon encoding the Ig heavy chain variable region is generated by the joining of
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Antigen Independent Lymphoid Pro-B Progenitor Cell
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CD34 CD10 CD19 CD21 CD24
FIGURE 9.1 Human B cell development. The antigen-independent stages of B cell development occur in the primary lymphoid organs, the fetal liver, and adult bone marrow. In the secondary lymphoid organs, the mature B cells may encounter cognate antigens and, usually with the help of T lymphocytes, undergo proliferation and differentiation to antibodysecreting plasma cells and memory B cells. The diagram depicts the stages of B cell development and several markers that help define these stages: CD19 ( ), Ig heavy chains ( ), Iga/Igb (||) the surrogate light chain peptides l5/14.1 and VpreB ( ), and conventional Ig light chains ( ). The configuration of the IgH and IgL chain genes during development is illustrated and is the predominant sequence of IgH prior to IgL rearrangement, although the order may sometimes be reversed. The phenotypic markers shown are a selection of those that have proven useful in identifying B cell developmental stages. The phenotype of the lymphoid progenitor that directly precedes the pro-B cell is unknown, but may be a common lymphoid progenitor that expresses a somewhat different cell surface phenotype depending on the tissue source (i.e., fetal or adult bone marrow, or cord blood) being analyzed. There are also some differences between the cell surface phenotype of early B lineage cells in adult bone marrow, depicted here, and fetal bone marrow (LeBien, 2000). See color insert.
mast cells, bone marrow stromal cells, and most endothelial cells. A population of CD34+ Lin- CD45RA+ adult bone marrow cells expressing CD10 appear to be CLP, since they lack erythroid, myeloid, and megakaryocytic potential but can give rise to T, B, NK, and lymphoid DC (Galy et al., 1995). These are CD38+/HLA-DR+ cells that do not express significant levels of Thy-1 or c-kit. A somewhat different CLP phenotype with similar developmental potential can be identified in cord blood, a hematopoietic stem cell source widely used in clinical transplantation. A cord blood CD7+ population that is CD34+, CD38-, HLA-DR+, CD45RA+, thy-1neg/lo, c-kitneg/lo, and IL-7Ra- was shown to generate B cells, NK, and dendritic cells, but lacks myeloid or erythroid potential (Hao et al., 2001). CD10 expression in cord blood, in contrast to bone marrow, marks a population of CD38cells with myelo-erythroid potential. The phenotype and developmental plasticity of CLP may therefore differ between individual lympho-hematopoietic sites. CD34+ pro-B cells express CD19, the earliest definitive B cell surface marker and one whose expression continues
until the plasma cell stage of development (Poe et al., 2001). However, the initial stage of IgH rearrangement, the DH to JH step, begins in CD34+ cells that are CD10+/CD19(Bertrand, III et al., 1997; Davi et al., 1997). CD10, originally identified on pre-B leukemic cells as common acute lypmphoblastic leukemia antigen (CALLA), is a transmembrane ectopeptidase that cleaves small peptides like substance P on the amino terminus of hydrophobic residues (LeBien and McCormack, 1989). In addition to its expression on many types of bone marrow cells, CD10 is found on nonhematopoietic cells, including intestinal and renal epithelia. The dearth of cell surface markers that unambiguously define each B cell differentiation stage reflects the fact that development proceeds as a continuum rather than in quantum leaps. Although CD34 and CD19 co-expression is a useful marker for bone marrow pro-B cells, productive VDJH rearrangements are detectable in 5 to 10% of cells of this phenotype (Dittel and LeBien, 1995). These preBCR+/CD34+ cells are present in highest frequency during fetal life (Wang et al., 2002b). Most pre-B cells are CD34/CD19+ and, by definition, all of them have intracellular m heavy chains. Nevertheless, they may either express preBCR in very low levels or not at all (discussed in more detail below). Clonal expansion, which occurs at several stages of B cell development, plays an important role in generating a diverse antibody repertoire. After the initial DJH rearrangement in a pro-B cell, proliferation generates multiple progeny with the potential to rearrange different VH gene segments to the original DJH. The pre-BCR+ cells then undergo proliferative expansion prior to IgL gene rearrangement, which may utilize different VJk or VJl in the generation of B cell progeny (Wang et al., 2002c). An on-and-off regulation of RAG1 and RAG2 expression controls the intermittent V(D)J rearrangement process after the successful VDJH rearrangement. The expression of both genes is downregulated during the proliferative phase of pre-B cell differentiation (Ghia et al., 1998). Pre-BCR expression is subsequently extinguished by the downregulation of the SLC receptor components, VpreB and l5. This leads to an exit from the cell cycle, reactivation of RAG1 and RAG2 expression, and IgL chain gene rearrangement in the quiescent small pre-B cells (Ghia et al., 1996; Grawunder et al., 1996; Wang et al., 2002c). The successful rearrangement of an IgL chain gene allows the expression of the BCR on the immature B cell. Each BCR is composed of an IgM monomer associated noncovalently with an Iga/Igb signaling module (Schamel and Reth, 2000). The assembly of pre-BCR and BCR and their association with key signaling elements constitute important quality control checkpoints during B cell development.
9. The Development of Human B Lymphocytes
SITES OF HUMAN B CELL DEVELOPMENT The relatively widespread distribution of pro-B and preB cells in early fetal tissues suggests a multifocal origin of human B lineage cells during embryonic development (Solvason and Kearney, 1992; Nunez et al., 1996). The liver is the principal site of embryonic B cell generation, and preB cells can be found there by 8 weeks gestation. Immature sIgM+ B cells appear by week 9, and mature sIgM+/sIgD+ cells appear by week 12 (Cooper, 1987). From midgestation onward, the bone marrow is the primary site of B cell generation. A relatively constant ratio of B cell precursors to B cells of immature phenotype (IgM+IgDCD24highCD10+CD20low) is maintained from mid-gestation through the eighth decade of life (Nunez et al., 1996; Rossi et al., 2002). Recombinase gene transcripts in the bone marrow pro-B cells of aged donors further attest the sustained production of B cells, albeit at lower levels with increasing age. A subpopulation of B cells with mature phenotype (CD24lowCD10-CD20highIgD+) begins to accumulate in the bone marrow during childhood and becomes the predominant B cell subpopulation in adult bone marrow. This mature population of bone marrow B cells represents a subpopulation of memory B cells that have undergone selection in the periphery, as indicated by CD27 expression and somatically mutated VH genes (Paramithiotis and Cooper, 1997; Rossi et al., 2002). The bone marrow is also a site in which long-lived plasma cells reside (Manz and Radbruch, 2002). The specificity of the BCR is monitored during clonal Bcell generation in hematopoietic tissues. Immature B cells expressing receptors with high affinity for self-antigens are either salvaged by receptor editing to change the BCR specificity or else eliminated. The IgM B cells exiting the bone marrow begin to express a second isotype, IgD, on their cell surface (Preud’homme et al., 2000). The variable regions of the m and d heavy chains are identical, ensuring that both IgM and IgD BCR have the same specificity. This is accomplished by the alternative splicing of a primary RNA transcript with the structure 5¢–VDJH–Cm–Cd–3¢. Although the molecular mechanism resulting in IgM/IgD co-expression on the mature B cell has been known for decades, the biological value of the IgD BCR remains unknown.
HUMAN IMMUNOGLOBULIN GENES Immunoglobulins are encoded in three unlinked loci. The H chain gene locus is located on chromosome 14q32, and the k and l L chain gene loci are on chromosomes 2p12 and 22q11, respectively. As in the adaptive immune systems found in all other jawed vertebrates, humans do not inherit
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intact Ig genes through the germ line, but instead have segmental genes that require somatic recombination to become functional during B cell development. The IgH and IgL loci were among the earliest regions in the human genome to be targeted for a comprehensive sequencing effort because this information was essential for determining the contribution of the germline variable region genes to the antibody repertoire, a fundamental issue in immunology. There are 123 VH gene segments in the originally sequenced Ig locus, but 79 are pseudogenes, leaving at most 44 functional genes (Matsuda et al., 1998). Genetic polymorphisms can result in Ig haplotypes with expansion or contraction of these VH gene numbers (Cook and Tomlinson, 1995). The 27 DH and 6 JH gene segments are located centromeric of the VH locus and are followed by the Cregion gene segments: telomere VH–DH–JH–Cm–Cd–Cg3– Cg1–yCe–Ca1–yCg–Cg2–Cg4–Ce–Ca2— centromere. The constant region exons can encode two forms of each heavy chain isotype, an integral membrane protein that is the anchoring element of the pre-BCR (m H chain) and BCR (m, d, g, e, or a H chain), and a soluble protein secreted as antibody by plasma cells. The choice of a transmembrane or secretory C-terminal exon is regulated at the transcriptional level by termination and RNA processing or polyadenylation events (Staudt and Lenardo, 1991). The expression of IgD as a BCR component is similarly regulated, whereas expression of the isotypes further downstream requires an additional DNA rearrangement event called class switch recombination (CSR). In humans, much more so than in mice, IgD is found in serum (~30 mg/ml), and, in the cells secreting IgD atypical CSR occurs that may involve homologous recombination between two direct repeats upstream of Cm and Cd to delete the Cm gene (White et al., 1990). CSR is dependent on the activation induced deaminase (AID) gene (Muramatsu et al., 2000; Revy et al., 2000), which appears to initiate the DNA cleavage required for CSR by deaminating the DNA at cytosine residues (Di Noia and Neuberger, 2002; Petersen-Mahrt et al., 2002). In the k locus, the duplication of a primordial VL gene cluster has resulted in two copies of the Vk locus located upstream of five Jk gene segments and a single Ck exon that encodes the entire constant region of the kL chain. The potential Vk repertoire consists of 32 functional gene segments among a total of 76 Vk genes (Thiebe et al., 1999; Kawasaki et al., 2001). The Vl locus spans nearly 1-MB of DNA and contains 36 potentially functional Vl genes and 56 Vl pseudogenes (Kawasaki et al., 2000; Williams et al., 1996). The number of Cl genes varies among individuals ranging from 7 to 10, and each of which is preceded by a single J gene segment. The nonrearranging genes that encode the pre-BCR components VpreB and l5 (termed 14.1 in humans based on the
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size of the EcoRI restriction fragment that contains the gene) are located within and downstream of the l locus on chromosome 22. The single VpreB gene is located within the Vl cluster, approximately 620-kb centromeric of the Jl–Cl pairs, whereas the 14.1 gene is found ~650 kb telomeric of this region (Bauer et al., 1988; Kawasaki et al., 1997; Bauer, Jr. et al., 1993; Tapper et al., 2001). The considerable polymorphism found in the l5/14.1 gene may result from gene conversion events involving the three closely related pseudogenes, 16.1, 16.2, and Gl1 (Conley et al., 1999). Two lymphocyte-restricted recombination activating genes (RAG1 and RAG2) are essential for the process of Ig V gene assembly, as are several ubiquitously expressed DNA repair genes, including the DNA-dependent protein kinase/Artemis complex that is important for nonhomologous end joining and V(D)J recombination (reviewed in Bassing et al., 2002). The rearrangement of Ig genes is an ordered sequential process, usually commencing at the H chain locus, DH Æ JH followed by VH Æ DJH joining. Rearrangement activity then shifts to the L chain loci, first k then l, leading to the eventual production of cell surface IgM by the newly formed B cell. The Ig gene rearrangement order IgH Æ Igk Æ Igl is not inviolate, and Igk chain gene rearrangement can precede IgH rearrangement in both humans and mice (Kubagawa et al., 1989; Chen et al., 1993; Ehlich et al., 1993; Novobrantseva et al., 1999). Thus the production of a functional m chain is not an absolute prerequisite for L chain gene rearrangement, nor is the failed rearrangement of both k alleles a prerequisite for l light chain gene rearrangement. The joining of Ig (and T cell receptor) gene segments is an imprecise process and most rearrangements are nonfunctional. These nonproductive rearrangements most often result from a shift in the translational reading frame due to the random addition and deletion of nucleotides at the site of joining. A translation stop codon is typically encountered shortly downstream of most reading frame shifts, resulting in a truncated, nonfunctional protein. Although a perilous strategy, junctional imprecision is an important source of antibody diversity in the hypervariable third complementarity determining regions (CDR3) of Ig heavy and light chains, which are encoded at the sites of VDJH and VJL recombination. The insertion and deletion of nucleotides prior to the ligation of the rearranging gene segments results from the activity of the enzyme terminal deoxynucleotidyl transferase (TdT) and other unidentified exonucleases. Short and long isoforms of TdT are generated by alternate splicing and, in mice, these have been found to add and delete nucleotides, respectively, at the site of gene segment joining (Thai et al., 2002). The relative activity of each isoform at the time of rearrangement governs the length of the CDR3 region. TdT activity is primarily restricted to B cell developmental stages during which IgH rearrangements occur, consequently the nontemplated (N) nucleotides added by TdT are frequently
present in heavy chain V genes and are less common among light chains. However, this skewing is less prominent in human than in mouse V genes. Since TdT expression is initiated after embryonic B lymphopoiesis begins, N sequences are limited or absent in the first B cells to be generated during ontogeny. The B lineage cells that fail the rearrangement process undergo apoptosis and are rapidly engulfed by resident macrophages in sites of B-cell generation (Osmond et al., 1994). The immune system is tolerant of this considerable wastage and, by producing large numbers of B cells daily, can maintain an adequately protective repertoire of B-cell specificities. Moreover, mechanisms exist to repair nonfunctional variable region genes so that the number of failed B cells is probably smaller than would be anticipated.
THE ROLE OF SURROGATE LIGHT CHAINS IN HUMAN B CELL DEVELOPMENT The m chains synthesized by pre-B cells are destined for intracellular degradation unless released from their noncovalent association with BiP and other endoplasmic reticulum chaperones that monitor the assembly and folding of multisubunit proteins. Conventional k or l light chains carry out this rescue mission in B cells, and the surrogate light chain (SLC) plays a similar role earlier in B cell differentiation. The SLC is composed of two noncovalently associated polypeptides encoded by the nonrearranging l5 (14.1) and VpreB genes (Karasuyama et al., 1996). In pre-B cells, the SLC is disulfide bonded via the l5 element to the CH1 domain of the m heavy chain. The SLC-m chain association is inefficient compared to that of k/l-m chain association, thus liberating only a fraction of the pre-B cell m chains to be expressed with the Iga/Igb heterodimer as the cell surface pre-BCR (Lassoued et al., 1993). The pre-BCR preferentially resides in lipid raft microdomains, where it constitutively associates with protein tyrosine kinases syk and lyn, the B cell linker protein BLNK, and PI-3 kinase signaling elements (Guo et al., 2000). The pre-BCR signaling event is essential for normal development as illustrated by the severe B cell immunodeficiency that results from mutation in genes encoding any of the receptor components (see below). The receptor is expressed at very low levels on normal pre-B cells, compared with cell lines at an equivalent differentiation stage. This feature has made it difficult to analyze the developmental regulation of pre-BCR expression on primary cells (Wang et al., 2002b). The low level of pre-BCR expression may have several explanations, including inefficient assembly and receptor downregulation as the consequence of binding to its ligand(s). Following the initial discovery of the pre-BCR, an immediately appealing idea was that the cell surface–expressed
9. The Development of Human B Lymphocytes
pre-BCR would interact with a stromal cell ligand. This would signal the cell of a successful H chain gene rearrangement and result in termination of further rearrangement at the H chain locus. Many futile attempts to identify the putative pre-BCR ligand then led to the view that there is no ligand. In this scenario, cell surface pre-BCR expression per se would be sufficient to signal in a ligand independent fashion. Recently, however, two candidate pre-BCR ligands of stromal cell origin have been identified. A soluble Fablike pre-BCR was used to identify a 135-kDa protein on murine bone marrow stromal cell lines that support B lymphopoiesis in vitro (Bradl and Jack, 2001). The function of this molecule is unknown. The second candidate is galectin1 (Gauthier et al., 2002). Galectins are a family of secreted, calcium-independent, S-type lectins. In this study, galectin was proposed to act as a supramolecular organizer that clusters the pre-BCR with counterreceptors on stromal cells, culminating in transduction of a signaling event in pre-B cells. These two reports are likely to foster a new round of investigation focusing on the role of the pre-BCR in promoting the survival signals to pre-B cells. Pre-BCR expression is limited to a subpopulation of preB cells that are relatively large and cycling, and have reduced expression of both RAG-1 and RAG-2 genes. A direct role for pre-BCR expression in clonal expansion at this stage of development is indicated by studies of a transgenic mouse model in which m chain expression could be induced at will (Hess et al., 2001). The downregulation of the pre-BCR in these cells is linked with exit from the cell cycle, increased RAG expression, and light chain gene rearrangement (Figure 9.1).
REPERTOIRE DIVERSIFICATION VIA RECEPTOR EDITING AND VH REPLACEMENT B cell development can fail for several reasons. First, the random joining of the coding segments during V(D)J rearrangement theoretically will generate two thirds of VH Æ DJH and VL Æ JL joints as out-of-reading frame nonfunctional products. B lineage cells with nonproductive rearrangements are unable to develop further. Even after generating a functional VDJH open reading frame, the expressed m heavy chains may fail to pair with surrogate light chain or with conventional light chains to form the functional pre-BCR or BCR needed for further differentiation. Moreover, B cells that possess self-reactive antigen receptors must alter their antigen specificities or be eliminated before their release into the periphery. In all of these situations, early B lineage cells may retain the capability to alter the initially generated Ig V gene exons, a process known as receptor editing (reviewed in Nussenzweig, 1998; Nemazee and Weigert, 2000).
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The organization of the VL and JL gene segments within the Igk and l loci allows secondary rearrangement by simply joining an upstream VL and a downstream JL gene segment using the same recombination elements used for the primary rearrangement (Figure 9.2). Thus, if upstream VL and downstream JL gene segments are available, the process can continue as long as the recombination machinery is operative and the locus remains accessible (reviewed in Radic and Zouali, 1996). With each round of rearrangement, a new VL-JL coding joint is formed and the previous VL-JL coding joint is deleted, leaving no trace of the initial rearrangement (Nussenzweig, 1998; Nemazee and Weigert, 2000). Consequently, the contribution of light chain gene editing to the normal repertoire can only be inferred by biased usage of the more 3¢ Jk gene segments or elevated Igl usage, since cells with two nonfunctional k rearrangements still have the option to rearrange their l light chain genes de novo (King and Monroe, 2000; King and Monroe, 2001; Nemazee and Weigert, 2000). However, in transgenic mice carrying a knockin human Ck marker, light chain editing was estimated to occur in nearly 25% of the B cell population (Casellas et al., 2001). In contrast to the relative ease of secondary rearrangement in the light chain loci, the secondary rearrangement of an upstream VH to a preformed VDJH rearrangement entails a more complex recombinatorial process. The intervening DH segments, which are flanked by the necessary recombination signal sequences (RSS), are deleted during the initial V Æ DJH rearrangement event (Figure 9.2) (reviewed in Nussenzweig, 1998; Nemazee and Weigert, 2000). Nonetheless, in mouse pre-B cell lines with nonfunctional IgH rearrangements, functional IgH genes appeared to arise through a secondary rearrangement involving a cryptic RSS (cRSS) sequence located within the third framework region of the VH germline gene segments (Kleinfield et al., 1986; Reth et al., 1986; Covey et al., 1990; Usuda et al., 1992). The biological importance of this type of VH gene replacement was suggested by gene knockin experiments. Self-reactive IgH transgenes were artificially inserted into the germline JH locus, leaving the upstream DH and VH gene segments intact. The self-reactive VDJH genes in these mice could be altered by secondary rearrangements, including VH replacement (Chen et al., 1995; Chen et al., 1997). In humans, 40 out of 44 functional VH germline genes contain cRSS motifs within the third framework regions (reviewed in Radic and Zouali, 1996). However, the potential function of these cRSS sites in RAG-mediated secondary recombination, and the possible contribution of VH replacement to the primary human B cell repertoire, have only recently been elucidated through studies of a suitable in vitro model of human B cell development, the EU12 cell line derived from a child with acute lymphoblastic leukemia (Wang et al., 2003). EU12 is remarkable among acute lymphoblastic leukemia–derived cell lines in containing cells representa-
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Igk Rearrangements VL
JL
//
// VL ¨ JL rearrangement
//
// Secondary VL¨JL rearrangement //
//
IgH Rearrangements VH
DH
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// DH ¨ JH rearrangement
//
// VH ¨ DHJH rearrangement
//
// CDR3
1st
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2nd VH replacement
//
VH replacement //
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// CDR3
FIGURE 9.2 Receptor editing and VH gene replacement. Following the initial rearrangement (red oval), subsequent rearrangements (blue oval) of the Igk chain locus can occur by RSS-mediated recombination of an upstream Vk to a downstream Jk gene segment. A similar process can occur in the Igl locus, but the organization of the gene segments is different. In the heavy chain locus, the rearrangement sequence is usually DH Æ JH, followed by VH Æ DJH (red ovals). A secondary VH Æ VDJH rearrangement utilizes the RSS of the incoming VH gene together with a cryptic RSS found in the 3¢ end of most germline VH genes to accomplish recombination. See color insert.
tive of multiple stages of B cell differentiation. Cellular subcloning studies indicate that the EU12 cells are capable of ongoing in vitro B cell development, from pro-B to pre-B to sIgM+sIgD+ B cells (Wang et al., 2003). During the proliferation and differentiation process, the EU12 pro-B cells generate progeny B cells with multiple VH and VL gene segment rearrangements. Through analysis of the IgH repertoire, VH gene replacement was shown to occur in a serial fashion (Zhang et al., 2003). Beginning with a nonfunctional VDJH joint, the continuous serial VH replacement generates a diversified VH repertoire. The cryptic RSS site embedded within the third framework region mediates the VH gene replacement reaction. In vitro protein binding and DNA cleavage assays indicate that the cryptic RSS sites found in almost all VH germline genes can be used in RAG-mediated recombination. One important feature of the serial VH gene replacement reaction distinguishes it from light chain receptor editing. Whereas the latter leaves no trace, with each round of VH replacement the resulting IgH gene renews the entire VH coding region, but also retains a short stretch of 3¢ nucleotides in the VH-DH junction from the replaced VH gene. This residual sequence serves as a diagnostic marker
that can be used to search for potential VH replacement products in primary B cells. Through an analysis of IgH gene sequences derived from normal individuals of different ages, potential VH replacement products could be identified in 5 to 12% of analyzed sequences, depending on the stringency used in the sequence comparisons (Zhang et al., 2003). If 1 in 20 B cells undergoes a VH replacement event, this would represent a significant contribution to the B cell repertoire. The true frequency of VH replacement may be higher, since the footprints of this reaction can be obscured by subsequent genetic changes in the CDR3 region, for example somatic hypermutation. VH replacement could occur at any stage in B cell development when the recombination machinery is still active and the locus remains accessible (Monroe et al., 1999; Yu et al., 1999). VH replacement may occur during the pro-B cell stage to rescue cells carrying nonfunctional IgH rearrangements, or during the pre-B or B cell stages when the IgH gene encodes m heavy chains failing to pair well with surrogate or conventional light chains, or which possess self-reactivity. The biological consequences of VH gene replacement and its potential contribution to autoimmune diseases remain to be elucidated.
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9. The Development of Human B Lymphocytes
REGULATION OF ANTIBODY PRODUCTION BY B CELL RECEPTORS The B cell response to antigen is regulated by cognate interactions between T cells and B cells that determine the extent of the B cell proliferative response prior to differentiation into plasma cells and memory B cells. These interactions also influence the qualitative features of the antibody response, including the CSR and the expression of particular switched isotypes. Cytokines produced by T cells, together with the T cell surface molecules CD28, CTLA4, and CD40 ligand, are important regulators of the humoral immune response. The response of a naive B cell often involves a low-affinity interaction between an antigen and the germline encoded IgM/IgD BCR and is antigen dose–dependent. An interaction between the BCR and the CD19/CD21/CD81 complex on the B cell provides an enhancing mechanism to allow B cell responses to low antigen doses (Carter and Fearon, 1992; Fearon and Carroll, 2000). Complement deposition on the antigen promotes the simultaneous binding of antigen to the BCR and the complement cleavage product C3d to its receptor CD21. The co-ligation of the two receptor complexes results in a CD19 mediated enhancement of BCR signaling, lowering the threshold antigen concentration required for B cell stimulation by 100 fold, and functionally links the innate and adaptive immune systems. In a primary immune response in which there is no pre-existing antibody, the necessary activation of complement most likely occurs via the alternate or mannose-binding lectin pathways in response to conserved molecular patterns on pathogens (Gadjeva et al., 2001; Janeway and Medzhitov, 2002). Intrinsic B cell regulatory networks also help initiate and terminate B cell responses. Antibodies secreted by their plasma cell progeny play a role in regulating B cell responses to antigen. Passively administered IgM antibodies enhance subsequent antibody responses, whereas IgG antibodies are immunosuppressive (e.g., Harte et al., 1983). The mechanism of IgM-mediated enhancement involves the classical activation pathway of complement and the complement receptors on B cells and follicular dendritic cells. The inhibitory effect of IgG antibodies arises via the formation of antigen–antibody complexes that bind both the BCR and the FcgRIIB expressed on B cells. (Ravetch and Bolland, 2001; Heyman, 2000). Following the co-ligation of these receptors, an immunoreceptor tyrosine based inhibitory motif (ITIM) in the FcgRIIB cytoplasmic domain is phosphorylated and recruits the SH2 domain-containing inositol 5-phosphatase (SHIP) and SH2 domain containing protein tyrosine phosphatase SHP-2. These two phophatases then dephosphorylate essential substrates and interrupt BCR-mediated signal transduction and B cell activation. The current view of the regulatory roles of Fcg receptors on B cells has been expanded recently with the identifica-
tion of a large family of FcR related genes (Figure 9.3). Many of their protein products are preferentially expressed on B cells or, in one case, within them. The first five Fc receptor homologs (FcRH 1–5) were identified through a search of the human genome using an FcR consensus sequence (Davis et al., 2002a; Davis et al., 2001) and through sequencing the breakpoint of a t(1;14)(q21;q32) translocation in a myeloma cell line (Hatzivassiliou et al., 2001; Miller et al., 2002). The human FcRH are considered homologs of the FcgR based on predicted amino acid sequence homology and mapping of these genes to the chromosome 1q21–22 region that also contains the FcgRI, RII, and RIII and the high affiinty FceRI genes. The FcRHs are likely to be important immunoregulatory molecules for B cells since they contain potential ITIM, ITAM (immunoreceptor tyrosine based activation motif), or both in their cytoplasmic domains. FcRH2, FcRH3, FcRH4, and FcRH5 appear to have the potential to bind IgG. How the products of the FcgRIIB gene and the multiple FcRH genes interact in regulating B cell homeostasis is currently under study. Another FcR relative identified by the bioinformatics approach is termed FcRL [Fc receptor-like (Mechetina et al., 2002)], FREB [Fc receptor homologue expressed in B cells (Facchetti et al., 2002), and FcRX (Davis et al., 2002b)]. FcRX has no transmembrane region or N-linked glycosyla-
BCR
FcRH1 FcRH2
FcgRIIB
FcRH3 FcRH4 FcRH5
FcRX/L FREB
B Cell FIGURE 9.3 Fc receptor and Fc receptor related genes expressed by human B cells. The BCR, composed of membrane Ig and the Iga/Igb heterodimer with cytoplasmic ITAM (green boxes) is shown at the top of the B cell. The ITIM-containing (red box) FcgRIIB inhibits B cell activation when antigen–antibody complexes crosslink it to the BCR. Also illustrated are members of a recently discovered family of FcR-related genes expressed on B cells (FcRH). These are Ig-like domain proteins that have ITIM, ITAM, or both in their cytoplasmic tails, suggesting a role in regulating B cell responses. The Ig domains (ovals) are color coded to indicate their homology to each other and to the Ig domains in the other FcR. FcRX is an intracellular FcR-related protein expressed in germinal center B cells. See color insert.
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tion sites and appears to be an intracellular protein in B lineage cells, principally in germinal center cells. The function of an intracellular receptor with the potential to bind Ig is unknown. Perhaps during the somatic hypermutation reaction in germinal centers, when B cells become surface Ig negative (presumably to prevent the expression of multiple, possibly conflicting specificities), FcRX may serve as a molecular chaperone that retains Ig intracellularly. Failure to do so could result in inappropriate apoptosis of B cells bearing useful specificities, or might allow the survival and escape of autoreactive B cell clones.
IMMUNODEFICIENCY DISEASES The resemblance between B lineage differentiation in humans and mice is clearly indicated in the patterns of immunodeficiency that result from mutations in some of the essential B lineage genes (Figure 9.4). Function-loss mutations in the recombination activating genes, RAG1, RAG2, and Artemis lead to combined B and T cell deficiencies in humans and mice (Mombaerts et al., 1992; Shinkai et al., 1992; Schwarz et al., 1996; Corneo et al., 2001). Likewise, deficiencies in the pre-BCR components and their intracellular signaling partners interrupt B lineage differentiation at the pro-B cell stage (Kitamura et al., 1992; Minegishi et al., 1998; Minegishi et al., 1999b; Pappu et al., 1999). Mutations preventing the expression of either m heavy chains or Iga result in a complete block at the pro-B cell stage in both species (Kitamura et al., 1991; Yel et al., 1996; Torres et al.,
Antigen Independent Lymphoid Progenitor
Pro-B
Pre-B Cells
VpreB/ l 5 m HC, Ig a /b, BTK, BLNK D RAG 1/2 D PU.1 E2A D IKAROS D EBF D
1996; Minegishi et al., 1999a; Milili et al., 2002; Wang et al., 2002a; Pelanda et al., 2002). AID mutations completely prevent Ig class switching and somatic hypermutation in humans and mice (Muramatsu et al., 2000; Revy et al., 2000a). For other gene defects, the completeness of the differentiation block may differ significantly between humans and mice. For example, l5 deficiency consistently interferes with pre-B cell differentiation, but the block is incomplete in mice; within a few months after birth, the level of splenic B cells reaches approximately 50% of normal levels (Kitamura et al., 1992). In contrast, a decisive block in pro-B cell to pre-B cell differentiation was found in a boy with l5 deficiency, who had not generated any B cells by 8 years of age (Minegishi et al., 1998). BTK deficiency also leads to a much more severe blockage in human B lineage differentiation than in mice (Conley and Cooper, 1998; Desiderio, 1997; Conley et al., 2000; Satterthwaite and Witte, 2000). Boys with X-linked agammaglobulinemia due to function-loss mutations of the BTK gene have very few B cells, whereas mice with Btk deficiency generate nearly normal numbers of B cells, albeit with significant functional impairment (Maas and Hendriks, 2001; Fischer, 2001). BLNK deficiency in humans also results in a complete block at the pro-B cell stage. In contrast, a leaky block is seen in Blnk-deficient mice (Minegishi et al., 1999b; Pappu et al., 1999). One of the most intriguing differences in mouse and human B cell development is the requirement for interleukin 7 (IL-7) as an essential B lymphopoiesis growth factor in
Antigen Dependent B Cells
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AID CD19, CD21, CD40/CD40L, CD45 Ltab /LTBR, BTK, Lyn Irf4, Oca-B, Oct-2 D Syk D
PAX 5 D
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Stromal Cell FIGURE 9.4 Genetic defects in B cell development. The model of B cell development in Figure 9.1 is recapitulated here to illustrate gene defects that affect this process. Mutations identified in humans and mice are indicated in red and discussed in the text. The other defects shown in black have only been identified by gene targeting in mice, but may be found in humans as more immundeficient patients are studied. The human diseases IgAD and CVID affect antibody production, but the predisposing MHC-linked susceptibility gene(s) have not yet been identified, and there is no mouse model. See color insert.
9. The Development of Human B Lymphocytes
mice, but not humans (LeBien, 2000). Although human proB cells express the IL-7 receptor, IL-7 does not support robust proliferation. Moreover, individuals with functionloss mutations in either the ligand binding IL7Ra chain or the signal transducing gc gene have normal numbers of B cells (Puel et al., 1998; Leonard, 1996; Sugamura et al., 1996). In common with their mouse mutant counterparts, however, these individuals exhibit a severe block in T cell development. These observations highlight the need to identify the essential growth factor(s) for human B lymphopoiesis. The most frequently occurring immunodeficiency in humans is IgA deficiency (IgAD), which is characterized by a severe deficiency of both IgA1 and IgA2 isotypes (Burrows and Cooper, 1997; Hammarstrom et al., 2000; Schroeder, Jr., 2000; Cunningham-Rundles, 2001; Schroeder, Jr. et al., 1998). This heritable disorder is related to common variable immunodeficiency (CVI), characterized by a deficiency of all immunoglobulin isotypes. Members of the same family may have IgA deficiency, CVI, or intermediate patterns of immunoglobulin isotype deficiency. The extent of Ig deficiency is variable with age, and affected individuals may convert from isolated IgA deficiency to a CVI phenotype, or vice versa. The genetic basis for this spectrum of immunoglobulin deficiencies is still unknown, although there is good evidence indicating that the susceptibility gene(s) may lie within or near the MHC region. ICOS gene mutations have been associated with the CVI phenotype in one family, but this appears to be a rare gene defect among individuals with the IgAD/CVI immunodeficiency spectrum (Grimbacher, B., personal communication). The identity of the immunoglobulin insufficiency gene(s) in most IgAD/CVI patients remains elusive, and it is even unclear whether the defect involves the B cell, the helper T cell, or their interaction with antigen-presenting cells.
B LINEAGE LEUKEMIA Acute lymphoblastic leukemia (ALL) of B lineage origin is the most common type of cancer in children. Bone marrow leukemic blasts from approximately 75% of pediatric ALL patients have a pattern of gene expression generally consistent with the pro-B or pre-B stages of B cell development shown in Figure 9.1. B-lineage ALL is universally characterized by the expression of CD19 (and/or CD10) and varying degrees of IgH or IgL rearrangement. The disposition of the IgH locus ranges from both IgH alleles in germline configuration to functional rearrangements leading to the expression of cytoplasmic m H chains and cell surface pre-BCR. Functional IgL rearrangements leading to cell surface BCR expression are exceedingly rare in B-lineage ALL, and cases where this has been reported may represent occult lymphomas that have metastasized to
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the bone marrow. B-lineage ALL generally reflects the maturational arrest of a dominant subclone accompanied by a degree of apoptotic resistance that exceeds the sensitivity of normal B cell precursors. The molecular genetic abnormalities in B-lineage ALL include nonrandom chromosomal translocations that give rise to fusion genes such as TELAML1, MLL-AF4, and E2A-PBX (Look, 1997). How these distinct translocations (alone or in concert with a multiple additional karyotypic abnormalities and specific mutations) subvert the normal developmental program of B-lineage cells is presently unknown. The target of neoplastic transformation in B-lineage ALL (i.e., a cell that acquires a leukemia-disposing chromosomal translocation and/or additional mutations) could be a lymphoid progenitor or an earlier hematopoietic stem cell. Cytogenetic analysis (Quijano et al., 1997) and analysis of TCRd gene rearrangements (George et al., 2001) have suggested that at least some CD19+ B-lineage ALL originate in a CD19- progenitor. In contrast, examination of CD34+/CD19- cells yielded no evidence for the presence of the TEL-AML1 fusion gene in CD19- progenitors in patients with B-lineage ALL (Hotfilder et al., 2002). The cytogenetic abnormalities in CD19+ B-lineage ALL blasts do not appear to be present in other lymphohematopoietic cell lineages. Despite the uncertainty regarding the precise transformation target in B-lineage ALL, compelling evidence exists for an in utero origin in some cases (Wiemels et al., 1999), whereas cases with pre-B ALL expressing the E2A/PBX1 fusion protein appear to have a postnatal origin (Wiemels et al., 2002). A cDNA microarray analysis was used to compare gene expression profiles in the leukemic cells from four patients with B-lineage ALL with their normal bone marrow counterparts. Approximately 330 of 4,000 named human genes were found to be overexpressed in B-lineage ALL vis-à-vis normal CD19+/CD10+ cells (Chen et al., 2001). The elevated expression of the products of several of these genes, CD58, ninjurin1 (an adhesion molecule), creatine kinase B, and Ref1, was verified by immunofluorescence analysis. Importantly, cell surface CD58 emerged as a potential marker for minimal residual disease in B-lineage ALL since it could not be detected on normal CD19+ bone marrow cells. Another recent survey of gene expression in CD34+ hematopoietic stem cells and normal pre-B cells (Muschen et al., 2002) indicated that the number of unique genes expressed in preB cells was less than that expressed in hematopoietic stem cells. Greater than 10% of the genes expressed in pre-B cells encoded pre-BCR subunits or components of the pre-BCR signaling pathway, underscoring the critical role of the preBCR checkpoint in B cell development. In addition, a number of genes were unexpectedly upregulated in pre-B cells. These included ATM and genes encoding molecules that regulate apoptosis (TNFR2, FADD, TRAF1). Other groups (Moos et al., 2002; Yeoh et al., 2002) have utilized
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gene chips to analyze gene expression in a large number of newly diagnosed B-lineage ALL. In these studies, gene expression profiling was useful in identifying known subtypes of leukemia based on immunophenotype and cytogenetics, and identified unanticipated differences in individual patients that may eventually allow for the development of more tailored therapy. Like any other area of modern biology, the challenge for the future is how to harvest the huge amount of information generated by gene profiling to further elucidate the developmental biology of normal and leukemic B-lineage cells. The role of the bone marrow stromal cell microenvironment in regulating the balance between survival, proliferation, differentiation, and death of normal and leukemic B-lineage cells is an area of continuing investigation (Figure 9.5). Stromal cell cultures and the NOD-SCID mouse have been utilized to evaluate the survival and proliferation requirements of B-lineage cells (LeBien, 2000). Many leukemic cell lines have been established from patients with B-lineage ALL, and most are easily maintained in standard suspension tissue culture conditions (Matsuo and Drexler, 1998). B-lineage ALL cell lines have also been developed that retain varying dependencies on human bone marrow stromal cells for survival and proliferation (Shah et al., 1998; Bertrand et al., 2001; Shah et al., 2001). The BLIN-2 cell line, for example, has maintained a strict requirement
P P D m Normal BCP
D
Leukemic BCP
P
on human bone marrow stromal cells for optimal survival and proliferation (Shah et al., 1998). BLIN-2 cells express the pre-BCR, undergo intrinsic cell death (i.e., mitochondrial-dependent) 2 to 3 days following removal from bone marrow stromal cells, and survival and proliferation is inhibited by IL-7. A second cell line, designated BLIN-3, bears the t(4; 11)(q21; q23) chromosomal translocation that encodes the MLL-AF4 fusion gene (Bertrand et al., 2001). Similar to the EU12 B-lineage ALL cell line (Wang et al., 2003), BLIN-3 cells can make functional IgH rearrangements and express the pre-BCR. They also undergo intrinsic cell death 2 to 3 days following removal from bone marrow stromal cells, but their survival is promoted by IL7. These biological characteristics of BLIN-3 are remarkably similar to the developmental characteristics of normal B-cell precursors. A third cell line in this series, BLIN-4 (Shah et al., 2001), consists of two predominant subclonesBLIN-4E and BLIN-4L. BLIN-4E and BLIN-4L have identical clonal IgH rearrangements, express a pro-B phenotype (i.e., no pre-BCR expression), but show major differences in their dependency on bone marrow stromal cells for optimal survival and proliferation. The different stromal cell requirements may recapitulate a type of leukemic cell progression that occurs as B-lineage ALL undergo clonal evolution in vivo. The stromal cell–independent ALL-derived cell line EU-12 follows an intrinsic differentiation program in vitro. The CD34+ cells, which appear to provide the stem cell source of the leukemic population, can spontaneously differentiate into pre-B and B cells with intraclonal diversification of their VH and VL gene repertoires. Analysis of the ALL cell lines of various phenotypes should continue to provide insight into normal human B lymphopoiesis and leukemagenesis.
m
P
P = IL-7/? P = proliferation D = differentiation
Surv/Prolif Cytokines
P
Stromal Cell
FIGURE 9.5 Bone marrow (BM) stromal cells synthesize cytokines essential for the survival and proliferation of normal and leukemic B cell precursors (BCP). In this model, IL-7 constitutes a survival signal, and an unknown cytokine (or cytokines) constitutes a proliferative signal for normal and leukemic BCP. However, the responses to these cytokines differ. Normal BCP (light green) undergo a limited proliferative response. The predominant normal BCP undergoes proliferation and expresses cell surface pre-BCR. Normal proliferating pre-BCR+ cells subsequently differentiate into small pre-B cells (red) expressing cytoplasmic mH chains. BM stromal cell–dependent leukemic BCP (dark green) undergo a robust and continuing proliferative response that is independent of pre-BCR expression. Subsequent mutations can give rise to leukemic subclones that are no longer dependent on BM stromal cells. See color insert.
CONCLUSION The development of human B lineage cells closely parallels that of mice, but the differences that exist are significant. Particularly notable is the disparity in the requirement for interleukin 7. Failure to identify the functionally equivalent human cytokine that provides a bona fide proliferation stimulus to pro-B and large pre-B populations is an important deficiency in our understanding of human B cell development. Identification of this key cytokine (or cytokines) may also reveal how B-lineage ALL subverts the normal developmental program of B-lineage cells, perhaps through a mechanism in concert with specific chromosomal translocations and mutations that render a state of heightened apoptotic resistance in the leukemic clone. Recognition of a significant role for VH gene replacement in shaping the human antibody repertoire requires a search for the mechanisms that control this process, as this will be important in understanding antibody diversification and perhaps
9. The Development of Human B Lymphocytes
autoimmunity. Defining additional genetic defects in human immunodeficiency diseases, combined with studies of Blineage ALL, will continue to be useful for elucidating the developmental biology of normal B-lineage cells.
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10 Development and Function of B Cell Subsets JOHN F. KEARNEY Division of Developmental and Clinical Immunology, Department of Microbiology, University of Alabama at Birmingham, Birmingham, Alabama, USA
Precursor B cells differentiate into B lymphocytes after expressing a functional surface immunoglobulin receptor (sIgM). These newly formed B cells are then subject to further selection steps during their entry into the mature, long-lived pool of peripheral B lymphocytes (Rolink and Melchers, 1996). These steps involve a series of developmental programs and checkpoints and eventually result in the production of a diverse, complete repertoire reactive to almost all potential pathogens (Goodnow, 1997; Rolink and Melchers, 1998; Osmond et al., 1998). Phenotypic, topographic, and functional characteristics have been used to delineate subsets of mature B lymphocytes. Based on such criteria, these subsets have been shown to have different developmental programs as well as generation and maintenance requirements (Kantor et al., 1992; Stall et al., 1996). The most prevalent of these subsets is the mature B2 cell population, which is also heterogeneous: these recirculating cells locate predominately in the B lymphoid follicles (FO) of spleen and lymph nodes, while a special population of mostly nonrecirculating cells enrich primarily in the marginal zone (MZ) of the spleen (Gray et al., 1982; Oliver et al., 1997). B1 B cells are self-renewing cells with cell cycle and activation properties different from the bulk of recirculating B2 cells (Stall et al., 1996). These predominate in the peritoneal and pleural cavities. MZ B and B1 cells are characterized by their ability to respond early and rapidly in immune responses. These properties appear to be related to the apparent lower threshold of MZ and B1 cells for activation, proliferation, and differentiation into antibodysecreting cells than recirculating or immature B cells. In contrast, FO B cells recirculate rapidly and appear to be involved in interactions with T cells and respond to T-dependent antigens (Martin and Kearney, 2000a, 2000b, 2002).
Molecular Biology of B Cells
B cell subsets, as well as being functionally different, have preferences for particular niches in the immune system. Similar to the restricted TCR expression of g/d T cells and ab NK T and their particular geographical preferences (Bendelac et al., 1997, 2001), B1 lymphocyte subpopulations reside in the peritoneal and pleural cavities, and are also clear examples of the differential distribution of lymphocyte subsets (Hayakawa et al., 1999; Arnold et al., 1994; Bendelac et al., 1997). Their differential distribution in characteristic microenvironments is likely to be at least partially receptor driven, given the canonical receptors used by some of these cells (Hu et al., 2002). Phenotypic, microanatomical localization and functional differences characterize the splenic MZ and FO B cell subsets. The compartmentalization of each of these B cell subsets is suggestive of specialized functions linked to the niches within the spleen in which they reside. It has been proposed that the MZ B cells are involved in the initiation of a rapid first line of defense against blood-borne particulate antigens, hence their position in the marginal zone. Immune cells in this microenvironment are constantly bathed in blood and its associated contents. They are also intimately associated with metallophilic and marginal sinus-associated macrophages richly endowed with innate receptors involved in scavenging foreign and self-antigens. MZ and B1 cells share functional characteristics, suggesting that they may be selected similarly. B2 cells constitute the numerically preponderant B cell subset and are exemplified by FO B cells of the spleen and the majority of B cells in lymph nodes. This subset recirculates extensively and participates later in the T-dependent Ab responses. However, this subset resides and recirculates within lymphoid tissues in a microenvironment that is separated from direct blood contact by cellular and connective
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tissue barriers (Oliver et al., 1999; Martin and Kearney, 2002). All B cell subsets must be derived from newly formed (NF) B cells traveling from bone marrow to appropriate sites in the periphery under the influence of some known molecules (Martin and Kearney, 2002). However, the mechanisms producing the enrichment of B cell subsets and the relative roles of self and environmental antigen signals and survival signals are largely unknown. Multiple hypotheses propose BCR signaling to be crucial in the enrichment of FO, B1, and MZ B cells in their independent niches since this process is impaired in xid, CD19-/-, CD45-/-, aiolos-/-, and other genetically manipulated mice (Okamoto et al., 1992; Cariappa and Pillai, 2002; Amano et al., 2003). Needless to say, this is a controversial area and is discussed later. Recent studies in which the B cell receptor was replaced by an EBV-derived membrane protein bearing multiple B cell activating motifs showed normal B cell subset formation and lymphoid tissue localization. These findings would seem to implicate signaling molecules other than the BCR in determining the fate of newly formed B cells in the periphery (K. Rajewsky, personal communication). Irrespective of which mechanism is dominant in this positioning effect, it does not alter the proposition that the immune system causes clones to be sequestered in strategically located sites where their BCR-induced functional capabilities are suited for a particular set of environmental antigens associated with a given “geographical location” (Kipps et al., 1998). It is clear from multiple experiments that Notch and downstream sigaling pathways may also play a role in establishing a functional MZ B cell subset, although it cannot be entirely excluded that this may have been the result of secondary effects accompanying the conditional knock-out (KO) of RBP-J (Kuroda et al., 2003; Tanigaki et al., 2002).
SELECTION AND DIFFERENTIAL SURVIVAL MECHANISMS—B CELL RECEPTOR SIGNALING Conditional KO of BCR shows that all B cells appear to be constantly in need of some kind of BCR-mediated signal from their microenvironment not only for clonal selection but for their continued survival (Schattner et al., 1995; Fagarasan et al., 2000b). Evidence obtained by several different experimental approaches suggests that BCR specificity is critical for clonal development into B1, FO, or MZ subsets. Studies in several independent Ig transgenic mice show that the density of surface BCR also may be involved in this decision by specifically modulating the amount of clonal signaling. In anti-DNA heavy chain transgenic mice, normally deleted B cells enrich in the MZ but this rescue is affected by the expression of two light chains. (Li et al.,
2002). Likewise, when surface expression of a B1-type receptor is reduced through the expression of a second heavy chain, B cell development proceeds towards the B2 compartment (Watanabe et al., 2000). Similarly, the size of the B1 compartment is larger in homozygous anti-RBC transgenic mice than in heterozygous mice (Ohdan et al., 2000). Mechanisms regulating B cell density through surface BCR density not only play a role in the B1 versus B2 decision but also are in effect at checkpoints that act to prevent selfreactivity by editing and deletion (Boes et al., 1998; Ochsenbein et al., 1998a). Modulation of BCR activity in concert with several co-receptors and downstream molecules such as CD5, CD19, CD22, CD21, CD45, btk, lck, and SHP-1, clearly affects the outcome of microenvironmental signals that affect B cell development and the maintenance of B cells within the immune system (Su and Tarakhovsky, 2000; Okamoto et al., 1992; Martin and Kearney, 2002). Knowledge of the retention and migration signals for B cell subsets to and from these sites is a key step in understanding why an anatomical separation of B cell subsets occurs and how these cells home to these distinctive sites. The B1 as well as MZ B cell populations appear to be enriched in clones that are self-reactive but also react with bacterial antigens (Okamoto et al., 1992; Garrone et al., 1995). The recruitment and enrichment of specific clones may depend on their selective activation and survival in the specialized niche in which they reside. Canonical MZ B cell clones survive preferentially over other clones in vivo and in vitro (Okamoto et al., 1992), similar to the receptor-driven selection of B1 cells (such as the VH11Vk9 clone, which survives in culture better than B2 cells). Thus, both MZ and B1 cells may owe their enrichment to preferential survival mechanisms (Guinamard et al., 2000). It has been previously shown that another clone with anti-PtC activity (VH12-Vk4) has a selective advantage in vivo over competitors at multiple checkpoints (Baumgarth et al., 1999; Baumgarth et al., 2000). Although microanatomical localization and phenotypic markers were first used to define B cell subsets, the molecular basis for the characteristic localization is now beginning to unravel. Rapid progress in the fields of chemokines and G-protein coupled receptors (GPCRs) have revealed complex mechanisms of retention, migration, and function. In the spleen, the chemokine BLC is clearly responsible for the development of B cell follicles (Gunn et al., 1998; Cyster et al., 1999). More recently, a novel chemokine receptor has been identified on MZ B cells that may be responsible for MZ B cell retention (Behrens, personal communication). This receptor may be involved with the chemokine-driven generation and maintenance of MZ B cells. Gene-targeting of pyk-2 (Guinamard et al., 2000), DOCK2 (Fukui et al., 2001), and lsc (Girkontaite et al., 2001), potential signaling pathways downstream of chemokine receptors, results in a
10. Development and Function of B Cell Subsets
drastic reduction or absence of the MZ B cell compartment. Pyk-2, a tyrosine kinase, may mediate signals from GPCR for chemokine, lipids, integrins, and antigen receptors, and clearly plays a major role in the generation of MZ B cells and the ability to respond to TI antigens.
COMPARTMENTALIZATION OF B CELL SUBSETS Although the various mechanisms described are important in the development and function of B cell subsets, the pathways by which they enter their characteristic niches have been unclear. Recent elegant work has shed light on the role of integrins and chemokines on the entrance pathways of B cells into the spleen and peritoneal cavities. It was recently shown that all B cells entering the spleen likely do so by the involvement of integrins LFA-1 and a4b1, binding to ICAM-1 and VCAM-1 respectively, and may also involve fibronectin. Antibodies to these integrins administered together prevent the entry of B cells into both the MZ and follicles (Lu and Cyster, 2003). MZ B cells express higher levels of these integrins, which may account for their increased binding to the ICAM-1 and VCAM-1 ligands on a variety of endothelial and hematopoietic cells in the MZ, thus preventing their passing through the endothelial lining of the MZ in the resting state. They also showed that antiintegrin antibodies caused the dissolution of MZ B cells, as did the inhibition of chemokine signaling. Antigen-induced migration of FO B cells into the T cell areas did not depend on these integrins; however, LPS-induced relocalization of MZ B cells was accompanied by integrin downregulation (Lo and Cyster, 2002). Thus, the possible scenario exists that environmental signaling to MZ B cells or to lipid receptors from unique ligands existing in the MZ microenvironment may lead to intrinsic upregulation of these integrins and/or chemokine receptors preferentially on these MZ B cells via downstream signaling. Decreased expression of either chemokine receptors or integrins alters the positioning of MZ B cells and, under the influence of antigen, permits their entry into the follicular area and access to the T–B border (Balazs et al., 2002). Cyster’s group also has shown that in mice lacking the chemokine, CXCL13, B1 cells are deficient in peritoneal and pleural cavities but not in spleen. They further showed that cells in the omentum and peritoneal macrophages produce CXCL13. In adoptive transfers, B1 cells home to the omentum and the peritoneal cavity in a CXCL13-dependent manner. CXCL13-/- mice are also deficient in the characteristic natural phosphorylcholine (PC)-specific antibodies and in their ability to mount an anti-PC response to peritoneally administered pneumococci. These clonally restricted antibody responses are produced by B1 cells (Ansel et al., 2002). Their findings provide the
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first insight into the mechanism of B1 cell homing and compartmentalization in the body cavities and re-emphasizes the critical role for the B1 cell in the production of natural antibodies (Benedict et al., 2001). Other factors may also be involved in the establishment and maintenance of B cell subsets. Recently, a new member of the TNF family, BAFF has been implicated in the survival of peripheral B cell subsets (Ochsenbein et al., 1999b; Macpherson et al., 2000; Ochsenbein et al., 2000; Zeng et al., 2000). Expanded B cell compartments occur in transgenic mice expressing BAFF, which is associated with enhanced B cell survival and the expansion of particular B cell subsets and autoimmune phenomena. The exact outcome of transgenic BAFF expression depended on the promoter–enhancer combinations used (Macpherson et al., 2000); in one, the autoimmunity was associated with increased splenic B1 B cells. Another, using liver-specific surface and generalized soluble expression (Ochsenbein et al., 1999), favored the transitional and MZ B cell compartments (Zeng et al., 2000). With ubiquitous expression (b-actin promoter), the autoimmune manifestations were preceded by a generalized B cell expansion (Ochsenbein et al., 2000). The functional sites of interaction between BAFF, expressed mostly by macrophages and dendritic cells, and its receptors (BCMA and TACI) are not known but these and other like molecules play a key role in the development, maintenance, and activation of B lineage cells.
OTHER FACTORS INVOLVED IN FORMATION OF B CELL SUBSETS If differential responsiveness and tonic signaling through the Ig receptor is necessary for B cell subset development, what are the unique mechanisms that permit B1 cells with higher affinity self-interactions to survive? B1 cells are less susceptible than both FO and MZ splenic B cells to antiIgM–induced apoptosis in vitro. In parallel with T cells, where CD5 is involved in downregulatory functions, CD5 may also be involved in decreasing B cell receptor–induced cell death in B1a cells (Wang et al., 1996; Fredrickson et al., 1999). CD5 expression on CLL and MZ lymphomas may reflect a relationship between self-renewing activated B1 cells and these neoplastic B cells (Rothstein et al., 1995; Lagresle et al., 1996; Hirose et al., 1997). The maintenance of peripheral tolerance also involves the elimination of activated T and B cells by Fas-mediated apoptosis (Batten et al., 2000). Although multiple pathways are involved in the apoptosis of B cells, Fas-triggered apoptosis eliminates activated B cells, including bystander B cells (Cook et al., 1999; Tachibana et al., 1999). B2 cell susceptibility to Fas-mediated apoptosis is enhanced by CD40mediated upregulation of Fas, whereas Fas susceptibility is
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decreased without a concomitant reduction of surface Fas expression (by signaling through the BCR) (MacKay et al., 1999; Smith et al., 1995; Rathmell et al., 1996; Dighiero and Borche, 1990; Fagarasan and Honjo, 2000). Recently, we found, in B1 compared to B2 cells with B1 cells, that proliferative responses after CD40 ligation are comparable, but Fas upregulation is impaired in B1 cells, thus making them more resistant to Fas-mediated apoptosis (unpublished). In NZB/W F1 mice, CD40--activated CD5+ B cells contained both Faslo and Fashi subsets; however, only the Faslo B cells were Fas resistant. Additionally, the IgG anti-DNA antibody was synthesized by splenic Faslo subpopulations in aged NZB/W F1 mouse (Won et al., 2000). The impairment of Fas induction in B1 cells after CD40 ligation is likely responsible for the maintenance of self-reactive B cells in this subset and their tendency to give rise to CLL-like B cell tumors, a proportion of which make autoimmune antibodies (Le Naour et al., 2000). An alternative model has been proposed to explain the regulation of B1 and MZ B T–independent antibody responses in the absence of T cell activity (Miyado et al., 2000). In this model, a balance arises between negative signals derived from P1/PDL-1 interactions and positive signals mediated through BLyS/TACI interactions by BCRactivated B1 and MZ B cells. Such a regulatory network may explain how the upregulation of survival signals on B1 and MZ B cells with BCR of low avidity to self antigens may prevent their maturation into active antibody-secreting cells and promote their maintainance and/or expansion and “self renewal.” CD5 expression by B1a B cells may be associated with BCR–self antigen interactions. However, the developmental stage or microenvironmental sites at which B1 cells receive these proposed signals are not known. More likely is a mixed hypothesis that B1 and B2 cells have separate precursors and that antigenic induction of the CD5hi B1 cells is pre-programmed for a given set of precursors (Kipps et al., 1998). The accumulation of self-reactive B1 cells then occurs in the peritoneal and pleural cavities, with small populations in other tissues, including the spleen. By flow cytometry CD5-/- mice have a lower apparent intensity of CD5 staining of B cells compared to CD5+/+ littermates, suggesting that all B cells may constitutively express low levels of CD5 (Fredrickson et al., 1999). Indeed, in some other species, all B cells may express CD5 under appropriate conditions (Jurgens et al., 1995; Knabel et al., 1993; Raman and Knight, 1992). The functional deletion of CD5 does not result in dramatic abnormalities in the immune system as a whole nor in B1 cell functions. However, just as CD5 may downregulate T cell activities, there is evidence that in B cells, a similar function for CD5 may be operative (Wang et al., 1996). The “activated” phenotype of the B1a subset, similar to that of the MZ B cell subset, may result from the BCR self-reactive specificities of these cells. Additionally,
the microenvironment in which B1 cells are located maintains them in state ready to react rapidly to potentially infectious organisms or gut-associated antigens and (Gross et al., 2000).
CONCLUSION This chapter is not meant to be an exhaustive review of the development and function of B cell subsets. More comprehensive reviews have recently been published in this area (Martin and Kearney, 2000b, 2001, 2002). Future research will be directed at the elucidation of clonal signals and co-signals and the miroenvironments within which B cell subsets receive these developmental guides. Knowledge of the chemokines and adhesion molecules that are involved in the direction of and retention of B cells within these microenvironments will be forthcoming. A closely associated field will involve the identification of resident cell types within the characteristic environment for each B cell subset and the functional interactions that occur between these cells during normal development, and in immunological functions and disease.
Acknowledgments We thank Ann Brookshire for editorial help and members of the Kearney lab for comments and discussions. This work was supported by NIH grants AI 14782 and CA13148.
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11 Structure and Function of B Cell Antigen Receptor Complexes MICHAEL RETH
JÜRGEN WIENANDS
Biologie III, University of Freiburg and Max Planck Institute for Immunobiology, Freiburg, Germany
Department for Biochemistry & Molecular Immunology, University Bielefeld, Bielefeld, Germany
The B cell antigen receptor (BCR) controls the development, activation, and maintenance of B lymphocytes. Despite extensive efforts over the last 10 years, the exact structure and activation mode of this receptor is only partly understood. Indeed, it is difficult to study this multicomponent transmembrane protein complex through biochemical or genetic means. A recently developed system for the reconstitution of BCR signaling helps to gain more information about the activation mode of this receptor. A unique feature of the BCR is that it can be activated by many structurally different ligands that immunologists summarize by the word antigen. Antigens recognized by B cell are in most cases foreign substances and comprise a heterogeneous group of molecules including proteins, DNA, polymeric sugars, or other polymeric molecules. The ability of the BCR to become activated upon binding to such a structurally diverse array of antigens indicates that the activation mechanism of the BCR must be different from that of other receptors that have only one or a limited set of ligands (Reth et al., 2000). For example, upon binding to its cognate ligand, namely the erythropoietin (EPO) molecule, the EPO receptor is fixed in an active conformation that allows signaling (Livnah et al., 1998, 1999). Other molecules can bind the EPO receptor without achieving this goal. The BCR, however, does not require a precise antigen structure for activation.
the mIg molecule and the Ig–a/Ig–b heterodimer in the membrane of the endoplasmic reticulum (ER) is a prerequisite for the transport of the BCR to the cell surface. All five major classes of mIg (mIgM, mIgD, mIgG, mIgA, and mIgE) are associated with the Ig–a/Ig–b heterodimer, presumably with a 1 : 1 stoichiometry (Schamel and Reth, 2000). Evidence for an oligomeric organization of the IgMBCR and IgD-BCR has been found in a study involving native gel seperation analysis (Schamel and Reth, 2000). These data led to the model that a conformational change of the oligomeric antigen receptor leads to activation of the BCR (see below). The mIg molecule is a tetramer consisting of two identical heavy (H) chains and two identical light (L) chains. The mIgM and mIgD molecules do not have a large cytosolic part that can interact with intracellular proteins. Thus, the signaling function of these mIg classes relies mostly on the Ig–a/Ig–b heterodimer. Ig–a and Ig–b share many structural features. Both proteins carry a glycosylated extracellular Ig domain, a linker region with the heterodimer-forming cysteine, one transmembrane part, and a cytoplasmic tail sequence of either 61 (Ig–a) or 48 (Ig–b) amino acids. These cytoplasmic sequences are the evolutionarily most conserved part of these transmembrane proteins, indicating that they have an important cellular function. The cytoplasmic tails of Ig–a and Ig–b contain a consensus sequence called immunoreceptor tyrosine-based activation motif (ITAM), which is also found in other receptors of the multicomponents immune receptor family (MIRR) (Cambier, 1995; Reth, 1989). The sequence of this is D/Ex7D/ExxYxxLx7YxxL/I. The two tyrosines in the ITAM sequence are phosphorylated during the activation of the BCR and become a binding target for signal transducing elements. The cytoplasmic tail of Ig–b carries only the two
STRUCTURE OF THE BCR COMPLEX The BCR comprises the membrane-bound immunoglobulin (mIg) molecule and the Ig–a/Ig–b heterodimer mediating antigen binding and signaling, respectively (Kurosaki, 1998; Wienands and Engels, 2001). The proper assembly of
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ITAM tyrosines, whereas that of Ig–a carries two additional tyrosines, the most C-terminal of which (Y204) also becomes phosphorylated during receptor activation (see below).
COUPLING BETWEEN THE BCR AND SYK The engagement of the BCR results in the activation of protein tyrosine kinases (PTK) and the rapid phosphorylation of several PTK substrate proteins. The PTKs Syk, Lyn, and Btk are involved in this process. Until recently, it was thought that the Src-family kinase Lyn was the first kinase to interact with the BCR and phosphorylate the two ITAM tyrosines, thereby allowing Syk recruitment and activation. In a new reconstitution system based on inducible coexpression of the murine BCR and its signaling elements in Drosophila S2 cells, it was found, however, that only Syk but not Lyn phosphorylates both ITAM tyrosines (Rolli et al., 2002), thus confirming studies obtained by in vitro kinase assays (Flaswinkel and Reth, 1994). It is now clear that the initiation of signaling at the BCR involves a positive feedback between Syk and the ITAM sequences, resulting in rapid amplification of the BCR signal. An important feature of this signal amplification process is the regulation of the kinase activity of Syk. Syk is an allosteric enzyme, whose activity is regulated by its tandem SH2 domains (Rowley et al., 1995; Shiue et al., 1995). In the absence of an ITAM sequence, Syk mostly exists in a closed conformation where the two SH2 domains block the kinase domain of Syk. Alone, Syk has therefore only a low kinase activity. However, once it meets and phosphorylates both ITAM tyrosines, it can bind to the phosphorylated tyrosines via its tandem SH2 domains. This binding fixes Syk in an open, active conformation at the inner leaflet of the plasma membrane. Here the active Syk can rapidly phosphorylate neighboring ITAM sequences, thus resulting in more Syk recruitment, activation, and the rapid amplification of the signal. This Syk activation model is supported by the phenotype of a Syk mutant carrying a binding-deficient C-terminal SH2 domain. This mutant is nearly as deficient in ITAM tyrosine phosphorylation as a kinase-negative mutant of Syk. In contrast, a Syk mutant with a deletion of both SH2 domains is constitutively active but no longer preferentially phosphorylates the ITAM tyrosines of Ig-a (Rolli et al., 2002). This mutant analysis demonstrates that the tandem SH2 domains of Syk have a dual role. In resting B cells, they lock Syk in its inactive conformation. Upon BCR activation, they allow Syk to bind to phosphorylated ITAM sequences. The intramolecular regulation of the Syk kinase and its activation by the ITAM sequence ensures that Syk is only active at the right place inside the cell, namely the BCR.
REDOX REGULATION OF BCR SIGNALING The positive Syk/ITAM feedback loop allows a rapid amplification of the BCR signal. However, to prevent hyperactivity, such positive feedback loops must be tightly controlled inside the cell. In the case of the Syk/ITAM loop, this control is efficiently exerted by protein tyrosine phosphatases (PTP) (Neel and Tonks, 1997; Pani et al., 1995). Indeed we found that in the Drosophila S2 cell system the Syk-mediated signal amplification at the BCR is abolished by co-expression of the PTP SHP-1 (Rolli et al., 2002). Interestingly, the target of SHP-1 seems not to be Syk directly, but rather the two phosphorylated tyrosines of the ITAM, which regulate Syk activity. In general, a PTP has a 10- to 100-fold higher turnover rate than a PTK, because a PTP simply removes a phosphate from a PTK substrate using the abundant water molecules as a donor in this reaction, whereas a PTK has to bind simultaneously ATP and the substrate protein to catalyze the phosphate transfer. A race between an active PTP and PTK for the respective dephosphorylation and phosphorylation of a substrate protein is therefore always won by the PTP. Thus the detection of an increased PTK substrate phosphorylation in activated B cells is not only due to PTK activation but also to PTP inactivation. Recent studies on several receptors found that signal transduction from these receptors requires kinase activation as well as phosphatase inhibition (Bae et al., 1997; Meng et al., 2002; Xu et al., 2002). How then are phosphatases inhibited inside the cells? All phosphatases carry an invariant, reactive cysteine (C-SH2) in their catalytic center that takes part in the removal of phosphate groups. In the presence of H2O2, this cysteine is reversibly oxidized (C-SOH), thus preventing all phosphatase activity (Lee et al., 1998; Meng et al., 2002). By inhibiting phosphatases, H2O2 can function as a secondary messenger in signal transduction (Baeuerle et al., 1996; Finkel, 1998; Reth, 2002; Rhee et al., 2000). H2O2 is indeed produced in stimulated B and T cells via the activation of the membrane-bound NADPH–oxidase complex, producing superoxide anions (O2-) that react with water to yield H2O2 and singlet oxygen (1O2) (Devadas et al., 2002; Qin et al., 1999). However, H2O2 is a short-lived molecule that exists inside the cell only close to its site of production. Interestingly, it was recently found that the BCR and TCR forms a catalytic center in the V : V interphase. This center catalyzes the conversion of singlet oxygen to H2O2, thus increasing the production of hydrogen peroxide in the vicinity of these receptors (Datta et al., 2002; Wentworth et al., 2001). In summary, the following scenario of BCR activation seems plausible (Figure 11.1). On resting B cells, the BCR forms an ordered oligomeric structure (Figure 11.1A) and the protein–protein interaction inside this oligomer may set
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FIGURE 11.1 New model for the antigen dependent activation of the BCR. A. On resting B cells the BCR forms an oligomeric complex of defined stoichiometry. Signal transduction from the BCR is inhibited by presence of PTP. B. Exposure to antigen results in the opening of this oligomeric complex and the targeting of the BCR to membranes containing an active NADPH-oxidase. The increased H2O2 production by the NADPH-oxidase inhibits the PTP around the BCR allowing the rapid amplification of the BCR signal through a positive Syk/ITAM feedback loop. See color insert.
critical thresholds for the activation of this receptor (Batista and Neuberger, 1998). The binding to antigen disturbs the oligomeric BCR in such a way that sequences situated either in the transmembrane or cytosolic part become exposed and target the BCR to a different membrane compartment. Note that according to this model many different antigens can disturb the BCR, as long as they are polyvalent structures, and thus BCR activation becomes independent of the structure of the antigen (Reth et al., 2000). It is further assumed that upon antigen binding the BCR is localized to membranes where the NADPH-oxidase resides (Figure 11.1B). Due to the H2O2 production in the vicinity of the NADPHoxidase, the co-localization of the BCR with this enzyme complex results in PTP inhibition at the BCR and, subsequently, signal amplification through the positive Syk/ITAM feedback loop. In resting B cells, the NADPH-oxidase seems to be not very active. However, signals through costimulatory receptors like CD19 and CD40 or through Toll-like receptors are able to activate this enzyme. Thus, T cells and the receptors of the innate immune system may participate in the antigen specific activation of B lymphocytes. In activated B cells, BCR signals processed through Lyn and Syk result directly in NADPH-oxidase activation, and this may be one of the reasons for the observed synergy of these two PTKs in BCR signaling (Kurosaki et al., 1994; Qin et al., 1996; Takata et al., 1994).
FIGURE 11.2 Known protein:protein interaction at the adapter protein SLP-65. Upon BCR activation the adaptor is phosphorylated by Syk on several critical tyrosines (72–189) which become a binding target of the indicated intracellular signaling molecules. The SH2 domain of SLP-65 has also two known interaction partner namely hematopoietic progenitor kinase (HPK1) and the tyrosine Y204 in the Ig-a sequence of the BCR.
ITAM- AND NON-ITAMCONTROLLED SIGNALING PATHWAYS TO SLP-65 The Syk/ITAM positive feedback loop leads to the activation of many Syk molecules, which then phosphorylate the intracellular adaptor protein SLP-65 (also known as BLNK or BASH) (Fu et al., 1998; Goisuka et al., 1998; Wienands et al., 1998). Phosphorylated SLP-65 nucleates the formation of the Ca2+ initiation complex (Figure 11.2) by providing docking sites for the SH2 domains of BTK (Hashimoto et al., 1999; Su et al., 1999) and phospholipase C (PLC)-g2 (Fu et al., 1998; Ishiai et al., 1999a, 1999b). Site-directed mutagenesis of BLNK/SLP-65 and peptide
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binding studies recently identified three tyrosine residues on human BLNK (Y84, Y178, Y189) as being responsible for the recruitment of PLC-g2 (Chiu et al., 2002). The subsequent one-by-one loss of these tyrosines gradually reduced the intensity of the Ca2+ signal. The mutation of tyrosine Y96 on human BLNK, or its equivalent Y115 on chicken BLNK, prevents the binding of Btk to the adaptor. When expressed on a SLP-65–deficient background, SLP-65 mutants carrying solely the PLC-g2 or Btk binding site still display BCR-induced SLP-65 phosphorylation and binding of either PLC-g2 or Btk, but were incapable of fully restoring Ca2+ mobilization and NFAT transcriptional activation. The same result was obtained when both SLP-65 mutant proteins were expressed simultaneously in one cell, showing that the two mutants cannot complement each other in trans. This analysis demonstrates that the components of the Ca2+ initiation complex have to be assembled in cis on the same SLP-65 molecule in order to achieve coordinate enzymatic activation. Conformational changes, together with multiple trans- and autophosphorylation steps activate Btk (Afar et al., 1996; Baba et al., 2001; Mahajan et al., 1995; Rawlings et al., 1996). Dual phosphorylation of PLC-g2 by Btk and Syk fully activates its ability to generate inositol-trisphosphate (IP3) and diacylglycerol (Fluckiger et al., 1998; Takata and Kurosaki, 1996; Takata et al., 1994). These second messengers trigger Ca2+ mobilization and protein kinase C (PKC) activation, respectively. The Ca2+ channel activity of the IP3 receptor in the ER membrane is triggered upon IP3 binding. Its activity is further increased upon tyrosine phosphorylation and the formation of a trimolecular complex between the IP3 receptor, the B cell scaffold protein with ankyrin repeats (BANK), and Lyn (Yokoyama et al., 2002). The defect in the Ca2+ response in Lyn-deficient DT40 cell may be due in part to the role of Lyn in the IP3 receptor activation, rather that ITAM phosphorylation as thought previously (Takata et al., 1994). The increased intracellular Ca2+ concentration is maintained by import of Ca2+ from the extracellular medium through Ca2+ channels in the plasma membrane, and transient receptor potential (TRP) channel proteins maybe involved in this activity (Mori et al., 2002). The importance of the SLP-65–controlled Ca2+ initiation complex for B cell function is evident from gene targeting experiments in the DT40 B cell line (Ishiai et al., 1999a, 1999b; Takata and Kurosaki, 1996; Takata et al., 1994) and in mice (Hashimoto et al., 2000; Hayashi et al., 2000; Jumaa et al., 1999; Pappu et al., 1999; Wang et al., 2000). Loss of one of the components of this complex reduces or prevents the Ca2+ release upon BCR signaling and severely impairs B cell development and function. In humans, mutations in the btk gene almost abrogate B cell development and result in X-linked agammaglobulinemia (XLA) (Fruman et al., 2000). The same clinical features are described for immunodeficient patients with a splicing defect in slp-65 causing
lack of SLP-65 expression (Minegishi et al., 1999). What remains to be solved is how the Ca2+ initiation complex is tethered to the plasma membrane, specifically to the lipidraft fraction, in order to provide PLC-g2 with access to its phospholipid substrate. One mechanism involves binding of the pleckstrin homology (PH) domains of PLC-g2 and Btk to phosphatidylinositol-3,4,5-trisphosphate (PtdIns-3,4,5P3), a product of activated phosphoinositide 3-kinase (PI3K). This interaction appears to be tightly controlled at different levels: first, by a multistep regulation of PI3K action and second, by newly discovered Btk-binding proteins. PI3K is a dimeric enzyme complex comprising the SH2 domain-containing p85 regulatory subunit and the p110 catalytic subunit. Upon B cell activation, PI3K is targeted to the plasma membrane by SH2-mediated binding of p85 to phosphotyrosine residues in the cytoplasmic tail of the BCR co-receptor subunit CD19 (Kurosaki and Okada, 2001). However, proper localization to the lipid-raft fraction requires additional tyrosine phosphorylation of cytoplasmic adaptor molecules like the B cell adaptor of PI3K (BCAP) (Okada et al., 2000; Yamazaki et al., 2002), the B cellassociated adaptor of 32 kDa (Bam32) (Marshall et al., 2000; Niiro et al., 2002), and the Grb2-associated binding protein 1 (Gab1) (Ingham et al., 2001). Enzyme activity of PI3K is positively controlled by the small GTPase Rac1, which itself is regulated by phosphorylated GDP/GTP exchange factors of the Vav family; that is, Vav 1–3 (DeFranco, 2001). As Vav adaptors are activated by PI3K, the three signaling elements (Rac, PI3K, Vav) may form a positive feedback loop inside the cell. The molecular details of PI3K regulation remain to be elucidated. The complex control of PI3K activity reflects the fact that the metabolism of membrane phospholipids not only affects localization of the Ca2+ initiation complex but is critical for many cellular responses, such as vesicular transport or survival. Multiple regulatory circuits also operate at the PH domain of Btk to control BCR-induced Ca2+ flux and perhaps other Btk effector functions (for example, WASP-mediated reorganization of the cytoskeleton). Recently, several binding proteins of the Btk PH domain have been identified, some of which seem to attenuate the membrane attachment and/or enzymatic activity of Btk. The inhibitor of Btk (IBtk) has an apparent molecular weight of 26 kDa and is restricted to cells of the hematopoietic lineage (Liu et al., 2001). The binding of IBtk to the Btk PH domain downregulates kinase activity, which might be decreased further by the more ubiquitously expressed SH3-domain binding protein SAB (Yamadori et al., 1999). However, it remains to be shown that either of these molecules is released from Btk upon B cell activation. A stimulation-dependent association has been reported between the Btk PH domain and the PKC-b isoform (Kang et al., 2001). After being activated by Btk, PKC-b is suggested to phosphorylate a negative-regulatory serine residue in the adjacent Tec homology region of Btk. The abrogation of this
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negative feedback loop may explain the increased and prolonged Btk kinase activity in immunodeficient PKC-b-/mice (Leitges et al., 1996). Collectively, the importance of the PH domains of Btk and also PLC-g2 for Ca2+ mobilization is well documented. However, their binding to phospholipids seems to play a more prominent role for the maintenance of the Ca2+ initiation complex at the plasma membrane rather than for its initial translocation from the cytosol (Rawlings, 1999). In activated T lymphocytes, a tyrosine-phosphorylated transmembrane adaptor protein called linker of activated T cell or LAT performs the latter function by recruiting PLC-g1, SLP-76, and the BtK-related LAT also functions as membrane anchor for a Ca2+ initiation complex in pre-B cells (Su and Jumaa, submitted) but not in mature B cells. Recently, a second member of the LAT family of transmembrane adaptor proteins was identified in mature B cells and termed non-T cell activation linker (NTAL) (Brdicka et al., 2002) or linker for activated B cells (LAB) (Janssen et al., 2003). The genes for both proteins, LAT and NTAL/LAB show the same exon–intron organization, indicating that they have evolved from a common ancestor gene. Also, the overall structure of the proteins is very similar in that they both possess multiple tyrosine phosphorylation sites and a cysteine-based motif for the fatty acid modifications of their N-terminus. The latter feature is responsible for the constitutive localization of LAT and NTAL in lipid rafts. Apart from its detection in B cells, NTAL is also expressed in macrophages, mast cells, and NK cells. NTAL expression in LAT-deficient Jurkat T cells reconstitutes some aspects of TCR signaling, but whether in B cells NTAL functions similarly to LAT in T cells awaits further analysis. A direct mechanism by which SLP-65-containing signaling complexes can translocate from the cytosol to the plasma membrane involves the activated BCR. Affinity-purification experiments revealed a stimulation-dependent association of the SLP-65 SH2 domain with phosphorylated Ig-a. Surprisingly, the SLP-65 binding site turned out to be not one of the ITAM phosphotyrosines but the C-terminally located phosphotyrosine 204, which is separated from the last ITAM tyrosine by eleven amino acids (Engels et al., 2001b; Kabak et al., 2002). This spacing is identical to that of the two ITAM tyrosines. The detailed functional role of this organization is not known but may be important for the phosphorylation of Y204. Phosphorylation outside of the ITAM was unexpected, because no Ig-a phosphorylation could be detected upon mutation of the ITAM tyrosine to phenylalanine (Flaswinkel and Reth, 1994). It now appears that the dual ITAM phosphorylation is a prerequisite for Y204 phosphorylation. It is thus likely that Syk that requires an ITAM sequence for its activation is phosphorylating Y204. However, note that the kinase domain of Syk is oriented towards the plasma membrane, whereas Y204 is facing the cytosol. The evolutionary conservation of Y204 and its posi-
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tion relative to the Ig-a ITAM (Sayegh et al., 2000) indicate that recruitment of its ligand, SLP-65, serves an important role for B cell signaling. A possible LAT-like function of Y204 in recruiting the Ca2+ initiation complex was suggested by experiments with Ig-a transmembrane chimeras (Kabak et al., 2002). This model could, however, not be confirmed in the context of the complete multimeric BCR. Reconstitution of SLP-65–deficient DT40 B cells with an SH2 domain mutant of SLP-65 rescued BCR-induced Ca2+ mobilization to an expect similar to that observed in wild-type cells (Dittmann, Engels, Kurosaki and Wienands, unpublished results). Another report proposes a function of Y204 for targeting the antigen-ligated BCR to the MHC IIloading compartment (Siemasko et al., 2002). Whatever the role of the non-ITAM phosphotyrosine might be, it is unique for Ig-a, as none of the known ITAM-containing immunoreceptor signaling subunits possess tyrosines outside their ITAM. Consistent with this, Ig-a and Ig-b perform distinct signaling functions in vitro (Choquet et al., 1994; Kim et al., 1993) and in vivo (Kraus et al., 2001; Reichlin et al., 2001; Torres et al., 1996; Torres and Hafen, 1999; Tseng et al., 1997), both of which are mandatory for the proper development and function of mature B cells (for review see Wienands and Engels, 2001). The phosphorylated hematopoietic progenitor kinase (HPK) 1 has been recently described as a second ligand of the SLP-65 SH2 domain and this complex formation modulates BCR-induced activation of the NF-kB pathway (Tan et al., 2001; Tsuji et al., 2001). Collectively, the above findings underline a fundamental difference between the SLP effector molecules, SLP65 and SLP-76, in B and T cells, respectively. The ligand for the SH2 domain of SLP-76 is the SLP-76–associated protein of 130 kDa (SLAP130), which couples TCR signaling to integrin function and results in altered adhesion properties of activated T cells (Yablonski and Weiss, 2001). SLAP130 is not expressed in B cells, but the amino acid sequence of the reported SLP-76 binding site in SLAP130 (pYDDV) is very similar to that of the SLP-65 binding site in Ig-a (pYQDV) and identical to that of HPK1 (pYDDV). Whether there is a B cell counterpart of SLP130 is another key objective in the ongoing research in B cell signal transduction.
ITAM-INDEPENDENT SIGNALING AND FINE-TUNING The quantity and quality of BCR signal output is modulated by several transmembrane proteins, which can be either co-stimulatory, like the co-receptor subunit CD19 (see above) or inhibitory, like CD22 and CD72. Several lines of evidence suggest that CD22 and CD72 participate in a negative regulatory feedback loop through coupling to the SH2 domain-containing protein tyrosine phosphatase 1 (SHP-1).
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The mechanism has been worked out in great detail using biochemical and genetic approaches (Vivier and Daëron, 1997). After being phosphorylated by activated Lyn, the immunoreceptor tyrosine-based inhibitory motifs (ITIM) in the cytoplasmic tails of CD22 and CD72 provide binding sites for SHP-1, which subsequently becomes activated and localized to the plasma membrane. By dephosphorylating BCR effector proteins, such as SLP-65, SHP-1 then contributes to signal termination (Mizuno et al., 2000). It was recently found that this control mechanism is employed in an isotype-specific manner. A first clue came from the observation that engagement of the IgG-BCR triggers a more robust signaling than the IgM-BCR in vitro and in vivo. This phenomenon was investigated further in K46 B lymphoma transfectants and found to be dependent on the long cytoplasmic tail of the gm heavy chain (Wakabayashi et al., 2002), which is conserved among the IgG subtypes but absent in IgM or IgD. The presence of the g2am cytoplasmic tail prevented the ITIM phosphorylation of CD22 and the subsequent recruitment of SHP-1. By contrast, signal inhibition by CD72 was similar for all BCR classes tested. Expression of a chimeric IgM-BCR containing the g2am cytoplasmic tail recapitulated IgG-specific hyperresponsiveness to antigen stimulation. These results are in agreement with the previous finding that transgenic mice expressing the IgM-g2am chimera show a phenotype similar to IgG-transgenic mice (Martin and Goodnow, 2002). Hence, the cytoplasmic tail of IgG-containing BCR reduces the signaling threshold by protecting from CD22- but not CD72-mediated signal inhibition. Moreover, a direct association of CD22 and CD72 with the IgM-BCR has been reported (Jamin et al., 1997; Peaker and Neuberger, 1993). Collectively, the resulting increased antigen sensitivity of IgG-positive B cells may confer a growth advantage over IgM/IgD-positive cells with the same antigen specificity and may play a role in the preferential selection and activation of switched memory B cells. The mIgD molecule has also developed an isotype-specific signaling mechanism. This is based on the ability of mIgD molecule to be expressed on the cell surface independently of its association with the Iga/Ig-b heterodimer (Venkitaraman et al., 1991; Wienands and Reth, 1991). The transport of mIgD molecules to the cell surface of Ig-a-negative J558L transfectants is due to an exchange of the dm transmembrane region with a glycosyl-phosphatidyl-inositol (GPI) anchor (Wienands and Reth, 1992). In the presence of Ig-a and Ig-b in naïve B cells, only 5% of the surface IgD contains a GPI moiety (Chaturvedi et al., 2002). However, the GPI-linked IgD fraction constitutively localizes to lipid rafts, where it can activate cyclic AMP- and protein kinase A-dependent signaling pathways. Removal of GPI-linked IgD reduced the inducible upregulation of multiple activation markers on the treated B cells and the number of germinal centers upon reinjection of the cells into BALB/c mice. The responses could be restored by
incubation of the cells with db-cAMP to mimic increased cyclic AMP signaling (Chaturvedi et al., 2002). The data suggest that the immunological function of GPI-linked IgD signaling is to optimize germinal center reactions under conditions of limited BCR occupancy.
CONCLUSION Despite substantial progress during the last decade in identifying more and more of the BCR signaling molecules, the understanding of intracellular signaling networks under various immunological conditions remains a challenge. However, it is likely that we will learn more about these pathways by studying pathogens such as the B-lymphotropic Epstein-Barr Virus (EBV), which reorganizes critical effector molecules like SLP-65 to establish a delicate signaling balance between activation and repression and allows a latent persistence of the virus (Engels et al., 2001a).
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12 Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22 LARS NITSCHKE
DOUGLAS T. FEARON
Institute of Virology and Immunobiology, Universität Würzburg, Würzburg, Germany
Sheila Joan Smith Professor of Immunology, University of Cambridge School of Clinical Medicine, Cambridge, United Kingdom
Membrane immunoglobulin (mIg), the antigen receptor on B cells, is a central regulator of B cell fate. Antigen binding to mIg triggers signaling pathways such as Ca2+ mobilization or MAP kinase activation. These signals may induce proliferation, differentiation, or functional inactivation and apoptosis, depending on the cellular context and microenvironment. Accessory transmembrane molecules or co-receptors on the B cell surface, which are constitutively or inducibly associated with mIg, modulate the B cell signaling. Regulation of both the strength and quality of the B cell mIg signal through co-receptors is accomplished by the recruitment of additional intracellular signaling molecules. Important examples of co-receptors are CD19, which enhances B cell signaling, and CD22, which inhibits this signaling. CD19 is associated with CD21 and can couple the recognition of microbial antigens by complement to the activation of B cells via mIg. Also, the inhibitory function of CD22 is controlled by ligand binding. The mechanisms of signal enhancement by CD19 and signal inhibition by CD22 and other inhibitory molecules are discussed in this chapter.
able in the formation of the complex but CD81, which interacts with CD19 through extracellular domains, is required for the optimal expression of CD19 (Tsitsikov et al., 1997). The CD19/CD21/CD81 complex and the role of CD19 as a stimulatory co-receptor of the B cell were discovered when a mechanism was sought that could account for the ability of CD21 that had been co-ligated or cross-linked to mIg (as would occur with antigen bearing C3d) to lower by several orders of magnitude the threshold for mIg-dependent increases in intracellular [Ca2+] (Fearon and Carroll, 2000). The short the 34 amino acid cytoplasmic tail of CD21 was considered unlikely to be able to recruit the necessary intracellular signaling proteins. The solubilization of B cell membrane proteins with various detergents was carried out in an attempt to preserve the association of other membrane proteins with CD21. This was accomplished using digitonin, and the co-immunoprecipitating proteins were identified as CD19, CD81, and Leu-13 (Fearon and Carroll, 2000). The large cytoplasmic domain of CD19, of approximately 230 amino acids, focused attention on this component of the complex as an important signal transducing element, and this view was reinforced when it was found that its coligation to mIg also enhanced the response of B cells with respect to both intracellular [Ca2+] (Carter et al., 1991) and proliferation (Carter and Fearon, 1992). Subsequent studies have identified aspects of biology of the B cell that are dependent on CD19 in vivo, and have begun to unravel the signaling pathways that CD19 recruits to modify the response of the B cell to ligating mIg.
CD19 CD19 is a component of a membrane protein complex on B cells that has the function of enhancing signaling by the antigen receptor, mIg. The other components are the receptor for the C3d fragment of the complement system (CR2 or CD21), TAPA-1 (also designated CD81), and Leu-13. Each component has a unique function: CD19 is the B cellspecific signal transducing element; CD21 mediates the binding of antigens that have activated the complement system to become coated with C3d; and CD81 is a tetraspanin that may promote the association of the complex with integrins and specialized lipid domains (Horvath et al., 1998); the role of Leu-13 is not known. CD21 and Leu-13 are dispens-
Molecular Biology of B Cells
CD19 and Development of the B Cell Transcriptional Regulation of CD19 Expression CD19 is expressed at the pro-B cell stage of B cell development, and its transcription is regulated by Pax-5 (also
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known as BSAP) (Kozmik, Z. et al., 1992), a major determinant of the commitment of the lymphoid progenitor to the B cell lineage (Nutt et al., 2001). A high-affinity Pax-5 binding site is located in the promoter region upstream of a cluster of transcription start sites. This binding site is occupied by Pax-5 in a CD19-expressing B-cell line but not in plasma or HeLa cells that do not express CD19. With respect to the plasma cell, it has been shown that Blimp-1, a transcription factor with an essential role in the terminal differentiation of the B cell, suppresses Pax-5 expression (Lin et al., 2002). Two additional sites in the promoter region, the PyG box that binds unknown nuclear proteins and the GC box that binds SP1 and Egr-1, have also been mapped (Riva et al., 1997). Mutation of the PyG box markedly reduces the activity of a CD19 reporter construct in B cells, whereas mutation of the GC box has less effect. B1 Cells The creation by two groups (Rickert et al., 1995; Engel et al., 1995) of mice in which the CD19 gene has been interrupted has provided the essential tool for the analysis of the role of CD19 in the development of all three types of B cells—B1, B2, and marginal zone B cells. CD19-/- mice have a deficiency in peritoneal B1 cells (Rickert et al., 1995; Engel et al., 1995; Sato et al., 1996), indicating an important function for CD19 in the development and/or maintenance of these cells. A role for CD19 in the maintenance of B1 cells also was demonstrated when downregulation of CD19 in adult mice, through the chronic administration of monoclonal anti-CD19, suppressed their incorporation of BrdU, gradually leading to a deficiency in these but not B2 cells (Krop et al., 1996). Variable levels of overexpression of CD19 in different founder lines of mice expressing transgenic CD19 demonstrated a direct correlation between the level of CD19 on B1 cells and their prevalence in the peritoneum (Sato et al., 1996). Interestingly, overexpression of CD19 in these mice diminished the development of B2 cells, thus suggesting that B1 and B2 cells fundamentally differ in the means by which they undergo positive and negative selection. For example, the ability of CD19 to promote signaling by mIg may enhance the development of B1 cells because they are positively selected by certain selfantigens (Hayakawa et al., 1999). However, this activity of CD19 might lower the threshold for deletion of B2 cells by self-antigens (Zhou et al., 1994). The absence of either of two other components of the complex, CD21 or CD81 (Horvath et al., 1998), also is associated with diminished numbers of B1 cells. Curiously, IgA-secreting B1 cells in Peyer’s patches are not dependent on CD19 for their development (Gardby and Lycke, 2000), perhaps reflecting the effects of other co-stimulatory receptors that might be stimulated directly or indirectly by microbial antigens at this site.
B2 Cells Although no major abnormalities were noted in B2 cell development in the bone marrow in initial studies using CD19-/- mice, when development was studied in a model system using mice expressing the 3–83 Tg Ig reactive with the mouse class I MHC antigens Kk and Kb, CD19 was found to promote the positive selection of B2 cells (Somani et al., 2001). Immature 3–83 Tg CD19-/- B cells were developmentally arrested in the bone marrow and matured only when the compromised receptor was compensated for by elevated levels of expression. The developmentally arrested 3–83 Tg CD19-/- B cells failed to impose L chain allelic exclusion, and they continued V(D)J recombination to edit their Ig. The immature 3–83 Tg CD19-/- B cells also failed to select positively and to survive when adoptively transferred into normal recipients. Elevation of mIg expression levels by transgene homozygosity restored mIg-mediated increase in intracellular [Ca2+], allelic exclusion, and positive selection. These in vivo studies extend an earlier report that CD19 co-stimulated signaling by the pre-B cell receptor in vitro (Krop et al., 1996), and provide an explanation for the finding that overexpression CD19 in B2 cells is associated with decreased levels of mIgM (Engel et al., 1995; Zhou et al., 1994; Sato et al., 1997), as this may have been the means by which these cells had survived negative selection during development. Marginal Zone B Cells Like B1 cells, marginal zone B cells require CD19 for their development (Martin and Kearney, 2000). However, despite also resembling B1 cells in being a source of “natural” IgM, the development of marginal zone B cells is not impaired in mice lacking CD21 (Cariappa et al., 2001), although CD81 is required (Tsitsikov et al., 1997), undoubtedly because of its effect on the expression of CD19. This finding implies either that a means is available for ligating CD19 that is independent of complement during marginal zone B cell development, or that ligation is not necessary to recruit CD19 function, an issue that will be discussed later in this chapter. In summary, CD19 promotes the development of all three types of B cells and presumably affects the Ig repertoire of these B cell sets by lowering thresholds for antigen receptor signaling to influence both positive and negative selection.
CD19 and the B Cell Response to Antigen Thymus-Independent Antigens Three reports investigating the role of CD19 in type 2 thymus-independent responses (TI-2) found no impairment in mice lacking the co-receptor (Rickert et al., 1995; Sato et al., 1995; Fehr et al., 1998), and two of these actually
12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22
found heightened responses. However, a fourth study evaluated this function of CD19 in transgenic mice expressing high or low affinity antibody specific for the hapten (4hydroxy-3-nitrophenyl)acetyl (NP) and found that CD19 deficiency did influence a TI-2 response to NP-Ficoll by increasing the affinity threshold for the response (Shih et al., 2002). Thus, a positive role for CD19 in TI-2 responses was revealed when the analysis was performed against the background of homogeneous mIg, presumably by eliminating the compensating effects of a diverse, polyclonal Ig repertoire. This function of CD19 may be related to its ability to promote the proliferative response of B cells in vitro to antiIgM antibodies (Engel et al., 1995) and is consistent with its role in the development of B1, B2, and marginal zone B cells, all of which involve signaling by mIg. Furthermore, the impaired response to trinitrophenyl-lipopolysaccharide (TNP)-LPS, a TI-1 antigen, in CD19-/- mice may have a similar basis (Engel et al., 1995). However, it is difficult to understand why CD19-deficient B cells respond less well to the mitogenic effects of LPS in vitro than do wildtype B cells, because it does not involve mIg (Engel et al., 1995). Perhaps CD19 also augments signaling from TLR4 or other innate immune receptors that respond to LPS. Thymus-Dependent Antigens The most striking and consistent effect of a CD19 deficiency in the mouse is the impairment of the response to thymus-dependent (TD) antigens. The three usual outcomes of the primary immunization of mice with protein antigens—germinal center formation, persistently elevated titers of high affinity antibody, and memory B cell development— are all absent in CD19-/- mice (Engel et al., 1995; Rickert et al., 1995; Sato et al., 1996). The defective response to protein antigens was anticipated by the finding in CD19-/mice with constitutively low serum levels of IgM and presumably is caused by the diminished numbers of B1 cells and marginal zone B cells, and of the IgG1, IgG2a, and IgG2b isotypes that may result from responses to environmental antigens (Engel et al., 1995). It had been concluded that the essential abnormality in the CD19-deficient B cell is an inability to differentiate into a germinal center B cell, since it is the developmental precursor of memory B cells and long-lived plasma cells. However, an additional observation requires a modification of this view; CD19-/- mice infected with vesicular stomatitis virus form germinal centers, but do not develop B cell memory or maintain high titers of high affinity antibody thus indicating a probable absence of long-lived plasma cells (Fehr et al., 1998). Also, germinal centers have been observed in Peyer’s patches CD19-/- mice, although these mice fail to produce specific antibody following oral immunization (Gardby and Lycke, 2000). Therefore, even though the requirement for CD19 for the development of germinal center B cells can be circum-
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vented by antigens that are present in abundance and associated with vigorous T cell help, CD19 remains necessary for the further differentiation of the germinal center B cell to a memory B cell or a long-lived plasma cell.
Signal Transduction by CD19 CD19 Ligands The co-receptor function of CD19 was discovered in relation to CD21 (Fearon and Carroll, 2000) and, because CD21 has its own ligand, C3d, a question has arisen concerning whether there is a need for a separate ligand that interacts directly with CD19. However, the more severe immunodeficient phenotype of CD19-/- mice, as compared to CD21-/(Fearon and Carroll, 2000) or C3-/- mice (Hasegawa et al., 2001), has strongly suggested either that there is a CD19 ligand or that ligation of the CD19/CD21 complex is not necessary for enhancing mIg signaling. There are three issues with respect to CD19 ligands: whether they are necessary to elicit the co-receptor function of CD19, whether ligands other than C3d of the complement system can recruit the function of the CD19/CD21 complex, and whether the complex must be crosslinked to the antigen receptor for the amplification of mIg signaling. CD19 can enhance mIgM signaling even when it is not ligated. This has been shown not only by the enhanced proliferation when mIgM is crosslinked alone in normal versus CD19-/- B cells (Engel et al., 1995; Buhl et al., 1997), but also when signaling has been assessed by more biochemical techniques, such as the intensity of tyrosine phosphorylation of CD19 (O’Rourke et al., 1998), the threshold at which mIg induces increased intracellular [Ca2+] response (Carter et al., 1991), and the activation of downstream enzymes, such as MAP kinases (Li and Carter, 2000; Li et al., 1997; Weng et al., 1994; Li and Carter, 1998; O’Rourke et al., 1998). A molecular basis for the recruitment of CD19 function by mIgM without intentional co-ligation has been suggested to be the weak, constitutive association of CD19 with the antigen receptor that enables crosslinking of the latter to induce tyrosine phosphorylation in the former (Carter et al., 1997). It is not known whether CD19 associates directly with a component of the mIg complex, or indirectly, perhaps by promoting the partitioning of CD19 into lipid rafts (Phee et al., 2001; Cherukuri et al., 2001a,b). If such a complex does exist, one can predict at least two consequences of CD19 being a close neighbor of mIgM: promoting the participation of CD19 in mIgM signaling when no ligand is present for the CD19/CD21 complex, and facilitating the crosslinking of the CD19/CD21 complex to mIgM when a CD19 ligand is physically associated with antigen. Despite the ability of CD19 to modestly augment mIg signaling in vitro without ligation, ligands were sought for the co-receptor because cross-linking CD19 to mIg with
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monoclonal antibodies enhanced B cell activation by up to 1000-fold, as measured by proliferation or by biochemical assessment (Carter et al., 1991; Carter and Fearon, 1992). Recombinant fusion proteins containing either the entire extracellular domain of CD19 or its membrane proximal Iglike domain bound specifically to IgM, but not to any other antibody isotype, and to heparin and heparan sulfate, but not to other glycosaminoglycans (de Fougerolles et al., 2001). It was proposed that the localization of antigen–antibody complexes containing IgM to follicular dendritic cells, and the expression by these cells of proteoglycans containing heparan sulfate would enable CD19 and mIg on B cells within the germinal center to be co-ligated. Consistent with this possibility was the binding of the fusion proteins to follicular dendritic cells in germinal centers (de Fougerolles et al., 2001), but whether CD19 on the surface of a B cell can interact with these potential ligands and costimulate signaling by the antigen receptor has not been determined. Several considerations support the proposal that the CD19/CD21 complex amplifies signaling by mIg most effectively when its is co-ligated, that is, crosslinked, to the antigen receptor. First, in one early description of the co-receptor function of CD19, the 100-fold enhancement of mIg-induced B cell proliferation occurred only when antiCD19 and anti-IgM bound to the B cells could be co-ligated by Fc receptors expressed on L cells (Carter and Fearon, 1992). In fact, if CD19 was ligated independently of mIgM in this system, proliferation was suppressed. Second, the tyrosine phosphorylation of CD19 is augmented over 10fold when it is crosslinked to mIgM relative to the phosphorylation that is observed when either of the two receptors is individually ligated (O’Rourke et al., 1998). This likely reflects the increased efficiency with which the tyrosine kinases that are activated by mIg can phosphorylate nearby CD19. Since the contribution of CD19 to signal transduction is dependent, at least in part if not primarily, on its tyrosine phosphorylation, it is more reasonable to consider co-ligation as the physiological means for eliciting CD19 function. Third, the only proven means for ligating CD19 in vivo, through C3d interacting with CD21, necessarily offers the opportunity for co-ligation to mIg because C3d will be covalently attached to antigen. For these reasons, the coligation of mIg and CD19 is considered an important mechanism by which the full potential of CD19 co-stimulatory activity is realized. The Biochemistry of Signal Transduction by CD19 The cytoplasmic domain of CD19 contains nine tyrosines (Figure 12.1), at least some of which are phosphorylated following the ligation of mIg. This phosphorylation is augmented by co-ligating CD19 to mIg, suggesting either that the tyrosine kinases that mediate this phosphorylation are
associated with the antigen receptor complex, or that they are associated with CD19 but are activated by the antigen receptor complex. Studies seeking the molecular explanation for the ability of CD19 to augment the signaling of mIg have focused on the phosphorylation of these tyrosines because early findings indicated that two of these, Y482 and Y513, become phosphorylated and bind phosphatidylinositol 3-kinase (PI 3-kinase) in a manner analogous to growth factor receptors (Tuveson et al., 1993). Three general issues must be resolved in the analysis of this process: the identity of the tyrosine kinase(s) responsible for the phosphorylation of CD19, the downstream effectors with which specific phosphotyrosines interact, and the relevance of these interactions to the in vivo functions of CD19. The tyrosine phosphorylation of CD19 can be induced by ligating mIg or CD19, but is greatest when the two receptors are cross-linked. This may indicate either that tyrosine kinases associated with either receptor complex can phosphorylate CD19, but that those activated by mIg are more effective and can act on CD19 only when it is juxtaposed to the mIg complex. One of the kinases activated by the mIg complex and reported to associate with CD19 is the src kinase, Lyn, which has been suggested to phosphorylate CD19. First, Lyn kinase activity was decreased in CD19-/B cells, and in vitro kinase assays using purified CD19 and purified Lyn revealed that the kinase activity of Lyn increased when it was co-incubated with CD19 (Fujimoto et al., 1999). Second, Lyn expression was reported to be required for CD19 tyrosine phosphorylation in primary B cells (Fujimoto et al., 2000; Somani et al., 2001). Tyrosine513 of CD19 was the first site of Lyn kinase activity. After this activity, it bound Lyn, which then phosphorylated Y482, which bound a second Lyn molecule, causing transphosphorylation and amplification of Lyn activation. Since Lyn was found in other studies to suppress B cell activation, this view of a Lyn-dependent positive feedback loop of CD19 phosphorylation was refined by the suggestion that CD19 amplified B cell activation by sequestering Lyn (Fujimoto et al., 2001). However, a more recent study using B cells from Lyn-/- mice found that CD19 phosphorylation following mIg ligation was not diminished by the absence of Lyn, but did require other Src-family kinases (Xu et al., 2003). Moreover, the ability of CD19 to recruit PI 3-kinase and to enhance intracellular [Ca2+] responses and MAP kinase activation after co-ligation with mIg was Lyn-independent. Conversely, the increase in Lyn activity following mIg ligation, and the inhibition of mIg signaling by CD22 and FcgRII were normal in CD19-/- B cells. This study concluded that the unique functions of Lyn and CD19 are independent, and that other Src kinases were involved in the tyrosine phosphorylation of CD19, which is in accord with the finding that CD19 deficiency suppressed the hyper-responsive state of Lyn-/- B cells (Hasegawa et al., 2001).
12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22
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FIGURE 12.1 Intracellular signaling by CD19. The tyrosines of the cytoplasmic domain of murine CD19 and the intracellular signaling proteins that these have been reported to interact with after phosphorylation. In vivo studies of mutant forms of CD19, in which specific tyrosines have been replaced with phenylalanines, have validated roles only for Y482 and Y513 in mediating the functions of CD19 for B cell development and responses to antigens (Wang, et al., 2002). See color insert.
The downstream effectors mediating CD19 signaling have been studied mainly by in vitro techniques, with some being identified through the analysis of cytosolic proteins that bind to particular phosphotyrosines, and others by analysis of known signaling cascades. The co-ligation of CD19 to mIg has been shown to positively affect many intracellular signaling pathways, including the generation of inositol 3,4,5 trisphosphate (Carter et al., 1991), presumably reflecting the increased activation of PLCg2; the activation of Btk (Buhl et al., 1997; Buhl and Cambier, 1999; Fujimoto et al., 2002; Li and Carter, 2000), which may be involved in PLCg2 activation; the activation of a phosphatidylinositol 4phosphate 5-kinase for the synthesis of phosphatidylinositol 4,5-bisphosphate (O’Rourke et al., 1998) to provide substrate both for PLCg2 and PI 3-kinase; the activation of PI 3-kinase (Buhl et al., 1997; Buhl and Cambier, 1999; Tuveson et al., 1993; O’Rourke et al., 1998); the stimulation of three MAP kinases, ERK, JNK, and p38 (Li and Carter, 2000; Li and Carter, 1998; Brooks et al., 2000; Tooze et al., 1997); the activation of STAT1 (Su et al., 1999); the tyrosine phosphorylation of a complex containing Shc (Lankester et al., 1994); and the activation of Akt (Otero et al., 2001). To mediate this diversity of effects, CD19 is thought to be coupled to multiple signaling pathways by tyrosines in its cytoplasmic domain which, following their phosphorylation, interact with specific signaling proteins. In this sense, CD19 serves as an adaptor protein whose
function can be modulated by extracellular ligands. As shown in Figure 12.1, seven proteins have been found to bind to distinct phosphotyrosines of CD19, either by examining the effects of substituting phenyalanine for specific tyrosines in the cytoplasmic domain or by determining which proteins bound to synthetic phosphopeptides. These are: • Grb2 and Sos to Y330 (Brooks et al., 2000), • Vav1 and possibly Vav2 to Y391 (Li et al., 1997; Weng et al., 1994; Sato et al., 1997; Doody et al., 2001; O’Rourke et al., 1998), • PLCg2 to Y391 and 403 (Brooks et al., 2000), • Fyn to Y403 and Y443 (Fujimoto et al., 2000; Chalupny et al., 1995), and • PI 3-kinase and Lyn to Y482 and Y513 (Fujimoto et al., 2000; Tuveson et al., 1993). Many in vitro studies have attempted to determine how these various pathways linked to these proteins may interact, as for example the apparent relationship between the recruitment of PI 3-kinase by phosphorylated Y482 and Y513, and Vav by phosphorylated Y391 and the intracellular [Ca2+] response (Buhl et al., 1997). However, the interpretation of these studies is difficult because we do not know which genes CD19 regulates to promote the development of B1 and marginal zone B cells or B cell responses to TI-2 and TD antigens. In other words, we do not know the ulti-
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mate targets of the signal transduction. Therefore, more informative assays reporting the expression of these genes, such as Bcl-2 (Roberts and Snow, 1999), cannot be designed. Until such assays are developed, the least ambiguous experimental approach is that taken in a recent study that analyzed the role of these various pathways that can be potentially recruited by CD19 in vivo. In this study (Wang et al., 2002), CD19-/- mice were reconstituted with transgenes encoding forms of CD19 that had pairs of cytoplasmic tyrosines substituted with phenylalanines, so that taken together all tyrosines except Y490 had been mutated. When these transgenic/knockout mice were examined for B cell development and responses to TI2 and TD antigens, only those in which the tyrosines that bind PI 3-kinase had been substituted were abnormal and indistinguishable from CD19-/- mice. Thus, PI 3-kinase is a critical pathway downstream from CD19, and the biological relevance of the other cytoplasmic domain interactions that have been found is unclear. Furthermore, despite the crippled in vivo function of the Y482F/Y513F CD19, in vitro analyses of B cells with this form of CD19 showed that its co-ligation to mIg caused activation to ERK that was equivalent to that obtained with wildtype CD19, and that ligation of mIg alone on these cells induced a [Ca2+] response that was the same as in B cells with wildtype CD19. Thus, these in vitro assays do not measure the signaling functions of CD19 that are relevant to its in vivo roles. Only one response, a co-stimulatory intracellular [Ca2+] increase induced by the ligation of both mIg and CD19, was impaired in B cells with the Y482F/Y513F CD19. These mice emphasize the need to develop in vitro assays that accurately reflect in vivo physiology. CD19 and Antigen Processing The more stringent requirement for CD19 in TD than in TI-2 B cell responses may indicate either that the relatively more oligovalent antigens associated with the latter require greater co-stimulation by CD19 than do the former, that CD19 is required for the effective processing of and presentation of antigen to class II restricted CD4 T cells, or possibly both. In support of the latter two possibilities, several studies have shown that CD19 may have a role in promoting effective interaction with T cells. Co-ligating CD21/CD35 to mIg enhanced the expression of B7.1 and B7.2 (CD80 and CD86) on primary B cells (Kozono et al., 1998). These membrane proteins are the counterligands for CD28, the ligation of which is required for the optimal stimulation of T cells by antigen-presenting cells and for reciprocal stimulation of the presenting cell by, for example, the CD40 ligand that is required for a germinal center reaction. The CD19/CD21 complex has also been found to enhance the speed and efficiency with which B cells produce
MHC class II-peptide complexes (Cherukuri, A. et al., 2001), an effect that may be especially important in the germinal center, where competition for limiting amounts of antigen may be intense among B cell variants expressing somatically mutated mIgs. Finally, signaling of the B cell by ligating class II, which would occur uniquely in TD responses, is enhanced by co-ligating CD19 to the class II (2). Taken together, these studies would support an important role for CD19 in promoting the interaction between B and T cells that may take place in the gerninal center reaction. Interactions of CD19 with the Inhibitory Receptors CD22 and FcgRII The co-stimulatory functions of CD19 in promoting mIg signaling can be reversed or blocked by the additional coligation of CD22 or FcgRII to mIg. With respect to FcgRII, this inhibitory effect appears to be mediated by the hydrolysis of phosphatidylinositol 3,4,5-phosphate (PIP3) that is catalyzed by the SHIP that is associated with the Lynphosphorylated FcgRII. This would negate those positive signaling effects of CD19 that are mediated by the binding and activation of PI 3-kinase (and possibly by Vav) because of their roles in the biosynthesis of PIP3. The reported dephosphorylation by FcgRII of CD19 (Hippen et al., 1997) must be indirect since SHIP is not a protein phosphatase. The ability of CD22 to suppress co-stimulation by CD19 (Tooze et al., 1997; Fong et al., 2000) is mediated by the SHP-1 that is recruited by Lyn-phosphorylated CD22, which suppresses the tyrosine phosphorylation of CD19, presumably by suppressing the activation of the tyrosine kinases linked to the mIg complex. The report that SHP-1 suppresses CD19 phosphorylation by inhibiting Lyn (Somani et al., 2001) must be considered in relation to the finding that Lyn is not required for this modification of CD19 (Xu et al., 2002). The functions of these inhibitory receptors are discussed in other sections of this chapter. Summary CD19 is the major stimulatory co-receptor of B cells. It is required for the normal development of subsets of B cells, and for the response of mature B cells to both TI-2 and TD antigens. The phosphorylation of tyrosines in its cytoplasmic domain by kinases activated by mIg enables CD19 potentially to recruit several enzymes to the larger signaling complex being assembled by mIg, but the relationship of these to the biological functions of CD19 in vivo require further definition, perhaps by determining the genetic targets of CD19 co-stimulation. The clinical relevance of understanding the basis of CD19 function is underscored by its possible participation in human autoimmune disease (Sato et al., 2000).
12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22
INHIBITORY CO-RECEPTORS ON B CELLS Inhibitory receptors are important for controlling the equilibrium of high reactivity and quiescence of the B cell. The B cell has a number of such regulatory receptors that seem to act by recruiting negative intracellular proteins, such as phosphatases. These phosphatases counteract the activatory signaling cascades triggered by tyrosine phosphorylation of immunoreceptor tyrosine-based activation motifs (ITAMs) of Iga and Igb. How inhibition is achieved in the different mechanisms are discussed for four mouse B-cell inhibitory co-receptors, with some emphasis on the important inhibitory receptor CD22.
CD22 Inhibits Intracellular Signaling CD22 is a member of the Siglec family, a family of inhibitory adhesion receptors on leukocytes. CD22 is expressed in a B-cell lineage–specific fashion, starting at the pre-B cell stage. By immunoprecipitation, a small percentage of CD22 molecules can be co-precipitated with surface IgM, so that a fraction of CD22 is constitutively associated with the mIg (Leprince et al., 1993; Peaker and Neuberger, 1993). After stimulation of the mIg, CD22 is quickly tyrosine phosphorylated on its cytoplasmic tail (Doody et al., 1995). The tyrosine kinase mainly responsible for CD22 phosphorylation is Lyn, a member of the Src kinase family, as was demonstrated by reduced CD22 phosphorylation in Lyn-deficient mice (Chan et al., 1998; Smith et al., 1998). The cytoplasmic tail of CD22 contains six tyrosines, three of which belong to the consensus of the ITIM (immunoreceptor tyrosine-based inhibiton motif) sequences with the consensus (Ile/Val/Leu/Ser)-x-Tyr-x-x-(Leu/Val). The phosphorylated ITIM motifs of CD22 recruit the tyrosine phosphatase SHP-1 (Doody et al., 1995), an important negative regulator of many signaling pathways in hematopoetic cells. SHP-1 is the most prominent intracellular binding partner of CD22, which binds via its tandem SH2 domains. The inhibitory role of CD22 was clearly demonstrated by analysis of CD22-deficient mice that showed increased Ca2+ mobilization in their B cells after mIg crosslinking (Nitschke et al., 1997; O’Keefe et al., 1996; Otipoby et al., 1996; Sato et al., 1996). However, other proteins, which are normally positively involved in mIg signaling, are also recruited via their SH2 domains to the tyrosine-phosphorylated tail of CD22. These include Syk, PLCg2, PI3K, Grb-2, and Shc (Law et al., 1996; Poe et al., 2000; Yohannan et al., 1999). When analyzing these interactions in detail, it was shown that out of the three tyrosines comprising ITIMs (Y2, Y5, Y6, for the second, fifth, or sixth tyrosine of the CD22 tail) Y5 and Y6 are sufficient to recruit SHP-1 (Blasioli et al., 1999). Another group showed that at least two of the three
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phosphorylated ITIM tyrosines must be present in order to bind SHP-1 in vivo (Otipoby et al., 2001) (Figure 12.2). Grb-2 binds to another phosphorylated tyrosine of CD22 (Y4), distinct from SHP-1 binding (Otipoby et al., 2001; Yohannan et al., 1999). An inhibitor for Src kinases could not inhibit Grb-2 binding to CD22, although it could inhibit SHP-1 binding. This and genetic data suggest that Lyn may not be crucial for tyrosine phosphorylation of Y4, but this may be achieved by Syk (Otipoby et al., 2001). By phosphopeptide mapping, the binding sites for the other factors were shown to be Y6 for PLCg and PI-3 kinase; or Y2, Y5, and Y6 for Syk. Therefore, these three intracellular proteins have overlapping binding sites with SHP-1. To confuse things even more, the CD22 tail can form a quaternary complex with the lipid phosphatase SHIP, Grb-2, and Shc (Poe et al., 2000). SHIP cannot bind directly to phosphopeptides of CD22 but requires both Grb-2 and Shc for binding (Figure 12.2). Thus, the cytoplasmic portion of CD22 acts as a multiple docking site for negative regulators of signaling, such as SHP-1 and SHIP, and for several proteins that are positively involved in B-cell signaling. What are the functions and relative importance of these binding partners? From CD22deficient mice made by four independent groups it is clear that the overall function of CD22 is inhibitory (Nitschke, 1997; O’Keefe et al., 1996; Otipoby et al., 1996; Sato et al., 1996). CD22-deficient B cells showed a strongly increased Ca2+ mobilization after mIg crosslinking. How is this negative regulation of the Ca2+ signaling achieved? Either SHP1 or SHIP could potentially inhibit Ca2+ mobilization. SHIP is the crucial phosphatase for the FcgRII pathway in B cells (see below). However, B cells of SHIP-deficient mice do not show increased Ca2+ mobilization when the mIg is stimulated alone (without co-crosslinking to the FcgRII) (Brauweiler et al., 2000). In contrast, it was demonstrated that moth-eaten mice (which carry a spontaneous mutation of SHP-1) show increased Ca2+ after antigenic stimulation of the mIg (Cyster and Goodnow, 1995). This was confirmed recently by a conditional B-cell specific SHP-1 knock-out mouse. These conditional knock-out mice have B cells with a similar B1-like phenotype in the periphery as moth-eaten mice. The mIg-induced Ca2+ mobilization of these SHP-1-/B1 cells was increased when compared to B1 cells of normal mice (L. Pao, L. Nitschke, K.P. Lam, M.L. Thomas and K. Rajewsky, unpublished). Additionally, higher tyrosine phosphorylation in CD22-deficient B cells of Vav-1 (Sato et al., 1997), CD19 (Fujimoto et al., 1999), and SLP65/BLNK (Gerlach and Nitschke, unpublished) all positively involved in Ca2+ signaling, indicated a decreased tyrosine phosphatase activity. Together, this clearly suggests that SHP-1 is the crucial downstream phosphatase in CD22 signaling. However, SHP-1–deficient mice develop a stronger phenotype than CD22-/- mice, demonstrating that SHP-1 is
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FIGURE 12.2 Intracellular signaling by CD22. All known intracellular binding proteins of CD22 are shown. The tyrosines that have been mapped as interaction sites are indicated. A clear function has only been demonstrated for SHP1 binding. SHP-1 is most likely the phosphatase responsible for the CD22-mediated inhibition of Ca2+ mobilization. See color insert.
involved in several other signaling pathways in B cells and other cells. What about the other signaling proteins binding to the phosphorylated tyrosines of CD22? Does CD22 have a dual role, both as a negative and positive regulator of mIg signaling? If so, then CD22-deficient mice should also show impaired signaling pathways. Overall, B cells of CD22deficient mice show a (mildly) activated phenotype, such as a higher proportion of mature B cells and upregulation of MHC class II or higher responsiveness to LPS (Nitschke, 1997; O’Keefe et al., 1996; Otipoby et al., 1996; Sato et al., 1996). At older age, CD22-/- mice develop high affinity autoantibodies (Mary et al., 2000; O’Keefe et al., 1999). This all confirms the negative role of CD22 in B cell signaling and B cell activation. Two findings do not fit the picture so well: impaired responses of CD22-/- mice to TI2 antigens and impaired proliferation of CD22-/- B cells after anti-IgM stimulation. The impaired response of CD22-/- mice to TI-2 antigens can be explained by the recent finding that their marginal zone B cell numbers are reduced (Samardzic et al., 2002). Marginal zone B cells are a B cell subpopulation that is crucial for TI-2 responses. The impaired proliferation was taken as a hint that CD22, maybe via Grb-2 activation, directly stimulates mitogenic signaling
pathways. Crosslinking of CD22 alone can induce signals such as the stimulation of the JNK pathway (Tuscano et al., 1999). CD22-deficient B cells have no strong impairment of the ERK MAP kinase pathway, while at least one study showed impaired JNK phosphorylation (Poe et al., 2000; Otipoby et al., 2001). So, CD22 could directly induce the JNK pathway. An alternative interpretation of the role CD22 may play in “positive” signaling in B cells comes from experiments in which CD22 was crosslinked by anti-CD22 beads on the surface. This separate ligation of CD22, or sequestration of CD22 from the mIg, led to higher proliferation or MAP kinase activation when the B cells were stimulated with anti-IgM. In contrast, when CD22 was co-ligated to the mIg, MAP kinase activation was inhibited (Tooze et al., 1997). There seem to be different “compartments” of CD22 on the B-cell surface. Proximity to the mIg apparently gives the strongest inhibition, whereas separation from the mIg releases the surface Ig from this inhibition. Ligand binding may control this membrane localization of CD22 (see below). Crosslinking of CD22 by antibodies may not trigger signals directly, but removal of CD22 thereby releases the mIg from constitutive inhibition.
12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22
The Extracellular Domain of CD22 Controls Signaling The Siglec CD22 has a high specificity for Neu5Aca26Galb1-4Glc(NAc) or a2–6 linked sialic acid (2,6Sia) (Kelm et al., 1994; Powell et al., 1995), a common structure on N-linked glycans. This structure is abundantly expressed on the surface of lymphocytes or other cells, such as cytokine-activated endothelial cells, but is also present on soluble plasma proteins such as haptoglobin or IgM (Engel et al., 1993; Hanasaki et al., 1995a,b). The extracellular portion of CD22 consists of seven Ig-like domains. The first N-terminal V-set domain binds the ligand sialic acid (van der Merwe et al., 1996). This was demonstrated by the X-ray crystallographic structure of the V-set domain of the CD22-homolog sialoadhesin (Siglec-1) with the ligand a2,3 sialyllactose (May et al., 1998). By solving this structure, it became clear that most molecular contacts occur with the sialic acid, rather than with the attached sugar units. Particularly, one Arg residue (Arg130 in murine CD22) and two aromatic amino acids are involved in molecular interactions to sialic acid. Molecular modeling and site-directed mutagenesis predict a similar sugar binding site for CD22 as for sialoadhesin (van der Merwe et al., 1996). The affinity of CD22 for free sialic acid is very low (10-4 M) (Bakker et al., 2002). CD22 can bind to a number of sialylated proteins on the cellular surface, among them prominently CD45, as was demonstrated by CD22-Fc binding and protein precipitation (Sgroi and Stamenkovic, 1994). However, a recent plasmon resonance study showed that the CD22 extracellular portion displayed a similar affinity for native CD45 as it did for a synthetic 2,6Sia-carrying glycoconjugate (Bakker et al., 2002). Thus, the protein backbone of the glycan does not contribute to ligand binding of CD22, and it is only the presence and density of 2,6Sia that determines binding. How does the ligand-binding of CD22 control its inhibitory signaling function in the B cell? This question has puzzled many researchers interested in CD22 function. It is now evident that CD22, like most other Siglecs, is bound to ligands in cis; that is, to ligands on the same cellular surface, on the majority of B cells (Floyd et al., 2000; Razi and Varki, 1998). This is concluded from experiments in which B cells were stained with a polyacrylamide-based 2,6Sia carrying glycoconjugate as synthetic ligand for CD22. This synthetic ligand could not bind to most B cells unless they were pretreated with sialidase to remove the cis ligands. However, small subpopulations of B-cells can bind the probe, hence these carry “unmasked” CD22 (Collins et al., 2002; Floyd et al., 2000). The expression of 2,6Sia on the cell surface is controlled by an a2,6sialyltransferase and by sialidases. These enzymes can be regulated, for example, by cytokines (Braesch-Andersen and Stamenkovic, 1994; Hanasaki et al., 1995b). For CD22-mediated cell–cell interactions, 2,6Sia-
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carrying ligands in trans would have to compete with ligands in cis. Nevertheless, a cell–cell adhesion function for CD22 has been demonstrated in the bone marrow. Bone marrow sinusoidal endothelium is unique in expressing 2,6Sia constitutively on the surface (Nitschke et al., 1999). This ligand expression has been directly implicated in the bone marrow homing of recirculating B cells, which are strongly reduced in CD22-/- mice. This homing may be possible because a population of B cells with “unmasked” CD22 is enriched in the bone marrow (Floyd et al., 2000). Recently, two sets of experiments have demonstrated that cis-interactions of CD22 also control signaling. In one approach, a CD22 protein with a mutated 2,6Sia-binding domain was expressed in a B cell line (Jin et al., 2002); in another approach, a CD22-specific sialic acid analog that inhibits ligand binding with high affinity was used (Kelm et al., 2002). In both cases, CD22 was less tyrosine phosphorylated, recruited less SHP-1 protein, and the Ca2+ mobilisation was increased after mIg stimulation. Thus, ligand-binding in cis stimulates CD22 tyrosine phosphorylation and signal inhibition. What are the crucial ligands for CD22 on the B cell surface? The recent results question the model that the main function of ligand binding is sequestration of CD22 from the mIg, because destroying ligand interactions would then result in increased CD22 tyr phosphorylation. Instead, it must be assumed that those transmembrane glycoproteins positively involved in signaling and activating Lyn are the crucial CD22 ligands. Candidates are the mIg itself or CD45, which dephosphorylates and activates Lyn (Figure 12.3). Indeed, initial results indicate that CD22 makes a 2,6Sia-dependent interaction with IgM (J. Gerlach, S. Ghash, and L. Nitschke, unpublished), but there may also be other ligands involved. A recent report demonstrated that ligand interactions of CD22 in trans can also control the signaling strength of the B cell. This study showed that B cell activation by antigen displayed on the surface of a target cell was depressed, if the target cell co-expressed 2,6Sia (Lanoue et al., 2002). An interpretation of this is that by ligand-binding on the target cell, more CD22 (and SHP-1) is moved into the cell–cell contact site where the mIg is clustered. As suggested by Lanoue et al., the CD22/2,6Sia trans interaction could be physiologically relevant to dampen the B-cell response to self-antigens displayed on the neighbor cell (Lanoue et al., 2002) (Figure 12.3). This could be important in certain microenvironments where there is close contact between B cells and other cells. One such site maybe the densely packed primary follicle, with a possible contact of B cells to themselves. The concept that CD22 interaction with sialic acid on other cells could suppress B cell reactivity and dampen autoimmunity is also supported by the fact that microorganisms do usually not express sialic acid on their surfaces (Crocker and Varki, 2001).
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FIGURE 12.3 Model for regulation of CD22 inhibition by ligand binding. (a) Ligand binding in cis increases tyrosine phosphorylation and SHP-1 recruitment to CD22. First evidence indicates that CD22 binds directly to 2,6Sia on IgM, but other ligands may also be involved. (b) A pool of CD22 exists on the cellular surface, which is not bound to endogenous ligands, but most CD22 is “masked” by ligands in cis. (c) When the mIg binds to self-antigens on other cells, additional CD22 molecules may be recruited by trans interactions into the cellular contact zone, thus resulting in a stronger CD22 inhibition of the mIg signal (indicated by “flash arrow”). In contrast, microorganisms usually do not display sialic acid on the surface, thus resulting potentially in a higher mIg response. See color insert.
In summary, increasing evidence suggests that the crucial regulation of CD22 inhibition is not through intracellular events but through ligand binding. The availability of the CD22 ligand 2,6Sia on the B cell surface leads to prominent CD22 binding in cis and supports tyrosine phosphorylation of CD22 ITIMs. CD22 binding to ligands on other cells in trans may help to suppress B-cell autoimmunity. CD72 CD72 is a type II transmembrane protein of the C-type lectin family. CD72 is expressed on B cells, but also on DCs, macrophages, and subpopulations of T cells (Kumanogoh and Kikutani, 2001). The cytoplasmic domain of CD72 contains two ITIMs. It has been shown that cross-linking of the mIg induces the phosphorylation of tyrosines on CD72 and its association with SHP-1 (Adachi et al., 1998). However, direct association of CD72 with the mIg has not been demonstrated so far. Establishment of CD72-deficient mice showed an inhibitory role for CD72. However, compared to CD22-/- mice, the increase of the Ca2+ response in CD72-/- B cells was very mild. At low concentrations of anti-
IgM antibodies, CD72-/- B cells showed a hyperproliferative response (Pan et al., 1999). In normal mice, anti-CD72 antibody treatments can activate some signaling pathways, such as tyrosine phophorylation of PLCg and CD19 and activation of Lyn, Blk, and Btk kinases (Wu et al., 2001). However, many of the effects by anti-CD72 antibodies are relatively weak. Thus, an additional positive role of CD72 in B cell signaling was proposed, similar to that of CD22. Recently, a ligand for CD72 was identified to help explain these findings. This ligand is CD100, a transmembrane protein that belongs to the semaphorin family (Kumanogoh et al., 2000). The semaphorin family is primarily expressed in the nervous system. However, new functions in the immune system are emerging. CD100 is expressed abundantly on resting T cells, but upregulated after cellular activation. It is also expressed weakly on B cells and DCs. CD100 can bind to two receptors, to plexin-B1 with high affinity (an interaction with unknown physiological significance) and to CD72 with lower affinity. Interestingly, when expressed transiently in COS7 cells, CD72 is constitutively tyrosine-phosphorylated and associated with SHP-1. CD100 can induce the dephosphorylation
12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22
of CD72 and dissociation of SHP-1 in these cells (Kumanogoh et al., 2000) (Figure 12.4). These findings suggest that CD100 turns off the negative signaling effects of CD72 and thereby enhances B cell responses. This was supported by the phenotype of the CD100-deficient mouse line, which was almost the opposite of CD72deficient mice (Shi et al., 2000). CD100-deficient mice displayed several immunological defects, including hyporesponsiveness of B cells. Thus the CD100–CD72 interaction seems to be a rare example, demonstrating that the binding of a ligand to an inhibitory receptor can create positive signals in B cells. PIR-B The paired immunoglobulin-like receptors (PIR) were cloned in an attempt to identify the mouse homolog to FcaR (Kubagawa et al., 1997). Yet, when expressed ectopically on cells they turned out to bind neither to IgA nor to other immunoglobulins. Instead, the PIR proteins comprise a new gene family having unidentified ligands (Takai and Ono, 2001). PIR-B is a 120- to 130-kD type-I transmembrane protein with six Ig-like extracellular domains. It contains three ITIM motifs in its intracellular tail (Blery et al., 1998).
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In contrast, the PIR-A proteins are a subfamily with several members and are characterized by a charged amino-acid (Arg) within the transmembrane region and only a short cytoplasmic sequence. Similar to FcgRs, which also carry a charged amino-acid in their transmembrane domain, PIR-A requires the gamma chain (FcRg) for expression on the surface. The activating role of PIR-A is thought to result from the immunoreceptor tyrosine-based activating motif (ITAM) of the FcRg chain. PIR-A and PIR-B exhibit pairwise expression, as implicated by their name, on B-lineage and myeloid cells, such as macrophages, mast cells, and dendritic cells (Takai and Ono, 2001). PIR-B molecules in B cells and macrophages are constitutively phosphorylated (Ho et al., 1999). The tyrosine kinase responsible seems to be Lyn and, as for CD22 and CD72, SHP-1 is the main phosphatase bound to the PIR-B ITIMs in vivo. When PIR-B is crosslinked to the mIg in B cells or to the FceRI receptor on mast cells, it can inhibit the Ca2+ response by the triggered activating receptor (Blery et al., 1998; Yamashita et al., 1998). However, similar to CD72, PIR-B seems not to be directly associated to the mIg. Nevertheless, ligation of PIRB on the chicken B cell line DT40 inhibits the mIg-induced tyrosine phosphorylation of Iga, Igb, Syk, Btk, and PLCg2 (Maeda et al., 1998) (Figure 12.4).
FIGURE 12.4 Additional inhibitory receptors on the B cell. CD72 is constitutively associated with SHP-1 bound to its tyrosine-phosphorylated ITIM motifs. CD100 binding reduces the tyrosine phosphorylation of CD72. PIR-B constitutively binds SHP-1. The ligands are not known yet. The FcgRII receptor is recruited via immune complexes to IgM. In this case, SHIP binds and inhibits sustained Ca2+ signaling by catalyzing dephosphorylation of phosphatidyl inositol (3,4,5) triphosphate (PtI-(3,4,5) P3) into PtI-(3,4)P2. Other functions of FcgRII are described in the text. See color insert.
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Since the ligands for PIR-A and PIR-B have not been identified yet, the physiological role of the proteins is still unknown. However, the closest homologs for PIRs in the human are members of the Ig-like transcript (ILT)/leukocyte Ig-like receptor (LIR)/myeloid Ig-like recptor (MIR) family, which exhibit 50 to 60% sequence similarity. The human ILT/LIR/MIR receptors show expression profiles similar to the murine PIRs, and some of their members have been shown to bind classical or nonclassical MHC class I alleles (Takai and Ono, 2001). Whether the PIRs can bind to similar self ligands remains to be demonstrated. Recently established PIR-B–deficient mice showed an increased B cell proliferation upon anti-IgM stimulation. Similarily to CD22-deficient or CD72-deficient animals, B-cell development was not grossly disturbed, with the exception of increased numbers of B1 cells in older animals (Ujike et al., 2002). Impaired maturation of DCs, as well as enhanced TH2 responses also indicated a physiological role for PIRB in other cell types. FcgRIIB Receptors for IgG are good examples of the coordinated and opposing roles displayed by activating and inhibitory receptors. IgG immune complexes were recognised as potent inhibitory ligands for B cells a long time ago. The two low affinity IgG receptors, FcgRIIB and FcgRIII, have very similar extracellular IgG binding domains, but differ in their intracellular domains. Although the FcgRIII gives an activatory signal via its associated g chain (FcRg) containing an ITAM sequence, the FcgRIIB has a cytoplasmic domain carrying an ITIM and acts as an inhibitory receptor (Ravetch and Lanier, 2000). Although the two IgG receptors are co-expressed on several cell types, thus suggesting that the ratio of expression may control the balance of activation or inhibition, B cells only express FcgRIIB. IgG immune complexes can co-ligate the FcgRIIB to mIg. This coligation leads to inhibition of mIg-induced Ca2+ and proliferation (Muta et al., 1994). The FcgRIIB ITIM motif is required and sufficient for this inhibition. The phosphorylated ITIM is the binding site for SHIP (Ono et al., 1996). From several studies using genetically modified mice and cell lines, it is evident that the FcgRIIB does not constitutively inhibit mIg signaling, but requires co-ligation by immune complexes (Brauweiler et al., 2000; Liu et al., 1998; Ono et al., 1997). The inhibition of Ca2+ signals by SHIP is caused by the phosphorylysis of PtI(3,4,5)P3, resulting in the dissociation of PH domain–containing proteins like Btk and PLCg2 (Figure 12.4). The inhibition of cellular proliferation by FcgRIIB seems to involve the activation of the adaptor protein Dok and subsequent inactivation of MAP kinases (Ravetch and Lanier, 2000). SHIP is required in this process, but the exact mechanism is not known.
When ligated separately from the mIg, the FcgRIIB can induce an apoptotic response. This signal is not only independent of SHIP but is increased when SHIP or the binding site for SHIP is deleted (Pearse et al., 1999). Thus, FcgRIIB seems to have a dual role depending on whether it is coligated to the mIg. These two types of signals may be crucial in germinal center B cell selection when FDCs display immune complexes either to cognate or noncognate B cells.
CONCLUSION In summary, B cells constitutively express a set of inhibitory receptors that are regulated by surprisingly different mechanisms. Generally, inhibition relies on the presence of ITIM motifs and on the recruitment of either SHP-1 or SHIP. Some receptors are constitutively tyrosine phosphorylated (CD72, PIR-B) and in others tyrosine phosphorylation is induced (CD22, FcgRII). The crucial regulation seems to be achieved by ligand binding, which can switch on inhibition (FcgRII), enhance inhibition (CD22), or even turn off inhibition (CD72). These different strategies enable the B cell to tightly control the strength of the mIg signal in response to the microenvironment.
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phosphoinositide hydrolysis and Ca2+ mobilization is integrated by CD19 dephosphorylation. Immunity 7, 49–58. Ho, L. H., Uehara, T., Chen, C. C., Kubagawa, H., and Cooper, M. D. (1999). Constitutive tyrosine phosphorylation of the inhibitory paired Ig-like receptor PIR-B. Proc Natl Acad Sci U S A 96, 15086–15090. Horvath, G., Serru, V., Clay, D., Billard, M., Boucheix, C., and Rubinstein, E. (1998). CD19 is linked to the integrin-associated tetraspans CD9, CD81, and CD82. J Biol Chem 273, 30537–30543. Jin, L., McLean, P. A., Neel, B. G., and Wortis, H. H. (2002). Sialic acid binding domains of CD22 are required for negative regulation of B cell receptor signaling. J Exp Med 195, 1199–1205. Kelm, S., Pelz, A., Schauer, R., Filbin, M. T., Tang, S., de Bellard, M. E., Schnaar, R. L., Mahoney, J. A., Hartnell, A., Bradfield, P., et al. (1994). Sialoadhesin, myelin-associated glycoprotein and CD22 define a new family of sialic acid-dependent adhesion molecules of the immunoglobulin superfamily. Curr Biol 4, 965–72. Kelm, S., Gerlach, J., Brossmer, R., Danzer, C. P., and Nitschke, L. (2002). The ligand-binding domain of CD22 is needed for inhibition of the B cell receptor signal, as demonstrated by a novel human CD22-specific inhibitor compound. J Exp Med 195, 1207–1213. Kozmik, Z., Wang, S., Dorfler, P., Adams, B., and Busslinger, M. (1992). The promoter of the CD19 gene is a target for the B-cell-specific transcription factor BSAP. Mol Cell Biol 12, 2662–2672. Kozono, Y., Abe, R., Kozono, H., Kelly, R .G., Azuma, T., and Holers, V. M. (1998). Cross-linking CD21/CD35 or CD19 increases both B7-1 and B7-2 expression on murine splenic B cells. J Immunol 160, 1565–1572. Krop, I., de Fougerolles, A. R., Hardy, R. R., Allison, M., Schlissel, M. S., and Fearon, D. T. (1996). Self-renewal of B-1 lymphocytes is dependent on CD19. Eur J Immunol 26, 238–242. Krop, I., Shaffer, A. L., Fearon, D. T., and Schlissel, M. S. (1996). The signaling activity of murine CD19 is regulated during cell development. J Immunol 157, 48–56. Kubagawa, H., Burrows, P. D., and Cooper, M. D. (1997). A novel pair of immunoglobulin-like receptors expressed by B cells and myeloid cells. Proc Natl Acad Sci U S A 94, 5261–5266. Kumanogoh, A., and Kikutani, H. (2001). The CD100-CD72 interaction: a novel mechanism of immune regulation. Trends Immunol 22, 670–666. Kumanogoh, A., Watanabe, C., Lee, I., Wang, X., Shi, W., Araki, H., Hirata, H., Iwahori, K., Uchida, J., Yasui, T., Matsumoto, M., Yoshida, K., Yakura, H., Pan, C., Parnes, J. R., and Kikutani, H. (2000). Identification of CD72 as a lymphocyte receptor for the class IV semaphorin CD100: A novel mechanism for regulating B cell signaling. Immunity 13, 621–631. Lankester, A. C., van Schijndel, G. M., Rood, P. M., Verhoeven, A. J., and van Lier, R. A. (1994). B cell antigen receptor cross-linking induces tyrosine phosphorylation andmembrane translocation of a multimeric Shc complex that is augmented by CD19co-ligation. Eur J Immunol 24, 2818–2825. Lanoue, A., Batista, F. D., Stewart, M., and Neuberger, M. S. (2002). Interaction of CD22 with alpha2,6-linked sialoglycoconjugates: Innate recognition of self to dampen B cell autoreactivity? Eur J Immunol 32, 348–355. Law, C. L., Sidorenko, S. P., Chandran, K. A., Zhao, Z., Shen, S. H., Fischer, E. H., and Clark, E. A. (1996). CD22 associates with protein tyrosine phosphatase 1C, Syk, and phospholipase C-gamma(1) upon B cell activation. J Exp Med 183, 547–560. Leprince, C., Draves, K. E., Geahlen, R. L., Ledbetter, J. A., and Clark, E. A. (1993). CD22 associates with the human surface IgM-B-cell antigen receptor complex. Proc Natl Acad Sci U S A 90, 3236–3240. Li, X., and Carter, R. H. (2000). CD19 signal transduction in normal human B cells: linkage to downstream pathways requires phosphatidylinositol 3-kinase, protein kinase C and Ca2+. Eur J Immunol 30, 1576–1586. Li, X., Sandoval, D., Freeberg, L., and Carter, R. H. (1997). Role of CD19 tyrosine 391 in synergistic activation of B lymphocytes by coligation of CD19 and membrane Ig. J Immunol 158, 5649–5657.
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12. Regulation of Antigen Receptor Signaling by the Co-Receptors, CD19 and CD22 Pan, C., Baumgarth, N., and Parnes, J. R. (1999). CD72-deficient mice reveal nonredundant roles of CD72 in B cell development and activation. Immunity 11, 495–506. Peaker, C. J., and Neuberger, M. S. (1993). Association of CD22 with the B cell antigen receptor. Eur J Immunol 23, 1358–1363 Pearse, R. N., Kawabe, T., Bolland, S., Guinamard, R., Kurosaki, T., and Ravetch, J. V. (1999). SHIP recruitment attenuates Fc gamma RIIBinduced B cell apoptosis. Immunity 10, 753–760. Phee, H., Rodgers, W., and Coggeshall, K. M. (2001). Visualization of negative signaling in B cells by quantitative confocal microscopy. Mol Cell Biol 21, 8615–8625. Poe, J. C., Fujimoto, M., Jansen, P. J., Miller, A. S., and Tedder, T. F. (2000). CD22 forms a quaternary complex with SHIP, Grb2, and Shc. A pathway for regulation of B lymphocyte antigen receptor-induced calcium flux. J Biol Chem 275, 17420–17427. Powell, L. D., Jain, R. K., Matta, K. L., Sabesan, S., and Varki, A. (1995). Characterization of sialyloligosaccharide binding by recombinant soluble and native cell-associated CD22. Evidence for a minimal structural recognition motif and the potential importance of multisite binding. J Biol Chem 270, 7523–7532. Ravetch, J. V., and Lanier, L. L. (2000). Immune inhibitory receptors. Science 290, 84–89. Razi, N., and Varki, A. (1998). Masking and unmasking of the sialic acidbinding lectin activity of CD22 (Siglec-2) on B lymphocytes. Proc Natl Acad Sci U S A 95, 7469–7474. Rickert, R. C., Rajewsky, K., and Roes, J. (1995). Impairment of T-celldependent B-cell responses and B-1 cell development in CD19deficient mice. Nature 376, 352–355. Riva, A., Wilson, G. L., and Kehrl, J. H. (1997). In vivo footprinting and mutational analysis of the proximal CD19 promoter reveal important roles for an SP1/Egr-1 binding site and a novel site termed the PyG box. J Immunol 159, 1284–1292. Roberts, T., and Snow, E. C. (1999). Cutting edge: Recruitment of the CD19/CD21 coreceptor to B cell antigen receptor is required for antigen-mediated expression of Bcl-2 by resting and cycling hen egg lysozyme transgenic B cells. J Immunol 162, 4377–4380. Samardzic, T., Marinkovic, D., Danzer, C. P., Gerlach, J., Nitschke, L., and Wirth, T. (2002). Reduction of marginal zone B cells in CD22-deficient mice. Eur J Immunol 32, 561–567. Sato, S., Steeber, D. A., and Tedder, T. F. (1995). The CD19 signal transduction molecule is a response regulator of B-lymphocyte differentiation. Proc Natl Acad Sci U S A 92, 11558–11562. Sato, S., Ono, N., Steeber, D. A., Pisetsky, D. S., and Tedder, T. F (1996a). CD19 regulates B lymphocyte signaling thresholds critical for the development of B-1 lineage cells and autoimmunity. J Immunol 157, 4371–4378. Sato, S., Jansen, P. J., and Tedder, T. F. (1997). CD19 and CD22 expression reciprocally regulates tyrosine phosphorylation of Vav protein during B lymphocyte signaling. Proc Natl Acad Sci U S A 94, 13158–13162. Sato, S., Miller, A. S., Inaoki, M., Bock, C. B., Jansen, P. J., Tang, M. L., and Tedder, T. F. (1996b). CD22 is both a positive and negative regulator of B lymphocyte antigen receptor signal transduction: altered signaling in CD22-deficient mice. Immunity 5, 551–562. Sato, S., Steeber, D. A., Jansen, P. J., and Tedder, T. F. (1997). CD19 expression levels regulate B lymphocyte development: Human CD19 restores normal function in mice lacking endogenous CD19. J Immunol 158, 4662–4669. Sato, S., Hasegawa, M., Fujimoto, M., Tedder, T. F., and Takehara, K. (2000). Quantitative genetic variation in CD19 expression correlates with autoimmunity. J Immunol 165, 6635–6643. Sgroi, D., and Stamenkovic, I. (1994). The B-cell adhesion molecule CD22 is cross-species reactive and recognizes distinct sialoglycoproteins on different functional T-cell sub-populations. Scand J Immunol 39, 433–438.
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13 The Dynamic Structure of Antibody Responses IAN C. M. MACLENNAN AND DEBORAH L HARDIE MRC Centre for Immune Regulation, University of Birmingham Medical School, Birmingham, United Kingdom
In antibody responses, B cells are induced by antigen to proliferate and differentiate into antibody secreting cells. There are several steps in this process, in which external control is exerted in a series of microenvironments. How B cells pass successively through these microenvironments is considered here, together with the regulatory signals they receive in each site. The main focus is on antibody responses elicited with CD4 T cell control. These responses will be considered from the time B cells first encounter antigen until they differentiate into mature antibody secreting cells (Figure 13.1).
days, the plasmablasts come out of cell cycle and become plasma cells. A proportion of these die early but the spleen has the capacity to sustain a finite number of plasma cells for much longer (Sze et al., 2000). Extrafollicular antibody responses provide the most rapid route to adaptive antibody production from conventional (non-B1) B cells. The speed of the response may be critical in controlling the spread of infection. These responses produce both switched antibody and IgM, but are not associated with affinity maturation of the antibody response through hypermutation and selection, or memory B cell formation.
GC and Affinity Maturation
THREE ROUTES TO ANTIBODY PRODUCTION
The final source of antibody is from plasma cells derived from GC. The delay before the onset of antibody production is longer than in extrafollicular antibody responses (Smith et al., 1996). Plasma cells derived from GC can be very long lived (Manz et al., 1999; Slifka and Ahmed, 1998). In addition, the affinity of the antibody they produce is augmented through Ig V-region hypermutation (Jacob et al., 1991b) and the selection of high affinity mutant B cells (MacLennan, 1994). Memory B cells and long-lived B cell clones characterize these responses (Askonas and Williamson, 1972; Coico et al., 1983; Klaus and Humphrey, 1977; MacLennan et al., 1990).
Three main sources of antibody production exist. First, a subset of B cells known as B-1 cells can mature to become antibody-secreting cells, without apparent activation by external antigen. Second, B cells can be induced by Tdependent and T-independent antigens to grow in extrafollicular sites as plasmablasts. These give rise to the plasma cells responsible for early switched and nonswitched antibody production. Finally T-dependent antigens also induce the formation of germinal centers (GC); these are required for sustained high affinity antibody responses. This review will focus on follicular and extrafollicular adaptive responses.
STAGES OF ADAPTIVE ANTIBODY RESPONSES
Adaptive Extrafollicular Antibody Responses The second source of antibody results from the growth of antigen-activated B cells as plasmablasts. This occurs in extrafollicular foci in the spleen (Jacob et al., 1991a; Toellner et al., 1996) and the medullary cords of lymph nodes (Luther et al., 1997). After proliferating for 2 to 3
Molecular Biology of B Cells
The changes that occur in B cells during adaptive Tdependent antibody responses can be divided into four main phases, each with its own microenvironment (Figure 13.1). The phases are:
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where the B cells proliferate as plasmablasts, or in follicles where the B cells form GC. • Plasma cells, or their immediate precursors, locating at sites that sustain plasma cell survival.
HOW AND WHERE B CELLS ENCOUNTER ANTIGEN
FIGURE 13.1 The phases of T cell-dependent antibody responses. An initial common path of antigen capture by the B cells occurs followed by cognate interaction with the T cells. Responses then diverge, with B cells growing in follicles and extrafollicular sites. The color code identifies the stages in which Ig heavy chain gene switch recombination, variable region hypermutation, and secretion of antibody occur. See color insert.
• Antigen entrapment by naïve or memory B cells through their antigen-specific receptors (BCR): This initiates a Tdependent antibody response if BCR engagement, together with accessory signals, is sufficient to provoke antigen internalization and processing and induces changes that allow B cells to find and interact with primed T cells. • The interaction of naïve or memory B cells that have taken up antigen with primed T cells: This occurs in the T cell-rich areas of secondary lymphoid tissues. • B cell proliferation and subsequent differentiate into plasma cells: This occurs either in extrafollicular foci,
The potential for and consequences of antigen encounter differs with the B cell type. Recirculating B cells are in constant migration between the follicles of the secondary lymphoid tissues via blood and lymph (Nieuwenhuis and Ford, 1976). They can pick up antigen when they are in the blood. This characteristically results in them migrating to the T zones of the spleen (Toellner et al., 1996). Recirculating B cells enter the T zones of lymph nodes by passing across high endothelial venules. They then migrate to the follicles via the walls of intranodal lymphatics (MacLennan and Gray, 1986). There they have access to antigen in the lymph. Dendritic cells have been described that transport intact antigen to naïve B cells in lymph nodes (Wykes et al., 1998). Similarly, dendritic cells have recently been identified in the blood, which transport antigen to splenic marginal zone B cells (Balazs et al., 2002). These are CD11clow dendritic cells that probably correspond to CD45RA- CD11clow CD11b+ blood monocytes with immediate dendritic cell precursor potential (pDC1) (Shortman and Liu, 2002). Antigen in the form of immune complex is held on follicular dendritic cells (FDC) (Brown et al., 1970; Tew et al., 1984). Recirculating B cells in the follicular mantle might be expected to have access to this antigen, but experimental evidence suggests this is not the case (Gray, 1988; MacLennan et al., 1990; Vonderheide and Hunt, 1990). Conversely, memory B cells in parallel transfer experiments do respond to antigen held on FDC. Newly produced naïve B cells, which have not been selected to enter either the recirculating or marginal zone pools, are also capable of eliciting T cell help after engaging antigen (Cook et al., 1998). These cells are more readily induced into apoptosis or receptor editing by engagement of antigen (Brink et al., 1992; Retter and Nemazee, 1998). This may be associated in part with their lower level of CD21 expression compared to that of recirculating B cells, which in turn have lower levels of CD21 than marginal zone B cells (Oliver et al., 1997; Timens et al., 1989). Cross-linking the BCR with CD21 markedly reduces the threshold for B cell recruitment into T-dependent antibody responses (Dempsey et al., 1996). Splenic marginal zone B cells are perfused by a blood sinusoidal network and consequently are well placed to pick up antigen from the blood. They are able to mount an
13. The Dynamic Structure of Antibody Responses
extrafollicular antibody response to polysaccharide antigens without eliciting T cell help (Lane et al., 1986; Martin and Kearney, 2000) and are also capable of mounting extrafollicular T-dependent and T-independent type 1 antibody responses (Liu et al., 1991b). Cells with a phenotype similar to that of splenic marginal zone B cells are found in the crypt epithelium of tonsil (Liu et al., 1995), beneath the dome epithelium of Peyer’s patches (Spencer et al., 1985), and on the inner wall of the subcapsular sinus of lymph nodes, particularly the mesenteric nodes (Stein et al., 1980). It will be appreciated that the locations of these marginal zone–like cells are favorable for encountering antigen in lymph or antigen that has crossed epithelial surfaces. Marginal zone B cells do not recirculate but equivalent cells circulate in the blood (Klein et al., 1998; Thorley-Lawson, 2001). M cells of Peyer’s patches and tonsil crypts transport antigen across epithelial surfaces (Brandtzaeg and Bjerke, 1989) and the pDC1, discussed above, transport bacteria to marginal zone B cells (Balazs et al., 2002).
PRIMARY COGNATE INTERACTION OF B CELLS WITH PRIMED T CELLS In vivo, naïve recirculating B cells pass primed T cells, like ships in the night, as they migrate through the outer T zone. After engagement of their BCR, the same B cells rapidly make cognate interaction with primed T cells (Liu et al., 1991b; Toellner et al., 1996). Human tonsillar B cells, on engaging antigen, temporarily lose responsiveness both to the chemokine CXCL13 (BLC or BCA-1), which is expressed in follicles, and to the crypt epithelial chemokines CXCL12 (SDF-1) and CCL3 (MIP-3a). At the same time, their migratory response to CCL4 (MIP-3b), which is produced in the T zone, is reinforced (Casamayor-Palleja et al., 2002). B cells make cognate interaction with T cells at an early stage of T cell priming. Thus, following subcutaneous immunization with virus or alum-precipitated protein, previously naïve T cells start to proliferate in the draining nodes at around the same time that they induce B cells to produce switch transcripts (Cunningham et al., 2002; Toellner et al., 1998). B cells, therefore, may make cognate interaction with CD4 T cells while the T cells are still in contact with the dendritic cells that induced the priming process. This may not be the case in secondary responses, in which primed T cells are available for immediate cognate interaction with B cells that have engaged antigen (Toellner et al., 1996). Complex roles for dendritic cells in modifying B cell growth and differentiation have been proposed from studies in vitro (Fayette et al., 1998). The role of plasmablast-associated dendritic cells in plasmablast maturation is discussed in a later section.
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Commitment of Activated B Cells to Follicular or Extrafollicular Growth Following cognate interaction with primed T cells, B cells enter cell cycle in the T zone, but they soon move either to follicles, where they form GC, or the extrafollicular sites where the B cells grow as plasmablasts (Jacob et al., 1991a; Toellner et al., 1996). Much is known of the divergent phenotypic changes that occur in B cells growing in these two sites. These are discussed later in the context of plasmablast growth and GC formation. By contrast, there is little insight into the nature of signals that commit a B cell to grow in follicles, as opposed to an extrafollicular focus or vice versa. The ligation of B cell CD40 by T cells is critical in T cell–dependent antibody responses, both for GC formation and for extrafollicular growth of B cells (Castigli et al., 1994; Xu et al., 1994). Surprisingly, the CD40 signaling via TRAF2/3 or 6 does not appear to be required to induce B cells to grow in either of these sites (Ahonen et al., 2002; Jabara et al., 2002). Roles for CD40-directed TRAF signaling in switching to IgG1 and to bone marrow plasma cell formation are discussed later. The ligation of T cell CD28 appears to be required for T-dependent GC formation, whereas some T-dependent extrafollicular B cell growth is induced in the absence of CD28 signaling (Lane et al., 1994). Strong BCR-ligation is capable of inducing both follicular and extrafollicular B cell growth without T cells (Vinuesa et al., 2000). It is unclear if a single B cell, activated by cognate interaction with T cells, sends progeny both to form a GC and to grow as plasmablasts. The finding of ipsiclonal cells in both GC and adjacent red pulp plasma cells has been taken to support this concept (Jacob and Kelsoe, 1992). An alternative explanation is suggested by finding hapten-specific plasma cells with heavily mutated Ig V-region genes in the splenic red pulp within 5 days of immunizing carrier-primed mice with hapten-carrier (Sze et al., 2000). These mutated hapten-specific plasma cells are likely to be early emigrants from GC. The kinetics of GC formation and the oligoclonality of GC also argue against dual differentiation pathways for a single cell. On average, three B cells give rise to a single GC (Kroese et al., 1987; Liu et al., 1991b), and these proliferate to yield 104 - 1.5 ¥ 104 cells in 96 hours (Liu et al., 1991b; Toellner et al., 1996). The cell cycle time of these cells is estimated at 6 hours (Hanna, 1964; Zhang et al., 1988). If these estimates are correct, three B cells should yield twelve thousand cells in 96 hours. This would not be achieved if there were significant emigration, divergent differentiation, or cell death. There is no absolute requirement for a follicular microenvironment for B cells to adopt a GC B cell phenotype and grow exponentially. This is shown by the ectopic growth of GC. The signals committing B cells to acquire a GC
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phenotype are received as B cells first engage antigen and then make cognate interaction with primed T cells in the T zone. The same conclusion seems to fit with B cell growth as plasmablasts. This also can occur in ectopic sites (Vinuesa et al., 1999). Conversely, plasmablast differentiation to plasma cells and maintenance of fully developed GC depends on external signals (discussed in later sections).
Growth and Differentiation of CD4 T Cells During Antibody Responses After priming, CD4 T cells move and differentiate in a number of directions. Some accumulate towards the edge of the T zone, while others move to follicles (GulbransonJudge and MacLennan, 1996; Garside et al., 1998;). Others leave the lymphoid tissue either as memory cells (Rogers et al., 2000), which move to and through distant secondary lymphoid tissues, or as effectors, which enter sites of inflammation (Sallusto et al., 1999). Primed T cells do not migrate to the sites in the spleen and lymph node where B cells grow as plasmablasts (Gulbranson-Judge and MacLennan, 1996; Luther et al., 1997). The migration of T cells to follicles is discussed later.
EXPONENTIAL GROWTH OF ACTIVATED B CELLS Extrafollicular Growth of B cells as Plasmablasts Plasmablast growth is a feature of antibody responses in lymph nodes and the spleen (Fig. 13.2). It has not been identified in the lymphoid tissue associated with the walls of the alimentary, respiratory, and genital tracts. The tonsils, unlike Peyer’s patches, contain large numbers of mature plasma cells. The Ig isotypes produced by these reflect the relative concentration of Ig classes in the blood IgG > IgA > IgM. Both the switched and nonswitched tonsil plasma cells have heavily mutated IgV-region genes, indicating a GC origin (Yavuz et al., 2001). In lymph nodes, plasmablast growth classically occurs in the medullary cords, which expand as the number of plasmablasts increases. The cords contain distinctive CD11chigh dendritic cells, which are capable of proliferating as the plasmablasts grow (Vinuesa et al., 1999). They do not contain CD4 T cells (Gulbranson-Judge and MacLennan, 1996; Luther et al., 1997). In the mouse spleen, plasmablast growth classically occurs in foci that lie in the red pulp where it directly abuts the T zone. These foci have a similar mixture of cells to that of the medullary cords. The differentiation of a B blast to a plasmablast is associated with the upregulation of the transcriptional repressor BLIMP-1 (Mock et al., 1996; Shaffer et al., 2002); this
FIGURE 13.2 Diagrammatic representation of the stages of an extrafollicular antibody response: Antigen capture by B cells and T cell priming is followed by cognate T cell interaction with B cells. Some of the activated B cells migrate to extrafollicular foci in the spleen or the medullary cords of lymph nodes where they proliferate as plasmablasts. This growth is associated with CD11chigh dendritic cells. Plasmablasts that are not associated with these dendritic cells appear to die without making the transition to plasma cells. In the spleen, plasma cells produced in the extrafollicular responses and plasma cells that have been generated in follicles compete for space on stroma that supports long-term plasma cell survival. This stroma is associated with blood vessels and contiguous fibrous bands in the red pulp. See color insert.
downregulates the expression of genes involved in B cell receptor signaling and proliferation while allowing the expression of genes required for plasma cell development, such as XBP-1 (Reimold et al., 2001; Shaffer et al., 2002). Expression of Bcl-6, which is associated with B cell growth in follicles, represses these changes associated with B cell differentiation to plasmablasts (Fearon et al., 2001). Although T cells are required to induce these features of plasmablast differentiation in responses to T-dependent antigens, they develop perfectly well in mice devoid of T cells in responses to T-independent antigens. In addition, their induction does not require the medullary environment, nor that of an extrafollicular focus (Vinuesa et al., 1999). Con-
13. The Dynamic Structure of Antibody Responses
versely, the transition of plasmablast to plasma cell seems to depend on environmental signals, which are usually available in the medulla or extrafollicular foci. This is seen when the number of plasmablasts produced is very large. For example, in responses to NP-Ficoll in mice with an NPspecific transgenic BCR there is an impressive growth of plasmablasts. These fill the splenic red pulp, but most of the ectopic plasmablasts die early. The absolute number of plasmablasts that make the transition and survive for some days is similar to the number surviving in nontransgenic mice. This suggests that the spleen has a finite capacity to allow full maturation from plasmablast to plasma cell (Sze et al., 2000). Evidence points to a critical role for CD11chigh dendritic cells in this transition. The cells making the transition to plasma cells are seen to be adjacent to CD11chigh dendritic cells. This still applies where the location of CD11chigh dendritic cells is changed, as in T cell–deficient mice where they are focused in the T zone. When the number of CD11chigh dendritic cells is increased by activation through CD40, the number and the location of mature plasma cells increases in parallel with the expansion of CD11chigh dendritic cells (Vinuesa et al., 1999). A recent report suggests that the transition is associated with dendritic cell–derived TACI ligands (BAFF/BLyS, or APRIL, or both of these) (Balazs et al., 2002). The chemokine CXCL12 is prominently expressed in extrafollicular foci and medullary cords (Hargreaves et al., 2001). Plasmablasts deficient in CXCR4 (the receptor for CXCL12) fail to migrate to normal sites of antibody and CXCL12 production in the spleen (Hargreaves et al., 2001). It will be helpful to determine if CD11chigh dendritic cells in the spleen and lymph nodes are the main source of CXCL12. Recent studies indicate that plasmablasts having a defect in cell cycle arrest, through lack of the CDK inhibitor p18(INK4c), are unable to make the transition to high-level antibody secretor status (Tourigny et al., 2002). Switch recombination is triggered at the time of primary T cell interaction with B cells in the T zone (Toellner et al., 1996). This leads to heavy chain gene recombination when the activated B cells are growing as plasmablasts. The switched and nonswitched plasmablasts have equivalent chances of differentiating into plasma cells (Sze et al., 2000).
The Exponential Growth Phase of GC Formation Physiologically, GCs form when B cells activated by antigen and cognate interaction with T cells grow in follicles and modify their immunoglobulin V-region genes by an active process of hypermutation. The cells with altered BCR specificity only leave the GC following positive selection, which involves the B cells binding antigen and presenting this to local CD4 T cells. Cognate interaction with the T cells induces differentiation to plasma cells, or memory B cells, or to centroblasts (Figure 13.3, reviewed in MacLennan,
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FIGURE 13.3 A scheme suggesting how B cells proliferate and activate hypermutation before being selected and induced to differentiate in established GC. Centroblasts proliferate and mutate their Ig-V region genes. Periodically, they are subjected to selection. Successful selection depends on the B cells binding antigen, normally held on FDC, processing this, and presenting the resulting peptides to local T cells. Selected cells differentiate to become memory B cells, plasma cells, or centroblasts. The last remain in the GC and undergo further proliferation and V-region hypermutation. This regeneration of centroblasts is essential for maintaining the GC. Cells that fail selection die in situ by apoptosis. See color insert.
1994). Naïve recirculating B cells can be induced to form GC; it is less clear whether marginal zone or memory B cells can also form classical GC. B1 cells are the main B cell population present in human infants during the first year of life. Neonates can mount T-dependent antibody responses with affinity maturation (Anderson, 1983; Eskola et al., 1990). It is unclear whether these antibodies originate by the recruitment of B1 cells, or a minority recirculating B cell subset, into the response. Memory B cells can be reactivated by antigen on FDC (Gray, 1988; MacLennan et al., 1990; Vonderheide and Hunt, 1990), and memory B cell clones have been reactivated and maintained through seven successive transfers through syngeneic recipients (Williamson and Askonas, 1972). It would be useful to revisit this model of persistent B cell clones to look for evidence of ongoing intraclonal Ig V-region mutation and selection in these clones.
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The induction of B cell growth by uptake of antigen and cognate interaction with primed T cells was described earlier. Here, the kinetics of GC formation is considered. This is conveniently studied when the response is synchronized and only a single cohort of B cells is recruited into the response. It is possible to achieve this in responses to hapten-carrier conjugate of rodents primed with the protein carrier. Carrierspecific T cell help and carrier-specific antibody are present at the time of intravenous challenge with hapten-carrier. These factors ensure that cognate T cell interaction with B cells in the spleen occurs rapidly and that antigen is swiftly cleared from the circulation. As a result, only a single cohort of B cells is recruited into the response. (Toellner et al., 1996). The majority of the B cells recruited are carrier-specific memory cells, which dominate the extrafollicular response, but the minor naïve hapten-specific component is mainly responsible for the GC formed (Liu et al., 1991b; Toellner et al., 2002). Once triggered to form, GCs B cells upregulate Bcl-6 and go through approximately twelve cell cycles in a 3 to 4 day period when the three or so founding cells have become twelve thousand cells. This marks the end of the phase when the number of B cells in the GC grows exponentially. After this there is a major change in the GC, and a phase ensues in which there is a balance between growth and cell loss (this second phase is discussed in the next section). The oligoclonality of GC was first identified by Kroese et al. (1987) in mixed irradiation chimeras and was confirmed in nonirradiated rats responding to two haptens conjugated to the same protein (Liu et al., 1991b). Around 25% of the GC were specific for one hapten, whereas the others were of mixed specificity. This is consistent with an average of only three B cells founding a single GC. The oligoclonality of the GC persists following a single immunization until the GC reactions ends after about a month (Liu et al., 1991b). The persistent oligoclonality of GC has been confirmed repeatedly by the analysis of V-region genes from single GC (Jacob et al., 1991b; Küppers et al., 1993). The exponential growth phase of GC can be induced without T cell help (Lentz and Manser, 2001; Vinuesa et al., 2000) and it can occur outside the follicular environment and in the absence of FDC (Futterer et al., 1998; Weih et al., 2001). Thus, the induction, maintenance, and termination of the exponential growth phase has no absolute dependence on signals from these elements. This also applies to the onset of expression of Bcl-6, as well as the molecule identified by the monoclonal antibody GL7 and the molecule(s) that binds peanut agglutinin (Vinuesa et al., 2000).
PROLIFERATION, HYPERMUTATION, AND SELECTION IN GC The exponential growth phase of GC formation ceases by 96 hours after the primary induction of B cells to grow in
follicles (Liu et al., 1991b; Toellner et al., 1996). By this stage, hypermutation is well underway and hapten-specific GC B cells have already produced second-generation Ig V region mutants (Toellner et al., 2002). By 96 hours, memory B cells have also started to colonize the marginal zone (Liu et al., 1991b; Toellner et al., 1996) and hapten-specific plasma cells with mutated Ig V region genes are detectable by 5 days after challenge (Sze et al., 2000). The switch from B cell growth without death or selection to one in which growth is balanced by death and emigration from the GC represents a massive transition of B cell behavior. In late 2002, there is still remarkably little insight into the way this transition is achieved at the molecular level.
The Organization of Established GC The compartmentalization of GC in human secondary lymphoid tissue into a dark and light zone was recognized long before the function of GC was identified (reviewed in Nieuwenhuis and Opstelten, 1984) (Figure 13.4). This is also apparent in rat (Zhang et al., 1988, Liu et al., 1991b) and sheep (Blacklaws et al., 1995) GC. It is less obvious in the early stages of GC formation in mice (Camacho et al., 1998), but the recognition of compartmentalization has contributed considerably to developing a working hypothesis for GC function, which is equally tenable in mice (MacLennan, 1994; MacLennan and Gray, 1986). Primary Follicles B cell follicles that contain GC are known as secondary follicles, whereas those without GC are primary follicles. Primary follicles comprise small recirculating lymphocytes and FDC. The presence of recirculating cells is necessary for the differentiation of FDC. There is an absence of FDC in animals congenitally deficient in B cells (EnriquezRincon et al., 1984; MacLennan and Gray, 1986). Recirculating cells home to follicles in rats that have previously lacked B cells and are devoid of FDC (Bazin et al., 1985). The arrival of recirculating B cells in the follicles results in the appearance of FDC within 2 to 3 days (MacLennan and Gray, 1986). The B cell influence on the differentiation of FDC from their still unidentified radiation-resistant stromal cell precursors is via the production of lymphotxin-a1b2 and TNF-a by the B cells (Endres et al., 1999). The FDC precursors require the presence of lymphotoxin-b receptor (Endres et al., 1999) and the TNF-aR (Matsumoto et al., 1997; Tkachuk et al., 1998). This appears to direct the production of the transcription factor composed of the heterodimer of RelB (Weih et al., 2001) and NF-kB/2 from their precursors (Franzoso et al., 1998). Both components of the heterodimer are required for FDC to differentiate from their precursors. Recirculating B cells are attracted to follicles by the chemokine CXCL13 (BLC or BCA-1), which
13. The Dynamic Structure of Antibody Responses
FIGURE 13.4 Histological sections of the light zone (above) and the dark zone (below) of a GC from a human tonsil. The section is stained with pyronin, which stains RNA magenta, and methyl green, which stains DNA blue/green. In the dark zone, pyroninophilic centroblasts are closely packed. There are many mitoses (marked M). Tingible body macrophages appear as pale islands in the continuum of centroblasts. Occasional apoptotic debris (tingible bodies) in these macrophages is arrowed. In the light zone, only occasional cells are pyoninophilic. The centrocytes are spaced by the presence of the follicular dendritic cell network. Apoptotic nuclear fragments are arrowed. See color insert.
they respond to through their CXCR5 (Okada et al., 2002). There is evidence that CXCL13 induces recirculating B cells to produce lymphotoxin-a1b2, whereas GC B cells constitutively produce this FDC differentiation factor (Ansel et al., 2000). Secondary Follicles In established GC, the recirculating B cells are largely displaced from the FDC network. They form a mantle, which surrounds most of the GC. The follicular mantle is thickest at the apical pole (light zone) of the follicle and is thin or absent from the base of the follicle where the dark zone is located. The line of demarcation between the GC and the follicular mantle is usually distinct in human (Figure
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FIGURE 13.5 Three-color fluorescence of a tonsil GC to show the zonal pattern of this structure. The section is stained for Ki67 nuclear expression by cells in cell cycle (red); these are most abundant in the dark zone (DZ). CD23 expression is shown in blue. This stains the FDC of the apical light zone (ALZ) and B cells in the follicular mantle (FM). CD21 (green) is expressed by a broader network of FDC than CD23; the CD21+ CD23- FDC network below the CD23+ network is termed the basal light zone (BLZ), and that between the apical light zone and the follicular mantle the outer zone (OZ). See color insert.
13.5) and rat GC, but in mice the border between the two is often indistinct with IgD+ recirculating cells mixed with GL7+, Bcl6+, and peanut agglutinin-binding GC B cells. Compartments of Secondary Follicles The dark and light zones initially were defined morphologically in conventionally fixed histological preparations. The dark zone contains closely packed blasts that have a relatively narrow rim of cytoplasm that is strongly pyroninophilic, reflecting its abundance of RNA. The chromatin of the blasts is open and mitotic figures are plentiful (Figure 13.4). The sheets of blasts are broken only by palestaining large macrophages, each of which forms an island in the continuum of blasts. The dark zone macrophages contain variable numbers of basophilic (tingible) bodies, which are apoptotic nuclear fragments.
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In the light zone (Figure 13.4), the B cells are more widely spaced, reflecting the presence of the pale-staining FDC and macrophages and dendritic cells. Classically, in human GC many of the B cells in the light zone are out of cell cycle, and most of these have much less pyroninophilia than the centroblasts of the dark zone. These light zone B cells are termed centrocytes (Lennert, 1978). Even in human GC, a significant proportion of B cells in the light zone are in cell cycle, as judged by Ki67-staining, but the proportion of proliferating cells is greater towards the dark zone. It will be appreciated from this that it is difficult to draw a precise line where the dark zone ends and the light zone begins. In the human tonsil, the FDC network is clearly defined by its high-level expression of CD21. A smaller part of the FDC network also expresses CD23. Tonsil GC have been divided into four zones on the basis of CD21 and CD23 expression by the FDC network (Hardie et al., 1993) (Figure 13.5). At the base of the GC is an area that has little or no FDC. This includes part of the conventional dark zone. Next is a zone with CD21+, CD23- FDC. This has been termed the basal light zone, although it does contain a substantial number of blasts and mitotic figures and largely falls within the conventional dark zone. The next zone is termed the apical light zone. This is defined by the CD21+, CD23+ FDC network. In this area are few mitotic figures. Finally, between the apical light zone and the follicular mantle, lies the outer zone. This, like the basal light zone, has a CD21+, CD23- FDC network. The outer zone has the highest concentration of T cells (Fig. 13.6). Although this zonal pattern is consistent in tonsils from one individual to another it does not apply in human lymph nodes, in which CD23 is expressed by most of the CD21+ FDC network (Brachtel et al., 1996). As indicated above, the proportion of proliferating cells in the FDC network is higher in mice (Camacho et al., 1998). Nevertheless, the dense FDC network is localized in the apex of GC and is relatively deficient at the base (see Figure 1C&E in Koni and Flavell, 1999).
the dark zone. These are seen in HIV-associated lymphadenopathy, but are usually CD8+. Although T cells are found throughout the light zone, they are particularly focused at the junction of the follicular mantle and light zone (Hardie et al., 1993). They are CD4+ and express CXCR5 (Ansel et al., 1999; Schaerli et al., 2000). In humans, they are uniformly CD45RO+, about half contain preformed CD40 ligand (Casamayor-Palleja et al., 1995), while most but not all contain CTLA-4 and some 20 to 30% express CD57 (Hardie et al., 1993). The CD57+ GC T cells are more frequent in the body of the light zone than at its rim (Figure 13.6). In mouse, and to a smaller extent in human, T cells are found in the follicular mantle (Gulbranson-Judge and MacLennan, 1996). GC T cells, in common with centrocytes and centroblasts, express minimal levels of Bcl-2. Although these phenotypic features characterize GC T cells, none is an absolute indicator of GC location, since similar effector CD4 T cells are seen in the outer T zone. The migration of T cells to follicles is driven by cognate interaction with dendritic rather than B cells and seems to require CD40 ligation (Fillatreau, 2002). This is consistent with the finding that in the absence of B cells T cells upregulate CXCR5 during primary T cell-dependent immune responses (Toellner, personal communication). The number of T cells colonizing follicles is substantially augmented in mice, with overexpression of OX40L on dendritic cells (Brocker et al., 1999). Situations have been identified in which T cells are induced by cognate interaction in the T zone to migrate to follicles some days before GC form (Luther et al., 1997).
Centroblasts and Centrocytes Pulse chase experiments in mice (Hanna, 1964) and rats (Liu et al., 1991b) indicate that centroblasts are precursors of centrocytes. These suggest that most of the non-dividing cells in the light zone are derived from precursors that were in S phase of the cell cycle some 9 hours previously (Liu et al., 1991b). It is plausible that the labeled but nondividing cells in the light zone are derived from centroblasts of the dark zone. Evidence for the formation of centroblasts from selected centrocytes is discussed later. T cells in Secondary Follicles In addition to the differences listed above, there is clear polarization of T cells in GC. Very few T cells are found in
FIGURE 13.6 Three-color fluorescence of tonsil GC. On the left the expression of CD3 by T cells is shown green. CD74 (invariant chain) expression by B cells but not FDC is stained blue, while CD21 expression by FDC is stained red. On the right, CD3 again is stained green; IgD (red) is expressed by follicular mantle B cells. CD57 (purple) is expressed by a minority of GC T cells. The CD57+ve T cells tend to be located in the center of the light zone whereas CD57-ve GC T cells are clustered along the junction of the follicular mantle and the light zone. See color insert.
13. The Dynamic Structure of Antibody Responses
The Role of FDC in Centrocyte Selection and the Maintenance of Antibody Responses Follicular dendritic cells characteristically localize antigen on their surface in the form of antigen–antibody complex. The antigen can persist on these cells for extended periods (Szakal et al., 1989; Tew and Mandel, 1979). Both complement (Klaus and Humphrey, 1977) and antibody (Nossal, 1965) are required for antigen localization on FDC, and this is associated with functional roles for FCgIIB receptors (Qin et al., 2000) and complement receptors on the FDC (Carroll, 1998; Fischer et al., 1998). The localization appears to involve an active cellular transport mechanism, with a different cell being responsible for localization in lymph nodes and the spleen. In the former, the transporting cell is radiation resistant (Mandel et al., 1980) while the localization on splenic FDC is highly radiation sensitive (Brown et al., 1973). The splenic antigen-transporting cells appear to be marginal zone B cells (Brown 1970; Gray et al., 1984; Oldfield et al., 1988). The antigen associated with FDC is in native form. Shortly after localization it is sometimes taken into the cell in vesicles. These vesicles, which have been termed iccosomes, appear to be extruded from the FDC, but remain attached to their surface (Szakal et al., 1989; Tew and Mandel, 1979). B cells can take up antigen from FDC and present this to T cells (Kosco et al., 1988). In addition to binding antigen, FDC passively acquire a number of molecules such as MHC class II molecules, which are not produced by the FDC themselves. This is clearly seen in bone marrow chimeras where donor class II MHC differs from the recipient (Gray et al., 1991). The lack of synthesis of class II MHC molecules by FDC is underscored by the lack of invariant chain expression by FDC, while this is strongly expressed by centrocytes, perhaps reflecting active antigen processing (Figure 13.6). Recently, electron microscopic analysis of class II molecules held on FDC suggests that this is held in exosomes, which are secreted internal vesicles from multivesicular endosomes of other cells (Denzer et al., 2000). Contact with FDC in vitro seems to inhibit GC B cell apoptosis (Koopman et al., 1991; Kosco et al., 1992; Lindhout et al., 1993). Disruption of adhesion of the B cell to the FDC via LFA1/CD54 and VLA4/VCAM1 results in B cell apoptosis (Koopman et al., 1991, 1994). The signaling pathways by which FDC inhibits B cell apoptosis have recently been reviewed (van Eijk et al., 2001). Although some affinity maturation may be achieved with little or no antigen localized on FDC (Hannum et al., 2000), several studies indicate this is suboptimal. This is elegantly shown in a recent study of mouse chimeras where antigen localization on FDC was impaired through lack of the complement receptors CD21 (CR2) and CD35 (CR1) (Barrington et al., 2002). Importantly, the lymphocytes were transferred
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from congenic mice without these deficiencies, for when cross-linked with the BCR, CD21 enhances B cell activation (Dempsey et al., 1996). These chimeras had a relatively unimpaired short-term antibody response, but those with a CD21/35-deficient background were less efficient than wild-type mice at maintaining serum antibody titers, or antibody secreting cell numbers in the spleen and bone marrow. In addition, they had reduced numbers of functional memory B cells. These studies confirm early reports that indicate that memory B cells are not sustained efficiently in the absence of antigen (Askonas and Williamson, 1972; Gray and Skarvall, 1988; Karrer et al., 2000). Thus, although some bone marrow and splenic plasma cells can survive for long periods (Manz et al., 1999; Slifka and Ahmed, 1998) B cell memory and long-term antibody titers are enhanced by persistent antigen on FDC. The studies considered in the previous paragraph indicate that in the short term GC can function, although probably inefficiently, in the absence of antigen held on FDC. Evidence for an additional role for FDC in selection comes from the studies of mice, mentioned earlier in which GC fail to develop in follicles, but form in the T zone. This happens in mice deficient in CXCR5 and CXCL13, but in both these strains of mice FDC form in the ectopic GC, and affinity maturation and B cell memory formation occurs (Voigt et al., 2000). Mice deficient in lymphotoxin-bR (Futterer et al., 1998) or TNF-aR (Endres et al., 1999) produce ectopic GC that lack FDC, and these do not appear to support FDC or memory production. This applies equally to NF-kB2deficient mice (Hsu, Caamaño, and MacLennan, unpublished data). The defects in these mice cannot be restored with wildtype B cells. Nevertheless, transfer of B and T cells from these mice to lymphocyte-deficient mice will generate GC with FDC that produce memory B cells (Endres et al., 1999; Matsumoto et al., 1997). Lymphotoxin-b-deficient, lymphotoxin-a and TNF play important roles in FDC formation and the organogenesis of secondary lymphoid tissue. Deficiency in any one of the cytokines is not associated with complete loss of FDC networks (Alexopoulou et al., 1998).
T Cells in Centrocyte Selection and the Maintenance of GC The popular and tenable hypothesis for selection in GC proposes that B cells that have undergone hypermutation enter a phase in which they must take up antigen and use this to make cognate interaction with local T cells if they are to survive (Figure 13.3). The selected B cells appear to be induced to differentiate in one of three directions. They can leave the GC to become memory B cells (Coico et al., 1983; Klaus and Humphrey, 1977) or plasma cells (Benner et al., 1981; Smith et al., 1996). Alternatively, they may stay within the GC and undergo further proliferation and
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hypermutation before again being subject to selection (Casamayor-Palleja et al., 1996; Vinuesa et al., 2000). Evidence for the death of GC B cells if T help is not available comes from studies of GC induced without T cells. The GC undergo involution on the fifth day after induction of growth. This is associated with massive B cell apoptosis and no output of memory B cells or plasma cells (Vinuesa et al., 2000). By contrast, normal GC, with CD4 T cells that can recognize processed antigen presented by centrocytes, do not undergo early involution. They retain a significant growth fraction and generate memory B cells and plasma cells. The death of cells in mature GC in the absence of T cells reflects a default mechanism where cells die for lack of signals that override apoptosis. GC B cells have an exceptional tendency to enter apoptosis when they are cultured (Liu et al., 1989). This, in part, is associated with their very low level of expression of the anti-apoptotic protein Bcl-2 (Liu et al., 1991a; Pezzella et al., 1990) and partially with the expression of the pro-apoptotic molecules FAS, c-MYC, and BAX (Martinez-Valdez et al., 1996). Although physiological GC only slowly wane in size, B cell apoptosis is a constant feature of all GC. The exponential growth phase of GC formation is not associated with B cell death; for the cell production achieved would be impossible if there were significant attrition at this stage. The transition to the selection phase is associated with an increased susceptibility to apoptosis. The conditions triggering this change appear to be obscure. During the selection phase, B cells can be prevented from entering apoptosis by T cell–derived survival signals. CD40 ligation is a powerful inhibitor of apoptosis in these cells (Liu et al., 1991b), and this appears to act by maintaining levels of cFLIPL inhibitory protein, which blocks the apoptotic cascade by inhibiting caspase 8 activation (Irmler et al., 1997; Tschopp et al., 1998). cFLIPL is constitutively expressed in GC B cells, but its levels rapidly decline when the cells are cultured (Hennino et al., 2001). It appears that signals delivered from FDC as well as CD40-dependent signals can inhibit apoptosis in GC cells (van Eijk et al., 2001). There may be additional mechanisms preventing apoptosis in the centroblasts in the dark zone, many of which do not have direct contact with FDC. In many in vitro experiments designed to probe the mechanisms of GC B cell selection, CD40-ligand or agonistic anti-CD40 antibodies are continuously present. The use of CD40 ligation in these cultures is valuable for keeping centrocytes alive in vitro, but the time scale for differentiation to putative memory cells in these cultures (Arpin et al., 1995) does not reflect the rapid in vivo transition from proliferating GC B cell to memory cell (Chan and MacLennan, 1993). Further, the phenotypes of cells generated following sustained CD40 ligation in vitro have many differences from those of freshly isolated memory B cells or plasma cells (Casamayor-Palleja et al., 1996). Physiologically, CD40-
ligand is only transiently expressed during cognate T cell–B cell interaction (Yellin et al., 1994). The ligand is present as a pre-formed intracellular protein in about 50% of GC T cells and is rapidly expressed on the cell surface on T cell receptor ligation (Casamayor-Palleja et al., 1995). It is rapidly lost from the T cell surface once it binds to CD40 on B cells (Yellin et al., 1994). GC T cells with induced CD40-ligand expression on their surface form conjugates with autologous GC B cells and rapidly induce about half of these to differentiate into cells with a plausible memory B cell phenotype (Casamayor-Palleja et al., 1996). Although CD40-ligation is necessary to achieve this effect, it is not sufficient, for CD45RA CD4 T cells from the same tonsil that have been induced to express equivalent levels of CD40-ligand do not protect from apoptosis. Recent data show that prolonged CD40 ligation inhibits GC formation in vivo and the production of long-lived bone marrow plasma cells (Erickson et al., 2002). Evidence that CD40 ligation is important for the maintenance of GC is provided by studies in which CD40L blockade caused rapid involution of established GC (Han et al., 1995). Blocking CD86 binding to CD28 or CTLA-4 did not have this effect but was reported to impair memory cell output from the GC. A recent study confirms that CD80 and CD86 signaling to T cells is not required to sustain established GC (Walker et al., 2003), although it is essential for their T-dependent induction (Lane et al., 1994). In this study, GCs were induced by T-dependent antigen in CTLA-4-Ig transgenic mice when an agonistic anti-CD28 antibody was administered. The GC persisted despite the continued presence of CTLA4-Ig. A possible regulatory role for CTLA-4 in established GC is suggested by the finding that these GC are substantially larger than those of wildtype mice given the same dose of anti-CD28 (Walker et al., 2003). Recent studies on mice deficient in the TRAF6 signaling domain of the cytosolic tail of CD40 show that GC induction, and probably maintenance, is achieved in these mice, but that they fail to produce long-lived bone marrow plasma cells (Ahonen et al., 2002). Another study confirms GC formation in mice deficient in the TRAF6-binding domain of CD40. This deficiency had little effect on in vitro or in vivo induced antibody levels, whereas mice deficient in the TRAF2/3-binding domains of CD40 have a selective loss of switching to IgG1 (Jabara et al., 2002). Further studies are required in this critical area of CD40 signaling. To summarize the data on the requirements for the induction and maintenance of GC in vivo: • T-independent GCs can be formed but are not sustained in the absence of GC T cells and appear to be nonproductive. • Ectopic GCs are produced in mice deficient in TNFaR, LTbR, NF-kB2, or RelB, but appear to be
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• •
•
•
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nonproductive. This might be due to the complete absence of FDC in these mice, a failure to attract T cells into the GC, or both. Each of these strains of deficient mice has B and T cells that can form productive GC in RAG-deficient mice. T cell-dependent GCs require CD40 and CD28 signaling for their induction. CD40 ligation, but not CD28 signaling, is required to sustain GC, probably reflecting a role in selecting centrocytes and inducing these to readopt a centroblast phenotype. CD40-activated TRAF-6 signaling may be required in the induction of centrocytes to become bone marrow plasma cells; neither this nor signals triggered through TRAFF-2/3 are required to induce or sustain GC. CD28 signaling may be important in inducing memory formation. There is evidence from short-term studies of GC T and B cell interaction in vitro that transient CD40ligation is necessary but not sufficient to induce memory B cell formation. CTLA-4 signaling may moderate GC size, perhaps by regulating the number of selected cells that stay within the GC.
There is a shortage of data that identify the signaling involved in centrocyte selection in vivo, or the signals that induce plasma cell, memory, or centroblasts differentiation in the selected cells. Future studies will also have to consider the differences between the various destinations of plasma cells leaving GC, for example, the gut, bone marrow, and tonsil, and the signals inducing class switching to particular isotypes. Models for plasma cell and memory B cell formation from centrocytes have been described in vitro, but these require rigorous correlation with events in vivo to test if they represent physiological signaling.
SUSTAINED SURVIVAL OF MEMORY B CELL CLONES AND PLASMA CELLS Memory B cell clones formed in GC responses can persist and produce antibody during the life of an animal or even in successive generations on cell transfer (Askonas and Williamson, 1972; MacLennan et al., 1990). This applies to inert antigens like tetanus toxoid or hapten protein, as well as to renewable sources of antigen such as viruses. In the former case, GC lasts for only a few weeks but antibody production can last indefinitely. As discussed earlier, this is in part attributable to long-lived plasma cells (Manz et al., 1999; Slifka and Ahmed, 1998) or committed post-GC plasma cell precursors (O’Connor et al., 2002). Nevertheless, in the absence of antigen localized on FDC, neither memory B cells nor antibody levels are sustained at normal
levels (Barrington et al., 2002). In addition, there is evidence that memory B cells will respond to antigen localized on FDC and mature to antibody-producing cells (Gray, 1988; MacLennan et al., 1990; Vonderheide and Hunt, 1990). Small numbers of transferred high-affinity memory B cells generate plasma cells that compete with and displace lower affinity host plasma cells (MacLennan et al., 1990). Indirect evidence suggests that T-dependent memory B cell activation, driven by antigen on FDC, continues at low levels for extended periods. This may be important for sustaining high levels of antibody production and both B and T cell memory, but direct evidence on this process is required. The transition of plasmablast to plasma cell in extrafollicular responses was considered earlier. Although CD11chigh dendritic cells appear important for this process the longterm survival of plasma cells in the spleen appears to occur adjacent to red pulp blood vessels and contiguous fibrous bands. The nature of signals that sustain the long-term survival of plasma cells from follicular or extrafollicular origin in these sites is unclear. There is a considerable volume of published work about the homing of plasmablasts emigrating from follicles to the lamina propria of the gut and the bone marrow. This large subject and the nature of the stroma in these sites that sustains antibody production is not considered in this review.
CONCLUSION In response to antigen, B cells move through multiple microenvironments on their way to becoming plasma cells. In each site they come in contact with distinct cells and stroma. Our understanding of the signals that influence B cells on this journey is far from complete. These influence the amount, affinity, and class of antibody that is produced and the length of time antibody is available. Consequently, understanding these processes is important for the control of clinical situations in which either insufficient or too much antibody is produced or the body is being harmed by autoantibodies. It is also critical if we are to understand the aberrant survival and expansion of neoplastic B cell and plasma cell clones and are to curtail by specific therapy the damage these cause.
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Qin, D., Wu, J., Vora, K. A., Ravetch, J. V., Szakal, A. K., Manser, T., and Tew, J. G. (2000). Fc gamma receptor IIB on follicular dendritic cells regulates the B cell recall response. J Immunol 164, 6268–6275. Reimold, A. M., Iwakoshi, N. N., Manis, J., Vallabhajosyula, P., Szomolanyi-Tsuda, E., Gravallese, E. M., Friend, D., Grusby, M. J., Alt, F., and Glimcher, L. H. (2001). Plasma cell differentiation requires the transcription factor XBP-1. Nature 412, 300–307. Retter, M. W., and Nemazee, D. (1998). Receptor editing occurs frequently during normal B cell development. J Exp Med 188, 1231–1238. Rogers, P. R., Dubey, C., and Swain, S. L. (2000). Qualitative changes accompany memory T cell generation: faster, more effective responses at lower doses of antigen. J Immunol 164, 2338–2346. Sallusto, F., Lenig, D., Forster, R., Lipp, M., and Lanzavecchia, A. (1999). Two subsets of memory T lymphocytes with distinct homing potentials and effector functions. Nature 401, 708–712. Schaerli, P., Willimann, K., Lang, A. B., Lipp, M., Loetscher, P., and Moser, B. (2000). CXC chemokine receptor 5 expression defines follicular homing T cells with B cell helper function. J Exp Med 192, 1553–1562. Shaffer, A. L., Lin, K. I., Kuo, T. C., Yu, X., Hurt, E. M., Rosenwald, A., Giltnane, J. M., Yang, L., Zhao, H., Calame, K., and Staudt, L. M. (2002). Blimp-1 orchestrates plasma cell differentiation by extinguishing the mature B cell gene expression program. Immunity 17, 51–62. Shortman, K., and Liu, Y. J. (2002). Mouse and human dendritic cell subtypes. Nat Rev Immunol 2, 151–161. Slifka, M. K., and Ahmed, R. (1998). Long-lived plasma cells: A mechanism for maintaining persistent antibody production. Curr Opin Immunol 10, 252–258. Smith, K. G., Hewitson, T. D., Nossal, G. J., and Tarlinton, D. M. (1996). The phenotype and fate of the antibody-forming cells of the splenic foci. Eur J Immunol 26, 444–448. Spencer, J., Finn, T., Pulford, K. A., Mason, D. Y., and Isaacson, P. G. (1985). The human gut contains a novel population of B lymphocytes which resemble marginal zone cells. Clin Exp Immunol 62, 607–612. Stein, H., Bonk, A., Tolksdorf, G., Lennert, K., Rodt, H., and Gerdes, J. (1980). Immunohistologic analysis of the organization of normal lymphoid tissue and non-Hodgkin’s lymphomas. J Histochem Cytochem 28, 746–760. Szakal, A. K., Kosco, M. H., and Tew, J. G. (1989). Microanatomy of lymphoid tissue during humoral immune responses: structure function relationships. Annu Rev Immunol 7, 91–109. Sze, D. M., Toellner, K. M., Garcia de Vinuesa, C., Taylor, DC. R., and MacLennan, I. C. (2000). Intrinsic constraint on plasmablast growth and extrinsic limits of plasma cell survival. J Exp Med 192, 813–821. Tew, J. G., and Mandel, T. E. (1979). Prolonged antigen half-life in the lymphoid follicles of specifically immunized mice. Immunology 37, 69–76. Tew, J. G., Mandel, T. E., Phipps, R. P., and Szakal, A. K. (1984). Tissue localization and retention of antigen in relation to the immune response. Am J Anat 170, 407–420. Thorley-Lawson, D. A. (2001). Epstein-Barr virus: Exploiting the immune system. Nat Rev Immunol 1, 75–82. Timens, W., Boes, A., Rozeboom-Uiterwijk, T., and Poppema, S. (1989). Immaturity of the human splenic marginal zone in infancy. Possible contribution to the deficient infant immune response. J Immunol 143, 3200–3206. Tkachuk, M., Bolliger, S., Ryffel, B., Pluschke, G., Banks, T. A., Herren, S., Gisler, R. H., and Kosco-Vilbois, M. H. (1998). Crucial role of tumor necrosis factor receptor 1 expression on nonhematopoietic cells for B cell localization within the splenic white pulp. J Exp Med 187, 469–477. Toellner, K. M., Gulbranson-Judge, A., Taylor, D. R., Sze, D. M., and MacLennan, I. C. (1996). Immunoglobulin switch transcript production in vivo related to the site and time of antigen-specific B cell activation. J Exp Med 183, 2303–2312. Toellner, K. M., Jenkinson, W. E., Taylor, D. R., Khan, M., Sze, D. M., Sansom, D. M., Vinuesa, C. G., and MacLennan, I. C. (2002). Lowlevel hypermutation in T cell-independent germinal centers compared
13. The Dynamic Structure of Antibody Responses with high mutation rates associated with T cell-dependent germinal centers. J Exp Med 195, 383–389. Toellner, K. M., Luther, S. A., Sze, D. M., Choy, R. K., Taylor, D. R., MacLennan, I. C., and Acha-Orbea, H. (1998). T helper 1 (Th1) and Th2 characteristics start to develop during T cell priming and are associated with an immediate ability to induce immunoglobulin class switching. J Exp Med 187, 1193–1204. Tourigny, M. R., Ursini-Siegel, J., Lee, H., Toellner, K. M., Cunningham, A. F., Franklin, D. S., Ely, S., Chen, M., Qin, X. F., Xiong, Y., MacLennan, I. C., and Chen-Kiang, S. (2002). CDK inhibitor p18(INK4c) is required for the generation of functional plasma cells. Immunity 17, 179–189. Tschopp, J., Irmler, M., and Thome, M. (1998). Inhibition of fas death signals by FLIPs. Curr Opin Immunol 10, 552–558. van Eijk, M., Defrance, T., Hennino, A., and de Groot, C. (2001). Deathreceptor contribution to the germinal-center reaction. Trends Immunol 22, 677–682. Vinuesa, C., Gulbranson-Judge, A., Khan, M., O’Leary, P., Cascalho, M., Wabl, M., Klaus, G. G., Owen, M. J., and MacLennan, I. C. (1999). Dendritic cells associated with plasmablast survival. Eur J Immunol 29, 3712–3721. Vinuesa, C. G., Cook, M. C., Ball, J., Drew, M., Sunners, Y., Cascalho, M., Wabl, M., Klaus, G. G., and MacLennan, I. C. (2000). Germinal centers without T cells. J Exp Med 191, 485–494. Voigt, I., Camacho, S. A., de Boer, B. A., Lipp, M., Forster, R., and Berek, C. (2000). CXCR5-deficient mice develop functional germinal centers in the splenic T cell zone. Eur J Immunol 30, 560–567. Vonderheide, R. H., and Hunt, S. V. (1990). Immigration of thoracic duct B lymphocytes into established germinal centers in the rat. Eur J Immunol 20, 79–86.
NOTE: Chapter 13 was submitted in November 2002.
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Walker, L. S., Wiggett, H. E., Gaspal, F. M., Raykundaalia, C. R., Goodall, M. D., Toellner, K. M., and Lane, P. L. (2003). Established T cell-driven germinal center B cell proliferation is independent of CD28 signaling but is tightly regulated through CTLA-4. J Immunol 170, 91–98. Weih, D. S., Yilmaz, Z. B., and Weih, F. (2001). Essential role of RelB in germinal center and marginal zone formation and proper expression of homing chemokines. J Immunol 167, 1909–1919. Williamson, A. R., and Askonas, B. A. (1972). Senescence of an antibodyforming cell clone. Nature 238, 337–339. Wykes, M., Pombo, A., Jenkins, C., and MacPherson, G. G. (1998). Dendritic cells interact directly with naive B lymphocytes to transfer antigen and initiate class switching in a primary T-dependent response. J Immunol 161, 1313–1319. Xu, J., Foy, T. M., Laman, J. D., Elliott, E. A., Dunn, J. J., Waldschmidt, T. J., Elsemore, J., Noelle, R. J., and Flavell, R. A. (1994). Mice deficient for the CD40 ligand. Immunity 1, 423–431. Yavuz, S., Grammer, A. C., Yavuz, A. S., Nanki, T., and Lipsky, P. E. (2001). Comparative characteristics of mu chain and alpha chain transcripts expressed by individual tonsil plasma cells. Mol Immunol 38, 19–34. Yellin, M. J., Sippel, K., Inghirami, G., Covey, L. R., Lee, J. J., Sinning, J., Clark, E. A., Chess, L., and Lederman, S. (1994). CD40 molecules induce down-modulation and endocytosis of T cell surface T cell-B cell activating molecule/CD40-L. Potential role in regulating helper effector function. J Immunol 152, 598–608. Zhang, J., MacLennan, I. C., Liu, Y. J., and Lane, P. J. (1988). Is rapid proliferation in B centroblasts linked to somatic mutation in memory B cell clones? Immunol Lett 18, 297–299.
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14 Dynamics of B Cell Migration to and within Secondary Lymphoid Organs JASON G. CYSTER
ULRICH H. VON ANDRIAN
Howard Hughes Medical Institute and Department of Microbiology and Immunology, University of California San Francisco, San Francisco, California, USA
The Center for Blood Research and the Department of Pathology, Harvard Medical School, Boston, Massachusetts, USA
The evolution of rearranging antigen receptors led to the conundrum that antigen-specific cells would be exceedingly rare. For these rare cells to be useful, they either needed to have antigens brought to them to sample, or they needed to survey the body for their specific antigen. Instead, the evolutionary outcome appears to be an intermediate between these extremes, with B lymphocytes surveying a subset of the body’s tissues, principally the secondary lymphoid organs, which themselves are specialized to concentrate and display antigens (Figure 14.1). Although each of the secondary lymphoid organs—the lymph nodes, spleen, and Peyer’s patches—filter antigens from only a portion of the body, B cells travel quickly between these organs and are able to survey most, if not all the lymphoid organs multiple times in their several month lifespan. In this chapter, we describe how naïve B cells migrate from the blood into the secondary lymphoid organs. We discuss what is known about their movement to the B cell zones, or follicles, within these tissues so that they can survey for antigen and how they relocate upon antigen encounter to favor their chance of interacting with helper T cells. Specialized subsets of B cells exist that do not follow the major migration pathways of conventional B cells, including marginal zone B cells in the spleen and B1 cells in the body cavities. Although these cell types will receive special attention in separate chapters, it will be useful to compare their migration properties in this chapter with those of follicular B cells. Following Tdependent immune responses, memory B cells and antibody secreting cells are produced. Differences in the trafficking properties of naïve and memory B cells is discussed. Differentiation into antibody-secreting cells leads to still further migrational reprogramming, and some of these cells localize in distinct subcompartments of secondary lymphoid
organs from B cell follicles, others go to mucosal surfaces, and others make a final journey back to the place where they were born, the bone marrow. The cues directing B-lineage cells on their final trek will be the subject of the last section in this chapter.
Molecular Biology of B Cells
LYMPHOID ORGAN ENTRY Following release into the blood from their site of production, the bone marrow, most newly produced B lymphocytes migrate first through the spleen, only later having a chance to experience the inside of a lymph node (LN) or a Peyer’s patch (PP). Contrary to this physiological ordering of events, we begin this section with a description of the steps involved in entry to LNs and PPs as our understanding of this process is more complete. This will be followed by a discussion of B cell entry into the spleen.
Entry via HEV into Secondary Lymphoid Organs The migration of naïve B cells from blood into LNs and PPs occurs via specialized postcapillary venules, known as high endothelial venules (HEVs) because of the thick, cuboidal shape of their endothelial cell lining (Butcher and Picker, 1996). The mechanism by which blood leukocytes attach to and transmigrate across endothelial cells has been worked out in most detail from studies of neutrophil attachment to inflamed endothelium and T cell attachment to HEVs, although in cases where B cells have been tracked, all the studies indicate that they abide by the same general rules as other leukocytes (Butcher and Picker, 1996). These
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FIGURE 14.1 Secondary lymphoid tissue organization and lymphocyte trafficking. Secondary lymphoid tissues function to bring together recirculating lymphocytes and antigen, with each lymphoid tissue sampling a different portion of the body’s fluids for the presence of antigen or antigen-presenting dendritic cells (DCs). The diagrams of a lymph node cross-section, a splenic white-pulp cord, and some surronding red pulp (accounting for about one fifth of a spleen crosssection), and a Peyer’s patch cross-section aim to show the themes common to all secondary lymphoid organs, with naïve lymphocytes gaining entry from the blood, and B cells and T cells quickly migrating into their separate subcompartments (dashed black arrows). B cells migrate to follicles in response to CXCL13 made by follicular stromal cells, whereas T cells localize within T zones in response to CCL21 and CCL19 made by T zone stromal cells. Within these compartments the cells undergo random walks to survey for intact antigen or MHC-peptide complexes, respectively. Each organ has areas rich in macrophages (indicated in purple shading) that capture and degrade antigen. The diagrams also illustrate key specializations of the tissues: the presence of a greater proportion of B cells (brown areas) than T cells (blue areas) in spleen and PPs, but not in lymph nodes; entry into LN and PP occurs via HEV, whereas entry into the spleen is by release from open-ended terminal arterioles (ta), many of which open into the marginal sinus (ms); antigen and antigen-bearing DCs arrive in LNs via afferent lymph fluid, whereas in the spleen antigens arrive via the blood, and DCs may arrive via this route. There is also a large population of immature DCs already present in the spleen (near the bridging zone); in PPs, antigen is transported by M cells directly to the subepithelial dome (sed), a region overlying the follicles that contains immature DCs and macrophages. Naïve B lymphocytes exit each of the lymphoid tissues (green arrows) after about one day, exiting via lymphatics from LNs and PPs or via red-pulp venous sinsusoids in the spleen. The lymphatics draining the PPs ferry cells to the mesenteric LNs. LN efferent lymphatics return cells to the blood via the thoracic duct, from where the cells can quickly gain entry to another secondary lymphoid organ in the ongoing process of lymphocyte recirculation. In addition to the populations of recirculating B cells, the spleen contains a more sessile population, the marginal zone B cells, located in the marginal zone. Intact antigen reaches lymphoid tissues in fluid phase and may also be carried in association with cells. Immune complexes can become trapped and displayed for long-periods on FDCs (a subset of follicular stromal cells), but other types of antigen transport cells (possibly DCs) may be involved in directly releasing antigen for recognition by B cells. Upon B cell activation by T-dependent antigens, germinal centers form within the B cell follicles, and antibody secreting cells (ASCs) migrate to the red-pulp of spleen or the medullary cords of LNs; in the case of PPs, many ASCs are released via the lymphatics and appear in the mesenteric LNs as well as homing to the gut. See color insert.
studies led to what is commonly referred to as the multistep model of leukocyte transmigration. In its simplest version (Figure 14.2), the model involves four steps: first, cells undergo low-affinity tethering interactions that are mediated by selectin–ligand, and in some cases integrin–ligand interactions. The shear force exerted on the cells by blood flow ensures that the weakly tethered cells roll along the endothelium. The rolling cell reaches sufficient proximity with the endothelium to receive a pertussis toxin (PTX) sensitive Gprotein coupled receptor (GPCR) signal that triggers integrin activation. Integrins engage ligands on the endothelium,
mediating the firm adhesion and arrest of the cell. Finally, the cell undergoes transmigration, or diapedesis, across the endothelium. Step 1: Rolling Interactions The major selectin involved in the initial tethering and rolling interaction of B lymphocytes, as for T cells, is Lselectin (CD62L). Selectins are calcium-dependent (C-type) lectins, and the ligands for L-selectin are mucin type glycoproteins that display highly modified carbohydrate groups.
14. Dynamics of B Cell Migration to and within Secondary Lymphoid Organs
FIGURE 14.2 Rolling, triggering, and adhesion requirements during B cell interaction with HEV in secondary lymphoid organs. Requirements are indicated for peripheral LN, mesenteric LN, and Peyer’s patches. The receptor–ligand pair that plays the dominant role at each step in each lymphoid organ is shown at the top of each list. Receptor–ligand pairs that make only minor contributions to an interaction are shown in smaller font size. In addition to CCL21, CCL19 may function as a triggering ligand for CCR7. Color code: brown, rolling cell; red, cell experiencing chemokine triggered integrin activation; blue, adherent cell. The corresponding molecular requirements for these steps are shown in the same color. See color insert.
Modifications necessary for L-selectin binding include sialylation, fucosylation, and sulfation (Rosen, 1999). Collectively, the principal ligands recognized by L-selectin are known as peripheral node addressin (PNAd). These Lselectin ligands are also recognized by an antibody that neutralizes L-selectin binding sites, MECA79. The molecules that carry the appropriately modified carbohydrates include CD34, glycam-1, podocalyxin, and Sgp200 and, in mucosal lymphoid tissues, MAdCAM1 (Rosen, 1999). The L-selectin–PNAd interaction has very fast on- and off-rates, a property that is important to the ability of this receptor–ligand system to mediate the tethering of fast moving cells and to subsequently support their rolling on the endothelium. Although PNAd expression is highest in peripheral lymph nodes, there is also expression in mucosal lymph nodes and weak expression in Peyer’s patches. Concordant with this expression pattern, short-term transfer experiments revealed that L-selectin–deficiency causes a 95% decrease in B cell entry to peripheral LNs, an 86% decrease in entry to mesenteric LNs, and an 80% reduction in homing to PPs (Tang et al., 1998). The importance of the appropriate carbohydrate modification of L-selectin ligands for normal lymphocyte trafficking is indicated by findings in mice lacking carbohydrate-modifying enzymes. Deficiency in high endothelial cell (HEC)-GlcNAc-6-sulfotransferase causes a marked reduction in lymphocyte trafficking to lymph nodes, although some L-selectin function is still observable in these animals (Hemmerich et al., 2001). Similarly, the importance of fucosylation in lymphocyte transmigration across HEV was demonstrated by the genetic disruption of two fucosyl (Fuc) transferases in mice, Fuc-T IV and Fuc-TVII (Homeister et al., 2001; Maly et al., 1996).
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B cells enter lymph nodes and Peyer’s patches with a lower efficiency than T cells, and this may in part be due to B cells having two-fold lower levels of L-selectin than T cells and undergoing fewer rolling interactions on HEVs (Okada et al., 2002; Tang et al., 1998). Studies in mice with a heterozygous L-selectin deficiency demonstrated that a two-fold reduction in L-selectin levels causes a 50 to 70% decrease in the efficiency of T cell homing to LNs (Tang et al., 1998). Although L-selectin is the major receptor on the lymphocyte required for tethering and rolling interactions, flow chamber and intravital studies have established that the a4-containing integrins, a4b7 and a4b1, are able to support rolling interactions on the ligands MAdCAM1 and VCAM1 (Alon et al., 1995; Berlin et al., 1995; Mazo et al., 1998; Sriramarao et al., 1996). Thus, the small number of remaining rolling interactions that occurred with L-selectin deficient cells were mostly blocked by antibodies to a4integrins (or to MAdCAM1). Analysis of wildtype cells that had been treated with a4-blocking antibodies, or of transferred b7 knockout cells, revealed that the average rolling velocity of cells within PPs was higher than with untreated or wildtype cells, thus demonstrating that a4b7 integrin–ligand interactions help to slow rolling cells, acting as a “bridge” between high-speed selectin-supported rolling and the triggering/firm adhesion steps (Bargatze et al., 1995; Berlin et al., 1995; Wagner et al., 1996). In contrast to a4 integrins, LFA1 does not appear to mediate this function on lymphocytes (Alon et al., 1995; Berlin et al., 1995; Warnock et al., 1998). One explanation for this difference is that a4integrins are localized to the tips of microvilli, together with L-selectin, whereas LFA1 is mostly concentrated on the cell body (Berlin et al., 1995). The contribution of a4integrin–ligand interactions to lymphocyte rolling is evident in mucosal LNs and PPs, but this pathway has not been shown to play a role in peripheral LNs, where PNAd is expressed at its highest levels (Hamann et al., 1988; Warnock et al., 1998). A small amount of L-selectin– independent rolling is observed in peripheral LNs, but this appears to be mediated by a second selectin, P-selectin, as it is completely blocked by treatment with anti-P selectin antibody (Diacovo et al., 1998). Step 2: Chemokine Triggering of Integrin Activation Following initial rolling interactions, a triggering event is necessary for cells to undergo firm integrin-mediated adhesion. The importance of this event is evident from the failure of lymphocytes treated with pertussis toxin (PTX), an inhibitor of gai signaling, to enter LNs and PPs (Huang et al., 1989). Intravital microscopy experiments of PTXtreated lymphocytes within murine Peyer’s patch or inguinal LN HEVs demonstrated that PTX did not affect the number of cells undergoing rolling events, but prevented the transi-
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tion from rolling to firm adhesion (Bargatze and Butcher, 1993; Warnock et al., 1998). Although these experiments did not distinguish between B and T cells, other studies measuring the frequency of B and T cell accumulation in LNs indicated that entry of both cell types was completely blocked by PTX treatment (Cyster and Goodnow, 1995a). The identification of a Gai signaling requirement for induction of firm adhesion implicated the involvement of chemokines at this step. In the case of T lymphocytes, a single major triggering chemokine receptor has been identified, CCR7; and in mice lacking either CCR7 or deficient in two CCR7 ligands, CCL21-ser/SLC-ser and CCL19/ELC (Table 14.1), few T cells enter LNs and PPs (Förster et al., 1999; Gunn et al., 1998; Nakano et al., 1997). Intravital microscopy experiments established that T cells with diminished ability to respond to CCL19 and CCL21 were strongly inhibited in their ability to undergo firm adhesion with the endothelium (Stein et al., 2000; Warnock et al., 2000). In the mouse, at least two genes have been identified that encode CCL21, CCL21-ser, and CCL21-leu (Nakano and Gunn, 2001; Vassileva et al., 1999). CCL21-ser is expressed by HEVs in LNs and PPs, as well as in the network of surrounding T zone stromal cells. CCL21 protein was detected on the lumenal surface of HEV as well as within the T zone (Gunn et al., 1998). CCL21-leu is expressed by lymphatic endothelium outside lymphoid tissues and is expressed little, if at all, within lymphoid organs (Vassileva et al., 1999). CCL19 is not expressed by HEV, but this CCR7 ligand is made by the surrounding stromal cells and can be displayed on HEVs in a functional form (Baekkevold et al., 2001; Luther et al., 2000; Ngo et al., 1998). The relative importance of CCL21 and CCL19 at the step of lymphocyte attachment to HEVs remains to be established.
The chemokine and chemokine receptor requirements for B cell entry to LNs and PPs is more complicated than for T cells. CCR7- or CCR7-ligand deficiency causes about a 50% reduction in B cell entry into LNs in short-term transfer experiments and has somewhat less effect on entry to PPs (Förster et al., 1999; Nakano et al., 1998; Okada et al., 2002). An examination of possible contributions made by other chemokine receptors expressed on B lymphocytes demonstrated that CXCR4 and its ligand, CXCL12/SDF1 (Table 14.1), contribute to B cell attachment to HEVs in LNs and PPs (Okada et al., 2002). Intravital microscopy of inguinal LN HEVs with B cells that had been treated with CXCL12 and CCL19 to desensitize their CXCR4 and CCR7 receptors, respectively, revealed that the cells underwent normal numbers of rolling interactions but were defective in their ability to undergo the transition from rolling to firm adhesion (Okada et al., 2002). Although CXCL12 does not appear to be expressed by HEVs, cells expressing this chemokine are present in close association with most HEVs in LNs and in the T zone of PPs, and CXCL12 protein can be detected on the lumen of HEVs (Okada et al., 2002). In contrast to the 90% inhibition of B cell homing to LNs, homing to PPs was only 50% affected by combined CXCR4deficiency and CCR7-ligand deficiency. A third chemokine receptor, CXCR5, was found to participate in B cell entry to PPs. The ligand for CXCR5, CXCL13/BLC (Table 14.1), was identified on HEV within PP follicles and within human tonsil, but not on HEV in T cell areas (Okada et al., 2002; Schaerli et al., 2000). Reciprocally, CCL21 was detected on PP T zone HEV but not on follicular HEV (Warnock et al., 2000). In accord with the pattern of ligand expression, B cells lacking CXCR5 fail to adhere to HEV within PP follicles while adhering with normal efficiency to T zone HEV
TABLE 14.1 Chemokines involved in directing B cell movements Chemokine† CXCL9 (MIG) CXCL10 (IP10) CXCL11 (ITAC)
Receptor
Chemokine distribution
Guidance function for B-lineage cells*
CXCR3
Sites of inflammation (IFNg induced), inflamed lymphoid tissue
Pre-pro-B cells; ASC homing
CXCL12 (SDF1)
CXCR4
BM, near HEV, RP, MCs, epithelium, other
BM retention, HEV attachment, ASC homing
CXCL13 (BLC, BCA1)
CXCR5
Follicles, body cavities
Follicular homing, body cavity homing/retention, HEV attachment
CCL20 (MIP3a, LARC)
CCR6
Inflamed epithelium, M cells
Memory B trafficking
CCL19 (ELC, MIP3b) CCL21 (SLC, 6Ckine)¥
CCR7
T zone, HEV (CCL21), lymphatics (CCL21)
HEV attachment, localization at T-B boundary
CCL25 (TECK)
CCR9
Epithelium of SI
Pre-pro-B in BM; IgA ASC homing to SI
CCL28 (MEC)
CCR10
Epithelium in stomach, intestine, salivary gland, mammary gland, trachea
IgA ASC homing
† Chemokines are shown by their standardized name and, in parentheses, by frequently used common names. Some of the chemokines have additional common names that could not be listed due to space limitations. * See text for details and citations. In addition: progenitor B cells have been reported to respond to CXCL9 and CCL25, and CCR9-deficient mice show reduced numbers of pre-pro-B cells in the BM (Bowman et al., 2000; Wurbel et al., 2001). ¥ Two CCL21 genes that encode proteins differing by a single amino acid have been defined in BALB/c mice, termed CCL21-ser and CCL21-leu; in some mouse strains there is an additional copy of the CCL21-leu gene (Vassileva et al., 1999; Nakano et al., 2001). Only a single CCL21 gene has been identified in humans. ASC, antibody secreting cell; RP, splenic red pulp; MCs, lymph node medullary cords; SI, small intestine.
14. Dynamics of B Cell Migration to and within Secondary Lymphoid Organs
(Okada et al., 2002). T cells, which are mostly CXCR5negative, fail to adhere in PP follicular HEV (Warnock et al., 2000). A further implication of these observations is that B cells will have a greater extent of HEV surface area by which they can enter PPs than T cells, and this might be expected to increase their efficiency of entry. Indeed, although T cells enter LNs and PPs with greater efficiency than B cells, the relative efficiency of B cell homing is greater in PPs than in LNs (Okada et al., 2002; Tang et al., 1998). Step 3: Integrin-Mediated Firm Adhesion Chemokine-triggered firm adhesion of rolling cells depends on interactions between integrins on the lymphocyte and ligands on the HEV. The integrin–ligand requirements for adhesion to HEVs have been explored in the Stamper-Woodruff frozen tissue-section adhesion assay, by in vivo antibody blocking experiments and, most recently, using cells from gene-targeted mice (Salmi and Jalkanen, 1997; von Andrian and Mackay, 2000). From these studies, three integrins have been shown to function in homeostatic trafficking of naïve lymphocytes, LFA1 (CD11a/CD18, aLb2), a4b1 (VLA4), and a4b7, although the contribution of each integrin differs in different types of secondary lymphoid organ. In peripheral LNs, LFA1 accounts for 80 to 95% of the integrin requirement for both T and B lymphocytes (Andrew et al., 1998; Berlin-Rufenach et al., 1999; Hamann et al., 1988). The LFA1 ligand ICAM-1 is highly expressed by HEV and functions in lymphocyte–HEV adhesion (Faveeuw et al., 2000; Lawrence et al., 1995; Schneeberger et al., 2000). A second LFA1 ligand, ICAM2, is expressed on vascular endothelium throughout the body, including HEVs, and recent experiments indicate that ICAM-2 works together with ICAM-1 to support lymphocyte adhesion and entry into LNs (Gerwin et al., 1999; Lehmann et al., 2003). An assessment of the integrins responsible for the remaining peripheral LN homing of LFA-deficient lymphocytes identified minor roles for a4b7 and a4b1, with VCAM-1 serving as the principal a4-integrin ligand (Berlin-Rufenach et al., 1999). In mesenteric LNs, LFA1 and a4b7 contribute almost equally to the integrin requirement for lymphocyte adhesion to HEV. MAdCAM1, rather than VCAM1, functions as the key a4-integrin ligand expressed on the HEV (BerlinRufenach et al., 1999) (Figure 14.2). In Peyer’s patches, a4b7-MAdCAM1 interactions are critical and account for the majority of the integrin–ligand requirement for lymphocyte homing, with LFA1 making ~30% of the integrin contribution and the a4-integrin ligand, VCAM1, playing a minor role (Bargatze et al., 1995; Berlin-Rufenach et al., 1999; Wagner et al., 1996). It should be kept in mind that these studies have mostly been performed with total lymphocyte populations, and the degree to which the higher total
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levels of a4-containing integrins and lower levels of LFA1 on B cells compared to T cells (Schmits et al., 1996; and unpublished observations) contribute to differences in B and T cell homing remains to be seen. Step 4: Transendothelial Migration The final steps in B lymphocyte entry to a lymphoid organ involve migration along the endothelium to a nearby border between endothelial cells and then squeezing of the lymphocyte between endothelial cells in an ameboid manner. The molecular events associated with this transendothelial migration step are only beginning to be defined. CD31/PECAM1, a homophilic adhesion molecule that is localized to tight junctions between endothelial cells and is also expressed on leukocytes, participates in some transendothelial migration events involving neutrophils, monocytes, and NK cells but has not been found to have a role in lymphocyte homing (Aurrand-Lions et al., 2002; Duncan et al., 1999). Recently, a new subfamily of Ig-superfamily molecules have been identified, known as the junctional adhesion molecules (JAMs) (Muller 2003). These may participate in transendothelial migration events. In particular, JAM-A can function as a ligand for LFA-1, and antibodies to JAM-A inhibit T cell and monocyte transmigration across endothelial cell layers in vitro (Martin-Padura et al., 1998; Ostermann et al., 2002). The expression of JAM-A, JAM-B, and JAM-C has been identified on HEV (AurrandLions et al., 2001; Palmeri et al., 2000). In vitro studies also suggest a role for human JAM-B in lymphocyte transendothelial migration, possibly through homophilic binding to JAM-B on lymphocytes or through heterophilic interactions with lymphocyte JAM-C or a4b1 (Arrate et al., 2001; Cunningham et al., 2002; Liang et al., 2002; JohnsonLeger et al., 2002). Conventional integrin–ligand interactions may also contribute to lymphocyte transendothelial migration. CD99, a heavily O-glycosylated molecule present on leukocytes and at endothelial cell junctions, functions as a homophilic adhesion molecule in monocyte transmigration, acting at a step following initial CD31 interactions (Aurrand-Lions et al., 2002; Schenkel et al., 2002). It remains to be established whether CD99 functions at HEVs in the process of homeostatic lymphocyte trafficking, although it is notable that CD99 is expressed on B and T lymphocytes (Park et al., 1999; Schenkel et al., 2002). Metalloproteases play important roles in neutrophil migration to inflamed tissues, and these enzymes also appear to have a role in lymphocyte homing because in vivo treatment with soluble metalloprotease inhibitors reduces the efficiency of lymphocyte transmigration across HEV (Faveeuw et al., 2001). One role of metalloprotease activity during lymphocyte homing is thought to be the cleavage of L-selectin (Faveeuw et al., 2001). Although it is often considered that a chemokine gradient across the endothelium
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may be needed to promote directed transmigration of a lymphocyte as it makes its way across a HEV, in vitro studies with T lymphocytes have shown that while a gai signal is needed for efficient transendothelial migration, a gradient is unnecessary (Cinamon et al., 2001). Instead, it seems that the information encoded by the endothelial junction provides sufficient directionality to the cell. Notably, exertion of shear forces on cells in vitro augments their ability to undergo transmigration, raising the possibility that mechanoreceptors are involved in this process (Cinamon et al., 2001).
Entry to the Spleen The spleen is a major secondary lymphoid organ and contains the largest single B cell population in the body and a unique reservoir of nonrecirculating B cells known as marginal zone B cells. In contrast to most other organs, the spleen has an open blood circulation. The large vessels that carry blood to the spleen rapidly branch to form arterioles, many of which are ensheathed by cords of lymphocytes to form areas known as white-pulp cords. Because of their location in the white pulp, these vessels are termed central arterioles (Figure 14.3). Terminal arterioles arise from central arterioles, and many of these vessels open into an
FIGURE 14.3 B cell distribution in the mouse spleen. Cryostat section of unimmunized mouse spleen stained in brown to detect IgD and in blue to detect IgM. Naïve, recirculating B cells appear brown (IgDhiIgMint) whereas marginal zone (MZ) B cells appear blue (IgDloIgMhi). The image encompasses about one fourth of the tangential cross-section and shows a large white-pulp cord centered around a central arteriole (ca) with two large B cell follicles (brown, labeled), two smaller follicles (brown), and central unstained T zones (white). The marginal zone (MZ) surrounds the B cell follicles, separated in the mouse by the marginal sinus (MS), a site where many small arterioles terminate. Gaps in the MZ are observed at the edges of the follicles, regions often referred to as MZ “bridging zones” (one of these is labeled). IgM ASCs (intense blue staining) can be seen in the bridging zones and also in clusters within the red pulp. The scattering of B cells (brown) within the red pulp may include recirculating B cells that are passing out of the spleen as well as cells resident in this area. See color insert.
area that immediately surrounds the follicular regions of the white-pulp cords, known as the marginal zone (Figures 14.1 and 14.3). Smaller numbers go beyond this zone and terminate within the splenic red pulp. Blood is released from the terminal arterioles, and many of the blood cells pass quickly from the site of release through the marginal zone or red pulp and into venous sinuses. These large, porous vessels anastomose to form splenic veins that then carry splenic blood back into the circulation. Through the process of acting as a blood-filtering device, the spleen contributes to the removal of effete red blood cells and serves as a site for bringing together lymphoid cells, antigen-presenting cells, and blood-borne antigens. Like other blood cells, many of the lymphocytes entering the spleen are released from terminal arterioles that open into the marginal zone (Brelinska and Pilgrim, 1982; Ford, 1969; van Ewijk and Nieuwenhuis, 1985) (Figure 14.3) and some of these cells pass to the outer region of the marginal zone and then to the red pulp or directly into venous sinuses. In contrast to all other blood cell types, a fraction of the lymphocytes take a different route and quickly begin appearing within the B and T cell areas of the white pulp cords (Nieuwenhuis and Ford, 1976). Entry into the white pulp is blocked by pertussis toxin (PTX) pretreatment (Cyster and Goodnow, 1995a; Lyons and Parish, 1995), establishing a requirement for gai signaling and implicating chemokines at this step. Deficiency in CXCR5 strongly reduces B lymphocyte accumulation within white-pulp cords, and CCR7deficiency reduces T cell accumulation in these areas (Förster et al., 1996, 1999). Small numbers of B cells do continue to appear within the white pulp of mice deficient in CXCR5, or its ligand, CXCL13 (Table 14.1), possibly due to their ability to respond weakly to the T zone chemokines CCL19 and CCL21 (Ngo et al., 1998). More recently, a requirement was identified for integrins in lymphocyte entry to splenic white-pulp cords. Combined inhibition of LFA1 and a4b1 was associated with greater than 90% inhibition in B cell migration into white-pulp cords (Lo et al., 2002). Blocking of LFA1 alone caused about 50% inhibition in B cell entry to the white pulp, whereas a4-blocking antibodies were insufficient to reduce entry. ICAM1 serves as a key LFA1 ligand involved in entry whereas the a4b1 ligand, VCAM1, accounts for part of the a4b1 ligand requirement. Both ICAM1 and VCAM1 are expressed at high levels throughout the splenic marginal zone (Lu and Cyster, 2002). MAdCAM1, a marker of the marginal sinus in the mouse spleen (Kraal et al., 1995), and a4b7 are not required for B cell migration into splenic white-pulp cords (Kraal et al., 1995; Lo et al., 2002). A comparison of early events following B cell transfer revealed that inhibition of Gai signaling with PTX and combined inhibition of LFA1 and a4b1 function blocked B cell homing at a similar early step: In both situations there was a reduction in the number of B cells associated with the inner edge of the marginal zone as
14. Dynamics of B Cell Migration to and within Secondary Lymphoid Organs
well as a block in their appearance within the white-pulp cords (Lo et al., 2002). Therefore, while lymphocyte entry into the spleen as a whole occurs by passive release from open-ended terminal arterioles, entry to the white-pulp cords is an active process that requires both Gai signaling and integrins.
COMPARTMENTALIZATION OF MATURE B CELLS Secondary Lymphoid Tissues Migration to Lymphoid Follicles Following entry from the blood into secondary lymphoid organs, many B cells migrate to lymphoid follicles. The chemokine CXCL13/BLC and its receptor, CXCR5, have been identified as an essential ligand–receptor pair necessary for this event (Table 14.1). CXCL13 and CXCR5 also function in an early step necessary for the development of many LNs and for the efficient generation of PPs (Ansel et al., 2000; Förster et al., 1996). However, the spleen and most mucosal LNs develop in the absence of this chemokine–receptor system, and anatomical characterization of these tissues in CXCR5- and CXCL13-deficient mice established that they lacked lymphoid follicles (Ansel et al., 2000; Förster et al., 1996). Furthermore, when CXCR5deficient B cells were transferred to wildtype recipients, the cells failed to localize within lymphoid follicles in spleen or lymph nodes. Within follicles, CXCL13 is made by radiation-resistant follicular stromal cells (Ansel et al., 2002; Cyster et al., 2000). In both mouse and human tissue, there is co-localization of follicular dendritic cell (FDC) markers and CXCL13, although this overlap usually does not appear complete. As CXCL13 is a secreted protein, it remains unclear whether FDC produce CXCL13 or whether they bind chemokine produced by other types of follicular stromal cell. In addition to the role of CXCL13/CXCR5 in B cell recruitment to follicles, CCR7 and its ligands have been suggested to influence the rate of B cell trafficking to follicles in the spleen, contributing to an initial tendency for cells to dwell in the outer regions of the T cell areas bordering with follicles (Förster et al., 1999), perhaps favoring early encounters between antigen-engaged cells and T cells. B lymphocytes are believed to migrate through lymphoid follicles primarily for surveillance purposes—to check for foreign antigen on the surface of FDCs. FDCs are able to capture and display antigen as C3d-antigen complexes and IgG-antigen complexes, via complement and Fc receptors, respectively. Two-photon microscopic analysis of B lymphocyte migration deep within intact lymph nodes revealed that B cells undergo continual migration within the follicle, following a random walk or roaming behavior (Miller
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et al., 2002). This extensive movement within the follicle is likely to facilitate the efficient surveillance of FDC processes for the presence of antigen. T cells undergo a similar behavior within the T zone, although the migration paths of T cells are longer and the cells move at approximately twice the speed of the B cells, perhaps reflecting differences in the requirements in surveying for MHCpeptide complexes versus intact antigens (Miller et al., 2002). A further compartment within the spleen, the marginal zone, contains a resident population of B cells with distinct properties to the major recirculating B cell population. This compartment is described later. In addition to follicles, lymph nodes contain B cell–rich areas that lack FDCs and CXCL13 expression, and that can be accessed by cells in a CXCR5-independent manner (Ansel et al., 2000). The function of these zones is not clear, although it is tempting to speculate that they favor interactions between B cells and antigen-bearing interdigitating dendritic cells or macrophages. On average, B lymphocytes spend about one day within a lymphoid tissue after which they exit and return to circulation (Ford and Simmonds, 1972). Relatively little is know about the pathways of lymphocyte exit, but in the spleen it is believed to involve transit to red-pulp venous sinuses, whereas in LNs, the cells most likely exit through medullary sinuses that then connect to efferent lymphatic vessels. Lymphocytes exiting Peyer’s patches travel via the lymphatics to the mesenteric lymph nodes before being returned again to the lymph and joining the blood circulation by way of the thoracic duct. During an immune response, lymphocyte transit through the responding lymphoid tissue is temporarily stopped and very few B or T lymphocytes appear within lymph during this “shut-down” period (Mackay et al., 1992). This process may contribute to the rapid enlargement of lymphoid tissues in the early phase of an immune response, a change that presumably helps increase the number of antigen-specific cells available in the lymphoid tissue to respond to the inflammatory stimulus. An immunosuppressive drug has been described, FTY720, that activates the shut-down process, preventing lymphocytes from exiting from lymph nodes and Peyer’s patches (Chiba et al., 1998). This drug may also affect lymphocyte homing at the level of HEV, since FTY720 treatment increases the frequency of CCR7deficient cells that enter LNs (Henning et al., 2001). The phosphorylated form of FTY720 has structural similarities to sphingosine-1-phosphate (S1P) and is active in stimulating four of the five known S1P receptors (Brinkmann et al., 2002; Mandala et al., 2002). It may act both to alter the properties of the lymphocytes and cause changes in the lymphatic endothelium and possibly in HEV. Its immunosuppressive effect may lie in its propensity to cause lymphocyte sequestration, thus limiting the ability of cells to attack transplanted tissues (Brinkmann and Lynch, 2002). In
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the limited studies so far performed, antibody responses appear to be largely unaffected by the drug, suggesting it does not affect the ability of lymphocytes to encounter antigen and undergo cognate interactions within lymphoid tissues (Brinkmann and Lynch, 2002).
body responses, and it is likely that these interactions are similarly favored by the antigen-induced changes in B cell localization and chemokine expression.
Marginal Zone and Memory B Cells Relocalization Following Activation
Marginal Zone B Cells
Following antigen engagement, B lymphocytes undergo rapid and marked changes in their migratory behavior, changes that are believed to favor their encounter with helper T cells and possibly also with accessory cells. Within 6 hours of B cell receptor (BCR) engagement, B cells move from follicular areas or possibly other sites of antigen encounter to the boundary of B and T cell zones in secondary lymphoid organs (Cyster and Goodnow, 1995b). This relocalization occurs whether the cell was naïve or memory (Liu et al., 1988) and seems to occur similarly in response to both T-dependent and T-independent antigens (Martin and Kearney, 2000) and also in response to self-antigens (Cyster et al., 1994; Mandik-Nayak et al., 1997). T cells are not required for the relocalization to occur (Schmidt and Cyster, 1999) and the mechanism involves BCR-induced changes in chemokine receptor level that cause a slight adjustment in chemokine sensitivity (Reif et al., 2002). Naïve B cells express both CXCR5 and CCR7 and exhibit a strong in vitro chemotactic response to CXCL13 and a weaker response to the CCR7 ligands (Gunn et al., 1998; Ngo et al., 1998). These in vitro observations agree well with the in vivo behavior of naïve cells in which, following entry into a region of the tissue near where the domain of CXCL13 expression abuts with the domain of CCR7 ligand expression, B cells preferentially migrate into the area of CXCL13 expression. Within hours of acute antigen exposure, B cells undergo a small increase in CCR7 expression. This confers an increase in the responsiveness of the cells to CCR7 ligands, a shift in the balance that appears to be sufficient for B cell relocalization to the outer T zone (Reif et al., 2002). Many other factors are likely to influence the efficiency of encounters with antigen-specific T cells. In particular, changes also take place in the chemokine responsiveness of activated helper T cells that help direct the cells towards B cell areas (Ansel et al., 1999; Breitfeld et al., 2000; Kim et al., 2001; Schaerli et al., 2000). Activated B cells produce several chemokines including CCL3/MIP1a, CCL4/MIP1b, and CCL22/MDC (Bystry et al., 2001; Glynne et al., 2000; Schaniel et al., 1998). These chemokines are efficacious attractants of activated T cells and may help promote encounters between B cells and T cells. Under some conditions, B cells may produce chemokines that favor the recruitment of regulatory T cells thus inhibiting or downregulating the B cell response (Bystry et al., 2001). Emerging evidence suggests that interactions between B cells and DCs are important during anti-
In addition to serving as a site of cell entry to the spleen, the marginal zone contains a population of resident B cells known as marginal zone B cells (Figure 14.3). These cells express a distinct pattern of cell surface molecules and are larger than follicular B cells. They respond more rapidly following exposure to antigen (Martin and Kearney, 2002). The MZ B cell repertoire is distinct from the follicular repertoire and is enriched in cells with germline encoded receptors specific for bacterial surface molecules, such as phosphorylcholine. Memory B cells generated during Tdependent antibody responses also contribute to the MZ B cell population (Liu et al., 1988; Shih et al., 2002). A striking feature of the MZ B cells, at least as studied in rodents, is that these cells do not recirculate but instead appear to be sessile within the MZ (MacLennan et al., 1982). The differentiation pathway of MZ B cells is not fully defined, and it is unclear what factors guide MZ B cells, or their precursors, to the MZ. CXCL13 is not expressed within the MZ and is not required for MZ B cell lodgement, but B cells can be displaced from the MZ by in vivo treatment with PTX (Guinamard et al., 2000), making it likely that a chemokine is involved. Within the MZ, integrin-mediated adhesion plays a critical role in B cell retention. MZ B cells express levels of LFA1 and a4b1 higher than follicular B cells, and antibodies that block the function of these integrins lead to displacement of MZ B cells from the MZ and their transient appearance in the blood (Lu and Cyster, 2002). ICAM1 and VCAM1, ligands for LFA1 and a4b1, respectively, are expressed within the MZ and both contribute to MZ B cell retention (Lu and Cyster, 2002). In addition to higher expression levels, other features of MZ B cells are likely to contribute to their elevated levels of functional integrins, such as their high expression of the integrininteracting 4-transmembrane protein, CD9 (Won and Kearney, 2002). Upon antigen-encounter in the MZ, memory B cells relocalize to the outer T zone in a similar fashion to the relocalization described for naïve B cells (Liu et al., 1988). By contrast, following exposure to LPS, MZ B cell migrate into the B cell follicle rather than to the outer T zone (MacLennan et al., 1982). The significance of this behavior is not fully established, but it has been suggested to serve as a mechanism for delivering antigens from sites of capture in the MZ into the B cell follicle for possible deposition on the FDC network and encounter by recirculating B cells (MacLennan et al., 1982). Migration into the B cell follicle
14. Dynamics of B Cell Migration to and within Secondary Lymphoid Organs
depends on CXCL13 and involves a decrease in integrinmediated adhesion (Lu and Cyster, 2002). However, the change in integrin-mediated adhesion is delayed compared to the rate of MZ B cell relocalization, and other changes in the cells are presumed to be required for the very rapid redistribution that takes place. Memory B Cells The splenic MZ of humans is anatomically more complex than its rodent counterpart (Satoh et al., 1997; Steiniger et al., 2001). Vh gene sequencing studies have established that many of the IgM+ cells in the human MZ are somatically mutated memory cells (Spencer et al., 1998; Tangye et al., 1998; Tierens et al., 1999). Studies in other human lymphoid tissues have identified MZ phenotype B cells (IgM+IgD-, complement receptor-positive, large size) distributed at the outer perimeter of follicles and extending into the dome region in PPs and beneath the subcapsular sinus of mesenteric LNs (Spencer et al., 1998; Tierens et al., 1999). Presently, it is unclear whether these cell populations are sessile, or whether they undergo some level of recirculation that keeps the various populations in communication. In keeping with the latter possibility, CD27 stains cells of the human MZ and is also a marker of memory B cells in the blood, and the IgM+ memory B cells in these two locations have similar extents of somatic mutations (Tangye et al., 1998). Perhaps the human MZ contains two types of MZ B cells, germline memory cells that don’t recirculate and classical memory cells that do. Although IgM+ B cells are identified within the MZ compartment, it is unlikely that all memory B cells are localized in this compartment. Many memory B cells express isotypes other than IgM, such as IgG or IgA, but there is little indication that these cells are concentrated within the MZ compartment. In human tonsil, isotype-switched memory B cells are identified in the subepithelial and intraepithelial areas (Liu et al., 1995). The mechanisms promoting this localization are not defined, although it is notable that CCL20/MIP3a (Table 14.1) is expressed in this region (Casamayor-Palleja et al., 2001), and the CCL20 receptor, CCR6, is expressed on memory B cells in a functional form (Krzysiek et al., 2000; Liao et al., 2002). CCL20 is also highly expressed in the M-cells associated with Peyer’s patches (Figure 14.1) and might be anticipated to influence memory B cell distribution in this compartment (Cook et al., 2000). In addition to CCL20, epithelial b-defensins can act as agonists for human CCR6, possibly also contributing to memory B cell accumulation near the epithelium (Yang et al., 1999). Experiments tracking the generation of long-term B cell memory following intestinal rotavirus infection of mice revealed that a4b7+ isotype switched memory B cells were concentrated in PPs (Youngman et al., 2002). a4b7 is
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uniformly expressed by naïve B cells, but expression on memory B cells is bimodal, consistent with the distinct trafficking patterns for different memory B cell subsets. Further evidence of memory B cell trafficking specialization comes from the discovery that a subset of isotype-switched memory B cells expresses E-selectin ligands (Rott et al., 2000; Yoshino et al., 1999). IgD- memory B cells from human tonsil exhibit upregulation of the fucosyltransferase Fuct-VII, an enzyme needed for the synthesis of E-selectin ligands (Maly et al., 1996; Montoya et al., 1999). Together, these observations support the view that, just as for memory/effector T cells (Kunkel and Butcher, 2002), memory B cells acquire a pattern of homing molecule expression that reflects their site of development and favors their accumulation in lymphoid tissues that collect antigens from similar anatomical compartments. Insight into the trafficking pattern of human memory B cells has come from a study tracking the distribution of Epstein Barr Virus (EBV)+ cells (Laichalk et al., 2002). Following EBV infection, latent virus is present in memory B cells but in few, if any, naïve B cells. The predominant site of B cell infection with EBV is in lymphoid areas of the oral cavity, known as Waldeyer’s ring and including the tonsil. EBV+ memory B cells recirculate from this area and are found in peripheral blood, spleen, and lymph nodes. However, long after infection, EBV-infected memory B cells are present at 20-fold higher concentrations in Waldeyer’s ring than in spleen or mesenteric LNs, providing evidence that memory B cells generated in Waldeyer’s ring preferentially home back to this compartment (Laichalk et al., 2002).
Body Cavity B Cells In addition to the major populations of B lymphocytes present in secondary lymphoid organs, small numbers of B cells are present in the peritoneal, pleural, and thoracic cavities (Hardy and Hayakawa, 2001). In mice, many of the body cavity B cells are of the B1 subset. B1 cells are a significant source of serum antibody, and they make a dominant contribution to the low-affinity IgM antibodies that are present in the serum of unimmunized mice, known as natural antibodies (Hardy and Hayakawa, 2001; Martin and Kearney, 2001). Studies in mice deficient in natural antibodies have established their critical role in providing early protection from a variety of pathogens (Hardy and Hayakawa, 2001; Martin and Kearney, 2001). The body cavities are lined by mesothelial cells, and the peritoneal cavity contains an additional bilayered mesothelial sheet known as the omentum (Williams and White, 1986). The omentum connects the spleen, pancreas, stomach, and transverse colon and is best characterized for its role in abdominal wound repair (Williams and White, 1986). Studies performed in the nineteenth century revealed the presence of cellular aggregates within the omentum
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and, because of their white appearance, these were termed “milk spots” (reviewed in (Williams and White, 1986)). These aggregates contain a mixture of macrophages and lymphocytes and smaller numbers of plasma cells and mast cells (Williams and White, 1986) (Figure 14.4). Surprisingly, the same chemokine that is needed for B cell lodgement in lymphoid follicles is also critical for B cell accumulation in the body cavities (Ansel et al., 2002). In mice lacking CXCL13, B1 and conventional B cell numbers in the peritoneum are more than tenfold and sixfold reduced, respectively (Ansel et al., 2002). CXCL13 is made by peritoneal macrophages and by radiation resistant cells within the omentum (Ansel et al., 2002). B1 cells express CXCR5 at somewhat higher levels than conventional B cells, and they are more responsive to CXCL13 (Ansel et al., 2002; Ishikawa et al., 2001). In transfer experiments, B1 cells showed a much greater propensity to home to the peritoneum than B2 cells, indicating that homing differences may contribute to the differential accumulation of B1 and B2 cells within the body cavities. In whole-mount microscopic analysis of the omentum taken from recipients early after transfer of fluorescently labeled B1 cells, the highest density of fluorescent cells was associated with vessels traversing omental milk spots (Ansel et al., 2002). A model has therefore been proposed in which B cells enter the omentum across vessels within milk spots and then some of the cells migrate from the milk spot via fenestrations in the overlying mesothelium into the peritoneal cavity (Ansel et al., 2002). Whether other mesothelial surfaces function in this way is not clear, although the detection of small numbers of milk spots within the diaphragm, the mediastinal pleura, and the pericardium make this a likely possibility (Doherty et al., 1995; Nakatani et al., 1988). Although B1 cells are predominantly located in the peritoneum, they are also found at low levels in the blood and spleen, thus suggesting that they undergo recirculation
FIGURE 14.4 Cross-sectional diagram of an omental milk spot. Milk spots lie in a double sheet of mesothelium and are made up predominantly of B cells and macrophages. They also contain fibroblasts and adipocytes. Mast cells and occassional T cells are also present (not shown). In the mouse, the majority of omental B cells are of B-1 phenotype. The capillary network within the milk spot is a site of attachment and entry of circulating B1 cells, and this depends on the chemokine CXCL13. B cells are likely to pass through the fenestrated mesothelium overlying milk spots to access the body cavity. The mesothelial basement membrane (not shown) is also discontinuous in areas overlying a milk spot. See color insert.
(Hardy and Hayakawa, 2001; Martin and Kearney, 2001). Indeed, several studies suggest that B1 cells participate in immune responses at sites outside the body cavities (Martin et al., 2001; Wardemann et al., 2002).They are also believed to give rise to antibody secreting cells in the gut (Fagarasan et al., 2001). In favor of the notion that B1 cells undergo recirculation, in parabiosis experiments where the blood circulation of pairs of mice were joined for a period of weeks, a gradual mixing of body cavity B1 cells occurred (Ansel et al., 2002). A well-developed lymphatic vasculature exists within the omentum, and in the diaphragm, and the vessels draining the peritoneal cavity carry lymph to the parathymic LNs en route to the thoracic duct. Consistent with the notion that B1 cells undergo some recirculation, B1 cells were detected in parathymic LNs, in contrast to other LN types, where few if any can be detected (Cyster et al., 2002).
Mature B Cells in the Bone Marrow In addition to serving as the site of B cell genesis, the bone marrow contains a population of mature, long-lived B cells. In the mouse, this typically corresponds to a few percent of total bone marrow cells or about 107 cells. Homing of transferred B cells to the bone marrow is dependent on the combined function of LFA1, a4b1, and a4b7 (Berlin-Rufenach et al., 1999). In mice with a conditional ablation of VCAM1, bone marrow homing of mature B cells is defective (Koni et al., 2001; Leuker et al., 2001). Accumulation of mature B cells in the bone marrow also depends on the chemokine–receptor pair CXCL12(SDF1)-CXCR4 (Ma et al., 1999; and unpublished observations). Mice lacking the B cell surface molecule, CD22, have a paucity of mature B cells in the bone marrow. When CD22-deficient B cells are transferred to wildtype recipients, they fail to accumulate in the bone marrow (Nitschke et al., 1997; Otipoby et al., 1996). CD22 is a B cell–specific member of the sialic acid binding immunoglobulin-like lectin (Siglec) family, and it preferentially binds sugars terminating in a2,6-sialic acid (the NeuNAc form for human CD22 and the NeuNGc-form for mouse CD22) (Nitschke et al., 2001). Staining with an Fc-fusion protein of CD22 reveals ligands on the bone marrow sinusoidal endothelium (Nitschke et al., 1999). CD22 can only bind to a2,6-linked sialic acids on target cells if the CD22 is not masked by a2,6-linked sialic acids on the B cell surface. Analysis using N-acetyl a2,6sialyllactose binding to B cells revealed that the frequency of cells able to bind was greater among mature bone marrow B cells than in other mature B cell populations (Nitschke et al., 1999). The factors promoting decreased a2,6-sialic acid production on B cells are not defined, but there is some indication that unmasking occurs during B-cell activation (Razi and Varki, 1998). Still less clear at this time is the purpose of mature B cell accumulation in the bone marrow. Given the highly vascular nature of the marrow, perhaps they serve
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a similar function to the marginal zone B cells within the spleen, responding to antigens that arrive in the bone marrow with the blood.
B CELLS AT SITES OF INFLAMMATION Although B cells are not typically thought of as inflammatory cells, they accumulate in surprising numbers in certain types of inflammation, particularly in chronic inflammatory diseases. This includes the autoimmune inflammation associated within the joint synovium of rheumatoid arthritis (RA) patients, in the thymus of myasthenia gravis (MG) patients, the thyroid of thyroiditis patients, the pancreas of type 1 diabetics, and the salivary glands of patients with Sjögren’s syndrome (Hjelmstrom, 2001). The extent to which local accumulation of B cells contributes to pathology is mostly unclear, but in some cases ectopic germinal centers are observed and are thought to be involved in autoantibody production. It is also notable that although the development of diabetes in NOD mice does not depend on autoantibody production, it does depend on B lymphocytes (Serreze et al., 1996). B cells might also contribute to pathology by serving as antigen-presenting cells and through expression of cytokines, such as LTa1b2 and TNF (Ansel et al., 2000; Endres et al., 1999; Harris et al., 2000). Transgenic studies established that ectopic expression of CXCL13 in the islet cells of the pancreas was sufficient to cause massive accumulation of naïve B cells (Luther et al., 2000). Several groups have tested for the expression of this chemokine at sites of autoimmune inflammation. Induction of CXCL13 occurs in the inflamed salivary gland of Sjögren’s syndrome patients (Salomonsson et al., 2002), in the ectopic follicles within the synovium of RA patients (Shi et al., 2001), in the lesions associated with ulcerative colitis (Carlsen et al., 2002), and in Helicobacter pylori–induced mucosa-associated lymphoid tissue (Mazzucchelli et al., 1999). Dendritic cells expressing CXCL13 have been identified in the thymus of lupus-prone mice, possibly contributing to B cell accumulation in the thymus and the development of disease (Ishikawa et al., 2001). CXCL12 can also promote transendothelial migration of B cells and might be expected to contribute to B cell accumulation at ectopic sites. Studies in rheumatoid arthritis synovium indicate notable expression of CXCL12 by synovial fibroblasts and possibly endothelial cells (Buckley et al., 2000; Nanki et al., 2000). In transgenic studies, CXCL12 was poor at recruiting lymphocytes to an ectopic site, but it seems likely that it could synergize with other factors induced at sites of inflammation to promote B and T cell accumulation (Luther et al., 2002). The ectopic expression of CCL21 causes a strong accumulation of T and B lymphocytes (Chen et al., 2002; Fan et al., 2000; Luther et al., 2002), and CCL21
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upregulation has been observed in the pancreas of pre-diabetic mice (Hjelmstrom et al., 2000; Luther et al., 2002). Indeed, the presence of CCL21 is sufficient to trigger integrin activation and extravasation of naïve T cells in noninflamed peripheral tissues (Weninger et al., 2003). In addition to chemokines, the upregulation of adhesion molecules that typically occurs at sites of inflammation, including PNAd, ICAM1, VCAM1, and MAdCAM1, is likely to contribute to the influx of B lymphocytes in chronically inflamed tissues.
HOMING OF ANTIBODY-SECRETING CELLS (ASCs) The differentiation of a B cell into an antibody-secreting cell is accompanied by a large number of gene-expression plasma changes and involves the cell transforming from a small blast expressing Ig as a surface molecule to a large cell that is full of rough endoplasmic reticulum and is manufacturing enormous amounts of soluble Ig (Calame, 2001). Plasma blasts, the immediate precursors of terminally differentiated plasma cells also secrete some antibody, and it has often been difficult to distinguish between these cells without performing ultrastructural studies. For this reason, many investigators refer to plasma blasts and plasma cells together as antibody secreting cells (ASCs). ASCs typically express high surface levels of the proteoglycan syndecan-1, and this molecule serves as a useful ASC marker (Calame, 2001). ASCs generated early during primary immune responses, prior to germinal center formation, or produced during T-independent responses, are typically short-lived and survive for only a few days (Ho et al., 1986; Smith et al., 1996). For the most part, these cells remain within the secondary lymphoid organ where they arose, contributing to a rapid burst of circulating Ig. Within the spleen, these rapidly produced ASCs migrate as large foci of blasts from the outer T zone of the white pulp, through the marginal zone bridging channels (Figures 14.3 and 14.5), to take up positions near vessels or collagenous fibers in the red pulp (Jacob et al., 1991; Liu et al., 1991; van Rooijen et al., 1986). In lymph nodes, newly produced ASCs migrate to the medulla and localize in medullary cords (Figures 14.1 and 14.5) (Kosco et al., 1989; Luther et al., 1997). ASCs arising later in the primary response, most likely emerging from GCs, and those generated from memory cells in secondary responses are often long lived, surviving in mice for weeks or months and probably even longer in humans (Manz et al., 1997; Slifka and Ahmed, 1998). Although some of these cells localize to the same locations as the short-lived ASCs, many travel to the bone marrow or, in the case of IgA secreting cells, to mucosal sites (Benner et al., 1977; Dilosa et al., 1991; Kosco et al., 1989; Lamm and Phillips-Quagliata, 2002).
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FIGURE 14.5 Principal migration pathways of B-lineage cells. Each arrow indicates an active migration event by a B-lineage cell (some arrows may incorporate more than one migration step). The principal type of Blineage cell at each location is indicated in parentheses (in blue). Green arrows indicate migration events that occur homeostatically or during development; red arrows refer to migration events that occur following antigen-encounter and B cell activation or differentiation. Distinct migration cues are required for cells to reach each of the indicated tissues or compartments. Note that the diagram emphasizes migration events and is not meant to be to scale or to represent anatomical organization. See color insert.
Analysis of IgM and IgG ASCs induced in the spleen or lymph nodes following various immunization protocols established that the cells downregulate CXCR5 and CCR7 and lose responsiveness to B and T zone chemokines (Hargreaves et al., 2001; Hauser et al., 2002; Wehrli et al., 2001). At the same time, they maintain expression of CXCR4 and exhibit increased chemotactic sensitivity to CXCL12 (Hargreaves et al., 2001; Hauser et al., 2002; Wehrli et al., 2001). CXCL12, originally called stromal cell derived factor (SDF)-1, was first characterized for its high expression within the bone marrow (Bleul et al., 1996), where it functions in precursor cell retention (Ansel and Cyster, 2001). Within secondary lymphoid tissues, CXCL12 is expressed at highest levels by cells within the red pulp of spleen and within the medullary cords of lymph nodes. CXCR4 is necessary for efficient migration of IgM and IgG ASCs into these areas (Hargreaves et al., 2001; Cyster, 2003). Consistent with a key role for CXCL12 in guiding ASC localization, IgM ASCs accumulate in the pancreatic islets of transgenic mice ectopically expressing CXCL12 under control of the rat insulin promoter (Luther et al., 2002). Another mechanism that is likely to influence ASC distribution is integrin-mediated adhesion. ASCs express high levels of a4b1 and LFA1, and IgG ASCs adhere strongly to the a4b1 ligand VCAM1 (Underhill et al., 2002), a ligand that is expressed by cells throughout the splenic red pulp (Lu and Cyster, 2002). VCAM1 is also constitutively expressed on bone marrow endothelial cells (Jacobsen et al., 1996; Mazo et al., 1998). Fibronectin, a second a4b1 ligand,
is also present in the red pulp and appears to be especially enriched near vessels and fibers, sites of ASC lodgement. Transit of ASCs from secondary lymphoid organs to the bone marrow requires that the cells pass into the blood and then attach to bone marrow endothelium and enter the marrow parenchyma. Considerable numbers of ASCs can be isolated from the blood during the early phase of a T-dependent antibody response (Benner et al., 1977), and immature ASCs have also been identified in human blood (Kawano et al., 1995). The factors determining whether an ASC exits a secondary lymphoid organ versus staying in the organ are poorly defined, but studies of the mouse response to sheep red blood cells have shown that the exit occurs as a synchronous wave of cells at about day 3 of the secondary response (Benner et al., 1977, 1981). In addition to its role in directing plasma cell localization in secondary lymphoid organs, CXCR4 is important for ASC homing to the bone marrow (Hargreaves et al., 2001). A kinetic analysis of antigen-specific ASCs appearing in the bone marrow during an immune response revealed that the cells lost their ability to chemotax to CXCL12 by day 12 of the response, while retaining CXCR4 expression (Hauser et al., 2002). It seems likely that CXCL12 plays dual roles in ASC homing to the bone marrow, helping promote adhesion and transmigration of immature ASCs across the bone marrow endothelium, and subsequently helping retain mature ASC within the bone marrow, in close contact with stromal cells, in a manner similar to that proposed for progenitor B cells (Ansel and Cyster, 2001). In addition to chemokines, ASC attachment to bone marrow endothelium is likely to require selectins and/or integrins. ASCs upregulate expression of P-selectin glycoprotein ligand (PSGL)-1, a protein that can be modified to display both P- and E-selectin binding sites (Xia et al., 2002). In in vitro assays, IgG ASCs undergo rolling interactions with E-selectin but not P-selectin (Underhill et al., 2002). E- and P-selectins are constitutively expressed by bone marrow endothelial cells (Frenette et al., 1998; Mazo et al., 1998; Schweitzer et al., 1996). The high expression of integrins a4b1 and LFA1 on ASCs (Underhill et al., 2002) may also be important for their homing to the bone marrow. VCAM1 is expressed constitutively in the bone marrow and (as discussed in an earlier section), a4b1 and VCAM1 function in B cell and progenitor cell homing to bone marrow (Berlin-Rufenach et al., 1999; Koni et al., 2001; Leuker et al., 2001), although a role for this integrin-ligand has yet to be directly demonstrated in vivo for ASCs. Plasma cells in humans express a5b1 in addition to a4-integrins, and this may contribute to enhanced binding to extracellular matrix proteins (Kawano et al., 1995). As noted earlier, CD22 functions in the homing of mature B cells to the bone marrow. CD22-deficiency is also associated with reduced accumulation of ASCs within the bone marrow (Nitschke et al., 1999). Although ASCs downregulate CD22 during
14. Dynamics of B Cell Migration to and within Secondary Lymphoid Organs
terminal differentiation (Calame, 2001), it seems likely that full downregulation may be delayed compared to the time of migration. IgG is the predominant Ig isotype in serum, but the major isotype synthesized in the body is IgA. Most of this production takes place through ASCs residing in mucosal and exocrine sites, especially along the intestinal tract. Much of this IgA is transported across epithelial surfaces without entering into circulation. Characterization of the chemotactic profile of IgA-producing cells from spleen, mucosal lymph nodes, and lamina propria revealed that, in contrast to IgM and IgG ASCs, IgA ASCs respond strongly to TECK/CCL25 (Table 14.1) and express mRNA for the CCL25 receptor, CCR9 (Bowman et al., 2002). CCL25 is constitutively expressed within the small intestine, especially in epithelial crypts, while being expressed more weakly or not at all in the colon and at other mucosal surfaces. Analysis of cells taken directly from the intestinal lamina propria (LP) revealed that B220intIgA+ ASC responded to CCL25, whereas the terminally differentiated B220-IgA+ ASC did not (Bowman et al., 2002). This is similar to the findings for CXCL12 responsiveness of bone marrow ASCs (Hauser et al., 2002) and suggestive of the conclusion that once plasma blasts reach their final destination and terminally differentiate into plasma cells they lose their ability to chemotax. In addition to CCL25, CXCL12 is expressed by gut epithelial cells and by cells in the LP (Agace et al., 2000). As IgA ASCs respond to CXCL12 as well as CCL25, these chemokines may work together to help ensure correct positioning of IgA ASCs in the small intestine (Bowman et al., 2002). ASCs in the intestine express a4b7 (Farstad et al., 1995). The a4b7 ligand, MAdCAM-1, is present on small venules in the gut (Briskin et al., 1997), making it likely that this integrin–ligand pair functions in ASC lodgement in the gut. In addition to LP IgA ASCs deriving from B cells activated in Peyer’s patches and mucosal LNs, some of the cells derive from IgM+ cells locally within the LP (Fagarasan et al., 2001). Up to half of the IgA produced in the gut is believed to be derived from B cells of B-1 origin (Kroese et al., 1989), and it has been suggested that the IgM+ LP cells that give rise to IgA ASCs are originally derived from body cavity B-1 cells (Lamm and Phillips-Quagliata, 2002). Whether the precursors of IgM+ LP cells express CCR9 and respond to CCL25 has not yet been investigated. Interestingly, when immune responses are induced by exposure to antigen in the colon or in the vaginal epithelium, there is greater accumulation of ASCs at these sites than at other mucosal surfaces such as the small intestine (Parr and Parr, 1998; Pierce and Cray, 1982). It can therefore be anticipated that further chemokine(s) operate to provide additional specificity to mucosal ASC homing. Indeed, recent studies provide evidence for CCR10 and its ligand, CCL28, playing a role in IgA ASC homing to mucosal sites, includ-
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ing colon, salivary glands, and the respiratory tract (Kunkel and Butcher, 2003). The lactating mammary gland is also an important site of IgA plasma cell accumulation. Expression of MAdCAM-1 has been detected on mammary gland endothelial cells (van der Feltz et al., 2001), but it is unclear which adhesion molecules and chemokines are involved in ASC recruitment to this site. IgM and IgG ASCs respond weakly to the CXCR3 ligands CXCL9/MIG, CXCL10/IP10, and CXCL11/ITAC, in addition to responding to CXCL12, and at least a fraction of the cells express CXCR3 (Bowman et al., 2002; Cyster et al., 2002; Hauser et al., 2002). As the CXCR3 ligands are all strongly induced by interferons (Cassese et al., 2001), they may contribute to the appearance of ASCs in some types of inflammation (Chvatchko et al., 1996; Kim and Berek, 2000).
CONCLUSION In summary, much has been learned about how B lymphocytes attach to endothelial cells and enter lymphoid tissues, about the integrins used by these cells to adhere to stromal cells or to other leukocytes, and about chemokines that direct the migration and adhesion of B lineage cells. Some of this knowledge is already being put to the test therapeutically as drug companies examine whether L-selectin inhibitors can reduce inflammatory diseases or whether lymphocyte-attracting chemokines can be used to improve adjuvants or serve as cancer immunotherapy agents. Current knowledge suggests further points for therapeutic intervention to diminish B cell–related immunological diseases, such as inhibiting CXCL12 function as a means of reducing plasma cell accumulation in the rheumatoid synovium or as an approach to displace and eliminate plasma cells in patients with lupus. The increasing number of examples in which the B cell attracting chemokine CXCL13 is expressed at sites of inflammation, together with the evidence that this chemokine can induce cells to express LTa1b2 and cause downstream effects including T cell recruitment, suggests there may be significant benefit in neutralizing this chemokine in people with inflammatory diseases. Similarly, remembering that CXCR5 was isolated because of its high expression in Burkitt’s lymphoma [and first given the name, Burkitt’s lymphoma receptor-1, BLR1 (Dobner et al., 1992)], CXCL13 could contribute to lymphoma cell clustering and to the induction of tumor supportive stromal niches in follicular and mantle zone lymphomas. The inhibition of CXCL13 function might be beneficial in these diseases. Marginal zone B lymphoma cells need to be tested for integrin expression profile and adhesiveness to determine whether they share with nontransformed marginal zone B cells the property of expressing high levels of functional integrins. The increasing evidence that memory B cells and
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plasma cells have highly specialized homing profiles reinforces the notion that it is best to vaccinate via the same route as the infection route of the pathogen. Although long-distance migration is a major functional requirement of a successful B cell, we know relatively little about the intracellular machinery regulating B cell movement. Impressive advances have been made in understanding how Dictostyelium cells and neutrophils chemotax, but lymphocytes appear to have unique specializations to support their highly motile lifestyle (Reif and Cyster, 2002). They use distinct phosphoinositide-3-kinase family members from other cells to couple chemokine receptors to downstream mediators of chemotaxis, and they have a unique requirement for the molecule DOCK2, a specialized type of RacGEF, for chemotaxis (Fukui et al., 2001). As the drug FTY720 is currently teaching us (Brinkmann and Lynch, 2002), the selective control of the migration of B and T lymphocytes has promise for the further development of new and improved immunoregulatory drugs.
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15 Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System PER BRANDTZAEG,1 H. CRAIG MORTON,1 AND MICHAEL E. LAMM2 1
Laboratory for Immunohistochemistry and Immunopathology (LIIPAT), Institute of Pathology, University of Oslo, Rikshospitalet, Oslo, Norway; 2 Department of Pathology, Case Western Reserve University, School of Medicine, Cleveland, Ohio, USA
1998), but this is potentially a proinflammatory reinforcement of the epithelial barrier function (Brandtzaeg and Tolo, 1977). The biological significance of both the unique inductive and the specific migratory properties of mucosal B cells is emphasized by the fact that more than 80% of all immunocytes are located in the gut and 80 to 90% of them normally produce pIgA (Brandtzaeg et al., 1999a). The mucosa and exocrine glands thus harbor by far the greatest activated Bcell system of the body. This chapter deals with the mechanisms involved in the differentiation of mucosal B cells and signals directing their preferential homing to secretory effector sites. Although the focus is on human mucosal tissues, much fundamental mechanistic information has to be extrapolated from animal experiments.
The first line of adaptive humoral defense depends on cooperation between mucosal B cells and exocrine epithelia to provide secretory immunity (Brandtzaeg et al., 1999a). Terminally differentiated B cells occur as immunoglobulin (Ig)-producing immunocytes (plasmablasts and plasma cells) at every secretory effector site where they normally produce dimers and some larger polymers of IgA (collectively termed pIgA). In addition to its light and heavy chains, pIgA contains a 15-kD polypeptide termed the “joining” or J chain (Mestecky and McGhee, 1987), which facilitates spontaneous noncovalent interactions with the polymeric Ig receptor (pIgR) (Brandtzaeg and Prydz, 1984; Johansen et al., 2001). This receptor is expressed basolaterally on secretory epithelial cells as a 100-kD glycoprotein, also called membrane secretory component (SC) (Brandtzaeg 1974a; 1985). By endocytosis and transcytosis pIgR exports pIgA and J chain-containing pentameric IgM with equal efficiency in humans, but there are considerable species differences with regard to transport of the latter ligand (Norderhaug et al., 1999). Although mucosal immunocytes of all Ig classes generally produce the J chain (Brandtzaeg, 1974b, 1985), it is linked only to the IgA and IgM subunits by covalent bonding to their C-terminal eighteen amino-acid-long heavy-chain tail-pieces (Johansen et al., 2000). Therefore, the consequence of the strong J-chain expression at secretory effector sites is abundant local formation of Ig polymers that can readily be subjected to pIgR-mediated epithelial transport. Secretory antibodies (SIgA and SIgM) are thereby provided at epithelial surfaces to perform immune exclusion (Figure 15.1) and noninflammatory clearance of antigens from the mucosa (Mazanec et al., 1993; Norderhaug et al., 1999). Locally or serum-derived IgG antibodies may contribute to external defense after paracellular leakage (Persson et al.,
Molecular Biology of B Cells
IMMUNE-INDUCTIVE TISSUE COMPARTMENTS Lymphoid cells are located in three histologically distinct tissue compartments at mucosal surfaces: immune-inductive organized mucosa-associated lymphoid tissue (MALT), the lamina propria or glandular stroma, and the surface epithelia. Peyer’s patches in the distal small intestine are typical MALT structures believed to be a main source of conventional (B2) surface (s)IgA-expressing primed mucosal B cells (Figure 15.2). The lamina propria is principally an effector site but is also important in terms of the expansion and terminal differentiation of B cells. MALT structures resemble lymph nodes, with B-cell follicles, intervening T-cell areas, and a variety of antigenpresenting cells (APCs), but they lack afferent lymphatics (Brandtzaeg et al., 1999a). All such structures therefore
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FIGURE 15.1 Model for external transport of J chain–containing dimeric IgA and pentameric IgM by the polymeric Ig receptor (pIgR), expressed basolaterally as membrane secretory component (SC) on glandular epithelial cells. The polymeric Ig molecules are produced with incorporated J chain (IgA + J and IgM + J) by mucosal plasma cells. The resulting secretory Ig molecules (SIgA and SIgM) act in a first line of defense by performing immune exclusion of antigens in the mucus layer on the epithelial surface. In addition, pIgR-mediated export of immune complexes from the lamina propria and epithelial compartment may contribute to noninflammatory mucosal defense (not shown). Although J chain is often (70–90%) produced by human mucosal IgG plasma cells, it does not combine with this Ig class, but is degraded intracellularly as denoted by (±J) in the figure. Locally produced (and serum-derived) IgG is not subjected to active external transport, but can be transmitted paracellularly to the lumen, as indicated. Free SC (depicted in mucus) is generated when pIgR in its unoccupied state (top symbol) is cleaved at the apical face of the epithelium, like bound SC in SIgA and SIgM. Although bound SC is covalently linked to one subunit in SIgA, providing protection against degradation, SIgM contains only noncovalently bound SC in dynamic equilibrium with free SC in the secretion.
sample exogenous antigens directly from the mucosal surfaces through a characteristic follicle-associated epithelium (FAE), which contains membrane (M) cells (Figure 15.2). These specialized thin epithelial cells have been shown to be effective in the uptake of live and dead (especially particulate) antigens from the gut lumen, and many enteropathogenic bacterial (e.g., Salmonella spp., Vibrio cholerae) and viral (e.g., poliovirus, HIV-1, reovirus) infectious agents use the M cells as portals of entry (Neutra et al., 2001).
Gut-Associated Lymphoid Tissue Gut-associated lymphoid tissue (GALT) includes Peyer’s patches, the appendix, and scattered solitary or isolated lymphoid follicles (ILFs). Early animal studies demonstrated that Peyer’s patches and mesenteric lymph nodes are enriched precursor sources for intestinal IgA immunocytes (Craig and Cebra, 1971; McWilliams et al., 1977; McDermott and Bienenstock, 1979), and that differentiation of sIgA+ B cells takes place during their dispersion to distant sites (Guy-Grand et al., 1974; Roux et al., 1981). Thus, the fraction with cytoplasmic IgA increased from an initial 2%
in Peyer’s patches to 50% in mesenteric lymph nodes and 75% in thoracic duct lymph, and finally 90% in the intestinal lamina propria (Parrott, 1976). Such seminal findings gave rise to the term IgA cell cycle (Lamm, 1976), but later studies showed that B cells of other Ig classes and T cells induced in Peyer’s patches also exhibit gut-seeking properties (Figure 15.2). Peyer’s patches occur mainly in the ileum (less frequently in the jejunum) and are defined to consist of at least five aggregated lymphoid follicles, but can contain up to 200 such structures (Cornes, 1965). Human Peyer’s patch anlagen, composed of CD4+ dendritic cells (DCs), can be seen at 11 weeks of gestation, and discrete T- and B-cell areas occur at 19 weeks. No germinal centers appear until shortly after birth, thus reflecting a dependency on antigenic stimulation (Figure 15.2), which also induces some follicular hyperplasia (Spencer and MacDonald, 1990). The number of macroscopically visible human Peyer’s patches increases from about 50 at the beginning of the last trimester to 100 at birth and 250 in the midteens, then diminishes to approximately 100 between 70 and 95 years of age (Cornes, 1965). Human intestinal mucosa harbors at least 30,000 ILFs (Figure 15.2), increasing in density distally (Trepel, 1974). Thus, the normal small intestine contains only 1 follicle per 269 villi in the jejunum, but 1 per 28 villi in the ileum (Moghaddami et al., 1998). In the normal large bowel, the density of ILFs is likewise relatively small—enumerated in tissue sections to increase from 0.02/mm muscularis mucosae in the ascending colon to 0.06/mm in the rectosigmoid (O’Leary and Sweeney, 1986). ILFs have recently been characterized immunologically in mice, showing features compatible with the induction of B cells for intestinal IgA responses (Hamada et al., 2002). Interestingly, the organogenesis of murine ILFs was found to commence after birth, in contrast to Peyer’s patches.
Nasopharynx-Associated Lymphoid Tissue Although GALT is the largest and best defined part of MALT, other potentially inductive sites for mucosal B-cell responses are bronchus-associated lymphoid tissue (BALT) and nasopharynx-associated lymphoid tissue (NALT). In humans, NALTis constituted mainly by the unpaired nasopharyngeal tonsil (often called adenoids) and the paired palatine tonsils (Brandtzaeg, 1987; Brandtzaeg and Halstensen, 1992; Perry and Whyte, 1998). These organs make up most of Waldeyer’s pharyngeal lymphoid ring and may play a major role for mucosal immunity in human airways because BALT structures are not present in normal lungs of adults and only in 40% of healthy adolescents and children (Tschering and Pabst, 2000). Rodents lack tonsils, whereas two paired NALT structures occur laterally to the nasopharyngeal duct dorsal to the
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
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FIGURE 15.2 Antigen-sampling and B-cell Ig class-switch sites for induction of intestinal antibody responses. Dots denote antigens. The classical inductive sites are constituted by gut-associated lymphoid tissue (GALT), which is equipped with antigen-sampling M cells, T-cell areas (T), B-cell follicles (B), and antigen-presenting cells (APCs). Switch of conventional B2 cells from surface (s)IgM to sIgA expression occurs in GALT and mesenteric lymph nodes; from here primed B and T cells home to the lamina propria (LP) via lymph and blood. T cells mainly end up in the epithelium (EP), whereas sIgA+ cells differentiate to LP plasma cells to produce dimeric IgA with J chain (IgA + J), which then is exported as secretory IgA (SIgA). Primed B cells may also migrate from Peyer’s patches and isolated lymphoid follicles directly into the LP as indicated, whereas those differentiating to plasma cells just outside a follicle often show reduced J-chain expression and a propensity for IgG production (IgG ± J). B2 cells also give rise to plasma cells producing pentameric IgM (IgM + J), which becomes secretory IgM (SIgM). B1 cells (CD5+) from the peritoneal cavity reach the LP by an unknown route (?), perhaps via mesenteric lymph nodes. These sIgM+ cells are particularly abundant in mice and may switch to sIgA within the LP, under the influence of APCs that have sampled microbial antigens as dendritic cells within the epithelium and become activated to secrete stimulatory factors (wavy arrow) such as BAFF and APRIL. The sIgA+ B1 cells differentiate to plasma cells that provide SIgA mainly directed against the commensal gut flora.
cartilaginous soft palate (Kuper et al., 1992). A regionalized protective IgA response has been shown to be induced by nasal vaccine application in mice (Yanagita et al., 1999). Indeed, murine NALT can drive an IgA-specific enrichment of high-affinity memory B cells, but gives additional rise to a major germinal center population of IgG-producing cells (Shimoda et al., 2001)—quite similar to the situation in human tonsils (Brandtzaeg, 1987; Brandtzaeg et al., 1999b). In contrast to tonsils, however, the anlagen of which appears at the same fetal age as that of Peyer’s patches (von Gaudecker and Müller-Hermelink, 1982), the organogenesis of murine NALT begins after birth, as does
murine ILFs (Fukuyama et al., 2002; Hamada et al., 2002; Mebius, 2003).
Other Sources of Mucosal B Cells In mice, proliferating T cells rapidly obtain gut-homing properties during antigen priming in mesenteric lymph nodes (Campbell and Butcher, 2002). Most likely, therefore, regional lymph nodes generally share immune-inductive properties with the related MALT structures from which they receive antigens via afferent lymph and antigentransporting DCs. Numerous DCs are found at epithelial
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surfaces, where they can pick up luminal antigens by penetrating tight junctions with their processes (Rescigno et al., 2001). Importantly, the human nasal mucosa is extremely rich in various DC types, both within and beneath the epithelium (Jahnsen et al., 2003), and a subepithelial band of putative APCs is seen below the surface epithelium and the FAE in the human gut (Rugtveit et al., 1997; Yamanaka et al., 2003). The peritoneal cavity is recognized as yet another source of mucosal B cells in mice, perhaps providing 40 to 50% of the intestinal IgA immunocytes (Kroese et al., 1989). The precursors are self-renewing sIgM+ B1 (CD5+) cells, and give rise to polyreactive (“natural”) SIgA antibodies (Figure 15.2), particularly directed against commensal bacteria as a result of T cell–independent responses (Macpherson et al., 2000). How and where this subset differentiates to the IgA phenotype remains uncertain, but the lamina propria has recently been suggested as an important class switch site (Fagarasan et al., 2001; Fagarasan and Honjo, 2003). Notably, though, no evidence exists to suggest that B1 cells are significantly involved in intestinal IgA production in man (Brandtzaeg et al., 2001; Boursier et al., 2002), despite considerable levels of polyreactive SIgA antibodies recog-
nizing both self and microbial antigens in human secretions (Bouvet and Fischetti, 1999).
CHARACTERISTICS OF B CELLS IN SECRETORY EFFECTOR TISSUES IgA-Producing Immunocytes Are Remarkably Abundant Secretory effector sites in normal human adults contain a striking preponderance (70–90%) of IgA-producing immunocytes (Figure 15.3), which in the normal gut amount to approximately 1010 per meter, or at least 80% of all Ig-producing cells of the body (Brandtzaeg et al., 1989). Thus, most large lymphoid cells dispersed from the lamina propria belong to the terminally differentiated phenotype (CD38+CD27+CD19+/-CD20-) with IgA on the surface and/or in the cytoplasm, whereas most small lymphoid cells are T lymphocytes (Table 15.1). This is in contrast to the flow-cytometric data obtained from GALT compartments such as Peyer’s patches and the appendix, where small B
FIGURE 15.3 Average percentage distribution of immunocytes (plasmablasts and plasma cells) producing different Ig classes in various human secretory tissues from healthy controls and subjects with IgA deficiency. Based on published data from the Brandtzaeg laboratory.
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
TABLE 15.1 Flow-cytometric analysis of the phenotypic distribution of B and T cells in two human gut compartmentsa Phenotype proportionb
Co-expression patternc Small cellsb
Large cellsb
Organized GALT B cells 50% CD19+CD38-a4b7int
25% L-selectin+ 50% sIgD+ (40% L-selectin+) 30% sIgA+ (15% L-selectin+) 14% sIgG+ (25% L-selectin+)
<5% CD19+/-CD38hia4b7int/hi
L-selectin-
T cells 45% CD3+a4b7int/hi
40% L-selectin+ 70% CD4+aEb720% CD8+aEb7-/+
Only few cells
B cells L-selectin>80% sIgA+
25% CD19+/-CD38hia4b7hi
L-selectin>90% s/cIgA+d
T cells 60% CD3+a4b7int
4% CD3+a4b7hi
Following CD40 ligation, these cells proliferate in vitro and constitutively secrete IgA, thus signifying a capacity for local recall responses (Farstad et al., 2000). Notably, lamina propria CD19+ cells are negative for CD5, which sup-ports the notion that B1 cells do not contribute significantly to the human IgA immunocyte population (Boursier et al., 2000). MALT-derived B cells also enter lactating mammary glands (Roux et al., 1977), and human colostrum contains 300 times more SIgA than stimulated parotid saliva. Nevertheless, the tissue density of IgA immunocytes is similar in human salivary and lactating mammary glands, and actually six to seven times less than in lacrimal glands and colonic mucosa. Therefore, the large organ size, combined with capacity for storage of locally produced pIgA in the epithelium and duct system of mammary glands, explains the striking output of SIgA during breast-feeding (Brandtzaeg, 1983a).
Disparate IgA Subclass Distribution
Mucosal lamina propria 10% CD19+CD38-a4b7int/hi
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L-selectin65% CD4+aEb7-/+ 30% CD8+aEb7+ L-selectin-aEb790% CD4+ <5% CD8+
a
Adapted from Farstad et al. (1995, 1996, 1997a) and unpublished data from the Brandtzaeg laboratory. b Dispersed mononuclear cells gated according to size and analyzed for surface markers and fluorescence intensity (+, positive; -, negative; lo, low; int, intermediate; hi, high). c Calculated from all B or T cells. d Cytoplasmic IgA (cIgA) determined by immunohistochemistry.
lymphocytes (CD19+) and T lymphocytes (CD3+) dominate. Importantly, these results accord with parallel immunohistochemical observations that show that small naïve B lymphocytes (sIgD+IgM+CD19+CD20+) in the human gut are almost exclusively present in follicular mantle zones of GALT (Farstad et al., 2000). The proportion of small B lymphocytes in dispersed lamina propria samples varies from 4 to 42%, and 5 to 50% of these cells show a naïve phenotype (sIgD+), thus reflecting a highly variable contamination from GALT structures such as ILFs. The human lamina propria contains only few and scattered sIgA+CD27+ memory cells bearing low levels of CD40.
Two IgA subclasses occur in humans, IgA1 normally constituting at least 85% of total serum IgA. A relatively large IgA2 proportion (29–64%) has been reported for IgA immunocytes in gut mucosa compared with peripheral lymphoid tissue and upper airways (7–25%), but IgA2 dominates (64%) only in the large bowel (Crago et al., 1984; Jonard et al., 1984; Kett et al., 1986; Burnett et al., 1987). A skewing towards SIgA2 may be important for the stability of secretory antibodies because this isotype is resistant to several IgA1-specific bacterial proteases (Kilian et al., 1996). The concentration ratio of the two SIgA subclasses in various secretions (Jonard et al., 1984; Müller et al., 1991; Feltelius et al., 1994) corresponds to the immunocyte proportions at the related secretory tissues, thus supporting the notion that both isotypes of pIgA are equally well exported by the pIgR (Brandtzaeg, 1977). The molecular events underlying preferential IgA1 or IgA2 responses remain unclear. Secretory antibodies to lipopolysaccharide (LPS) are generally of the SIgA2 subclass, whereas protein antigens stimulate predominantly SIgA1 (Mestecky and Russell, 1986; Tarkowski et al., 1990). The fact that jejunal IgA immunocytes are mainly of the IgA1 subclass (~77%), in contrast to the IgA2 dominance (~64%) in the colon (Kett et al., 1986), may therefore reflect the disparate luminal distribution of food antigens versus gram-negative bacteria. Bacterial overgrowth in bypassed jejunal segments alters the immunocyte composition, with an increase of IgA2 and a decrease of IgM production (Kett et al., 1995), thus suggesting LPS-induced direct isotype switching from Cm to Ca2 or progressive sequential downstream switching of the Ig heavy-chain constant region (CH) genes.
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Disparate Isotype Distribution of Other Immunocytes IgM-producing cells constitute a substantial but variable immunocyte fraction in the adult human gut (Figure 15.3). The relatively high proportion of this isotype (~18%) in the proximal small intestine may be related to the low levels of LPS (see above), and is in striking contrast to a much lower frequency in the upper aerodigestive tract (Brandtzaeg et al., 1979). This disparity is remarkably accentuated in IgAdeficient patients (Figure 15.3), who may have clinical problems due to lack of compensatory SIgM in their airways (Brandtzaeg et al., 1987). IgG-producing cells normally constitute only 3 to 4% of human intestinal immunocytes, but there is a considerably larger proportion in gastric and nasal mucosae (Figure 15.3), which often show some low-grade inflammation (Valnes et al., 1986). In inflammatory bowel disease (IBD), the IgG fraction is dramatically increased (Brandtzaeg et al., 1989, 1997). Moreover, although IgA immunocytes remain dominating in both ulcerative colitis and Crohn’s disease, the cells are aberrant in showing an increased IgA1 subclass proportion (Kett and Brandtzaeg, 1987) and decreased Jchain expression (Brandtzaeg and Korsrud, 1984; Kett et al., 1988). Immunohistochemical studies of human upper airways (Brandtzaeg et al., 1987) as well as normal jejunal (Nilssen et al., 1991), ileal (Bjerke and Brandtzaeg, 1990a), and colonic (Helgeland et al., 1992) mucosa have demonstrated that IgG1 is the predominating locally produced IgG subclass (56–69%), similar to its dominance in serum. However, IgG2 immunocytes are generally more frequent (20–35%) than IgG3 cells (4–6%) in the distal gut, whereas the reverse is often true in airway mucosae (Brandtzaeg et al., 1987). Such IgG-subclass disparity supports the idea that isotype switching pathways may differ in various body regions. Interestingly, the Cg2 and Ca2 genes are located on the same DNA segment (Flanagan and Rabbits, 1982), and many carbohydrate and bacterial antigens preferentially induce an IgG2 response in addition to IgA2, whereas proteins (which are clearly T cell-dependent antigens) primarily generate IgG1 responses together with IgA1 (Papadea and Check, 1989). Such response differences might be reflected in the variable intestinal IgA and IgG immunocyte subclass patterns. IgD-producing cells are only occasionally encountered in the human gut, whereas they normally constitute a significant fraction (3–10%) at secretory sites in the upper aerodigestive tract (Brandtzaeg et al., 1979, 1987; Korsrud and Brandtzaeg, 1980). In IgA deficiency, this disparity is even more striking for IgD than that noted for IgM immunocytes (Figure 15.3), which may reflect compartmentalized differences in immune regulation and homing mechanisms (see below).
IgE-producing cells are virtually absent from human mucosa, with rare exceptions only in allergic patients, whereas IgE-bearing mast cells are commonly found (Rognum and Brandtzaeg et al., 1989).
J-Chain Expression Is a Characteristic of Mucosal Immunocytes To support secretory immunity, MALT must favor the development and dispersion of B cells with prominent expression of J chain; this is a prerequisite for the production of pIgA and pentameric IgM that can be exported by the pIgR (Figure 15.1). Although this peptide is not absolutely required for IgA and IgM polymerization, the cellular expression level of J chain determines the production of dimers versus monomers of IgA, and pentamers versus J chain-deficient hexamers of IgM (Brandtzaeg, 1985; Brewer et al., 1994; Wiersma et al., 1998; Sørensen et al., 1999; Johansen et al., 2000; Braathen et al., 2002). Notably, only J chain-containing polymers show spontaneous noncovalent interaction with pIgR or its cleaved extracellular portion, the so-called free SC (Brandtzaeg, 1973, 1974a, 1985; Eskeland and Brandtzaeg, 1974; Brandtzaeg and Prydz, 1984; Johansen et al., 2001). Most IgA1 (~90%) and virtually all IgA2 immunocytes in normal gut mucosa express substantial levels of cytoplasmic J chain (Crago et al., 1984; Kett et al., 1988), and the same is true for IgA immunocytes in other secretory tissues (Brandtzaeg and Korsrud, 1984; Bjerke and Brandtzaeg, 1990b). Most mucosal IgM immunocytes likewise produce J chain (Brandtzaeg, 1983b, 1985). By contrast, IgA immunocytes present in mesenteric, and particularly in typical systemic-type lymphoid tissue such as peripheral lymph nodes, show a much lower expression level of J chain (Brandtzaeg et al., 1999b). Direct evidence that J chain–positive IgA immunocytes do in fact produce pIgA was first obtained by a binding test on human tissue sections performed with free SC (Brandtzaeg, 1973, 1974a, 1985). Subsequent immunoelectron-microscopical localization of J chain in intestinal IgA immunocytes suggested that the IgA dimerization process begins in the rough endoplasmic reticulum (Nagura et al., 1979), a notion that was supported by similar studies of transformed normal lymphoid cells (Moro et al., 1990). Interestingly, 80 to 90% of the IgG immunocytes in normal intestinal mucosa produce J chain (Brandtzaeg and Korsrud, 1984; Bjerke and Brandtzaeg, 1986, 1990b; Nilssen et al., 1992), although it is not secreted from these cells but degraded intracellularly (Mosmann et al., 1978). The same is probably true for J chain produced by mucosal IgD immunocytes, which are almost 100% positive for this polypeptide (Brandtzaeg et al., 1979; Korsrud and Brandtzaeg, 1980; Brandtzaeg, 1983b). We have proposed
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
that J chain-expressing mucosal IgG and IgD immunocytes probably represent “spin-offs” from MALT-derived, relatively immature B-cell effector clones during their class switch and differentiation to pIgA production (Brandtzaeg et al., 1999a,b). This notion is strongly supported by the observation (Figure 15.3) that J chain-positive IgM, IgG, and IgD immunocytes numerically replace the normal immunocyte population in the secretory tissues of IgAdeficient subjects (Brandtzaeg et al., 1979; Brandtzaeg and Korsrud, 1984). Thus, differentiation and homing properties reflecting a mucosal B-cell phenotype are more closely related to J-chain than to IgA expression per se. This notion is further in keeping with the fact that Jchain expression is dramatically decreased in nonsecretory tissues—the only exception being the mesenteric lymph nodes and germinal centers of MALT structures (Brandtzaeg 1974b, 1983a; Brandtzaeg and Korsrud, 1984; Bjerke and Brandtzaeg, 1990b; Brandtzaeg et al., 1999b). Downregulation of J chain in extrafollicular B cells therefore appears to be a sign of clonal maturation according to the “decreasing potential” hypothesis, involving an enhanced tendency for terminal differentiation and apoptosis (Ahmed and Gray, 1996). Notably, B cells that undergo terminal differentiation just outside of MALT follicles generally show reduced J-chain and increased IgG production (Figure 15.2), suggesting that they belong to relatively exhausted effector clones that have been through several rounds of stimulation in germinal centers (Korsrud and Brandtzaeg, 1981; Bjerke and Brandtzaeg, 1986; Brandtzaeg et al., 1999a,b).
Regulation of J-Chain Expression Little is known about the factors causing the high levels of J chain in B cells that home from MALT to secretory effector sites (Figure 15.2). Transcriptional regulation of the J-chain gene involving cytokines such as interleukin (IL)-2, IL-4, IL-5, and IL-6 has been described in mice, and both positive and negative regulatory elements appear to be present in the promoter region (Tigges et al., 1989; Takayasu and Brooks, 1991; Randall et al., 1992; Shin and Koshland, 1993; Kang et al., 1998; Rao et al., 1998; Turner et al., 1994). Contrary to the situation in mice, transcription of the human J-chain gene is apparently initiated during early stages of B-lineage differentiation, even before Ig production takes place (McCune et al., 1981; Hajdu et al., 1983; Kubagawa et al., 1988; Max and Korsmeyer, 1985). Altogether, therefore, how the human J-chain is induced and regulated to provide a mucosal B-cell phenotype remains an enigma.
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B-CELL STIMULATION IN MALT STRUCTURES Antigen Encounter via FAE The bell-shaped M cells characteristic of FAE can sample luminal antigens unspecifically or by receptor-mediated uptake (Neutra et al., 2001); their pockets represent an intimate interface between the external environment and the mucosal immune system (Figure 15.2). Although no evidence exists for an antigen-presenting function of this specialized epithelial cell type, T and B cells as well as various MHC class II-expressing putative APCs are present immediately underneath the FAE (Bjerke and Brandtzaeg, 1988; Bjerke et al., 1993). The M-cell pockets are dominated by memory T and B lymphocytes in approximately equal distribution (Farstad et al., 1994; Yamanaka et al., 2001). Interestingly, experimental results suggest that lymphoid interaction, particularly involving activated B cells, can induce the epithelial M-cell phenotype (Kernéis et al., 1997). Studies in germ-free and conventionalized rats have furthermore demonstrated that bacterial colonization drives the accumulation and differentiation of T and B cells in the M-cell pockets, apparently with an initial involvement of antigen-transporting DCs, followed by germinal center formation (Yamanaka et al., 2003). The M-cell pockets may in fact represent specialized germinal-center extensions designed for rapid recall responses (Figure 15.4). The most likely cell type to mediate MHC class II interaction with cognate T cells in these microcompartments is the long-lived sIgD-IgM+Bcl-2+CD27+ memory B cells that express co-stimulatory molecules (Yamanaka et al., 2001). In human tonsils, memory B cells have been shown to colonize the antigen-transporting reticular crypt epithelium; by rapid upregulation of co-stimulatory B7 molecules, they acquire potent antigen-presenting properties (Liu et al., 1995). Likewise, we have found that memory B cells present in the M-cell areas of human Peyer’s patches express B7.2 (CD86) relatively often and sometimes B7.1 (CD80), which would be a prerequisite for the stimulation of productive immunity (Figure 15.4).
Molecular Interactions in Germinal Center Formation Role of Lymphotoxins and FDCs Primary lymphoid follicles contain recirculating naïve B lymphocytes (sIgD+IgM+), which pass into the network formed by antigen-capturing follicular dendritic cells (FDCs). The origin of FDCs remains obscure (Kapasi et al., 1998), but both their development and the clustering that allows follicle formation depend on lymphotoxin (LT) signaling (Gommerman et al., 2002). Experimental evidence suggests that B cells are one important LT source (Fu et al.,
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FIGURE
FIGURE 15.4 Schematic depiction of the relationship between elements of secondary (activated) B-cell follicle and M-cell pocket in human Peyer’s patch. The germinal center (GC) is mostly surrounded by the mantle zone (MZ) and filled with sIgD-CD20+CD80/86hiBcl-2-CD10+CD27- B cells. The MZ consists of naïve sIgD+CD20+CD80/86-Bcl-2+CD10-CD27- B cells but is broken (thick arrows) beneath the M-cell pocket. This connected area (defined by reduced sIgD, strong CD20, and variable CD80/86 immunostaining) is only seen in a restricted part of the follicle. The follicular–dendritic cell network (defined as CD21+CD20- phenotype) shows a topographically similar extension towards the M cell but does not reach inside the pocket. The M-cell pocket contains both naïve sIgD+CD20+CD80/86-/loBcl-2+CD10-CD27- and memory (or recently stimulated) sIgD-CD20loCD80/86hiBcl-2+CD10-CD27+ B-cell phenotypes, the latter being predominant. Adapted from Yamanaka et al. (2001).
1998; Tumanov et al., 2002). Among the known actions of the soluble homotrimer LTa, previously termed tumor necrosis factor (TNF)-b, is the augmentation of B-cell proliferation and adhesion molecule expression. Knockout mice deficient in LTa virtually lack lymph nodes and have no detectable Peyer’s patches. A membrane-associated form of LT exists as a heterotrimeric complex containing LTa together with a transmembrane protein designated LTb (a1b2). Knockout mice deficient in LTb have no detectable FDCs, and they lack Peyer’s patches, peripheral lymph nodes, and organized splenic germinal centers (Chaplin and Fu, 1998). Primary follicles are turned into secondary follicles by the germinal center reaction. In humans, this process has been extensively studied in tonsils (MacLennan, 1994; Liu and Arpin, 1997), but much relevant mechanistic information relies on the observations of lymph nodes and spleen from immunized animals (MacLennan et al., 1997). Germinal centers are of vital importance for the T cell–dependent generation of conventional (B2) memory B cells, affinity maturation of the B cell receptor (BCR), and Ig class switching. It has been shown that naïve B cells are first stimulated
15.5 Schematic depiction of adhesion molecule- and chemokine-regulated steps of T- and B-cell migration to, and positioning within, organized gut-associated lymphoid tissue (GALT) compartments. Antigens (dots) are sampled from the gut lumen by M cells (M) in GALT, whereas mesenteric lymph nodes receive antigens via draining lymph (either in soluble form or carried by dendritic cells; not shown). Naïve T and B cells enter both GALT and mesenteric lymph nodes (left panels) via high endothelial venules (HEV) by interactions principally between Lselectin (L-sel.) and endothelial MAdCAM-1 or PNAd distributed as indicated. Primed (memory/effector) T and B cells may to some extent reenter these sites by leukointegrin a4b7-MAdCAM-1 interactions. The chemokines involved (right panel) at the level of HEVs are SLC (CCL21) and ELC (CCL19), provided by stromal cells and redistributed to the HEV endothelium as indicated to preferentially attract CCR7+ naïve T cells and, less actively, B cells (broken arrow); SLC may also be involved in the exit of lymphoid cells from GALT via draining lymphatics. Naïve B cells are CXCR5+ and extravasate mainly via modified HEVs presenting CXCL13 (called BCA-1 in humans) juxtaposed to, or inside of, the lymphoid follicles; they are next attracted to the mantle zone, where BCA-1 is deposited on dendritic elements such as the follicular-dendritic cell (FDC) tips. Also follicular B-helper T (TFH) cells (CXCR5+CD4+CD57+) are attracted to the follicle by similar interactions. B cells are primed just outside the lymphoid follicle by interaction with cognate T cells and antigen-presenting cells (APC); they then re-enter the follicle and end up as CCR7+ germinal center cells after interactions with FDCs and TFH cells. The B cells may thereafter leave the follicle as memory or effector cells.
at the edge of the primary follicle (Figure 15.5) by cognate interaction with activated CD4+ T cells that have previously been presented with processed antigen by MHC class IIexpressing interdigitating DCs (Garside et al., 1998). The B cells then re-enter the follicle to become proliferating sIgD+IgM+CD38+ germinal center “founder cells,” as described in human tonsils (Liu and Arpin, 1997; Lebecque et al., 1997). Such initially stimulated B cells produce unmutated IgM (and some IgG) antibody of low affinity that can bind circulating antigen; the resulting soluble immune complexes subsequently become deposited on the FDCs, where antigen is retained for prolonged periods to maintain B-cell memory (Ahmed and Gray, 1996; Lindhout et al., 1997; MacLennan et al., 1997). Such a role for IgM in the induction of secondary immune responses with antibody affinity maturation has been strongly supported by observations in knockout mice lacking natural (nonspecific) background IgM antibodies (Ehrenstein et al., 1998).
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
The complement receptors CR1/CR2 (CD35/CD21) are considered among the cell surface molecules that play a crucial role in the germinal center reaction. CD21 is expressed abundantly on both FDCs and B cells, and may function by localizing antigen to the FDC network and/or by lowering the threshold of B-cell activation via recruitment of CD19 into the BCR (Tarlinton, 1998). Activation of complement on FDCs is controlled by regulatory factors when these cells retain immune complexes, but some release of inflammatory mediators may cause edema that facilitates the dispersion of FDC-derived immune complex-coated bodies, or iccosomes, thereby enhancing the BCR-mediated uptake of their contained antigens by B cells (Brandtzaeg and Halstensen, 1992). Role of Chemokines and Their Receptors Several homeostatic chemokines have been identified as major cues for lymphocyte trafficking and positioning in organized lymphoid tissue (Cyster, 1999; Moser and Loetscher, 2001). The CXC chemokine BCA-1 (B cellattracting chemokine-1)/CXCL13 (CXC chemokine ligand 13) is an attractant for naïve human B cells in vitro and has been shown to be produced in follicles of human lymph nodes (Legler et al., 1998). This chemokine was concurrently described in mice and called BLC (B-lymphocyte chemoattractant) (Gunn et al., 1998a). Several lines of evidence suggest that CXCL13, and its receptor CCR5, are directly involved in the formation of organized lymphoid tissue of mice (Förster et al., 1996; Luther et al., 2000). Interestingly, CXCL13 upregulates LT a1b2 on B cells, and a positive feedback loop may thereby be established (Ansel et al., 2000). The follicular expression of murine CXCL13 is reportedly more consistent in murine Peyer’s patches than peripheral lymph nodes (Gunn et al., 1998a). Alternative B cell-attracting chemokines may also operate in human lymphoid tissue. Indeed, stromal cell-derived factor 1 (SDF1)/CXCL12, which appears to be produced by cells lining tonsillar germinal centers, has been shown by an in vitro assay to attract naïve and memory B cells expressing CXCR4 (Bleul et al., 1998). CXCL13 (BCA-1) and CXCR5 were found to be expressed in normal human GALT structures, both in the ileum (Peyer’s patches) and colon (ILFs), as well as in irregular lymphoid aggregates of IBD lesions (Carlsen et al., 2002). The general expression of CXCR5 seen in follicular mantle zones (Figure 15.5) agreed with the notion that CXCL13 is a selective chemoattractant for naïve B cells from human blood in vitro, although with moderate effect (Legler et al., 1998), thus paralleling the relatively low receptor level observed in the mantle zones. Scattered T cells with strong CXCR5 expression are present within GALT follicles (Carlsen et al., 2002), and the CXCR5+CD4+ phenotype has been functionally described as
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follicular B-helper T cells, or TFH cells (Figure 15.5). It shows all the characteristics required for efficient B-cell help (Schaerli et al., 2000; Breitfeld et al., 2000; Moser et al., 2002). Nevertheless, flow-cytometric analysis of lymphoid cells from murine Peyer’s patches and human tonsils has revealed a much higher proportion of CXCR5+ T cells, implying the presence of this phenotype in the extrafollicular areas (Schaerli et al., 2000; Breitfeld et al., 2000). However, a small TFH-cell subset, identified as CXCR5+CD57+ and termed germinal center T-helper (GCTh) cells, appears to be essential for B-cell differentiation and antibody production (Kim et al., 2001); its exclusive germinal-center localization agrees with our immunohistochemical observations in tonsils, normal GALT, and IBDassociated lymphoid aggregates (Carlsen et al., 2002). The partial overlap produced by immunostaining for CXCL13 and several traditional FDC markers in human tissues suggested that this chemokine is deposited on the peripheral extensions of FDCs (Figure 15.5) after secretion by another cell type (Carlsen et al., 2002). Indeed, the main source of CXCL13 appeared to be the previously identified germinal center dendritic cell (GCDC) reported to stimulate T cells in this compartment (Grouard et al., 1996). Interestingly, both GCDCs and large CXCL13-producing cells in IBD-associated B-cell aggregates were found to exhibit a phenotype compatible with macrophage derivation (Carlsen et al., 2003).
Differentiation and Dispersion of Germinal-Center B Cells Positive Selection and Plasma-Cell Induction Germinal centers can be divided into different compartments in which the antigen-dependent selection of B cells takes place (MacLennan, 1994). Stimulation in the dark zone produces exponential growth of B-cell blasts positive for the Ki-67 nuclear proliferation marker (Brandtzaeg and Halstensen, 1992). The resulting centroblasts somatically hypermutate their Ig variable (V)-region genes and give rise to sIgD-IgM+CD38+ centrocytes. This process changes the affinity as well as specificity of the BCR and will likely induce some self-reactivity. However, mechanisms exist to eliminate autoreactive B-cell clones (Liu and Arpin, 1997; Lindhout et al., 1997; Pulendran et al., 1997). Also centrocytes with specificity for exogenous antigens undergo apoptosis unless selected by high affinity binding to FDCs via their sIgM/BCR. The centrocytes may actually pick up antigen from iccosomes (Brandtzaeg and Halstensen, 1992), process it, and present foreign peptide to cognate CD4+ TFH cells (Figure 15.5). The importance of cognate interaction between B and T cells is documented by the fact that no germinal centers are formed when CD40-CD40L (CD154) ligation is exper-
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imentally blocked (Lindhout et al., 1997). Moreover, this ligation promotes a switching of the CH genes from Cm to downstream isotypes, while apparently representing a negative signal for terminal B-cell differentiation within the follicles (MacLennan et al., 1997; Randall et al., 1998). The mechanisms contributing to the decision whether primed B cells should continue down the memory pathway or leave it and differentiate along the effector pathway remain elusive (Ahmed and Gray, 1996; Arpin et al., 1997). However, interaction between CD27 on CD38+ germinal center B cells and CD27L (CD70) on T cells may be a decisive event (Agematsu et al., 2000; Jung et al., 2000). Exit of B Cells from Germinal Centers Emigration of activated B cells from germinal centers is most likely directed by chemokines, and the actual cues may be extrafollicular ligands for CCR7 (Figure 15.5). Thus, activated germinal center B cells downregulate CXCR5 and upregulate CCR7, which in animal experiments have profound consequences for their positioning (Reif et al., 2002). In fact, most MALT-induced sIgD-IgM+CD38- putative memory B cells migrate continuously out of the germinal centers to sites such as the tonsillar crypt epithelium (Liu et al., 1995) or Peyer’s patch M-cell pockets (Yamanaka et al., 2001), where they presumably present recall antigens to cognate memory T cells (Figure 15.4). Likewise, most plasma cell precursors (CD20-CD38hi) become rapidly dispersed in juxtaposed extrafollicular compartments or migrate via lymph and blood to distant effector sites, where they undergo terminal differentiation (Figure 15.2).
CLASS SWITCH AND IgA ISOTYPE PROMOTION The Switching Process Following activation, naïve B cells usually first change their BCR composition from sIgD+IgM+ to become sIgDIgM+ memory cells, and may then switch to another class, such as IgG or IgA. During plasma-cell differentiation, the BCR is gradually lost, together with several other B-cell markers, particularly CD20 and then CD19 (in mice also B220). Activation-induced cytidine deaminase (AID) plays an essential role in this process (Kinoshita and Honjo, 2001). This enzyme is present during class-switch recombination (CSR) and may link CSR to the somatic hypermutation of Ig V-region gene segments that takes place during the germinal center reaction. Fagarasan et al. (2001) used AID knockout mice with defective CSR to study the accumulated switching potential of intestinal B cells harvested outside of Peyer’s patches. Like IgA-deficient humans (Figure 15.3), AID-deficient mice had numerous lamina propria IgM-
producing immunocytes (B220-), which gave rise to abundant SIgM. When sIgM+B220+ intestinal B cells from these mice were transformed with retrovirus to overexpress AID, they displayed a strong IgA switch propensity after in vitro stimulation with LPS, transforming growth factor (TGF)-b, and IL-5. The same tendency was observed for similarly harvested sIgM+B220+ B cells from normal mice in conjunction with AID upregulation, whereas cells of the same phenotype obtained from Peyer’s patches showed a lower IgA-switch efficiency under identical conditions. During CSR, the DNA between the switch sites is looped out and excised, thereby deleting Cm, which is followed either sequentially or directly by loss of other CH genes (Kinoshita and Honjo, 2001). After a direct switch to IgA expression, the Ia-Cm circular transcripts (aCTs), derived from the excised recombinant DNA, are gradually lost through dilution from progeny cells during proliferation (Fagarasan and Honjo, 2003). Therefore, readily detectable aCTs are considered a marker of recent CSR. Interestingly, aCTs are detectable in murine intestinal sIgA+B220+ B cells located outside of Peyer’s patches, which might suggest that lamina propria IgA immunocytes are derived directly from sIgM+B220+ B cells in situ (Fagarasan et al., 2001). This CSR is not believed to require CD40–CD40L interaction, therefore most likely involving T cell-independent B1 cells in a process engaging the BLyS/BAFF (B cell-activating factor of the TNF family) receptor (Litinskiy et al., 2002; Fagarasan and Honjo, 2003). BAFF or other proliferationinducing ligands, such as APRIL (Mackay and Browning, 2002), are secreted by activated DCs (e.g., after LPS exposure in the gut) and may thus operate in the intestinal lamina propria (Figure 15.2). It is of note, however, that AID-deficient mice show a dramatic hyperplasia of ILFs in response to the intestinal overgrowth of the indigenous microbiota (Fagarasan et al., 2002). This could reflect an inadequate compensatory antibody repertoire in the gut due to lack of somatic hypermutation in the SIgM that replaces the missing SIgA in these mice. It cannot be excluded, therefore, that Fagarasan et al. (2001) actually observed a difference in IgA-switching capacity between Peyer’s patches and hyperplastic ILFs rather than CSR outside of such GALT structures (Brandtzaeg et al., 2001).
Isotype-Switch Mechanisms Differ The fact that the IgA1 subclass dominates IgA responses both in tonsils and the related exocrine tissues supports the notion that mucosal B-cell differentiation in this body region mainly takes place from sIgD-IgM+CD38+ centrocytes by sequential downstream CH gene switching (Brandtzaeg, 1987; Brandtzaeg et al., 1999b). Conversely, the relatively enhanced IgA2 expression in Peyer’s patches and the distal gut altogether, including the mesenteric lymph nodes (Kett
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
et al., 1986; Bjerke and Brandtzaeg, 1990a,b), could reflect direct switching from Cm to Ca2 with the excision of intervening CH gene segments. B cells from murine Peyer’s patches are able to switch directly from Cm to Ca, and in human B cells this pathway may preferentially lead to IgA2 production (Conley and Bartelt, 1984). Molecular evidence for autocrine TGF-b–mediated switch region (S) recombination, either direct (Sm Æ Sa) or sequential (Sm Æ Sg, Sg Æ Sa), has been obtained in naïve human B cells after engagement of CD40 (Zan et al., 1998). However, although it is known that IgA expression induced by TGF-b involves the mobilization of the transcription factor CBFa3 (AML2), the critical role of this pleiotropic cytokine in IgA regulation remains elusive (Cazac and Roes, 2000). The germinal center reaction generates relatively more intrafollicular J chain–positive IgA cells in human Peyer’s patches and appendix than in tonsils (Brandtzaeg et al., 1999b). Also, in juxtaposed extrafollicular GALT compartments, IgA immunocytes are equal to or exceed in numbers their IgG counterparts (Bjerke and Brandtzaeg, 1986; Bjerke et al., 1986), whereas in tonsils there is a more than two-fold dominance of IgG immunocytes outside of the follicles (Brandtzaeg, 1987). Therefore, the drive for switching to IgA and the expression of J chain is clearly much more pronounced in GALT than in tonsils. Perhaps GALT is at least partially distinct from other MALT structures because of special accessory cells or a particular cytokine profile. Alternatively, the continuous superimposition of new exogenous stimuli in the gut may enhance the development of early effector B-cell clones having an increased potential for IgA and J-chain expression (Brandtzaeg et al., 1999b). A regionalized microbial impact on mucosal B-cell differentiation may be exemplified by the unique sIgD+IgM-CD38+ subset identified in the dark zone of palatine tonsillar germinal centers (Liu and Arpin, 1997). These centroblasts show a deletion of the Cm and Sm gene segments, therefore selectively giving rise to IgD immunocytes by nonclassical switching (Arpin et al., 1998). We also have obtained molecular evidence for the preferential occurrence of B cells with Cm deletion in normal adenoids and secretory effector tissues of the upper aerodigestive tract, but virtually never in small intestinal mucosa (Brandtzaeg et al., 2002). Such compartmentalized B-cell dispersion explains the relatively high frequency of IgD immunocytes normally occurring in this upper region, and particularly the large IgD-positive replacement subset that is often seen at secretory sites in IgA deficiency (Figure 15.3). Most strains of Haemophilus influenzae and Moraxella catarrhalis, which are frequent colonizers of the nasopharynx, produce an IgDbinding factor (protein D) that can crosslink sIgD/BCR (Ruan et al., 1990; Janson et al., 1991). In this manner, it is possible that sIgD+ tonsillar centrocytes are stimulated to proliferate and differentiate polyclonally, thereby driving Ig
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V-region gene hypermutation and Cm deletion (Liu et al., 1996; Arpin et al., 1998). Such regional microbial influence on B-cell differentiation is supported by our observation that Cm deletion is more frequently detected in diseased than in clinically normal tonsils and adenoids, and an increased number of extrafollicular IgD immunocytes occurs in recurrent tonsillitis and adenoid hyperplasia (Brandtzaeg, 1987). Molecular evidence strongly suggests that these extrafollicular plasma cells are indeed derived from the sIgD+IgM- centroblast subset (Arpin et al., 1998). IgD-deficient mice are sensitive to tolerance induction because sIgD protects B cells against deletion (Carsetti et al., 1993), whereas sIgM is associated with prohibitin and a prohibitin-related protein that transduces negative signals (Terashima et al., 1994). Therefore, sIgD+IgM- cells could also have a particular proliferative advantage when stimulated through their BCR. Conversely, LPS that is abundantly present in the distal gut may inhibit selective expression of IgD (Parkhouse and Cooper, 1977).
Identified IgA-Promoting Stimuli DCs from murine Peyer’s patches and spleen were initially suggested to enhance IgA production in a microculture system based on cognate interactions between B and T cells (Schrader et al., 1990). A similar role for DCs was later observed in a human in vitro system not including T cells, but employing CD40-activated naïve B cells (Fayette et al., 1997). As mentioned earlier, TGF-b appears to be a crucial IgA switch factor for activated (AID+) B cells, whereas IL2, IL-5, and IL-10 may be important for their expansion and terminal differentiation, perhaps with support from IL-6 and interferon-g (Figure 15.6). All these cytokines are known to be produced by antigen-activated CD4+ T cells from human intestinal mucosa (Nilsen et al., 1995) and are also expressed in human Peyer’s patches (Hauer et al., 1998; MacDonald and Monteleone, 2001). IL-6 has furthermore been reported to preferentially enhance IgA production (IgA2 > IgA1) by human appendix B cells (Fujihashi et al., 1991). A central role for IL-10 is supported by the fact that this cytokine can release the differentiation block of IgA-committed B cells from IgA-deficient patients (Briére et al., 1994). Human naïve B cells activated through CD40 can be pushed towards IgA production by TGF-b and IL-10 in combination (Defrance et al., 1992). Interestingly, DCs synergistically enhance the effect of both TGF-b and IL-10 on IgA expression and, via unknown signals, may be essential for the IgA2 phenotype (Fayette et al., 1997). Furthermore, neuroendocrine peptides may be involved in mucosal B-cell differentiation. Thus, human fetal B cells activated in vitro through CD40 were shown to be selectively induced by vasoactive intestinal polypeptide (VIP) to produce both IgA1 and IgA2 (Kimata et al., 1995). Similarly treated sIgM-CD19+ pre-B cells from human fetal bone
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FIGURE 15.6 Model for regulation and differentiation of B cells (B) in inductive mucosa-associated lymphoid tissue (left) leading to generation of plasma cells that mainly produce dimeric IgA with J chain (IgA + J) at secretory effector site (right). Antigen (Ag) is processed by antigenpresenting cells (APCs) at inductive site and presented to naïve CD4+ T cells (T) in the context of MHC class II molecules (MHC-II); this step is highly dependent on the co-stimulatory molecules B7 (CD80/86) and CD28. Activated T lymphocytes and other cells in the microenvironment secrete immunoregulatory factors such as cytokines and vasoactive intestinal polypeptide (VIP), which are important for various steps in mucosal Bcell differentiation, as indicated (boxes at the bottom). The co-stimulatory molecules CD40 and CD40L are crucial for the initiation of the switching process of Ig heavy chain constant region (CH)-genes in sIgM+ cells to become IgA expressing by looping-out of Ia-Cm circular transcripts (aCT), which thereafter are gradually lost. The class switch is facilitated by activation-induced cytidine deaminase (AID), which is highly upregulated in activated B cells, as indicated (AID++). Information obtained in experimental animals suggests that productive transcription of the J-chain gene depends on IL-2, IL-5, and IL-6, whereas IL-4 may have an opposing effect.
marrow were likewise induced to produce the two IgA subclasses, in addition to IgM. These results suggested that VIP can act as a true switch factor (Figure 15.6), which is interesting in view of its relatively high concentration in the gut. In cultures of intestinal mononuclear cells, VIP was also reported to enhance the number of IgA precursors, increase the synthesis of IgA, and decrease IgG production (Boirivant et al., 1994). Finally, substance P has been shown to promote both IgA and IgM production by murine B-cell lines; the latter isotype particularly in the presence of LPS (Pascual et al., 1991).
Switch to IgA Outside of Germinal Centers Natural antibodies secreted by B1 cells are generally encoded in germline (unmutated Ig V-region genes), but when produced in response to commensal gut bacteria such murine IgA often shows somatic mutation, which suggests a germinal center event (Bos et al., 1996). Nevertheless, although microbial colonization is a prerequisite to induce SIgA antibodies in mice, implying an antigen-induced process, no clear dependency on germinal centers or T cells has been revealed (Fagarasan and Honjo, 2000; Macpherson et al., 2000). Under certain conditions, IgA differentiation
driven by gut bacteria may even bypass the usual sIgM (or sIgD) BCR requirement (Macpherson et al., 2001); and the intestinal lamina propria is suggested, but not directly proven (see earlier sections), to be a potent site for switch to IgA (Fagarasan et al. 2001). The possibility remains that this could be true for B1 cells derived from the murine peritoneal cavity (Figure 15.2), but it appears to be of little or no relevance to the human gut, in which both IgA and IgM immunocytes have highly mutated Ig V-region genes— consistent with precursor selection in germinal centers (Dunn-Walters et al., 1997a; Fischer et al., 1998). Sequences of heavy chain V gene segments from human Peyer’s patch B cells are in fact clonally related to ileal lamina propria immunocytes (Dunn-Walters et al., 1997b), in accordance with a predominant derivation from GALT (Figure 15.2). Conversely, in the human peritoneal cavity IgM genes are mostly unmutated, and the mutated ones exhibit fewer mutations than corresponding genes from intestinal B cells (Boursier et al., 2002). Likewise, the IgVH4-34 genes used by IgG and IgA in human peritoneal B cells show significantly lower numbers of mutations than their mucosal counterparts. Altogether, there is no reason to believe that switching to IgA takes place to any significant degree in the human lamina propria. Even for murine B1 cells, the possibility remains that their precommitment to IgA is induced in the peritoneal cavity, because freshly isolated sIgM+IgA- cells from this site are reportedly class-switched at the DNA level (Hiroi et al., 2000). Notably, although some studies have suggested that murine B1 cells may depend on the microenvironment of mesenteric lymph nodes for plasma cell differentiation, the actual route and speed of migration of such cells to the intestinal lamina propria remain elusive (Fagarasan and Honjo, 2000).
MECHANISMS DIRECTING HOMING AND RETENTION OF MUCOSAL B CELLS Adhesion Molecules and Chemokines Operating in GALT Certain adhesion molecules guiding immune-cell extravasation are more strongly expressed on naïve than on primed (memory/effector) subsets, and vice versa, and some are relatively tissue-specific in their function. Counterreceptors expressed by endothelial cells may likewise show tissue specificity (Butcher and Picker, 1996). Thus, in human GALT and mesenteric lymph nodes, but not in peripheral lymph nodes, mucosal addressin cell adhesion molecule (MAdCAM)-1 is abundantly expressed by high endothelial venules (HEVs) (Brandtzaeg et al., 1999a). However, the microenvironmental factors that explain such
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preferential expression in the gut remain elusive (Denis et al., 1996; Brandtzaeg et al., 1999c). Human MAdCAM1 has also been cloned and characterized (Shyjan et al., 1996), and it is well documented in mice that this complex multidomain adhesion molecule plays a major role in the intestinal extravasation of immune cells (Streeter et al., 1988). When MAdCAM-1 is expressed by HEVs in murine GALT, the glycosylation of its mucinlike domain promotes the binding of L-selectin (CD62L) that is present at a high level on naïve lymphocytes (Berg et al., 1993). This initial endothelial adherence (tethering), together with the binding of leukointegrin a4b7 to the two N-terminal Ig-like domains of MAdCAM-1, is crucial for the preferential emigration of naïve lymphocytes into GALT structures such as Peyer’s patches. In addition, mesenteric lymph nodes employ mucin-like domains on peripheral lymph node addressin, or PNad (Figure 15.5). The less prominent GALT endowment with primed immune cells (a4b7hiL-selectinlo) may be mediated selectively by MAdCAM-1, because its interaction with a4b7 also supports tethering (Berlin et al., 1995). Interestingly, under flow conditions, the secondary lymphoid tissue chemokine (SLC/CCL21) stimulates a4b7-mediated human lymphocyte adhesion to MAdCAM-1, in contrast to other CC chemokines tested (Pachynski et al., 1998). Regardless of tissue site, an additional contribution to the emigration of both naïve and memory cells is provided by other more generalized adhesion molecules, such as leukocyte functionassociated molecule (LFA)-1 (aLb2 or CD11a/CD18) that binds to intercellular adhesion molecule (ICAM)-1 (CD54) and ICAM-2 (CD120) on the endothelium (Butcher and Picker, 1996). The phenotype-related distribution of adhesion molecules has been analyzed in human Peyer’s patches and appendix both by flow cytometry and immunohistochemistry (Table 15.1). The naïve B cells constituting follicular mantle zones generally express abundant L-selectin but variable levels of a4b7 and usually no b1 (CD29). Also lymphocytes positive for L-selectin found around or within the parafollicular HEVs are generally weakly positive or negative for a4b7. Notably, they are mostly naïve T cells (CD3+CD45RA+)—only some are B cells, again usually of the naïve (sIgD+) phenotype (Farstad et al., 1995, 1996, 1997a). Therefore, these vessels do not appear to be a major entrance site for B cells, as discussed below (Figure 15.5). The initial tethering of leukocytes to the endothelium is relatively loose until they are stopped by chemokine signaling through G protein–coupled seven-transmembrane receptors (Baggiolini, 1998; Kunkel and Butcher, 2002). SLC/CCL21, as well as Epstein-Barr virus–induced molecule 1 ligand chemokine (ELC)/CCL19, are produced by stromal cells in secondary lymphoid tissue and become transcytosed by HEV cells for presentation at the vascular surface (Figure 15.5). Both chemokines preferentially attract
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CCR7-expressing T cells, which are retained in the parafollicular areas (Gunn et al., 1998b; Campbell et al., 1998; Willimann et al., 1998; Baekkevold et al., 2001). The chemokines responsible for B-cell recruitment via HEVs are unclear. A recent mouse study indicated that although CCR7 ligands operating together with the CXCR4 ligand SDF-1/CXCL12 are crucial for endothelial B-cell adhesion in lymph nodes, B-cell entry in Peyer’s patches significantly depends on CXCR5 (Okada et al., 2002). CXCR5–CXCL13 interaction appeared to mediate the extravasation of B cells directly into the follicles via HEVlike vessels and not into the parafollicular zone via ordinary HEVs. The importance of this alternative extravasation pathway was supported by intravital microscopy that demonstrated T- and B-cell positioning at various vascular levels in murine Peyer’s patches (Figure 15.5), with B cells mainly adhering to SLC-negative vessels near or within the follicles (Warnock et al., 2000). CXCL13-positive vessels are also present in human tonsils and GALT structures (Schaerli et al., 2000; Carlsen et al., 2002).
Traffic of Naïve and Primed B Cells from GALT Immune cells exit from GALT through draining microlymphatics (Figure 15.5). In human Peyer’s patches and the appendix, these vessels are seen as thin-walled spaces lacking the endothelial expression of von Willebrand factor (Farstad et al., 1997a,b). Similar lymph vessels have been described in human tonsils (Fujisaka et al., 1996). Draining microlymphatics are believed to start blindly with a fenestrated endothelium, and the lymphoid cells probably enter them by selective mechanisms. Thus, lymph endothelium shares with HEVs the expression of both SLC and certain adhesion molecules (Gunn et al., 1998b; Irjala et al., 2003). In human GALT, memory B (sIgD-) and T (CD45R0+) cells with strong expression of a4b7 are often located near the draining microlymphatics, and also within them together with some CD19+CD38hia4b7hi blasts (Farstad et al., 1997a,b). However, the lymph vessels contain mainly naïve lymphocytes with low levels of a4b7. Cytochemical and flow-cytometric analyses of human mesenteric lymph has provided similar marker profiles; notably, the small fraction of identified B-cell blasts (2–6%) contained cytoplasmic IgA, IgM, and IgG in the proportions 5 : 1 : <0.5 (Farstad et al., 1997b). Altogether, the a4b7hi subsets identified at exit through lymphatics in human GALT may be taken to signify the first homing step particularly to populate the intestinal lamina propria with primed lymphoid cells (Figure 15.2). Relatively few memory cells concurrently expressed high levels of Lselectin in intestinal and mesenteric lymph (Farstad et al., 1997b); those that did might likely either re-enter GALT or
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extravasate in mesenteric lymph nodes, peripheral lymphoid organs, or Waldeyer’s ring together with naïve cells, by binding to PNAd expressed by HEVs (Bradley et al., 1996). A flow-cytometric study of circulating human lymphocytes supported such a fundamental subdivision according to vascular adhesion properties (Rott et al., 1996).
Adhesion Molecules and Chemokines Operating in Gut Lamina Propria A general consensus exists that the homing of primed lymphoid cells to the intestinal lamina propria depends on their high level of a4b7 in the absence of L-selectin (Figure 15.7), which allows binding to unmodified MAdCAM-1 on the lamina propria microvasculature (Butcher and Picker, 1996; Brandtzaeg et al., 1999a,c). This phenotype is predominantly induced on antigen-specific B cells appearing in human peripheral blood after enteric stimulation, whereas such cells elicited by systemic immunization preferentially show L-selectin but relatively little a4b7 expression (Quiding-Järbrink et al., 1995a, 1997; Kantele et al., 1996, 1997). Although the interaction of MAdCAM-1 with Lselectin has apparently been explored only in mice, the virtual absence in human intestinal lamina propria of lymphoid cells bearing L-selectin (Table 15.1) strongly suggests that it does not bind to MAdCAM-1 outside of GALT (Brandtzaeg et al., 1999a,c). Interestingly, many large B
cells retain high levels of a4b7 after migration into the human intestinal lamina propria, despite abundant coexpression of CD38 and cytoplasmic IgA (Table 15.1). Therefore, it is possible that a4b7, in addition to mediating extravasation, together with CD44 contributes to local retention of effector cells (see below). The thymus-expressed chemokine (TECK/CCL25) appears to have a decisive role in the migration of both T and B cells into, and/or retention within, the small intestinal lamina propria (Figure 15.7). Notably, this chemokine, which interacts with CCR9, is selectively produced by the crypt epithelium in this part of the gut (Kunkel et al., 2000; Papadakis et al., 2000; Bowman et al., 2002, Kunkel and Butcher, 2002). In the large intestine, the mucosae-associated chemokine (MEC/CCL28) has recently been identified as a decisive cue for attracting IgA plasmablasts, which express high levels of the corresponding receptor, CCR10 (Kunkel et al., 2003). Surprisingly, T cells are not directed by this chemokine. Because the epithelial expression of MEC is much higher in the colon than in the appendix and small intestine (Pan et al., 2002; Wang et al., 2000), this chemokine probably plays a compartmentalized role in intestinal B-cell homing. The cues that determine extravasation of GALT-derived circulating B cells in gut mucosa, may also be involved in the migration of primed cells from GALT structures directly into the lamina propria. There are direct although limited vascular connections from Peyer’s patches to the villi immediately surrounding them, and these channels can be used for trafficking of B cells (Parrott, 1976). Such a mechanism could explain why the first IgA immunocytes occur around Peyer’s patches when germ-free mice are transferred to conventional conditions (Crabbé et al., 1970). It seems likely that there are similar direct pathways from ILFs to the surrounding lamina propria (Figure 15.2). This notion is strongly supported by the report that the intestinal immunocyte population was remarkably well retained in rats when the B-cell traffic through the thoracic duct was diverted by lymph cannulation (Mayrhofer and Fisher, 1979). However, as discussed earlier, many of those B cells that normally settle immediately adjacent to GALT follicles apparently belong to exhausted clones (Ahmed and Gray, 1996) with decreased J chain–expressing potential and a disproportionately increased class switch to IgG (Figure 15.2).
FIGURE 15.7 Schematic depiction of putative homing mechanisms that preferentially attract gut-associated lymphoid tissue (GALT)-derived T and B memory/effector cells to human small intestinal lamina propria. Interaction between unmodified (containing no L-selectin-binding O-linked carbohydrates) MAdCAM-1 expressed on ordinary flat lamina propria venules (LPV) is important for the targeting of primed a4b7-bearing cells to all segments of the gut. The level of this activated integrin is particularly high on lymphoblasts. Adherence to the endothelium is strengthened by interactions between generalized adhesion molecules such as LFA-1 and ICAM1/ICAM-2, as indicated. Selectively produced by the epithelium of the small intestine, the chemokine TECK (CCL25) attracts GALT-derived lymphoid cells that express CCR9 only to this segment of the gut.
Mucosal Homing Molecules Operating Beyond the Gut Although GALT-derived dissemination of secretory immunity to extraintestinal effector sites is well documented, migration to the gut of B cell induced in NALT or BALT appears to be quite limited, as revealed by immunization or infection experiments in rodents and pigs (McDermott and Bienenstock, 1979; Sminia et al., 1989;
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Van Cott et al., 1994). On the other hand, considerable indirect evidence summarized elsewhere suggests that the dispersion of primed pIgA precursor cells takes place from Waldeyer’s ring (human NALT) to regional secretory effector sites (Brandtzaeg, 1999). Even more convincing results have been obtained by the immunization of murine NALT (Yanagita et al., 1999) and rabbit palatine tonsils (Inoue et al., 1999). Such putative B-cell homing dichotomy between the upper and lower body regions is supported by the disparate dispersion of human tonsillar sIgD+IgMCD38+ centroblasts, identified by tracking of their Cm-gene deletion (Brandtzaeg et al., 2002). We believe that their distribution reflects the homing properties of all B cells with a mucosal phenotype (J chain–expressing) primed in Waldeyer’s ring. In keeping with this notion, activated human tonsillar B cells transferred intraperitoneally to mice with severe combined immunodeficiency (SCID) migrated to the lung but not to gut mucosa (Nadal et al., 1991). Several studies have suggested that a4b7 is not an important homing receptor for lymphoid cells in the airways of humans (Picker et al., 1994), mice (Wagner et al., 1996), or sheep (Abitorabi et al., 1996). Intranasal immunization in humans induced an insufficient level of a4b7 on specific B cells to make them gut-seeking, whereas antibody production was evoked in both adenoids and nasal mucosa (Quiding-Järbrink et al., 1995b). Notably, the circulating specific B cells showed substantial co-expression of Lselectin and a4b7, in contrast to the high level of a4b7 induced on antibody-producing cells by enteric immunization (Quiding-Järbrink et al., 1997). A nonintestinal homing receptor profile might also explain B-cell migration from NALT to the urogenital tract. This putative link is reflected by particularly high levels of specific IgA and IgG antibodies in the cervicovaginal secretions of mice and monkeys after intranasal immunization with a variety of antigens (Brandtzaeg, 1997; Johansson et al., 2001). A relatively consistent level of L-selectin on NALT-derived B cells, allowing them to bind to PNAd on lymph node HEVs, could furthermore explain the substantial integration between mucosal immunity in the upper aerodigestive tract and the systemic immune system (Rudin et al., 1998; Johansson et al., 2001). B-cell migration to secretory tissues outside the gut might also involve a4b1 (CD49d/CD29) interactions. The chief counter-receptor for this integrin is vascular-cell adhesion molecule (VCAM)-1, which is expressed on microvascular endothelium in human bronchial and nasal mucosa (Bentley et al., 1993; Jahnsen et al., 1995). However, no evidence exists to suggest that a high expression of b1 integrin consistently directs primed B cells to the upper aerodigestive tract, lungs, or urogenital tract. CCR10 appears to be a unifying chemokine receptor for primed mucosal B cells (but not T cells) by affording homing to intestinal as well as extraintestinal secretory
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effector sites. This receptor has recently been shown to be expressed by human IgA plasmablasts (and less so by plasma cells) at every studied mucosal effector site (Kunkel et al., 2003). The corresponding ligand MEC is produced by secretory epithelia all over the body and at relatively high levels in the upper aerodigestive tract (Pan et al., 2000; Wang et al., 2000). Interestingly, MEC (but not TECK) was shown to attract tonsillar IgA plasmablasts in an in vitro assay (Kunkel et al., 2003). Therefore, graded tissuedependent CCR10-MEC interactions, together with insufficient levels of classical gut-homing molecules, most likely explain the observed dispersion dichotomy for effector B cells derived from Waldeyer’s ring.
Signals for B-Cell Retention, Proliferation, and Terminal Differentiation Experiments in gene-manipulated mice have indicated that signaling through LTbR on lamina propria stromal cells is necessary for the presence of B cells and IgA immunocytes in gut mucosa (Kang et al., 2002; Newberry et al., 2002). Adhesion molcules as well as chemokines and chemokine receptors may also be involved in cellular retention. One interesting but unproven candidate in this respect is the extracellular matrix receptor CD44, which is expressed at high levels on post-germinal center B cells (Kremmidiotis and Zola, 1995). Little decisive knowledge likewise exists about factors triggering terminal B-cell differentiation at secretory effector sites, although IL-5, IL-6, and IL-10 have been suggested to be particularly important (Figure 15.6). Notably, topical exposure to antigen appears to have a marked impact on the site-specific accumulation of IgA-producing cells (see below), thereby influencing the observed homing pattern but without imposing any selectivity on the extravasation step. Thus, GALT-derived blasts home to presumably antigen-free neonatal intestinal mucosa (Halstead and Hall, 1972) and to fetal gut grafted under the adult kidney capsule of experimental animals (Guy-Grand et al., 1974; Parrott and Ferguson, 1974). The impact of exogenous antigens on the conventional B2 cell-dependent effector arm of the SIgA system is most likely mediated largely via “second signals” from activated cognate T cells. However, several other factors may contribute to the site-specific survival and restimulation of memory T cells (Bode et al., 1997). Compartmentalized variables could be a high density of MHC class II molecules (Matis et al., 1983) that allow only trace amounts of foreign antigens or anti-idiotypic antibodies to elicit sufficient second signals for B cells. Interestingly, in human salivary and lactating mammary glands, IgA immunocytes preferentially accumulate adjacent to HLA-DR-expressing epithelial ducts (Newman et al., 1980; Brandtzaeg, 1983a; Thrane et al., 1992).
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Site-specific survival differences of IgA immunocytes might directly or indirectly (via activated T cells) be influenced by similar variables, including rescue from apoptosis by contact with stromal cells (Merville et al., 1996). It has been suggested that the half-life of plasma cells varies from a few days to several months, and those ending up in the gut may be particularly short-lived (Ahmed and Gray, 1996; Slifka and Ahmed, 1998).
Role of Topical Antigen Exposure Substantial antigen-driven proliferation of IgA cells has been observed in the intestinal lamina propria of experimental animals (Pierce and Gowans, 1975; Lange et al., 1980), especially in the crypt regions (Husband, 1982). Most IgA immunocytes likewise occur in the human gut (Brandtzaeg and Baklien, 1976), around the crypt openings either by direct extravasation or as a result of a certain local proliferation of sIgA+ memory/effector cells (Farstad et al., 2002). Although a stimulatory effect of topical antigen outside Peyer’s patches has been demonstrated in terms of localization, magnitude, and persistence of human SIgA antibody responses (Ogra and Karzon, 1970), the role of ILFs is difficult to evaluate in such experiments. Thus, rectal immunization elicits particularly high levels of IgA antibodies in colorectal secretions and feces, both in experimental animals (Hopkins et al., 1995) and humans (Kantele et al., 1998), apparently reflecting an enhanced stimulation by combined exposure of ILFs and the lamina propria to the same antigen. Repeated vaginal or rectal immunization in monkeys has likewise demonstrated local accumulation of effector B cells at the respective sites (Eriksson, 1998). Altogether, it appears that primed immune cells tend to accumulate preferentially at effector sites that correspond to the inductive site where they were initially stimulated. Rapid and widespread dissemination of fed antigen (presumably carried by DCs) followed by extravasation and activation of specific T cells at sites of cognate antigen presentation has been observed in T-cell receptor transgenic mice (Gütgemann et al., 1998). This observation, together with previous results of adoptive B-cell transfer in syngeneic rats (Dunkley and Husband, 1990), supports the notion that local antigen-driven T-cell activation provides important second signals for the retention, proliferation, and terminal differentiation of Ig-producing immunocytes at secretory effector sites, even at a long distance from mucosal surfaces. Nevertheless, the density of immunocytes at a particular effector site generally reflects the level of topical antigen exposure, being seven times higher in human colonic mucosa (which has an enormous microbial load) than in parotid and lactating mammary glands (Brandtzaeg, 1983a). It is not likely that live or dead exogenous material normally gains direct access to the latter sites, whereas the lacrimal gland, which is connected by many short ducts to the
excessively aeroantigen-exposed conjunctiva, shows an IgA immunocyte density approaching that of the colon (Brandtzaeg, 1983a).
WHAT IS ACTUALLY KNOWN ABOUT HUMAN MUCOSAL B CELLS? It is well established that the human mucosal B-cell system responds to an infection with local IgA and IgM production (Söltoft and Söberg, 1972). The level of this response appears to determine whether clinical symptoms will occur (Agus et al., 1974; Brandtzaeg and Johansen, 2003). However, there are many open questions regarding the nature and regulation of this large antibody system. Mechanistic information about mucosal B cells is to a great extent based on animal experiments, with the inherent problem of species differences. The following facts and puzzles related to the human secretory antibody system can be listed on the basis of the above discussion: • Secretory immunity depends on an intimate cooperation between mucosal B cells and exocrine epithelia. The biological significance of the striking J-chain expression shown by MALT-derived immunocytes dispersed to secretory effector sites is thus that pIgA and pentameric IgM with high affinity for the pIgR are produced locally and become readily available for export to the mucosal surface. This important functional goal, in terms of clonal differentiation, appears to explain why J chain is also expressed by mucosal B cells terminating their differentiation with IgG or IgD production; such immunocytes may be considered as “spin-offs” from early effector clones that, through class switch, are on their way to pIgA production. • Considerable evidence supports the notion that intestinal immunocytes are largely derived from B cells initially induced in GALT. Nevertheless, insufficient knowledge exists concerning the relative importance of M cells, MHC class II-expressing epithelial cells, B cells, and other professional APCs in the transport, processing, and presentation of luminal antigens that take place in GALT to accomplish the extensive and continuous priming and expansion of mucosal B cells. Also, it is not clear how the germinal-center reaction in GALT so strikingly promotes class switch to IgA and expression of J chain. • Although the B-cell migration to the intestinal lamina propria is guided by rather well-defined adhesion molecules and chemokines and chemokine receptors, a better definition of chemotactic stimuli determining homing mechanisms in different segments of the gut is required. This is even more true for homing of mucosal B cells to secretory effector sites beyond the gut, and in
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this respect, the role of Waldeyer’s ring as a regional inductive lymphoid tissue needs further characterization. • In addition to homing molecules, the retention and accumulation of B cells extravasated at secretory effector sites is influenced by antigen-driven local proliferation and differentiation. However, the role of cognate T cells, MHC class II-expressing APCs, and epithelial cells in providing the necessary stimulatory signals remains poorly defined. • Compartmentalization of the mucosal immune system must be taken into account in the development of effective local vaccines to protect the airways, eyes, oral cavity, and urogenital tract. Even without employing the classical gut-homing receptors, selective migration of putative early effector B-cell clones with preferential expression of J chain and pIgA is just as remarkable to secretory tissues in the upper aerodigestive tract as to the intestinal lamina propria. Future studies will hopefully help to elucidate the complexity of molecular mechanisms underlying this basic principle of secretory immunity. • It is also important to point out that clinical observations in immunodeficient patients have shown that SIgA, SIgM, and IgG antibodies are not the only important components of the mucosal immune system. Evidence is accumulating to reveal that innate defense mechanisms are much more crucial and complex than previously believed. The cooperation between innate and adaptive immunity must be further explored to better understand how the homeostasis of mucous membranes is normally maintained.
Acknowledgments Studies in the Brandtzaeg laboratory are supported by the University of Oslo, the Research Council of Norway, the Norwegian Cancer Society, Anders Jahre’s Fund, and Rakel and Otto Kr. Bruun’s Legacy. Studies in the Lamm laboratory were supported by the National Institute of Allergy and Infectious Diseases. Hege Eliassen and Erik K. Hagen are gratefully acknowledged for excellent assistance with the manuscript.
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16 The Cellular Basis of B Cell Memory KLAUS RAJEWSKY
ANDREAS RADBRUCH
Center for Blood Research, Harvard Medical School, Boston, Massachusetts, USA
Deutsches Rheumaforschungszentrum Berlin, Berlin, Germany
If we define memory as “the capacity for retaining, perpetuating, or reviving the thought of things past” (The New Shorter Oxford Dictionary), then the central feature of immunological memory is the capacity of the immune system to retain, upon a first encounter of antigen but subsequently in its absence, the capability of sustained antibody production and an enhanced response to antigenic rechallenge. Memory formation is often discussed in the context of another mechanism of long-term immunity, namely the ability of the immune system to store antigen over long periods of time on the surface of follicular dendritic cells and to entertain a chronic immune response on this basis. Although there is good reason to believe that memory formation and reactivity to persisting antigen both contribute in a concerted and interwoven fashion to the protection of the organism against reinfection by pathogens and memory formation by itself may be often insufficient to confer protection (Ahmed and Gray, 1996; Ochsenbein et al., 2000), memory formation, cellular selection by persisting antigen, and protection against pathogens are all separate issues. In this chapter, we concentrate on mechanisms of (true) immunological memory in the humoral immune system of mouse and human. These are based on two specific modes of cellular differentiation, one resulting in the generation of long-lived, antigen-independent memory cells that respond to renewed antigenic challenge by the rapid differentiation into plasma cells and production of secreted antibodies, the other generating a compartment of longlived plasma cells dedicated to long-term antibody production independent of sustained antigenic challenge. Both types of cells are selected in the germinal center reaction for the production of high-affinity antibodies resulting from somatic hypermutation of rearranged antibody variable region genes. Affinity maturation and the germinal center
reaction are discussed in detail in a separate chapter of this book. Both memory and long-lived plasma cells are generated, and later reside, in defined cellular microenvironments whose integrity is critical for the survival of the cells. Upon antigenic re-encounter, memory cells recruit T cell help through efficient antigen presentation, which in turn drives their expansion and terminal differentiation, thus replenishing the plasma cell pool.
Molecular Biology of B Cells
GENERATION OF B CELL MEMORY AND MEMORY B CELLS IN T CELLDEPENDENT ANTIBODY RESPONSES Memory is a classical phenomenon in humoral immunity. Its two main features are enhanced and more rapid antibody formation upon re-exposure to antigen, in concert with an increase in antibody affinity for the antigen. In the late 1960s, it became clear that memory in the B cell compartment typically develops in T cell–dependent immune responses. A classical readout system was the adoptive secondary antibody response in mice, in which antigenprimed T helper cells and B cells were combined in irradiated recipients and stimulated by antigen (Mitchison, 1971). It soon became clear that the requirements for the activation of primed, as compared to naïve B cells, by antigen were different, the former responding to lower doses of antigen and not requiring adjuvants (see below). It also became clear that the production of high-affinity antibodies in secondary responses was a (memory) B cell–specific phenomenon. Soon thereafter, when the methods of molecular biology and of monoclonal antibody production became available, it was discovered that these high-affinity antibodies, as a rule,
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carried somatic point mutations in their variable (V) regions and that affinity maturation of antibodies was based on the gradual acquisition of such mutations and their selection, following primary immunization. A typical feature of primary, T cell–dependent immune responses is the formation (and later resolution) of germinal centers (GCs) in secondary lymphoid organs, sites of intense, oligoclonal B cell proliferation and long suspected to be responsible for the generation of B cell memory. When it was found that the B cells proliferating in the GC environment were undergoing stepwise somatic hypermutation of their rearranged V region genes and high-affinity antigen binding mutants were positively selected, a clear scenario of the generation of B cell memory was at hand: Following primary exposure to antigen, memory B cells expressing somatically mutated antigen receptors with high affinity for the immunizing antigen are positively selected by antigen presented to them in the GC environment in a process of rapid somatic evolution, and these cells subsequently persist in the immune system (for review see Rajewsky, 1996). Although this general scheme still holds, a few important additions must be made. Thus, recent evidence obtained in mouse mutants, in which the GC reaction is impaired, as well as in autoimmune mice, suggests that somatic hypermutation and memory cell generation may not be entirely restricted to the GC environment (Kato et al., 1998; Matsumoto et al., 1996; William et al., 2002) and, more important in the present context, that functional memory B cells can be generated in the absence of both GCs and somatic hypermutation (Toyama et al., 2002). The latter result is in accord with earlier findings that memory cell populations invariably contain a fraction of cells expressing unmutated antibodies. We also want to mention that, while the precursor– product relationship of naïve and memory B cells is undebated, there has been the notion that subsets of naïve B cells may be predetermined to either the production of primary or secondary responses (Linton et al., 1989). If such a distinction indeed exists, it is likely not an absolute one (Jacob and Kelsoe, 1992).
Subsets and Properties of Memory B Cells Because in mouse and human naïve B cells express IgD and/or IgM on the cell surface, the GC is a major site of isotype switching, and secondary antibody responses to T cell–dependent antigens are dominated by isotypes other than IgM and IgD, memory B cells were expected to express IgG, A, or E in their antigen receptors (BCRs), thus opening a way to their specific isolation. Indeed, when IgM-, IgDsplenic B cells were isolated on the basis of their reactivity with antibodies specific for a B lineage marker, they were found in their majority to express somatically mutated antibodies, in contrast to their IgM/IgD expressing counterparts,
thus qualifying as memory B cells (Schittek and Rajewsky, 1992). The search for other memory B cell-specific markers was not very successful (reviewed by Gray, 1993), until Liu and colleagues discovered a unique surface marker combination on human memory B cells (Liu et al., 1994; Pascual et al., 1994). It was found that in mouse and man, memory B cells express high levels of Fas (Liu et al., 1995; Takahashi et al., 2001), a property they share with GC B cells from which, however, they can be distinguished using other markers. Most important, however, human memory B cells appear to share surface markers that distinguish them from other B cell subsets. The most commonly used such marker is CD27, a member of the tumor necrosis receptor superfamily. Monoclonal antibodies against this protein allow a clean separation of all human B cells that carry somatically mutated immunoglobulin (Ig) gene rearrangements (Klein et al., 1998b; Tangye et al., 1998) and also resemble memory B cells in functional terms (see below). It should be kept in mind, however, that mature B cells begin to express CD27 already in the GC reaction and then continue to express this marker all through plasma cell differentiation (Jung et al., 2000; Odendahl et al., 2000; van Oers et al., 1993). Using the CD27 marker, an amazing heterogeneity of human memory B cells became apparent. Apart from the familiar isotype-switched cells, the B cell population circulating in the blood harbors a large fraction of CD27-positive B cells expressing IgM as the only Ig isotype as well as cells expressing IgM and IgD, such as naïve B cells and a minute, curious subset of IgD-only cells. The IgM-only cells may represent memory cells that have attempted, but failed, to switch isotype, because they are impaired in their ability to undergo switch recombination upon in vitro stimulation (Werner-Favre et al., 2001). Together, all these cells that express somatically mutated antibodies make up roughly 40% of the B cells in the peripheral blood of adult individuals (Klein et al., 1998a, 1998b). In contrast, in the mouse almost no memory cells (defined as isotype-switched cells) are detectable in the blood, and in the spleen these cells represent only 5 to 10% of the total B cells. This difference may have to do with the different lifespans of mouse and human, allowing for the accumulation of a large fraction of memory cells only in the latter. However, other factors may also be involved. Thus, because of the lack of appropriate surface markers, certain memory B cell subsets may have escaped detection in mice. This is clearly the case for the IgMexpressing memory cells that are known to exist in mice (Dell et al., 1989; Shinall et al., 2000), but for which no specific marker was available. In addition, recent work suggests that in mice additional subsets of memory B cells may exist that curiously lack typical B lineage markers like B220 and CD19 and may represent memory cells differentiating in the direction of plasma cells (Driver et al., 2001). It should also be kept in mind that the CD27-expressing B cells in the human may not necessarily altogether represent memory B
16. The Cellular Basis of B Cell Memory
cells resulting from an antigen-driven response. Somatic mutation has been identified in the sheep as a mechanism of primary repertoire diversification (Reynaud et al., 1995), and presently it cannot be excluded that a similar mechanism operates in human but not mouse. Evidence in favor of such a possibility is the presence of IgM+IgD+, CD27positive, and somatically mutated B cells in patients with hyper-IgM-syndrome, in which the GC reaction is impaired because of CD40 deficiency (Agematsu et al., 1998b). Arguing against this interpretation are the functional properties of these cells, which resemble those of “classical” memory B cells (see below). Memory in the compartment of IgD- and/or IgM-expressing cells is not a peculiarity of the human, but has also been observed in other species (Herzenberg et al., 1980; Schirrmacher and Rajewsky, 1970; White and Gray, 2000). The classical notion that memory B cells can be more easily and rapidly triggered to differentiate into antibodysecreting plasma cells than naïve B cells (for review see Vitetta et al., 1991) is amply borne out in a variety of more recent studies (Agematsu et al., 1995; Arpin et al., 1997; Maurer et al., 1992; Tangye et al., 2003). Significantly, the interaction of CD27 with its ligand, CD70, contributes to memory B cell differentiation into plasma cells (Agematsu et al., 1998a). CD70 is dominantly expressed by T lymphocytes such that CD70–CD27 interaction may be involved in the delivery of T cell help to memory B cells, a critical element in the activation of the latter in T cell–dependent antibody responses. Memory B cells are well equipped to recruit T cell help in that they efficiently present antigen in the context of MHC class II and are prone to receive T cell co-stimulation because of rapid upregulation of B7-1 and -2 upon or even before activation (Bar-Or et al., 2001; Liu et al., 1995). The higher levels of adhesion molecules expressed on the surface of memory (as compared to naïve) cells may make these cells additionally prone to engage in interactions with T cells (Maurer et al., 1990). Another factor likely contributing to the efficient recruitment of memory cells into secondary responses relates to the isotype switch that has taken place in a significant fraction of these cells during their differentiation in the GC. The IgG, -A, and -E antibodies expressed in the BCRs of those cells differ from the IgM and -D bearing receptors of naïve B cells in that their Ig heavy (H) chains possess evolutionary conserved, potentially signal-transducing cytoplasmic tails that might make the cells uniquely responsive to BCR-mediated signals (Wakabayashi et al., 2002). Indeed, using transgenic mouse models, evidence could be obtained that the cytoplasmic tail of the g1 H chain is not only required for the generation of IgG1-expressing memory cells (Kaisho et al., 1997), also shown for the cytoplasmic tail of the e H chain (Achatz et al., 1997), but also enhances their “burst size” upon antigenic stimulation, presumably by promoting cellular survival (Martin and Goodnow, 2002). Although this
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mechanism is specific for isotype-switched memory cells, recent evidence indicates that human memory cells are in general more prone to stimulation via the BCR than naïve B cells, because of the downregulation of genes that negatively control BCR signaling and upregulation of their counterparts (Feldhahn et al., 2002). Thus, analyzed from a variety of angles, memory B cells consistently display properties that distinguish them from naïve B cells and enable them to efficiently engage in secondary antibody responses upon contact with antigen.
Localization and Recirculation of Memory B Cells Memory B cells have been identified in mice, rats, and humans by functional and molecular criteria in the marginal zone of the spleen, an area surrounding B cell follicles (Dunn-Walters et al., 1995; Liu et al., 1988). In human tonsils, they are mainly associated with the mucosal crypt epithelium (Liu et al., 1995). In gut-associated lymphatic tissues (GALT) cells of a similar phenotype are found under the dome epithelium of Peyer’s patches (Spencer et al., 1985) and in the inner wall of the subcapsular sinus of mesenteric lymph nodes (Stein et al., 1980). Another site of memory cell accumulation may be the bone marrow (Manz et al., 1998; McHeyzer-Williams et al., 2000; O’Connor et al., 2002), where long-lived plasma cells are also localized (see below). Finally, a large fraction of the B cells in the blood of adult humans are memory cells that are either on the way from their site of production or recirculating (Fig. 16.1). B cell memory is thus exported from the site of its generation and established throughout the organism. However, it should be kept in mind that while there is some evidence for memory cell recirculation (Laichalk et al., 2002), its extent remains to be determined and memory cells may be rather sessile cells once they have reached their favored location and are not activated by antigen (Gowans and Uhr, 1966; Liu et al., 1988, 1991). The migratory pathways of memory cells likely depend on the expression of chemokine receptors (Bleul et al., 1998) and adhesion molecules. It is of interest in this context that human memory cells exhibit diversity in terms of the expression of adhesion molecules such that subsets of memory cells may be destined to settle in different parts of the body, such as IgA-producing cells in the GALT and nasal-associated lymphoid tissue (Rott et al., 2000; Shimoda et al., 2001) (Figure 16.1).
The Lifespan and Homeostasis of Memory B Cells As we have discussed above and will again discuss in the context of humoral memory for plasma cells, memory B cells occupy certain “niches” in the lymphatic system that
crypt epithelium
marginal zones
subcapsular sinus
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R
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Peyer's patches
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FIGURE 16.1 Schematic view of traffic and location of memory B and memory plasma cells.
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likely determine their lifespan and homeostasis (Freitas and Rocha, 2000). Therefore, a faithful analysis of lifespan and homeostasis depends on the histological integrity of the lymphatic tissue. In the case of memory B cells, this is highlighted by the failure of keeping these cells functionally alive upon transfer into irradiated recipients in the absence of added antigen (Askonas et al., 1972; Barrington et al., 2002; Celada, 1967; Gray and Skarvall, 1988). Whereas this finding was initially interpreted as an absolute dependency of memory cells on persisting antigen (inadvertently disqualifying these cells as carriers of memory), subsequent work has reinforced that true B cell memory indeed exists. This does not imply, of course, that persisting antigen does not also contribute to the maintenance of long-term immunity and immune protection. The capacity of the immune system to store for long periods antigen complexed with antibodies and components of the complement system on the surface of follicular dendritic cells (FDCs) (Mandel et al., 1980) surely has its purpose. In antigen-primed mice, memory B cells persist for long periods, probably for the lifetime of the animals (Schittek and Rajewsky, 1990; Sprent and Tough, 1994). Although these cells are generated from proliferating progenitors in the GC, they are largely in a resting state (Maruyama et al., 2000; Schittek and Rajewsky, 1992; Toyama et al., 2002). In one study, only 10% of the memory cell population in the spleen (characterized as antigen-binding, isotype-switched, functional memory cells) incorporated bromodeoxyuridine (BrdU) over a period of 18 days, with labeling starting 140 days after primary immunization (Schittek and Rajewsky, 1990). It is unclear whether this residual proliferative activity reflected a very slow self-renewal of the memory population or the recruitment of newly generated cells into the memory pool. As in the mouse, memory cells in the human are largely in a resting state (Bar-Or et al., 2001; Liu et al., 1995). Although these experiments established that most memory B cells are long-lived cells, they leave unanswered the question of their antigen independency. This problem is difficult to study in the intact animal in which the presence of persisting antigen can hardly be excluded beyond doubt. In a recent experiment, this problem was circumvented by generating a mutant mouse strain in which the BCR specificity of the memory cells could be inducibly changed through a genetic switch, such that the cells were unable to “see” the antigen in response to which they had originally been generated. These altered cells were maintained in the animals like the wildtype cells over the period of observation (106 days) (Maruyama et al., 2000). In line with these results are data showing that the persistence of memory B cells is at least largely independent of T cell help (Takahashi et al., 1998; Vieira and Rajewsky, 1990) and that memory B cells can be maintained in mouse mutants lacking the
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follicular dendritic cell network (Karrer et al., 2000; Matsumoto et al., 1996). There is, thus, strong evidence that long-lived, antigenindependent memory B cells indeed exist. After an initial, dramatic expansion in connection with the GC response, their frequency in the spleen of mice stabilizes over time at a value of roughly 1 in 10,000 for a given antigen (Hayakawa et al., 1987; Lalor et al., 1992; McHeyzerWilliams et al., 1993; Schittek and Rajewsky, 1990). Since the size of the overall memory compartment in the immune system is limited and kept rather constant, competition of memory cells for space must occur. Very little is known about this homeostatic control in which persisting antigen may play a role in promoting the long-term maintenance of the corresponding memory cells at the expense of others (Gray, 2002). The recent work of Barrington et al. (2002) supports this notion. One would also expect that the pool of plasma cells that provide humoral memory through long-term antibody production is replenished from the memory pool as needed. The ability to induce plasma cell differentiation of human memory cells in vitro through Toll-like receptors (like TLR9), which are prominently expressed on the surface of these but not naïve B cells (Bernasconi et al., 2003), has led to the concept that signals provided by the innate immune system might be involved in this homeostatic control (Bernasconi et al., 2002). This would necessitate some renewal in the memory cell compartment itself, but given that plasma cells often have very long lifespans, this renewal could be slow and perhaps compatible with the very low proliferative activity seen in the memory compartment. It is also possible that the replenishment of plasma from memory cells mainly involves one of the more recently identified subsets of memory cells, which indeed preferentially home to the bone marrow, one of the main sites of plasma cell residency, and exhibit a pattern of cell surface markers closer to that of plasma cells than that of the “classical” memory cells (McHeyzer-Williams et al., 2000; O’Connor et al., 2002). The proliferative properties of these cells have not yet been studied in detail, but O’Connor et al. report that the memory cells they identified are able to generate plasma cells through proliferative expansion, in the absence of antigen. The survival signals for memory B cells in the absence of antigen remain largely unknown, but should soon be elucidated. Nerve growth factor has been proposed as a specific memory B cell survival factor (Torcia et al., 1996). In the human, these cells express high levels of anti-apoptotic proteins of the Bcl-2 family (Bovia et al., 1998; Liu et al., 1995). It should soon become clear whether they depend on BCR expression and/or the B cell survival factor BAFF, as do naïve, mature B cells (Lam et al., 1997; Rolink et al., 2002), or whether they require signals from interleukins
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through common-g-chain–associated receptors, as do certain subsets of CD8 memory T cells (Sprent et al., 2002). It will be more difficult to answer whether these cells require low-level stimulation by cross-reacting (self-) antigens or through the innate immune system for survival.
MEMORY PLASMA CELLS Plasma Cells as Cellular Correlate of Humoral Immunity More than 100 years ago, the original observation that specific immunity to pathogens is based on and can be transferred with secreted antibodies contained in serum (Behring and Kitasato, 1890) identified antibody-secreting cells as the cellular correlate of “humoral” immunity. In 1948, Fagraeus showed that the presence of so called “plasma cells” in rabbit spleen was strictly correlated to antibody production, in vivo and ex vivo. In 1955, Coons and collaborators could show the presence of antibodies inside plasma cells by immunofluorescence. In 1959, Nossal provided the evidence that these cells were actually secreting antibodies (reviewed in Nossal, 2002). In her detailed analysis of the changes in cellular composition in the spleen of rabbits immunized with ovalbumin, Fagraeus had already obtained evidence that “transitional” (B) lymphocytes would develop into immature plasma cells and then into mature plasma cells, a supposed terminal stage of differentiation.
The Lifespan of Memory Plasma Cells In the early years, the lifetime of plasma cells had been a matter of debate. Through pulse-labeling proliferating cells with 3H-thymidine, and determining labeled plasma cells in the spleen at various time points afterwards, plasma cells of a given immune response could still be detected after several weeks (Ehrich et al., 1949). However, most of the plasma cells generated in a given immune reaction disappear within a few days from the secondary lymphoid organs—spleen and lymph nodes—and only a small fraction of less than 10% is still present in the secondary lymphoid organs for periods of up to 6 months (Schooley, 1961; Mäkelä and Nossal, 1962; Miller, 1964). Neglecting the option that plasma cells could have emigrated from the secondary lymphoid organs, their disappearance was taken as evidence that most plasma cells have a short lifespan. In accordance, when isolated from spleen or lymph nodes, plasma cells do not proliferate and die within days ex vivo (Smith et al., 1996). The concept of short-lived plasma cells as a terminal differentiation stage of activated B lymphocytes has to postulate the constant generation of new shortlived plasma cells from activated memory B lymphocytes, in order to explain the stable concentrations of secreted
antibodies, since estimated half lives of serum antibodies are only a few days. The constant generation of short-lived plasma cells could be based either on a low-key chronic immune reaction driven by persisting antigen (Ochsenbein et al., 2000), or on a bystander activation of memory B lymphocytes in immune reactions to other antigens—the memory B cells being activated via pattern recognition signals (Bernasconi et al., 2002). Both mechanisms have been demonstrated to exist, and may contribute to humoral immunity. By 1898, the antibody concentrations in various organs at various time points after immunization had been determined (Pfeiffer and Marx, 1898), and it had been postulated that antibodies would be secreted mostly in spleen, lymph nodes, lung, and bone marrow. Later work, as described, had focused mainly on spleen and lymph nodes, until it could be shown that, after immunization, antigen-specific plasma cells can be detected in the bone marrow in large numbers. However, such plasma cells would appear in the bone marrow later than in secondary lymphoid organs, at times when plasma cells were already disappearing from the secondary lymphoid organs (McMillan et al., 1972; Brenner et al., 1981; Tew et al., 1992). Could plasma cells be generated in the secondary lymphoid organs, then migrate to the bone marrow and survive there for extended periods of time? The persistence of plasma cells in the bone marrow was determined for 10 days following immunization by radioactive pulse-labeling. At that time, essentially two populations of plasma cells were found in the bone marrow, one with a short half life of a few days, and one with a lifespan of up to 3 weeks, extrapolating from the 10-day period of observation (Ho et al., 1986). Although this lifetime would be longer than that of plasma cells in the spleen and lymph nodes, such a longevity would still not suffice to explain the observed stability of specific serum antibody titers and numbers of specific plasma cells in the bone marrow. Meanwhile, however, two lines of experimental evidence suggest that most plasma cells survive in bone marrow for periods much longer than 3 weeks. Using bromo-deoxy-uridine pulse-labeling of DNA synthesizing cells, the presence of newly generated plasma cells could be followed over more than 100 days (Manz et al., 1997). Labeling dividing cells for the first 3 weeks after secondary immunization, about 30% of newly generated specific plasma cells appeared in the bone marrow between the fourth and sixth week. If extrapolated, this would indicate a half life of bone marrow plasma cells of about 5 weeks. However, after the sixth week, essentially no additional newly generated plasma cells were observed in the bone marrow. This allowed a consistent interpretation of the old and new labeling experiments: New plasma cells enter the bone marrow for a certain period after secondary immunization, but then persist there for long periods, without cell division but still secreting antibodies. The absolute numbers
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of antigen-specific plasma cells in the bone marrow can reach a stable plateau of about 50% of the plasma cells present in the spleen at the peak of the response (Manz et al., 1998). When transferred, bone marrow plasma cells could establish stable concentrations of specific serum antibodies in the host, although at a lower level than in the donor. This antibody secretion is independent of and not influenced by co-transferred antigen (Manz et al., 1998). A second line of evidence in support of an extended lifetime of bone marrow plasma cells is provided by experimental blocking of proliferation and differentiation of memory cells into plasma cells, by irradiation or mitomycin C treatment, in a transfer model of an established immune response to lymphocyte choriomeningitis virus (LCMV) (Slifka et al., 1998). Serum antibody titers and the numbers of specific plasma cells persisted for more than one year, without any apparent decay, in the absence of cellular proliferation. Thus, experimental evidence available so far supports the concept that in the bone marrow, antibody secreting plasma cells survive for long periods and provide stable concentrations of specific antibodies in the serum. These long-lived plasma cells account for humoral immunity and represent a cellular entity of B cell memory, the memory plasma cell.
Recruitment of Plasma Cells to the Memory Pool In chronic immune responses, for example to replicating pathogens, humoral immunity can also be provided by shortlived plasma cells that are continuously generated. Thus, in the immune responses of mice to LCMV and vesicular stomatitis virus (VSV), persistent serum antibody titers were achieved only with replicating, but not with inactivated virus (Ochsenbein et al., 2000). These experiments raise two questions: First, is there a difference in plasma cells generated in primary versus secondary (and chronic) immune responses? Second, which signals decide whether a plasma cell is allowed to emigrate from the secondary lymphoid organ and home to the bone marrow? The first question arises since immunization with inactivated virus induces a primary immune response, whereas replicating virus leads to a chronic immune response. It has been shown that plasma cells presumably derived from naïve B cells are not per se excluded from the pool of longlived plasma cells (Smith et al., 1997; Sze et al., 2000), but most of the plasma cells in the bone marrow secrete antibodies of switched isotype and of an even higher affinity than the corresponding memory B cells (Smith et al., 1997). Interestingly, the modulation of co-stimulation of B cell activation also can modulate the humoral memory provided by plasma cells. Plasma cells secreting low-affinity antibodies are not found in bone marrow from CD21/CD35-deficient mice, as compared to wildtype mice (Chen et al., 2000), and administration of type1 interferon in primary immunizations
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does induce persistent antibody secretion (Le Bon et al., 2001). In the absence of any detailed knowledge about the signals inducing plasma cell differentiation of naïve versus memory B cells in T cell–dependent B-cell activation, it remains unclear to what extent plasma cells generated from naïve B cells can enter the memory plasma cell pool (Smith et al., 1997). The observation that most long-lived plasma cells are generated in secondary lymphoid organs but later found in the bone marrow, implies that recruitment of a plasma cell to the memory cell pool is critically dependent on its ability, or the ability of its precursor, the plasma blast, to leave a secondary lymphoid organ and home to the bone marrow (Pihlgren et al., 2001). In humans, this migration can be readily observed. In the blood of healthy humans, plasma blasts can be identified according to high expression of CD27 and low expression of CD19 (Odendahl et al., 2000). These cells make up less than 1% among the CD19+ B cells. Their phenotype in terms of expression of CD20, CD22, CD45, HLA-DR, CD19, CD138, CD95, and Bcl-2 is intermediate between antibody-secreting cells from tonsils and from bone marrow (Medina et al., 2002). Plasma blasts or plasma cells secreting antibodies of a given specificity are extremely rare, if present at all (Bernasconi et al., 2002). Between days 6 and 8 after secondary immunization, a wave of plasma blasts is detectable in the blood, making up 5 to 40% of the peripheral B cells, and quickly disappearing again. The majority of these plasma blasts is antigenspecific (Bernasconi et al., 2002; Odendahl, Radbruch, Dörner, unpublished data). Thus the release of plasma blasts from the secondary lymphoid organs seems to be strictly regulated (Fig. 16.1). The ability of plasma blasts to home to the bone marrow depends on their expression of chemokine receptors. In CXCR4-deficient fetal liver chimeras, the frequencies of plasma cells in the bone marrow reached only about 30% of that of wildtype controls, identifying CXCR4 and its ligand SDF-1/CXCL12 as the major but not the only attractant of plasma cells to the bone marrow (Hargreaves et al., 2001). The second, redundant or complementary attraction may be conferred by CXCR3 and one or several of its ligands CXCL9, CXCL10, and CXCL11. When tested functionally in migration assays on day 6 after secondary immunization, specific IgG1-secreting cells from spleen and bone marrow migrated exclusively towards gradients of ligands for CXCR3 and CXCR4, but not any other known chemokine receptor (Hauser et al., 2002). In particular, such plasma blasts do not respond to the CCR7-addressing chemokines CCL19, CCL21, and CXCL13, which are expressed in secondary lymphoid organs and could have retained them there (Wehrli et al., 2001). It should be noted that IgA-, but not IgM- or IgG-secreting plasma cells generated from B-2 cells in the Peyer’s patches and other secondary lymphoid organs, express CCR9 and migrate towards its ligand
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CCL25/TECK, thus attracting them into the intestinal epithelium (Bowman et al., 2002; Lamm and PhilipsQuagliata, 2002). Their lifespan there, however, has not yet been determined precisely, and it thus remains to be clarified whether there is actually a mucosal memory plasma cell (Medina et al., 2003). The expression of the chemokines CXCL9, CXCL10, and CXCL11 is a hallmark of inflammation and thus also attracts newly generated CXCR3-expressing plasma blasts into inflamed tissue. This has been demonstrated for plasma cells of a secondary immune response to ovalbumin in NZB/W mice. These cells home to the inflamed kidneys of the animals and survive there for more than 50 days, in numbers roughly equivalent to the plasma cells of the bone marrow (Cassese et al., 2001). In immune reactions to pathogens, the migration of plasma cells into inflamed tissue would provide high concentrations of secreted pathogenspecific antibodies at the site of infection and inflammation. The inflammation will be transient, and it is unlikely that those tissue plasma cells will survive thereafter, for reasons discussed below. In chronic inflammation, however, the inflamed tissue apparently can host long-lived plasma cells of any specificity, and high local antibody concentrations may contribute essentially to the pathogenesis of the disease.
Survival Signals for Long-Lived Plasma Cells Apparently, the bone marrow can provide an environment having the appropriate survival signals, or survival niches. The number of survival niches seems to be limited, since the frequency of plasma cells in the bone marrow is 0.2 to 0.4% of all cells, both in mice and man (Haaijman et al., 1977; Brieva et al., 1991), independent of the genetic and ontogenetic heterogeneity within the human population. In mice, this frequency is reached at about 1 year of age. Even in NZB/W mice with chronic generation of plasma blasts in the secondary lymphoid organs, the frequency of plasma cells in the bone marrow is not enhanced (Cassese et al., 2001). In view of the limited capacity of the bone marrow for plasma cells, competition of newly generated plasma blasts with resident plasma cells is a critical parameter of the plasma cell memory. The molecular definition of the plasma cell survival niche is still lacking. If isolated from the bone marrow or secondary lymphoid organs, or generated ex vivo, plasma cells undergo apoptosis within a few days. Their survival in vitro is prolonged in the presence of bone marrow stroma fibroblasts (Merville et al., 1996), but also by secreted proteins, like IL-6, TNF-alpha, and SDF-1 (Cassese et al., unpublished observations). All these factors act in synergy, but even so cannot mimick the in vivo situation of extended survival of functional plasma cells over months and years. For short-term survival ex vivo, IL-6 is the most effective signal. However, IL-6–deficient mice, although suffering from
impaired immune reactivity, can generate long-lasting antibody responses (Eugster et al., 1998) and have the same frequencies of memory plasma cells in the bone marrow as their wildtype littermates (Cassese, Radbruch, Manz, unpublished observation). SDF-1/CXCL12, the ligand of CXCR4 and putative attractant of plasma blasts to the bone marrow, is also an effective survival factor for bone marrow plasma cells. Remarkably, resident bone marrow plasma cells that express CXCR4 and respond to CXCL12 by prolonged survival do no longer migrate in response to it (Hauser et al., 2002). This lack of motility may well be a selective disadvantage of resident plasma cells when competing with migratory plasma blasts. IL-7 does not seem to be a survival factor for plasma cells. In IL-7–deficient mice, however, in which B cell lymphopoiesis is disturbed, the numbers of bone marrow plasma cells increase (Carvalho et al., 2001). This may indicate that pre-B and plasma cells share components of their respective survival niches.
ADAPTIVE B CELL MEMORY The Interplay of Memory B and Memory Plasma Cells Memory plasma cells provide long-lasting protection against pathogens. This protection is slowly waning due to the competition of plasma blasts newly generated in response to more recent antigens. Plasma cells generated in a given immune reaction and not making it to survival niches will be short-lived and provide peak responses either in secondary lymphoid organs, draining to lymph and blood, or in transiently inflamed tissue. Eventually, somatic hypermutation and isotype switching will optimize the antibodies generated for the efficient elimination of the pathogen, and protection against reinfection. Once antigen is eliminated, long-lived memory plasma cells maintain protective memory in its absence. It is still not clear whether, and if so, how plasma cells secreting protective antibodies are selectively recruited to the pool of memory plasma cells. One option would be that plasma blasts might be retained in the secondary lymphoid organs, as long as an immune reaction is going on, through signals from the reactive lymphoid organs. After the successful elimination of the antigen and termination of the germinal center reaction, the secondary lymphoid organs might release the surviving plasma cells. These would be the plasma blasts generated in the final phase of the immune response and thus in all likelihood secreting “protective” antibodies. Such plasma blasts would then compete for bone marrow survival niches. It is not clear how this competition works. Do immune reactions mobilize resident bone marrow plasma cells and thus generate free survival niches? In any case, antigens not encountered for a long time will eventually be “forgotten,” because the
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resident plasma cells are competed out of the bone marrow. Memory B cells adjust humoral memory. When pathogens reach concentrations exceeding those at which they can be neutralized by the protective antibodies, memory B cells are reactivated and adjust the levels of circulating antibodies to that required for pathogen control. According to this concept, memory B and plasma cells together provide a memory that adapts to the antigenic environment of the immune system.
The Differentiation of Memory B versus Plasma Cells The choice of differentiation of activated B cells into memory B or plasma cells is a critical step in the generation of reactive versus protective memory. Differentiation of activated B cells into antibody-secreting plasma cells seems to be the default pathway, whereas the generation of memory B cells seems to be dependent on co-stimulation of antigenactivated B cells by CD40L (Arpin et al., 1995). When B cells are activated by antigen in the absence of CD40 signals, or by signals from pattern recognition receptors such as the LPS receptor or TLR9, they will differentiate into plasma blasts. Although the LPS-receptor is also functional on naïve B cells, reaction to TLR9 signals requires co-stimulation via antigen for naïve, but not for memory B cells (Bernasconi et al., 2003). It has been speculated that this dichotomy may serve to prevent antigen-independent activation of naïve B cells, while allowing the antigen-independent differentiation of memory B cells into plasma cells. This would allow the accidental and continuous replenishment of the plasma cell pool from memory B cells in the absence of antigen. Although this bystander activation has been demonstrated (Bernasconi et al., 2002), it remains to be shown that cells generated in this way migrate to the bone marrow, persist there, and contribute significantly to the maintenance of the pool of memory plasma cells. Other signals, from OX40L, CD27, or the cytokine receptors for TNF, IL-10, and IL-6 also support differentiation of activated B cells into plasma cells (Agematsu et al., 1999; Stuber and Strober, 1996; Choe and Choi, 1998). IL-6 is of particular interest because it induces p18INK4c, a cyclindependent kinase inhibitor acting on CDK6 (Morse et al., 1997). In the absence of p18INK4c, the formation of antibodysecreting cells is selectively blocked (Tourigny et al., 2002). Thus, cell cycle arrest seems to be a molecular prerequisite for terminal plasma cell differentiation. IL-6 also upregulates the expression of X-box binding protein 1 (XBP-1) in B cells. This requires the unfolded protein response, signaling activation-induced antibody synthesis and inducing splicing of the XBP-1 mRNA by the endonuclease IRE1 (Yoshida et al., 2001). The product of the spliced XBP-1 mRNA is necessary and sufficient to
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restore antibody-secretion in XBP-1 deficient B cells (Iwakoshi et al., 2003). Little is known about the target genes of XBP-1. One is probably p18INK4c (Wen et al., 1999). In a positive feedback loop, XBP-1 also induces the expression of IL-6, a cytokine which in turn induces XBP-1 expression. While XBP-1 serves to sense signals from the IL-6 receptor and the unfolded protein response, signals from the antigen-receptor apparently lead to a downregulation of BCL-6, a direct transcriptional repressor of the PRDM1 gene that encodes the B-lymphocyte–induced maturation protein 1 (Blimp-1) (Shaffer et al., 2000). Blimp-1 is a master gene of plasma cell development and the key antagonist of the B-cell lineage-specific activator protein (BSAP), encoded by the Pax-5 gene. Blimp-1-directly represses transcription of Pax-5. Since Pax-5 in turn represses XBP-1 transcription, Blimp-1–mediated downregulation of Pax-5 allows the induction of expression of XBP-1, and thus the expression of plasma cell–specific genes (Lin et al., 2002). By repressing Pax-5, Blimp-1 downregulates a large variety of B cell–specific genes, including genes for antigen-presentation, class switching, and somatic hypermutation. It also downregulates BCL-6, its own repressor, thus creating a negative feedback loop that stabilizes plasma cell differentiation (Shaffer et al., 2002). CD40 signaling downregulates Blimp-1 mRNA in activated B cells (Randall et al., 1998), offering a molecular explanation for the CD40dependent differentiation of activated B cells into memory B cells. Several other transcription factors also seem to be involved in the differentiation of activated B cells into plasma cells, such as IRF-4, NFATc1, and NFATc2, and octamer-binding proteins (Oct-1, -2, and -B), because their genetic inactivation and/or ectopic expression affects the generation of plasma cells. Their precise role in plasma cell differentiation remains to be determined (reviewed in Calame, 2001). In the decision between memory versus plasma cell differentiation, the differentiation of B cells into memory B cells is dependent on T cell–derived CD40L signals. In the absence of CD40-signaling, however, BCL-6 expression is downregulated by antigen-receptor signaling, its repression of Blimp-1 is released, and Blimp-1 in turn represses the transcription of Pax-5. The gene expression program of B cells is terminated, and XBP-1 induces a plasma cell–specific gene expression profile, including p18INK4c, which will arrest proliferation. XBP-1 is activated by signals of the unfolded protein response, triggered by enhanced antibody synthesis. This differentiation program is stabilized by positive and negative feedback loops, for example, induction of IL-6 by XBP-1 and repression of BCL-6 by Blimp-1. The fate of the memory B and plasma cells thus generated then depends on their ability to home to molecular niches that allow their survival.
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Acknowledgments We are grateful to C. Berek, S. Casola, T. Doerner, R. Kueppers, and M. Shlomchik for helpful discussion. Supported by the National Institutes of Health and Infectious Diseases, Grant # 1RO1A1054636-01, and the German Research Council.
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17 Immunoglobulin Assembly and Secretion LINDA M. HENDERSHOT
ROBERTO SITIA
St. Jude Children’s Research Hospital, Memphis, Tennessee, USA
Università Vita-Salute San Raffaele, DIBIT-HSR Scientific Institute, Milano, Italy
Immunoglobulin (Ig) molecules serve as both cell surface B cell antigen receptors (BCR) and secreted effector molecules (antibodies) that provide protection against infections and foreign antigens. In their simplest form, each molecule is comprised of two identical heavy chains (HC) and two identical light chains (LC). The building blocks of antibodies are provided by a homology unit termed the Ig domain. Seven types of C domain and nine types of V domain exist each of which is comprised of antiparallel ßsheets connected by loops and stabilized by a disulfide bond. Although the ß-sheets are subject to stringent structural constraints, the loops are free to vary, providing this unit with a unique versatility, both in phylogeny and ontogeny. The HC is comprised of an N-terminal variable domain (VH) followed by three to four constant domains (CH) depending on the isotype, and the LC comprises a VL and a single CL domain. It has been estimated that humans can synthesize between 107 and 109 different Ig molecules. Based on an average size of 50,000 to 70,000 daltons for the HC and 25,000 for the LC, it would take over three genome equivalents to encode a repertoire of this size, if conventional methods were used to encode antibodies! However, using a unique combination of gene rearrangement, the addition of nontemplated nucleotides to the cleaved ends of variable region gene segments, imprecise rejoining of these segments, and targeted hypermutation of the assembled variable region (which are the topics of other chapters in this book), the Ig repertoire is produced from less than 0.1% of the genome. However, this remarkable feat comes at a cost. The possibility is high of producing HC or LC with premature stops or mutations that prevent proper folding, assembly, transport, or interaction with signaling molecules. Through the ordered synthesis of HC and LC, coupled with intricate systems to examine and test their fitness, B cells express functional antigen
receptors that, upon binding to antigen, induce differentiation into plasma cells, the dedicated factories for producing tremendous quantities of effector antibodies.
Molecular Biology of B Cells
MECHANISMS OF IG SYNTHESIS AND ASSEMBLY Like all proteins destined to exocytic compartments, HC and LC are co-translationally translocated into the endoplasmic reticulum (ER). In this organelle, they undergo posttranslational modifications (i.e., folding, assembly, disulfide bond formation, and glycosylation) that play fundamental roles in controlling both intracellular transport and functional activities of antibodies.
Folding and Assembly The V region genes each encode a targeting sequence that directs the nascent chain to the ER (Milstein et al., 1972). As it enters this compartment, folding begins cotranslationally from the V to the C domains (Bergman and Kuehl, 1979). Sequential folding probably reduces the risk of aberrant interdomain disulphide bonds and helps to ratchet the nascent chain into the ER lumen, thus limiting the backward movement into the cytosol (Ooi and Weiss, 1992). N-linked glycans are added to the nascent HC and, with the notable exception of the CH1 domain (see below), intradomain disulfide bonds form co-translationally to stabilize the folding of each individual Ig domain (Bergman and Kuehl, 1979). Like folding, assembly with LC also begins on nascent HC. Curiously, in spite of the overall conservation of Ig structure, the order of assembly differs within classes. Thus, IgM follows the H-HL-(H2L)-H2L2 pathway, whereas in
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IgG1 HC dimerization precedes H-L assembly (Scharff et al., 1970). It appears that the CH3 domain may play a role in this assembly, as HC mutants that have deleted the CH3 domain do not form HC dimers readily and are instead secreted as HL “hemimers” (Yelton et al., 1982). In hybridomas or transfectomas producing more than one isotype, the promiscuous pairing of HC occurs only between subclasses. Hybrid mg molecules are rarely observed, whereas g1g2aLC2, g2ag2b are formed and secreted efficiently (Winter and Milstein, 1991 and references therein). Despite the fact that m and a chains do not pair directly with each other, cells producing both chains can release heteropolymers containing m2L2 and a2L2 subunits (Urnowitz et al., 1988). Ig Assembly in Pre-B Cells and Allelic Exclusion The production of antibody molecules begins at the preB cell stage, and the HC is normally restricted to the m isotype. However, unlike plasma cells, pre-B cells transcribe and translate very small quantities of HC. This is perhaps due to the fact that the HC must first be tested for its ability to form a functional Ig molecule, and the HC does not perform any effector functions at this point that require large amounts of it. In the absence of LC synthesis, HCs do not fold completely and are retained by ER chaperones and eventually degraded (see below). Although pre-B cells do not make conventional LC, they do synthesize a LC-like molecule called the surrogate LC (SLC) (Melchers & Kincade, this volume). If the HC is able to assemble productively with the SLC, a small amount of pre-B cell receptor (mm2-SLC2) will be transported to the cell surface, where it generates a signal to proliferate and differentiate. To produce this signal(s), the HC must be able to assemble with the SLC and fold properly, interact with the Ig accessory molecules, be transported through the Golgi to the cell surface, and engage the proper signaling proteins (Reth & Wienands, this volume). If a signal is generated, HC rearrangement on the second allele is halted, as the pre-B cell has successfully generated a functional HC. If, however, a signal is not generated, the first allele is silenced (a process known as allelic exclusion), rearrangement of the second HC allele is activated, and a second HC is synthesized. Again, the same processes and tests are directed towards this HC. If it is unable to successfully produce a signal, the preB cell will die, having missed its two opportunities to make a functional HC. If, however, either attempt to generate a signal is successful, SLC synthesis stops and the rearrangement and synthesis of a conventional LC begins. As was the case for the HC, the newly synthesized LC must be examined to ensure that a complete protein that is able to fold and assemble properly is made. However, the cell is given four opportunities to make a LC. Perhaps B cells have evolved in this way to provide the HC with a greater chance of producing a functional Ig protein. If a given LC is able to
assemble with the HC and complete its folding, the mm2-LC2 B cell receptor will assemble with the accessory proteins Iga/b, thus allowing them to be transported to the B cell surface and generating a poorly understood signal that stops further LC arrangements. Assembly and V Region Selection The rate of Ig synthesis remains relatively low in B cells. The CL domain folds very rapidly and stably (Hellman et al., 1999) and pairs with the CH1 domain, which otherwise remains unfolded in the absence of LC (Lee et al., 1999). Data suggest that domain pairing between HC and LC drives the final folding of the HC (Lee et al., 1999) and in some cases of the LC (Leitzgen et al., 1997) as opposed to proper folding of these domains being a requirement for assembly. This limitation would ensure that free HCs do not get transported without LCs. It is possible that similar help could be provided by domain pairing between variable domains, in which the greatest likelihood of folding difficulties is likely to occur due to the processes used to generate integrated variable regions. Although this would increase the opportunity for a relatively unstable variable region to fold, it would also put limitations on HC–LC pairing. This hypothesis predicts that a VH or VL region that folds stably by itself could pair with either a stable or unstable VL and VH respectively, while VH or VL that is unable to form a stable fold by itself could only pair with a stable partner (Figure 17.1). If variable domains make use of domain pairing–assisted folding as the CH1 domain does, it is clear this would affect the repertoire generation.
Glycosylation HCs contain variable numbers of N-linked glycans that are added co-translationally and then processed as the molecules proceed along the secretory route. The inhibition of N-linked glycosylation generally results in intracellular retention and degradation of HC, with the effects being greatest for IgM and least for the less glycosylated IgG, thus underscoring the role of sugar moieties in Ig folding, assembly, and stability (Hickman and Kornfeld, 1978). IgA1 and IgD also possess O-linked sugars attached to their extended hinge regions. Blocking their elaboration beyond the first N-acetylgalactosamine did not significantly affect intracellular assembly or secretion of either IgD or IgA1 (Gala and Morrison, 2002), suggesting that O-linked sugars probably have little influence on assembly. Genetically engineered Ig molecules lacking one or more N-glycans can be secreted, allowing a functional characterization of individual groups (Wright and Morrison, 1997; Jefferis et al., 1998; Rudd et al., 2001 and references therein). The conserved glycan in the tailpiece of m and a chains is essential for J chain binding, but not for oligomerization (Atkin et al.,
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FIGURE 17.1 Assembly-dependent folding resulting in restrictions on HC/LC pairing. Folding of the CH1 domain is dependent on binding to a folded CL domain. If V regions follow suit, a folded VH region could pair with a LC possessing either a folded or unfolded VL region and induce folding of the latter. However, a VH region unable to fold by itself could only pair with a LC containing a folded VL domain. See color insert.
1996). However, m chains lacking the Asn563 glycan are secreted as huge, precipitation-prone polymers (de Lalla et al., 1998), suggesting that the sugar moieties regulate polymerization, perhaps by recruiting lectin or chaperone molecules. In secreted IgM polymers, the Asn563 glycans are found in the high-mannose form typical of ER resident proteins (Davis et al., 1989a), probably because polymerization, an event thought to occur in the ER (Cals et al., 1996; Reddy and Corley, 1999) hinders their accessibility to downstream processing enzymes. Exposure of these high mannose groups upon antigen binding might be important for the clearance of immune complexes and antigen delivery to dendritic cells for Class II presentation.
Differential Fate of mm and ms During B Cell Differentiation In both lymphoid and nonlymphoid cells, the coexpression of HC and LC results in the efficient assembly of H2L2 complexes, implying that the chaperones and enzymes required to accomplish these processes are conserved. Important cases exist, however, in which cell speci-
ficity is evident. A striking one is the diverse fate of mm and ms chains in B and plasma cells. The physiology of the immune system requires that B cells express antigen receptors (BCR) on their surface, but secrete little if any Ig until they encounter antigen. Indeed, premature secretion might hinder antigen recognition via the BCR. In contrast, plasma cells are antibody producing factories, with little use for active BCR on their surface. This dramatic change in job description is in part explained by the differential splicing of the HC transcripts, resulting in the membrane or secreted forms being predominant in B and plasma cells, respectively. However, post-translational events also help prevent IgM secretion by B cells and BCR expression by plasma cells. Many B lymphoma cells synthesize the two forms of m chains in similar amounts and rapidly assemble them into mm2L2 and ms2L2 complexes. However, whereas the former exit the ER to reach the cell surface, most ms chains are retained and degraded intracellularly (Brooks et al., 1983; King and Corley, 1989). The reverse is true for myeloma cells, which efficiently secrete ms chains in the form of polymeric IgM but fail to express mm2L2 on the surface (Sitia et al., 1987, 1990).
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FIGURE 17.2 ER quality control checkpoints regulating Ig transport. (1) In the absence of SLC or LC assembly, the CH1 domain remains unfolded and is associated with BiP, which prevents its transport to the Golgi. (2) The transmembrane domain of mm contains a number of hydrophilic residues that prevent its transport until they are masked by Iga/b. (3) VpreB (green) is missing its ninth and final b strand and has a unique sequence (white box). It is unable to fold until l5 assembles and provides its missing strand. (4) The secretory IgM monomer possesses an unpaired cysteine in its Cterminal tailpiece (reddish) that is recognized by thiol retention mechanisms until it associates with J chain in IgM pentamers or with other monomers in IgM hexamers. See color insert.
MULTIPLE LAYERS OF QUALITY CONTROL EXIST TO AID AND MONITOR THE ASSEMBLY OF FUNCTIONAL IGS In mammalian cells, the ER provides a unique folding environment for secretory proteins and exerts tight quality control measures to ensure that only native proteins are secreted or deployed onto the cell surface (Figure 17.2). The mechanisms for generating diversity should give rise to many nonfunctional proteins, which must be detected and destroyed. Thus, given the critical reliance of the immune system on ER quality control, it is not surprising that many ER molecular chaperones, folding enzymes, and qualitycontrol mechanisms of the eucaryotic secretory pathway were first identified through their association with HC. The various features of Ig molecules that are subjected to quality control mechanisms are discussed here.
The CH1 Domain The faces of the VL and CL domains first form noncovalent interactions with the VH and CH1 domains of the HC (Chothia et al., 1985). An interchain disulfide between the penultimate cysteine of the LC and a cysteine in the Cm1 domain or the hinge region of most g HC subclasses stabilizes this assembly. Unlike the CH2, CH3, and CH4 domains, which pair with the corresponding domain on the other subunit of the HC dimer, the VH and CH1 domains remain unpaired in the absence of LC. It was long appreciated that the deletion of the CH1 domain allowed HC to be secreted without LC from cell lines (Morrison, 1978; Birshtein et al.,
FIGURE 17.3 Protein quality control in the ER. Following co-translational translocation into the ER lumen via the Sec61 channel (in blue), HC and LC fold, assemble, and when required form polymers with the assistance of many ER resident chaperones and enzymes before transport to the Golgi. Disulfide bonds are inserted and isomerized in this phase. Molecules that fail to attain their native structure within a given time are dislocated to the cytosol to be degraded by proteasomes. Disassembly and disulfide reduction precede dislocation (Fagioli and Sitia, 2001). The extraction of substrates across Sec61 is facilitated by p97 and ubiquitination (Tsai et al., 2002). See color insert.
1974) and from plasma cells of patients with HC disease (Seligmann et al., 1979). This suggested that unpaired CH1 domains were somehow recognized and used to retain the free HC (Figure 17.3). In 1983, the first eukaryotic ER chaperone to be identified was BiP (immunoglobulin HC binding protein), which
17. Immunoglobulin Assembly and Secretion
was found noncovalently associated with unassembled HC in an Abelson transformed pre-B cell line (Haas and Wabl, 1983). BiP also interacts with Ig assembly intermediates but not with completely assembled Ig molecules (Bole et al., 1986). Deletion of CH1 allows the secretion of free HC and Ig assembly intermediates, suggesting that BiP prevents the transport of incompletely assembled Ig molecules (Hendershot et al., 1987). BiP itself is localized in the ER via its C-terminal tetrapeptide sequence, KDEL. If BiP, along with any bound substrate protein, exits the ER, it is captured by the KDEL receptor in the downstream compartments and promptly retrieved to the ER (Munro and Pelham, 1987). BiP is not an Ig-specific chaperone and is, in fact, produced in all eukaryotic cells, where it binds unfolded nascent secretory pathway proteins and prevents their transport to the Golgi. BiP preferentially binds to unfolded regions on proteins containing hydrophobic residues (Blond-Elguindi et al., 1993). Although the CH1 domain binds stably to BiP, the other domains interact transiently with the chaperone as they fold. An algorithm generated to predict BiP binding sites (Knarr et al., 1995) identified multiple potential BiP binding sites in the sequence of all Ig domains. However, no data as yet demonstrates that any of these peptides mediates BiP binding in vivo. In fact, the stoichiometry of BiP to HC is ~1 : 1. Perhaps the most provocative data on BiP binding sites come from the analysis of two well-characterized LCs, LEN and SMA. A well-conserved, high affinity BiP binding peptide sequence was identified in each of the two b sheets of the variable region (Davis et al., 1999). These sequences are eventually buried in the hydrophobic core of the folded variable region, and presumably BiP binding prevents aggregation (Davis et al., 2000) and helps direct the folding of the nascent LC variable region. Unassembled HC remain substrates for BiP, because they are unable to fold completely in the absence of LC (Lee et al., 1999). This feature of CH1 domains is essential to ensure that only properly assembled H2L2 molecules are transported out of the ER. The binding of LC appears to trigger the release of BiP from the CH1 domain, thus allowing the intrachain disulfide to form in the CH1 domain (Vanhove et al., 2001), which could be critical to pass the ER quality control mechanisms that are based on the detection of free thiols. Clearly, the transport of partially assembled Ig molecules would be both wasteful and potentially damaging to the effectiveness of the immune response. Interestingly, camels do not synthesize LC and instead make HC without a CH1 domain (Hamers-Casterman et al., 1993)!
Monitoring the Assembly of Functional BCRs The BCR is a multimeric complex formed by at least four polypeptides, mm, LC, Ig-a, and Ig-b. The latter two proteins are involved in signal transduction upon antigen binding by mm2L2 (Reth & Wienands, this volume). Cells that do not synthesize Ig-a and Ig-b in sufficient amounts
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are unable to express surface BCRs. The presence of hydrophilic residues within the HC transmembrane region mediates the retention of incompletely assembled BCR (Williams et al., 1990). The mechanism bears some similarities to the TCR, in which charged residues are found in the transmembrane regions of several interacting subunits. Assembly with Ig-a and Ig-b masks these hydrophilic residues, allowing the transport of the complete BCR (see Figure 17.2). This implies that the retention of unassembled chains exploits interactions in a nonaqueous environment, although the mechanism or proteins are not clearly understood. The BCR-associated proteins BAP29 and BAP31 (Adachi et al., 1996) are candidates to either mediate this type of quality control or selectively promote the forward transport of assembled BCR (see next section). Ig-a and Igb interact with all Ig classes. Interestingly, their glycosylation patterns differ depending on the isotype with which they pair (Venkitaraman et al., 1991), thus suggesting that architectural modifications ensue upon association.
Regulation of Surrogate LC Assembly and Pre-B Receptor Transport Pre-B cells synthesize a “surrogate” LC, which can interact with mm to form the pre-BCR. The SLC is produced from Vpre-B and l5, which encode a variable-like and a constant-like domain, respectively (Sakaguchi and Melchers, 1986; Kudo and Melchers, 1987). However, unlike their counterparts in conventional LC, these two genes do not encode nine- and seven-strand Ig domains, and they both contain an additional sequence that has been termed the “unique” region. Vpre-B possesses an ER targeting signal sequence, eight of the nine strands normally found in a V region and a C-terminal extension of 24 amino acids that shows no homology to other Ig domains. Without its ninth b strand, it is unable to fold. l5 also has an ER targeting signal sequence, followed by a 50 amino acid unique region, and eight b strands instead of the usual seven found in other constant region domains. Interestingly, the unique region of l5 appears to act as an intramolecular chaperone or “pro” sequence to inhibit the folding of unassembled l5. Interaction of this sequence with the unique sequence of Vpre-B evidently allows the extra b strand of l5 to interdigitate with the incomplete Vpre-B protein and supply its missing b strand (Minegishi et al., 1999). Thus, folding of both SLC components is controlled by their assembly with each other (see Figure 17.3). It is unclear why such an unusual structure for these genes evolved and why the SLC is not simply synthesized as a single protein. It is interesting to speculate that interaction with the VH and CH1 domains of the nascent HC plays some role in the assembly of the SLC and that this serves as an indication of the HC ability to help a LC fold. However, there are no experimental data to support this idea.
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The SLC serves to test the ability of the newly rearranged HC to interact with LC, release BiP, fold properly, form the requisite inter- and intrachain disulfide bonds, and exit the ER before the conventional LCs are rearranged and synthesized. Pre-B cells have been identified in which two functional HC rearrangements occurred but where only one of the HCs was capable of combining with the SLC, suggesting that allelic exclusion was dependent on the assembly of the pre-B receptor and its expression on the cell surface. This is supported by data from mice, in which a targeted disruption of mm results in a block in pre-B cell differentiation and allelic inclusion (Melchers et al., 1993). Mice with targeted disruption of l5 or Vpre-B genes also show a defect in preB cell differentiation, although it is not as complete as that seen in mice with the mm disruption. This has led others to speculate that either other protein(s) can interact with the CH1 domain and induce its folding, or that compensatory premature rearrangements of conventional LC genes occur in some cells. This point remains controversial. There is no clear additional role for the SLC in generating the pre-B signal other than helping the CH1 domain to fold, because transgenic mice that express either a CH1 domain–deleted HC or a truncated HC comprised of only the CH4 and transmembrane domains do not require the surrogate LC to induce differentiation or conventional LC gene rearrangement (Shaffer and Schlissel, 1997; Muljo and Schlissel, 2002).
Developmental Control of IgM Secretion In line with the notion that exit from the ER is restricted to fully assembled proteins, only polymeric IgM (hexamers or pentamers containing a J chain) can be secreted (Cattaneo and Neuberger, 1987; Davis et al., 1989b; Sitia et al., 1990; Brewer et al., 1994). For unclear reasons, B cells are unable to polymerize IgM: As a result, they retain and degrade intracellularly virtually all ms2LC2 complexes (Sidman, 1981; Sitia et al., 1988). Retention is mediated by a conserved 20 amino acid tailpiece at the C-terminus of m and a chains. Within the tailpiece, a cysteine in the penultimate position (Cys575 in ms) forms the covalent bond linking HC2LC2 subunits to each other or to a J chain. This cysteine is essential for IgM polymerization and is also responsible for the selective retention of unpolymerized m2LC2 subunits. Appending the ms tailpiece causes retention and degradation or polymerization of the resulting chimeric molecules only if the critical cysteine residue is present (Sitia et al., 1990; Fra et al., 1993; Smith et al., 1995). Therefore, Cys575 in the ms tailpiece acts as a three-way switch, mediating assembly, retention, and degradation of ms2LC2 subunits (see Figure 17.4). This thiol-based quality-control mechanism is widely exploited to regulate the expression of secretory, transmembrane, and GPI-anchored proteins (Kerem et al., 1993; Capellari et al., 1999). Its stringency
FIGURE 17.4 Glycan processing diverts terminally misfolded glycoproteins to ERAD. To maintain homeostasis, proteins that fail to fold within a given time must be degraded. Removal of the terminal mannose from the central branch of N-glycans by ER mannosidase I targets terminally misfolded or orphan glycoproteins to dislocation, probably via interaction with EDEM (Ellgaard and Helenius, 2001). See color insert.
can be modulated by the amino acid context surrounding the unpaired cysteine(s) involved: For example, the presence of vicinal acidic residues weakens retention. This allows some unassembled LCs and monomeric IgA to be secreted by plasma cells (see below). The observation that monomeric IgA is efficiently retained by B cells and can be secreted by plasma cells implies that thiol-mediated retention is also regulated by cellular factors (Guenzi et al., 1994). The failure of B cells to polymerize and secrete IgM or IgA may reflect the differential expression of specific chaperones or redox enzymes.
Secretion of Free LC The presence of Bence-Jones proteins in the blood and urine of myeloma patients implies that LC can escape quality control to be secreted even in the absence of HC. Many Bence-Jones proteins are secreted as covalent or noncovalent LC2 homodimers (Leitzgen et al., 1997), a conformation that prevents detection by the ER quality control by masking the hydrophobic surfaces on each domain. Cys213, normally utilized to form the S-S bond with HC, avoids thiol-mediated retention by forming a disulfide bond with free cysteines present in the ER (Reddy et al., 1996). Weak LC retention seems to be an elegant solution to optimize Ig assembly and reduce the risks of intracellular accumulation. However, this requires that LC be synthesized in excess of HC in plasma cells to ensure that all HC can assemble rapidly. Estimates suggest that, in fact, plasma cells synthesize from one and a half to two times an excess of LC over HC (Baumal and Scharff, 1973; Bergman and Kuehl, 1979).
17. Immunoglobulin Assembly and Secretion
Role of J Chain in Polymerization and Transcytosis Although J chain is neither sufficient nor necessary for IgM secretion, it determines the type of IgM polymer produced (Niles et al., 1995; Reddy and Corley, 1999). In the absence of J chains, hexamers are the main form of IgM secreted (Cattaneo and Neuberger, 1987). Since hexamers have been shown to bind complement more efficiently than pentamers, J chain might therefore regulate the type of humoral responses. Studies on knockout mice confirm that a crucial function of J chain is mediating the delivery of polymeric Ig to external secretions (Hendricksson et al., 1995; Vaerman et al., 1998; Erlandsson et al., 2001). Transcytosis is mediated by the polymeric Ig receptor, which binds J chain–containing IgM or IgA at the basolateral surface of epithelial cells. Sequences in the cytoplasmic portion of the receptor drive internalization, intracellular transport, and release from the apical surface. During transcytosis, a fragment of the receptor becomes disulphidebonded to dimeric IgA. This fragment constitutes the “secretory component” (Mostov and Blobel, 1983).
TRANSPORT OF ASSEMBLED IG MOLECULES TO THE GOLGI Once a pre-BCR or BCR has folded completely, formed all disulfide bonds, and assembled with the Iga and Igb accessory molecules, it no longer binds to BiP nor is it retained by thiol-mediated mechanisms or by the recognition of hydrophilic residues in the transmembrane domain. It is ready to be transported to the cell surface. Several years ago, the prevailing wisdom held that in the absence of retention, a properly folded and assembled protein would move to the Golgi via bulk flow transport from one organelle to the other. In recent years, however, a number of transporter proteins have been identified that appear to specifically recognize some “cargo” proteins and deliver them to sites of exit from the ER. Disruption of these transporter proteins specifically hinders the transport of their target proteins. This suggests that instead of transport occurring automatically in the absence of specific retention, for some proteins transport is signal-mediated. A putative “signal patch” has been identified on VL and Vpre-B domains, which include highly conserved amino acids at positions 15, 59, 61, 62, and 82 that are contiguous on the folded structure. The mutation of these amino acids does not interfere with Ig assembly or chaperone release but does prevent the transport of the Ig molecules to the Golgi (Dul and Argon, 1990; Argon, personal communication), suggesting that this “patch” may represent a very late checkpoint. If indeed there is a signal-mediated transport of Ig molecules, it is clear that the transporter is not a B cell–specific protein, since trans-
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fected Ig molecules can be synthesized and secreted from a number of non-B cell lines. It is also unlikely to be dependent upon BAP29 or BAP31, as they appear to interact specifically with transmembrane residues to retain IgM and IgD (Adachi et al., 1996; Schamel et al., 2003).
DEGRADATION OF MISFOLDED AND UNASSEMBLED IG SUBUNITS The retention of folding and assembly intermediates within the specialized environment of the ER may facilitate their structural maturation. For instance, retention may increase the local concentration and favor HC–LC assembly or IgM polymerization. However, mutations (O’Hare et al., 1999) or unbalanced subunit synthesis (Kohler, 1980), can make maturation, and hence exit from the ER, impossible. An efficient ER-associated degradation (ERAD) pathway disposes to the proteasome those terminally misfolded or unassembled molecules that could otherwise accumulate (see below), aggregate, and become toxic to the cell. This implies that degradation substrates must be “retro-translocated” or “dislocated” across the ER membrane to reach the cytosol. Dislocation is thought to occur via Sec61, a protein complex also used by nascent proteins to enter into the ER (reviewed by Tsai et al., 2002). This unexpected “cytoplasmic connection” between quality control in the ER and cytosolic proteasomes explains how nascent unfolded proteins might coexist in the ER lumen with an aggressive proteolytic system. At the same time, it poses questions relating to the mechanisms underlying 1) the recognition of terminally misfolded proteins to be targeted for degradation, 2) their discrimination from newly made proteins that have not had time to fold, 3) their extraction from the ER lumen, and 4) their degradation. Work on Ig subunits has been instrumental in providing answers to some of these questions. A unifying concept to explain quality control and ERAD is that the systems are able to recognize the “unfoldedness” of some proteins as specifically different from nascent unfolded and unassembled proteins entering the ER. Since a limited set of chaperone molecules appear to play a role in both processes, it is not yet clear how the two are separated. The BCR and IgM polymers are clear examples in which assembly masks chaperone binding sites. In principle, prolonged binding to chaperones should divert proteins to ERAD, and thus the rate of assembly versus targeting for dislocation might control the fate of a HC. However, while unassembled m chains turn over quite rapidly, unassembled g chains enjoy quite long half lives, and this correlates with tighter BiP binding (Skowronek et al., 1998). In the case of orphan m and J chains, N-glycan processing, and in particular removal of the terminal mannose from the central branch, acts as a timer in diverting unassembled molecules to dislo-
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cation or degradation. Inhibitors of ER mannosidase I block m chain degradation as efficiently as inhibiting proteasomes (Fagioli and Sitia, 2001). A molecule endowed with lectinlike activity (EDEM, for ER degradation enhancing mannosidase-like) has been proposed to bind misfolded proteins with mannosidase I–processed glycans, thus providing an elegant way to target terminally misfolded proteins for degradation (Hosokawa et al., 2001; Jakob et al., 2001). It is not clear how proteins lacking sugar moieties are diverted for dislocation after their time has elapsed. Short-lived LC mutants remain bound to BiP until degradation ensues (Knittler and Haas, 1992). Mechanisms must exist that dissociate BiP, open the Sec61 gate, insert the substrate into the translocon, and activate dislocation (Tsai et al., 2002). Due to the size limitations imposed by the structure of the Sec61 channel, dislocation is thought to require disassembly and partial unfolding of the substrate proteins. Evidence has been provided that covalent complexes between transport-competent LC and short-lived HC are reduced prior to dislocation of the latter. Freed LC can reassemble with newly made HC, indicating that disulfides can be simultaneously formed and reduced in the same compartment. The reduction of interchain disulfides also takes place during the degradation of ms2L2 complexes by B cells (Fagioli et al., 2001). Reduction and dislocation seem to be coupled events, suggesting that a reductase activity is associated with active dislocons. It is unclear whether (and to what extent) unfolding is required to negotiate retrograde transport across the ER membrane (Tsai et al., 2002). Once initiated, the process of retrotranslocation requires energy for completion. Preventing proteasome function with specific inhibitors causes the accumulation of short-lived LC and m chains in the ER lumen (Chillaron and Haas, 2000; Mancini et al., 2000) suggesting that degradation and dislocation are coupled events as well. However, proteasome blockage has a wide range of effects on cell physiology, and dislocation could be affected indirectly, for example, by a reduction in the free ubiquitin pools. A member of the AAA ATPase family, p97/cdc48, seems to play a key role in extracting different substrates from the ER, including IgM subunits (Ye et al., 2001; Rabinovich et al., 2002). Further work is necessary to dissect the complex machinery that distinguishes proteins that are unable to fold from those that are in the process of folding and is responsible for maintaining homeostasis in the ER.
Russell Bodies When the synthesis of a protein exceeds the combined rates of transport to the Golgi and degradation, accumulation in the ER ensues. In Mott cell myelomas, this has spectacular consequences, with massive amounts of the monoclonal Ig concentrating in dilated ER cisterna, called Russell bodies (Russell, 1890). Russell bodies (RB) are
readily induced by transfecting into LC producing cells mutant HC lacking the CH1 domain (Valetti et al., 1991; Kaloff and Haas, 1995). Because the CH1 domain is the main BiP binding site in HC, condensation or aggregation in the ER could be caused by failure to engage the chaperone. In support of this idea, BiP seems to be excluded from RB. It is not clear whether RB are formed de novo, or if they represent the expansion of pre-existing ER subcompartments, perhaps one specialized in ERAD. Structures similar to RB are observed in many ER storage diseases (Kim and Arvan, 1998), with the common feature being the synthesis of abundant proteins that can neither be secreted nor degraded. In some patients with hereditary emphysema, the presence of a1 anti-trypsin–containing aggregates in the ER correlates with a severe hepatopathy. However, it is not clear if RBlike structures are toxic per se, or rather reflect a defensive cellular response meant to clear the exocytic pathway from aggregation-prone proteins and to spare essential chaperone molecules (Kopito and Sitia, 2000).
DIFFERENTIATION TO PLASMA CELL Upon encountering antigen, B lymphocytes undergo dramatic morphological changes to become the antibodyproducing machine known as plasma cells (Figure 17.5). This spectacular metamorphosis involves the massive development of the ER and other components of the secretory apparatus. It is worth seeing the changes in Ig production from a quantitative point of view, as the figures underscore the pressure for speed and efficiency during the immune response. It has been calculated that a single plasma cell is able to produce thousands of antibody molecules per second. For IgM secreting cells, this corresponds to forming 100,000 disulfides bonds and adding 50,000 N-linked glycans per second to cargo proteins. IgM polymers are planar molecules 36 nm wide and 4 nm thick (Perkins et al., 1991). There is only room for a few dozen of them in a transport vesicle with an 80-nm diameter (de Curtis and Simons, 1989; Malhotra et al., 1989) Therefore, about 100 vesicles per second must leave any donor compartment to fuse with the downstream station along the exocytic route. It follows that the entire ER would disappear in a few minutes if retrograde membrane transport did not proceed at a similar rate. Thus, membrane traffic must be extremely well organized in plasma cells to avoid exiting and returning vesicles from constantly bumping into each other. In addition, these large amounts of Igs are secreted under the constant inspection of the tight quality control schedule that has been outlined in previous paragraphs. Even in plasma cells, IgM polymerization is not efficient (Tartakoff and Vassalli, 1979). In addition, a considerable fraction of ms chains fail to be inserted into transport-competent polymers and are eventually degraded intracellularly by the proteasomal pathway.
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FIGURE 17.5 Delevopment of the ER in plasma cells. Electron micrographs showing a mature plasma cell (A). Note the parallel stacks of ER distributed within the entire cytosol (N, nucleus). In contrast, the cytosol of B cells (B) contains mostly free polysomes. Upon mitogen stimulation, B cells begin to develop ER cisternae (C). Courtesy of Prof. C.E. Grossi (University of Genoa, Italy). Taken with permission from Atlas of Blood Cells. D. Zucker-Franklin and C.E. Grossi (Eds). Edi.Ermes Milano-Italy.
The formidable secretory load may explain why plasma cells are particularly sensitive to proteasome inhibitors—so much so that these drugs are being considered for the treatment of multiple myeloma (LeBlanc et al., 2002).
Regulation of Ig Production The increase in Ig production during terminal B cell differentiation appears to be controlled at multiple levels. First, transcription of the Ig loci proceeds at a higher rate, due to the convergence of multiple regulatory pathways that involve Blimp1, Bcl6, XBP-1, and other transcription factors (Shaffer et al., 2002). The preferential processing of primary HC transcripts to generate the secretory forms of HC and a general increase in the stability of mRNA encoding secretory molecules (Mason et al., 1988, Hyde et al., 2002) further augments the pool of translatable molecules. Lastly, the spectacular development of the ER, coupled with increased amounts of its structural proteins, resident chaperones (i.e., BiP), and folding enzymes (i.e., proteindisulfide isomerase) (Stockdale et al., 1987) makes the plasma cell an extraordinarily efficient antibody factory.
Role of ER Stress Response The dramatic changes in cellular architecture that accompany the differentiation of a B cell to a plasma cells are
required to allow for the rapid synthesis and secretion of tremendous quantities of Ig. The mechanism(s) by which these changes are accomplished is not presently understood, but some recent data shed light on possible pathways. In all cells types, ER chaperones are transcriptionally upregulated by cellular conditions that affect protein folding in the ER and lead to the accumulation of unfolded proteins. Many of the signal transducers and downstream components of the unfolded protein response (UPR) pathway have recently been identified (Kaufman, 1999; Ma and Hendershot, 2001). Less is known about the ER overload pathway described to upregulate ER chaperones in response to increased synthesis of secretory proteins (Pahl and Baeuerle, 1995). These pathways are thought to monitor the physiological demands placed on the protein folding machinery and promptly adapt to novel developmental requests. Thus, since the production level of Ig-secreting cells represents a formidable task for the synthetic machinery, recent data implicating some UPR components in plasma cell differentiation do not come as a surprise (Calfon et al., 2002; Reimold et al., 2000). It is possible that increased Ig synthesis could drive the expansion of the ER via the UPR. However, an opposite scenario can be envisioned, in which the development of an efficient protein factory could precede the actual production and release of antibodies. Recent proteomics-based and northern analyses indicate that some ER expansion occurs prior to the upregulation of Ig
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synthesis in differentiating B lymphoma cells. Interestingly, mitochondrial proteins and enzymes involved in amino acid and membrane synthesis are also upregulated in this initial phase. Only later does the synthesis of Ig and J chains increase coordinately with the upregulation of additional UPR components (Gass et al., 2002; VanAnken et al., 2003). Another pathway has been described that controls ER chaperone levels in response to mitogenic growth factor signaling. While the specific components of this pathway have not been identified, it appears to be at least partially distinct from the UPR, since only some of the UPR targets are induced (Brewer et al., 1997). These observations raise the interesting possibility that in different cells, or in response to different signals, specific UPR components might be activated independently. It is clear that the induction of the complete classic UPR pathway would not be beneficial to the synthesis of large quantities of Ig by plasma cells. The various components and their possible benefit or detriment to the plasma cell are discussed below.
The UPR Three sensors of ER stress that signal the UPR have been identified, all apparently by monitoring BiP levels. These include the Ire1 a/b and PERK kinases and ATF6, an ERanchored transcription factor (Kaufman, 1999). Activation of Ire1 induces the endonuclease domain at its C-terminus, leading to the removal of 26 bp from the XBP-1 mRNA. Religation by non-spliceosome mediated mechanisms alters the reading frame, thus producing an XBP-1 protein with a novel C-terminus. The remodeled XBP-1 protein encodes a transactivation domain, which is tethered to the N-terminal DNA binding domain that was present in the original XBP1 protein (Yoshida et al., 2001). The larger, stress-induced XBP-1 protein is synthesized in LPS-induced plasmablasts (Calfon et al., 2002), suggesting that Ire1 might be activated at this stage; interestingly, XBP-1p is required for plasma cell differentiation (Reimold et al., 2001). The altered XBP1p can bind to the ERSE regulatory cassettes in the promoters of ER chaperones in vitro, which could provide the mechanism for the generalized upregulation of ER folding factors during plasma cell differentiation (Figure 17.6). In yeast, Ire1p is responsible for membrane biogenesis, and activation of the yeast UPR results in a greatly expanded ER (Chapman et al., 1998). However, to date no such role for mammalian Ire1 has been demonstrated. Activated PERK phosphorylates eucaryotic initiation factor 2a (eIF-2a), which inhibits the assembly of the translation initiation complex, thereby blocking the synthesis of most proteins during conditions of ER stress (Harding et al., 1999). Although this is important to limit the damage to cells undergoing stress until normal physiological conditions can be restored in the ER, it is clear that prolonged activation of PERK during plasma cell differentiation would not be ben-
FIGURE 17.6 Signal transducers of the mammalian UPR and their downstream effects. All transducers possess a lumenal stress-sensing domain and a cytosolic affector domain. Activated Ire1 has a C-terminal endonuclease activity that cleaves XBP1 mRNA. This is known to occur in plasma cell differentiation (Calfon et al., 2002). Activated PERK phosphorylates eIF-2a, thereby inhibiting protein translation, arresting cells in G1 and inducing the pro-apoptotic CHOP protein. No data suggest that PERK-/- mice have a defect in antibody production (Harding et al., 1999). Activation of ATF6 liberates the transcription factor domain, which induces the transcription of ER chaperones, XBP1, and CHOP. No data suggests a role for ATF6 in B cell differentiation. See color insert.
eficial to cells that are gearing up to produce very large amounts of antibody. ATF6 is an ER-localized transmembrane protein with a cytosolic domain that is a transcription factor and a lumenal domain that senses ER stress (Haze et al., 1999). During stress, ATF6 is transported to the Golgi, where it is cleaved sequentially by the S1P and S2P proteases (Ye et al., 2000). This liberates the transcription factor, which binds and activates sequences in the chaperone promoters and, interestingly, in the XBP-1 promoter. If the UPR were used to upregulate ER components during plasma cell differentiation, this would suggest a responsive activation as opposed to a preparatory response, the latter of which is more in keeping with plasma cell activation. Alternatively, it is possible that the various signaling molecules and downstream components of the UPR can be activated independently by different signals. A recent report demonstrated that BLIMP-1 can upregulate XBP-1 mRNA early in plasma cell differentiation (Shaffer et al., 2002), which, since Ire1 is activated (Iwakoshi et al., 2003), could provide a method for inducing ER chaperones without the activation of the complete UPR. However, at this time, it is not known which, if any, components of the UPR are used by plasma cells to increase the secretory capacity of the cell.
CONCLUSION As in the case of recombination, transcription, and splicing, studies on Ig synthesis have revealed basic principles
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of cell biology that control protein folding, transport, and degradation. These concepts have not only entered textbooks but have also contributed to biotechnology and medicine. We look forward to experiments aimed at dissecting the adaptive regulation of protein synthesis, cell morphology, and intercellular communication, as the results will surely offer new exploitable paradigms.
Acknowledgments This work is dedicated to the memory of César Milstein. We thank Drs. Tiziana Anelli, Carlo E. Grossi, Alexandre Mezghrani, and Jenny Woof for useful criticism and suggestions. Part of the work discussed here has been made possible through grants from Associazione per la Ricerca sul Cancro (AIRC), Italian Ministries of Health and Research (Center of Excellence in Physiopathology of Cell Differentiation and CoFin), and Telethon.
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FIGURE 3.1 Comprehensive map of the human immunoglobulin k locus, taken from Kawasaki et al. (2001). (a) Locations of the clones used in sequencing. Sequenced and unsequenced regions are depicted as red and green lines, respectively. (b) Locations of the genes. Vk (red), Jk1-5 (sky blue), and Ck (blue) genes with the same transcriptional polarity are indicated as small vertical lines on the same side of the horizontal line. Lines with full height, 2/3 height, and 1/3 height represent Vk genes with ORFs, pseudogenes with >200 bp, and relics with <200 bp in length, respectively. Relics consisting only of exon I are not included. The names of the Vk genes with ORFs are shown. Thick horizontal lines within the k locus represent duplicated regions; thin horizontal lines indicate regions, which exist in either the proximal or the distal unit. Wedge-shaped shadows indicate deletion events that happened after the inverted duplication. The 6-kb regions between the two wedges show sequence homology, but do not seem to be inverted duplication counterparts; rather they correspond to adjacent biock duplicates generated prior to the inverted duplication. (c) Six categories of interspersed repeats are indicated; Alu (green), MIR (blue), LINE L1 and L2 (red), LTR (yellow), DNA transposons (sky blue), and others (purple). (d) The GC content was plotted with a window size of 4,000 nt and with a sliding size of 2,000 nt. (e) Sequence identity without indels between the proximal unit and the distal unit was plotted with a window size of 10,000 nt and with a sliding size of 500 nt. Thirteen homology blocks (A–M) and the average sequence identities (red dashed lines) are indicated. The scale at the bottom of the figure shows the proximal unit; it does not take into account the gaps in the distal unit.
FIGURE 3.2 A researcher’s dream of the mouse immunoglobulin k locus, taken from Thiebe et al. (1999). The Jk–Ck and Vk genes are depicted as mice. Different gene families have different colors. Mice in full color designate potentially functional Vk genes, mice sketched only in outline are pseudogenes. Relics are not included. The 5¢, 3¢ direction of the Vk genes is indicated by the direction of the mice.
FIGURE 6.1 Summary of transcriptional regulatory elements in the immunoglobulin heavy chain locus. Boxes indicate protein binding sites that are thought to be functional. Where known, the proteins that bind these sites are shown with activators in green and repressors or inhibitors of the activators in red. The arrow indicates the transcription initiation site. Distances are not to scale.
FIGURE 6.2 Summary of transcriptional regulatory elements in the immunoglobulin light chain loci. Boxes indicate protein binding sites that are thought to be functional. Where known, the proteins that bind these sites are shown with activators in green and repressors or inhibitors of the activators in red. Parentheses indicate sites where proteins are presumed to bind but have not been shown experimentally. The arrow indicates the transcription initiation site. Distances are not to scale.
Antigen Independent Lymphoid Pro-B Progenitor Cell
Antigen Dependent
Pre-B Cells
B Cells M
IL-7
SDF-1
IL-7
Immature Mature
Stromal Cell D - JH CD34 CD10 CD19 CD21 CD24
D
Tdt
TdT SDF-1
M
Plasma Cell
V-DJH
V-JL
Activated Memory
FIGURE 9.1 Human B cell development. The antigen-independent stages of B cell development occur in the primary lymphoid organs, the fetal liver, and adult bone marrow. In the secondary lymphoid organs, the mature B cells may encounter cognate antigens and, usually with the help of T lymphocytes, undergo proliferation and differentiation to antibody-secreting plasma cells and memory B cells. The diagram depicts the stages of B cell development and several markers that help define these stages: CD19 ( ), Ig heavy chains ( ), Ig••/Ig•• (||) the surrogate light chain peptides ••5/14.1 and VpreB ( ), and conventional Ig light chains ( ). The configuration of the IgH and IgL chain genes during development is illustrated and is the predominant sequence of IgH prior to IgL rearrangement, although the order may sometimes be reversed. The phenotypic markers shown are a selection of those that have proven useful in identifying B cell developmental stages. The phenotype of the lymphoid progenitor that directly precedes the pro-B cell is unknown, but may be a common lymphoid progenitor that expresses a somewhat different cell surface phenotype depending on the tissue source (i.e., fetal or adult bone marrow, or cord blood) being analyzed. There are also some differences between the cell surface phenotype of early B lineage cells in adult bone marrow, depicted here, and fetal bone marrow (LeBien, 2000).
Igk Rearrangements VL
JL
//
// VL
JL rearrangement
//
// Secondary VL JL rearrangement //
//
IgH Rearrangements VH
DH
JH
//
// DH
JH rearrangement
//
// VH
DHJH rearrangement
//
// CDR3
1st VH replacement
CDR3
2nd
//
// VH replacement
//
// CDR3
FIGURE 9.2 Receptor editing and VH gene replacement. Following the initial rearrangement (red oval), subsequent rearrangements (blue oval) of the Igk chain locus can occur by RSS-mediated recombination of an upstream Vk to a downstream Jk gene segment. A similar process can occur in the Igl locus, but the organization of the gene segments is different. In the heavy chain locus, the rearrangement sequence is usually DH Æ JH, followed by VH Æ DJH (red ovals). A secondary VH Æ VDJH rearrangement (blue oval) utilizes the RSS of the incoming VH gene together with a cryptic RSS found in the 3¢ end of most germline VH genes to accomplish recombination. BCR
FcRH1 FcRH2
FcgRIIB
FcRH3
Fca/mR
FcRH4 FcRH5
FcRX
B Cell Antigen Independent Lymphoid Progenitor
Pro-B
Pre-B Cells
VpreB/ l 5 m HC, Ig a /b, BTK, BLNK D RAG 1/2 D PU.1 E2A D IKAROS D EBF D
FIGURE 9.3 Fc receptor and Fc receptor related genes expressed by human B cells. The BCR, composed of membrane Ig and the Iga/Igb heterodimer with cytoplasmic ITAM (green boxes) is shown at the top of the B cell. The ITIM-containing (red box) FcgRIIB inhibits B cell activation when antigen–antibody complexes crosslink it to the BCR. Also illustrated are members of a recently discovered family of FcR-related genes expressed on B cells (FcRH). These are Ig-like domain proteins that have ITIM, ITAM, or both in their cytoplasmic tails, suggesting a role in regulating B cell responses. The Ig domains (ovals) are color coded to indicate their homology to each other and to the Ig domains in the other FcR. FcRX is a cytoplasmic FcR-related protein expressed in the cytoplasm of germinal center B cells. The Fca/mR binds IgM and IgA and is an endocytic receptor in mice, but the fucntion of its human counterpart is unknown (Sakamoto et al., 2001; Shibuya et al., 2000; Shimizu et al., 2001).
Antigen Dependent B Cells
PAX 5 D
Tdt gc SDF-1 IL-7
Stromal Cell
Plasma Cell
AID CD19, CD21, CD40/CD40L, CD45 Ltab /LTBR, BTK, Lyn Irf4, Oca-B, Oct-2 D Syk D
IgAD/CVID
FIGURE 9.4 Genetic defects in B cell development. The model of B cell development in Figure 9.1 is recapitulated here to illustrate gene defects that affect this process. Mutations identified in humans and mice are indicated in red and discussed in the text. The other defects shown in black have only been identified by gene targeting in mice (Chapter ••), but may be found in humans as more immundeficient patients are studied. The human diseases IgAD and CVID affect antibody production but the predisposing MHC-linked susceptibility gene(s) have not yet been identified, and there is no mouse model.
P P D m Normal BCP
D
Leukemic BCP
P
m
P
Surv/Prolif Cytokines
P = IL-7/? P = proliferation D = differentiation
Stromal Cell
P
FIGURE 9.5 Bone marrow (BM) stromal cells synthesize cytokines essential for the survival and proliferation of normal and leukemic B cell precursors (BCP). In this model, IL-7 constitutes a survival signal, and an unknown cytokine (or cytokines) constitutes a proliferative signal for normal and leukemic BCP. However, the responses to these cytokines differ. Normal BCP (light green) undergo a limited proliferative response. The predominant normal BCP undergoes proliferation and expresses cell surface pre-BCR. Normal proliferating pre-BCR+ cells subsequently differentiate into small pre-B cells (red) expressing cytoplasmic mH chains. BM stromal cell–dependent leukemic BCP (dark green) undergo a robust and continuing proliferative response that is independent of pre-BCR expression. Subsequent mutations can give rise to leukemic subclones that are no longer dependent on BM stromal cells.
FIGURE 11.1 New model for the antigen dependent activation of the BCR. A On resting B cells the BCR forms an oligomeric complex of defined stoichiometry. Signal transduction from the BCR is inhibited by presence of PTP. Exposure to antigen results in the opening of this oligomeric complex and the targeting of the BCR to membranes containing an active NADPH-oxidase. The increased H2O2 production by the NADPH-oxidase inhibits the PTP around the BCR allowing the rapid amplification of the BCR signal through a positive Syk/ITAM feedback loop.
FIGURE 12.1 Intracellular signaling by CD19. The tyrosines of the cytoplasmic domain of murine CD19 and the intracellular signaling proteins that these have been reported to interact with after phosphorylation. In vivo studies of mutant forms of CD19, in which specific tyrosines have been replaced with phenylalanines, have validated roles only for Y482 and Y513 in mediating the functions of CD19 for B cell development and responses to antigens (Wang, et al., 2002).
FIGURE 12.2 Intracellular signaling by CD22. All known intracellular binding proteins of CD22 are shown. The tyrosines that have been mapped as interaction sites are indicated. A clear function has only been demonstrated for SHP1 binding. SHP-1 is most likely the phosphatase responsible for the CD22-mediated inhibition of Ca2+ mobilization.
FIGURE 12.3 Model for regulation of CD22 inhibition by ligand binding. (a) Ligand binding in cis increases tyrosine phosphorylation and SHP-1 recruitment to CD22. First evidence indicates that CD22 binds directly to 2,6Sia on IgM, but other ligands may also be involved. (b) A pool of CD22 exists on the cellular surface, which is not bound to endogenous ligands, but most CD22 is “masked” by ligands in cis. (c) When the mIg binds to self-antigens on other cells, additional CD22 molecules may be recruited by trans interactions into the cellular contact zone, thus resulting in a stronger CD22 inhibition of the mIg signal (indicated by “flash arrow”). In contrast, microorganisms usually do not display sialic acid on the surface, thus resulting potentially in a higher mIg response.
FIGURE 12.4 Additional inhibitory receptors on the B cell. CD72 is constitutively associated with SHP-1 bound to its tyrosine-phosphorylated ITIM motifs. CD100 binding reduces the tyrosine phosphorylation of CD72. PIR-B constitutively binds SHP-1. The ligands are not known yet. The FcgRII receptor is recruited via immune complexes to IgM. In this case, SHIP binds and inhibits sustained Ca2+ signaling by catalyzing dephosphorylation of phosphatidyl inositol (3,4,5) triphosphate (PtI-(3,4,5) P3) into PtI-(3,4)P2. Other functions of FcgRII are described in the text.
FIGURE 13.2 Diagrammatic representation of the stages of an extrafol-
FIGURE 13.1 The phases of T cell-dependent antibody responses. An initial common path of antigen capture by the B cells occurs followed by cognate interaction with the T cell. Responses then diverge, with B cells growing in follicles and extrafollicular sites. The color code identifies the stages in which Ig heavy chain gene switch recombination, variable region hypermutation, and secretion of antibody occur.
licular antibody response: Antigen capture by B cells and T cell priming is followed by cognate T cell interaction with B cells. Some of the activated B cells migrate to extrafollicular foci in the spleen or the medullary cords of lymph nodes where they proliferate as plasma blasts. This growth is associated with CD11chigh dendritic cells. Plasmablasts that are not associated with these dendritic cells appear to die without making the transition to plasma cell. In the spleen, plasma cells produced in the extrafollicular responses and plasma cells that have been generated in follicles compete for space on stroma that supports long-term plasma cell survival. This stroma is associated with blood vessels and contiguous fibrous bands in the red pulp.
FIGURE 13.3 A scheme suggesting how B cells proliferate and activate hypermutation before being selected and induced to differentiate in established GC. Centroblasts proliferate and mutate their Ig-V region genes. Periodically, they are subjected to selection. Successful selection depends on the B cells binding antigen, normally held on FDC, processing this, and presenting the resulting peptides to local T cells. Selected cells differentiate to become memory B cells, plasma cells, or centroblasts. The last remain in the GC and undergo further proliferation and V-region hypermutation. This regeneration of centroblasts is essential for maintaining the GC. Cells that fail selection die in situ by apoptosis.
FIGURE 13.4 Histological sections of the light zone (above) and the dark zone (below) of a GC from a human tonsil. The section is stained with pyronin, which stains RNA magenta, and methyl green, which stains DNA blue/green. In the dark zone, pyroninophilic centroblasts are closely packed. There are many mitoses (marked M). Tingible body macrophages appear as pale islands in the continuum of centroblasts. Occasional apoptotic debris (tingible bodies) in these macrophages is arrowed. In the light zone, only occasional cells are pyoninophilic. The centrocytes are spaced by the presence of the follicular dendritic cell network. Apoptotic nuclear fragments are arrowed.
FIGURE 13.5 Three-color fluorescence of a tonsil GC to show the zonal pattern of this structure. The section is stained for Ki67 nuclear expression by cells in cell cycle (red); these are most abundant in the dark zone (DZ). CD23 expression is shown in blue. This stains the FDC of the apical light zone (ALZ) and B cells in the follicular mantle (FM). CD21 (green) is expressed by a broader network of FDC than CD23; the CD21+ CD23- FDC network below the CD23+ network is termed the basal light zone (BLZ), and that between the apical light zone and the follicular mantle the outer zone (OZ).
FIGURE 13.6 Three-color fluorescence of tonsil GC. On the left the expression of CD3 by T cells is shown green. CD74 (invariant chain) expression by B cells but not FDC is stained blue, while CD21 expression by FDC is stained red. On the right, CD3 again is stained green, IgD (red) is expressed by follicular mantle B cells. CD57 (purple) is expressed by a minority of GC T cells. The CD57+ve T cells tend to be located in the center of the light zone whereas CD57-ve GC T cells are clustered along the junction of the follicular mantle and the light zone.
FIGURE 14.1 Secondary lymphoid tissue organization and lymphocyte trafficking. Secondary lymphoid tissues function to bring together recirculating lymphocytes and antigen, with each lymphoid tissue sampling a different portion of the body’s fluids for the presence of antigen or antigen-presenting cells (DCs). The diagrams of a lymph node crosssection, a splenic white-pulp cord, and some surronding red pulp (accounting for about one fifth of a spleen crosssection), and a Peyer’s patch cross-section aim to show the themes common to all secondary lymphoid organs, with naïve lymphocytes gaining entry from the blood, and B cells and T cells quickly migrating into their separate subcompartments (dashed black arrows). B cells migrate to follicles in response to CXCL13 made by follicular stromal cells, whereas T cells localize within T zones in response to CCL21 and CCL19 made by T zone stromal cells. Within these compartments the cells undergo random walks to survey for intact antigen or MHC-peptide complexes, respectively. Each organ has areas rich in macrophages (indicated in purple shading) that capture and degrade antigen. The diagrams also illustrate key specializations of the tissues: the presence of a greater proportion of B cells (brown areas) than T cells (blue areas) in spleen and PPs, but not in lymph nodes; entry into LN and PP occurs via HEV, whereas entry into the spleen is by release from open-ended terminal arterioles (ta), many of which open into the marginal sinus (ms); antigen and antigen-bearing DC arrive in LNs via afferent lymph fluid, whereas in the spleen antigens arrive via the blood, and DCs may arrive via this route. There is also a large population of immature DCs already present in the spleen (near the bridging zone); in PPs, antigen is transported by M cells directly to the subepithelial dome (sed), a region overlying the follicles that contains immature DCs and macrophages. Naïve B lymphocytes exit each of the lymphoid tissues (green arrows) after about one day, exiting via lymphatics from LNs and PPs or via red-pulp venous sinsusoids in the spleen. The lymphatics draining the PPs ferry cells to the mesenteric LNs. LN efferent lymphatics return cells to the blood via the thoracic duct, from where the cells can quickly gain entry to another secondary lymphoid organ in the ongoing process of lymphocyte recirculation. In addition to the populations of recirculating B cells, the spleen contains a more sessile population, the marginal zone B cells, located in the marginal zone. Intact antigen reaches lymphoid tissues in fluid phase and may also be carried in association with cells. Immune complexes can become trapped and displayed for long-periods on FDCs (a subset of follicular stromal cells), but other types of antigen transport cells (possibly DCs) may be involved in directly releasing antigen for recognition by B cells. Upon B cell activation by T-dependent antigens, germinal centers form within the B cell follicles, and antibody secreting cells (ASCs) migrate to the red-pulp of spleen or the medullary cords of LNs; in the case of PPs, many ASCs are released via the lymphatics and appear in the mesenteric LNs as well as homing to the gut.
FIGURE 14.2 Rolling, triggering, and adhesion requirements during B cell interaction with HEV in secondary lymphoid organs. Requirements are indicated for peripheral LN, mesenteric LN, and Peyer’s patches. The receptor–ligand pair that plays the dominant role at each step in each lymphoid organ is shown at the top of each list. Receptor–ligand pairs that make only minor contributions to an interaction are shown in smaller font size. In addition to CCL21, CCL19 may function as a triggering ligand for CCR7. Color code: brown, rolling cell; red, cell experiencing chemokine triggered integrin activation; blue, adherent cell. The corresponding molecular requirements for these steps are shown in the same color.
FIGURE 14.3 B cell distribution in the mouse spleen. Cryostat section of unimmunized mouse spleen stained in brown to detect IgD and in blue to detect IgM. Naïve, recirculating B cells appear brown (IgDhiIgMint) whereas marginal zone (MZ) B cells appear blue (IgDloIgMhi). The image encompasses about 1/4 of the tangential cross-section and shows a large white-pulp cord centered around a central arteriole (ca) with two large B cell follicles (brown, labeled), two smaller follicles (brown), and central unstained T zones (white). The marginal zone (MZ) surrounds the B cell follicles, separated in the mouse by the marginal sinus (MS), a site where many small arterioles terminate. Gaps in the MZ are observed at the edges of the follicles, regions often referred to as MZ “bridging zones” (one of these is labeled). IgM ASCs (intense blue staining) can be seen in the bridging zones and also in clusters within the red pulp. The scattering of B cells (brown) within the red pulp may include recirculating B cells that are passing out of the spleen as well as cells resident in this area.
FIGURE 14.4 Cross-sectional diagram of an omental milk spot. Milk spots lie in a double sheet of mesothelium and are made up predominantly of B cells and macrophages. They also contain fibroblasts and adipocytes. Mast cells and occassional T cells are also present (not shown). In the mouse, the majority of omental B cells are of B-1 phenotype. The capillary network within the milk spot is a site of attachment and entry of circulating B1 cells, and this depends on the chemokine CXCL13. B cells are likely to pass through the fenestrated mesothelium overlying milk spots to access the body cavity. The mesothelial basement membrane (not shown) is also discontinuous in areas overlying a milk spot.
FIGURE 14.5 Principal migration pathways of B-lineage cells. Each arrow indicates an active migration event by a B-lineage cell (some arrows may incorporate more than one migration step). The principal type of Blineage cell at each location is indicated in parentheses (in blue). Green arrows indicate migration events that occur homeostatically or during development, red arrows refer to migration events that occur following antigen-encounter and B cell activation or differentiation. Distinct migration cues are required for cells to reach each of the indicated tissues or compartments. Note that the diagram emphasizes migration events and is not meant to be to scale or to represent anatomical organization.
FIGURE 17.1 Assembly-dependent folding resulting in restrictions on HC/LC pairing. Folding of the CH1 domain is dependent on binding to a folded CL domain. If V regions follow suit, a folded VH region could pair with a LC possessing either a folded or unfolded VL region and induce folding of the latter. However, a VH region unable to fold by itself could only pair with a LC containing a folded VL domain.
FIGURE 17.2 Protein quality control in the ER. Following co-translational translocation into the ER lumen via the Sec61 channel (in blue), HC and LC fold, assemble, and when required form polymers under the assistance of many ER resident chaperones and enzymes before transport to the Golgi. Disulfide bonds are inserted and isomerized in this phase. Molecules that fail to attain their native structure within a given time are dislocated to the cytosol to be degraded by proteasomes. Disassembly and disulfide reduction precede dislocation (Fagioli and Sitia, 2001). The extraction of substrates across Sec61 is facilitated by p97 and ubiquitination (Tsai et al., 2002).
FIGURE 17.3 ER quality control checkpoints regulating Ig transport. (a) In the absence of SLC or LC assembly, the CH1 domain remains unfolded and is associated with BiP, which prevents its transport to the Golgi. (b) The transmembrane domain of mm contains a number of hydrophilic residues that prevent its transport until they are masked by Iga/b. (c) VpreB (green) is missing its ninth and final b strand and has a unique sequence (white box). It is unable to fold until l5 assembles and provides its missing strand. (d) The secretory IgM monomer possesses an unpaired cysteine in its C-terminal tailpiece (red) that is recognized by thiol retention mechanisms until it associates with J chain in IgM pentamers or with other monomers in IgM hexamers.
FIGURE 17.4 Glycan processing diverts terminally misfolded glycoproteins to ERAD. To maintain homeostasis, proteins that fail to fold within a given time must be degraded. Removal of the terminal mannose from the central branch of N-glycans by ER mannosidase I targets terminally misfolded or orphan glycoproteins to dislocation, probably via interaction with EDEM (Ellgaard and Helenius, 2001).
FIGURE 17.6 Signal transducers of the mammalian UPR and their downstream effects. All transducers possess a lumenal stress-sensing domain and a cytosolic affector domain. Activated Ire1 has a C-terminal endonuclease activity that cleaves XBP1 mRNA. This is known to occur in plasma cell differentiation (Calfon et al., 2002). Activated PERK phosphorylates eIF-2a, thereby inhibiting protein translation, arresting cells in G1 and inducing the pro-apoptotic CHOP protein. No data suggest that PERK-/- mice have a defect in antibody production (Harding et al., 1999). Activation of ATF6 liberates the transcription factor domain, which induces the transcription of ER chaperones, XBP1, and CHOP. No data suggests a role for ATF6 in B cell differentiation.
FIGURE 18.2 Signaling pathways triggered by BCR–CD19/21–FcgRIIB co-ligation. Cellular activation is inhibited by the recruitment of the inositol phosphatase SHIP to the FcgRIIB phosphorylated ITIM sequence.
FIGURE 18.3 A model for the role of FcgRIIB in affinity maturation of germinal center B cells. Higher affinity BCRs rescue somatically hypermutated B cells from FcgRIIB-triggered apoptosis and negative selection.
FIGURE 18.4 Complement and Fc receptors are important in the regulation of B lymphocyte responses at multiple stages of the peripheral response. Activation of naïve B cells by engagement of antigen coupled to complement C3d results in co-ligation of BCR and co-receptor and, in the presence of T cell help, leads to activation and expansion in absence of specific IgG. Activated cells initiate a germinal center within the splenic follicles that is organized by follicular dendritic cells (FDC). Complement and Fc receptors (FcRIIB) expressed on GC B cells have opposing roles in that CD21/CD35 acts to enhance BCR signaling whereas FcRIIB participates in dampening the BCR signal in the presence of specific IgG immune complexes. By contrast, both complement and Fc R expressed on FDC act to promote the antigen selection of high-affinity GC B cells. Whereas CD21/CD35 are critical in the retention of antigen complexes, FcRIIB seems to promote B cell selection by competing for Ig Fc ligand. Post GC B cells continue to require both complement and FcRIIB in continued antigen selection, at least within the spleen, as maintenance of long-term memory B cells is dependent on both receptor systems.
FIGURE 25.3 The repertoire of VH gene segments expressed by B cells from normal adult blood. Single CD19+ B cells from a normal adult donor were separated by FACS, and rearranged VH genes in seventy-five cells sequenced. The percentage of B cells with recombined individual and potentially functional VH gene segments is shown.
FIGURE 25.4 Ribbon diagram of a molecular model of an Ig Fv region showing VH (darker shading) and VL (paler shading). (a) The amino acids in FR1 identified as being involved in recognition of the I carbohydrate antigen by mutagenesis (H7trp, H22cys, H23ala, H24val, H25tyr) are indicated. (b) The space-filling representation of H7trp, H23ala, and H25tyr reveals a hydrophobic patch in FR1 that is a candidate for interaction with sugar residues.
FIGURE 25.5 Model of the route to development of autoantibodies characteristic of primary biliary cirrhosis (PBC). High-affinity somatically mutated IgG autoantibodies derived from patients with PBC bind to the inner lipoyl domain of the pyruvate dehyrogenase complex. Reversion of VH and VL sequences to the germline sequences of the naïve IgM-expressing B cell of origin leads to complete loss of reactivity to the autoantigen. This suggests that another antigen, possibly a pathogen, initiated the B-cell response. Reversion of VH to germline, with retention of mutations in VL, changes specificity from the inner lipoyl domain to a different part of the protein, the E1/E3 binding domain. Shifting of epitope specificity in vitro may reflect similar events in vivo.
Mouse L VH1
L VHn D1 D13 JH1 JH4 Cµ Cd Cg3 Cg1 Cg2bCg2a Ce Ca
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Eµ3’ FIGURE 27.4 Schematic comparison of the mouse and catfish immunoglobulin heavy chain gene loci indicating the positions of the enhancers (orange) in relation to the VH exons, D and JH segments, and CH exons. The mouse 3¢ enhancers are, from left to right: hs3a; hs1,2; hs3b; hs4.
FIGURE 29.1 Comparison of sites of primitive and definitive hematopoiesis during embryogenesis. Sites of primitive hematopoiesis are in red and definitive sites in green. The diagram for mouse is based on Dzierzak and Medvinsky, 1995; Godin et al., 1995; for chicken on Dieterlen-Lievre et al., 1993; and for frog on Turpen et al., 1997.
FIGURE 29.3 Sites of hematopoiesis in zebrafish embryo. Genes associated with hemangiogenic progenitors are expressed in the lateral mesoderm in two stripes. In the lateral mesoderm of the trunk (TLM), the stripes converge medially to form the intermediate cell mass (ICM). Primitive erythrocytes differentiate from the cells of the anterior ICM. Genes associated with hematopoietic progenitors are expressed in the posterior ICM. Embryonic macrophages are derived from the caudal lateral mesoderm (CLM) before 24 hpf. Definitive hematopoiesis is thought to originate in cells within or near the dorsal aorta (DA) from 24 to 72 hpf. Cells in the ventral tail (VT) express genes associated with hemangiogenic progenitors until day 4. Lymphoblasts are seen in the thymus (Th) from 65 hpf and express Ikaros from 72 hpf.
FIGURE 31.1 Structural overview of the murine IgG2a monoclonal antibody, Mab231. (a) Structure of the intact antibody, including two light chains each composed of a variable (VL) and a constant (CL) immunoglobulin (Ig) domain (red) and two heavy chains each composed of a variable (VH) and three constant (CH1, CH2, and CH3) domains (blue). The two hinge regions are highlighted within the dotted oval, revealing the source of structural asymmetry within the intact antibody. (b) Ribbon diagram of a single Ig domain, VL, of Mab231 highlighting its anti-parallel b-sheet secondary structure. The amino- and carboxy-termini are marked, as well as the complementarity determining region loops, CDR1 (yellow), CDR2 (blue), and CDR3 (green). (c) Molecular surface of the antibody combining site of Mab231 formed by the intersection of the apical regions of VL and VH. The CDR loops provide a nearly contiguous surface for antigen recognition. VLCDR1 (yellow); VLCDR2 (blue); VLCDR3 (green); VHCDR1 (magenta); VHCDR2 (cyan); VHCDR3 (red).
FIGURE 31.2 Conformational changes induced by antigen binding. (a) Concerted movement of the Fab8F5 CDR H3 loop induced upon binding its peptide antigen. The unbound Fab structure is in green, the bound Fab structure in blue, and the peptide antigen in yellow. VHSer101 in the bound form makes two hydrogen bonds to Lys157 of the peptide, and VHTyr102 is displaced by more than 7 Å between the unbound and bound Fab8F5 molecules. (b) Atomic rearrangement of the CDR H3 loop of the anti-peptide Fab17/9. The color scheme is the same as in (a). Side chains of residues in contact between the antibody and antigen in the bound complex are shown. Contrary to many anti-peptide antibodies, anti-protein antibodies generally exhibit relatively small conformational changes upon binding antigen as shown in panels (c) and (d) for the anti-HEL antibody FabHyHEL63. (c) Superposition of the CDR H3 loops of FabHyHEL63 bound to HEL (blue) and three different unbound forms: solved in the C2 spacegroup (green); one molecule from the asymmetric unit of the free antibody solved in the P1 spacegroup (red); and the second molecule of the asymmetric unit of the free antibody solved in the P1 spacegroup (yellow). (d) Superposition of the CDR H2 loops of FabHyHEL63 bound to HEL (blue) and three different unbound forms. The color scheme is the same as in (c).
FIGURE 31.3 Structure of antibody–antigen complexes. (a) Ribbon diagram of the FvD1.3–HEL complex. HEL (yellow), D1.3 VL domain (green), and D1.3 VH domain (blue). Residues of HEL and D1.3 involved in interactions in the antigen–antibody interface are cyan and red, respectively. Heavy (H) and light (L) chain CDRs 1–3 are numbered. (b) Diagram of the FvD1.3–FvE5.2 complex. D1.3 VL domain (green), D1.3 VH domain (blue), E5.2 VL domain (yellow), and E5.2 VH domain (gray). Residues of D1.3 and E5.2 in contact in the structure are red and cyan, respectively. D1.3 and E5.2 heavy (H) and light (L) chain CDRs are labeled 1–3, with an asterisk (*) denoting CDRs from E5.2.
FIGURE 31.4 Energetic maps of antigen–antibody interfaces. (a) Space-filling model of the surface of D1.3 (left) in contact with HEL and of the surface of HEL (right) in contact with D1.3. The two proteins are oriented so that they may be docked by folding the page along a vertical axis between the components. Residues are color-coded according to the loss of binding free energy upon alanine substitution: red, >4 kcal/mol; yellow, 2–4 kcal/mol; green, 1–2 kcal/mol; blue, <1 kcal/mol. VL and VH residues are labeled in white, and VL residues are denoted by an asterisk (*). (b) Model of the surface of D1.3 (left) in contact with E5.2 and of the surface of E5.2 (right) in contact with D1.3. Residues are colored and labeled as in (a).
FIGURE 31.6 Cross-reactivity of antibodies. (a) Interaction of FvD11.15 (VH domain in blue, VL domain in green) with PHL (yellow) and JEL (red). The left panel is a close-up view of the encircled region in the right panel, highlighting the relative displacement of the 100–104 loop region between PHL and JEL, resulting in a steric clash between JEL residues Val102 and His103 with the FvD11.15 VH domain. (b) Interaction between FvD11.15 [same color scheme as in (a)] with PHL (yellow) and HEL (red). The left panel is a close-up view of the encircled region in the right panel and highlights the productive interactions that are made between FvD11.15 VHTyr57 and PHL Lys113 (four hydrogen bonds, indicated by dotted lines). Conversely, productive interactions between FvD11.15 VHTyr57 and HEL Asn113 are largely absent (one hydrogen bond, not shown for clarity) and is likely the reason for the binding affinity discrepancy between the two antigens. (c) Hydrogen bonding between FvD1.3 residues VLTyr32, Phe91, Trp92, and Ser93 with HEL Gln121 (left panel) and TEL His121 (right panel), the only amino acid difference between these two antigens. HEL Gln121 makes three hydrogen bonds (indicated by dotted lines) to the main chain nitrogen atom of Ser93, the main chain oxygen atom of Phe91, and the phenyl ring of Tyr32. All three hydrogen bonds are lost in the FvD1.3–TEL complex; however, a peptide flip between FvD1.3 residues Trp92 and Ser93 results in a new hydrogen bond between the TEL His121 side chain and the main chain oxygen atom of Trp92.
FIGURE 31.7 Affinity maturation via preorganization of the antibodycombining site in Fab48G7. Superposition of the CDR loops of the free germline Fab48G7 (green), the antigen bound germline Fab48G7 (red), the unliganded mature Fab48G7 (blue), and the liganded mature Fab48G7 (yellow). The conformational changes invoked upon antigen binding by the germline antibody are commonly replicated in the mature antibody, especially for VLCDR1, VLCDR2, VLCDR3, and VHCDR3.
FIGURE 32.1 Generation of monoclonal antibodies using hybridoma technology and phage display. Hybridoma technology (Panel A, left): The naïve mouse generates a primary repertoire of more than 106 rearranged VH and VL genes (colored bars) in B cells, coding for antibodies that are displayed as membrane-bound molecules. Immunization causes antigen-driven proliferation and somatic hypermutation (“stars” within V genes). To make hybridomas, B cells are harvested from the spleen or marrow and fused with immortal myeloma cells (wrinkled edges) to generate immortalized, antibody-secreting hybridomas. Hybridomas are screened by ELISA for antigen binding and the monoclonal antibodies produced in tissue culture. Phage display (Panel B, right): For phage display, B cells are isolated from immunized mice (as in panel A) or naïve or immunized humans. Heavy and light chain V genes (shaded bars) are amplified by PCR and assembled as single-chain Fv antibody genes (scFv). Alternatively, rearranged V genes can be generated entirely in vitro from cloned V segments and synthetic oligonucleotides. The repertoire of scFv genes are cloned into a phage display vector, where the encoded scFv proteins (colored ovals) are displayed as fusion proteins to one of the phage coat proteins. The phage contain the appropriate scFv gene within. Multiple rounds of selection with immobilized antigen allows isolation of even rare antigen-binding phage antibodies, which are identified by ELISA. Native sdFv can be expressed from E. coli and purified for characterization and use in assays.
FIGURE 32.5 Different formats for antibody phage display. Wildtype phage (far left) has four coat proteins, pIII, pVIII, pVII, and pIX. pIII display (middle two phage). ScFv (red and yellow) are fused to the minor coat protein pIII. In true phage vectors (left), scFv are displayed on all three to five copies of pIII. In phagemid display, scFv display is typically single copy per phage particle, due to competition from wildtype pIII from the helper phage. pVIII display (far right phage) theoretically yields multiple scFv/phage particle. Wildtype pVIII must be supplied to make phage.
FIGURE 32.7 Yeast display of scFv antibody fragments. For yeast display (left panels), scFv genes are cloned as Cterminal fusions to the Aga2 gene, with a c-myc epitope tag at the scFv C-terminus for detection of expression. On the yeast cell surface, Aga2 disulfide bonds to Agal leading to scFv display on the cell surface. HA = HA epitope tag; cmyc = myc epitope tag. Binding yeast can be selected by flow cytometry (right panel). Binding of fluorescent antigen is displayed on the X-axis and binding of fluorescent anti-myc antibody (to quantitate expression) is displayed on the Y-axis. Populations of non-scFv expressing yeast, no antigen binding, low affinity antigen binding, and high affinity antigen binding can be clearly seen.
FIGURE 32.8 Ribosome display of scFv antibody fragments. ScFv DNA is generated by PCR and then transcribed in vitro to generate mRNA. mRNA is translated in vitro, the absence of a stop codon in the scFv mRNA results in continued attachment of the mRNA to the ribosome, along with the translated scFv. Ribosomes displaying antigen-binding scFv are isolated from nonbinders by affinity chromatography. The ribosome is then disrupted, freeing the mRNA, which is converted back to DNA by RT-PCR. This last step can introduce random mutations into the scFv gene, mimicking the random introduction of mutations by the somatic hypermutation machinery. These steps are repeated to isolate and affinity mature antigen binding scFv.
Figure 34.2 Two-color FISH analysis of tail fibroblasts prepared from a 4-week old double chromosomic mouse. Probes identified the human H chain locus (red) and k L chain locus (green). Permission for display of the figure (Tomizuka et al., 2000) was kindly provided by Prof. Ishida and Proc. Natl. Acad. Sci. USA.
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18 Fc and Complement Receptors JEFFREY V. RAVETCH
MICHAEL C. CARROLL
Laboratory of Molecular Genetics and Immunology, The Rockefeller University, New York, New York, USA
The CBR Institute for Biomedical Research and the Department of Pediatrics, Harvard Medical School, Boston, Massachusetts, USA
Although the Fc and complement systems have evolved distinct strategies for facilitating the interaction of environmental antigens with B cells to contribute to an effective antibody response, we have chosen to consider these two systems in parallel since they are interdependent in considering the fate of interaction of the immune complex with the adaptive immune response. In this chapter we first deal with the significance of these systems by reviewing the phenotypes of complement and FcR deficiencies in both mice and humans. Our understanding of the mechanisms that account for these phenotypes is discussed for each pathway, and the interactions that occur between these systems is summarized. For the Fc receptor system, we focus on FcgRIIB, the inhibitory receptor expressed exclusively on both B cells and FDCs, for which substantial data is now available. The early components of complement, notably C1q, C4, and C3 and the complement receptors CR1 and CR2 are particularly relevant to the role of complement in afferent responses. In vivo studies of complement and Fc receptor systems is our primary emphasis, although in vitro data is discussed where appropriate. Complement and FcR involvement in the efferent response is not discussed here; for discussion of those systems the reader is referred to several recent reviews [1, 2].
apparent on all genetic backgrounds investigated. Backcrossing onto the C57Bl/6 background further resulted in animals with severe autoimmune disease [4]. These animals displayed a spontaneous loss of tolerance to nuclear antigens, with high titers of IgG antibodies to dsDNA and histone H2A/2B by 6 months of age. Autoimmune disease was apparent in these animals, with prominent vasculitis and glomerulonephritis resulting from immune complex deposition. The defect in these animals appears to reside, in part, in the B-cell compartment, as determined by reconstitution of the autoimmune phenotype through the transfer of RIIB deficient bone marrow into B cell or T and B cell–deficient recipients who have otherwise normal RIIB expression on their myeloid cells. Further evidence that RIIB deficient animals have a defect in the maintenance of peripheral tolerance was demonstrated by the ability of these animals to develop autoantibodies to murine type II or type IV collagen after immunization with bovine type II or IV collagen, respectively, on a nonsusceptible H-2b background [5, 6]. The consequence of this induced breakage of tolerance to these autoantigens was the development of collagen-induced arthritis in nonsusceptible backgrounds in response to type II collagen immunization, or Goodpasture’s syndrome in the case of type IV collagen immunization. These data suggest that decreased levels of RIIB correlates with a susceptibility to the development of autoimmune diseases like SLE and is further supported by the reports of decreased RIIB levels on B cells in mouse strains predisposed to the development of antinuclear antibodies, such as NZB, BXSB, and MRL [7, 8] and by its tight linkage to SLE loci on chromosome 1 in both human and mouse studies [9]. These data indicate that RIIB expression on B cells contributes to the maintenance of peripheral tolerance to nuclear antigens and to the suppression of autoantibody production
CONSEQUENCES OF FCgRIIB DEFICIENCY The targeted disruption of FcgRIIB results in animals with amplified antibody responses to both TI and TD antigens, with Ig titers increased by five- to ten-fold [3]. Baseline serum immunoglobulin levels are unaffected and the mice display normal Ig half-life. This amplification is
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in response to cross-reactive, exogenous antigens. The mechanisms that account for these observations will be discussed in detail below.
CONSEQUENCES OF COMPLEMENT AND COMPLEMENT RECEPTOR DEFICIENCIES As is the case for FcgRIIB, complement and complement receptors are required for the maintenance of tolerance to autoantigens, although through quite different mechanisms, as will be discussed below. Deficiency in complement proteins C1q and C4 leads to an increased susceptibility to the development of lupus and the production of autoantibodies both in humans and murine models. This striking observation supports a critical role for complement and its receptors in the protection from maturation of self-reactive B cells. Deficiency in CD21/CD35, when combined with Fas deficiency, results in earlier onset and increased disease. Moreover, the identification of a mutant Cr2 allele with reduced C3d binding correlates with increased autoantibodies and suggests a role for CD21/CD35 in the regulation of selfreative B cells. The stage at which complement participates in negative selection is not clear, as it could affect immature B cells within the bone marrow, in the transitional stage in the spleen, or both. The development of knockout mice bearing targeted Cr2 loci provided animal models to not only directly test the importance of the two receptors but to dissect the mechanism of B cell regulation in vivo. Two lines of targeted mice have been reported, and in general they have a similar phenotype, although one line expresses a low level of truncated CD21. Both lines fail to develop secondary antibody responses to inert protein antigens administered in the absence of adjuvant; a reduced number and size of germinal centers characterize their response. Both the primary and secondary responses to certain T-D antigens are reduced, thus suggesting that complement can function both in the initial activation and expansion of antigenspecific B cells as well as their survival within GC. Given the similar impaired response of mice deficient in either C3 or C4 with those deficient in the receptor suggests that complement mediates its effects on B cell responses via CD21/CD35. The observation that co-cross linking of the BCR and coreceptor lowered the threshold for B-cell activation led to the important general concept that complement receptors link innate and adaptive immunity [10]. Thus, the innate immune system provides a novel mechanism of identifying pathogens by covalent attachment of C3 and enhancing the humoral response via CD21/CD35. This concept was tested by comparing the humoral response of mice deficient in complement proteins C3 or C4 or receptors CD21/CD35
following infection with type I herpes simplex virus (HSV-1). Significantly, all three groups of mice failed to develop a significant antibody response to the infectious virus despite multiple infections (Figure 18.1). Thus, complement and its receptors are essential in the formation of a memory B cell response to this important human pathogen. The defect was in the B cell compartment, because CD4+ T cells were activated normally and responded to viral antigens on secondary stimulation in vitro. In summary, complement receptors CD21 and CD35 mediate the enhancing effects of complement C3. Thus, the receptors link the innate and adaptive response that results in enhanced humoral immunity. Experiments involving chimeric mice support the overall conclusion that CD21/CD35 expression on B cells is essential for B cell activation in vivo but that FDC expression is also important in the localization of antigen within the follicles and the promotion of long-term B cell survival. Complement receptors CD21/CD35 also appear to be important in the selection or expansion and maintenance of B-1 cells, because Cr2-def mice have an altered repertoire of natural antibody to certain but not all self-antigens. Thus, Cr2-def mice are missing or have reduced levels of natural antibody involved in the induction of reperfusion injury (I/R)
Fc RECEPTORS Expression Pattern and Signaling Properties of RIIB The interpretation of the phenotypes displayed by RIIBdeficient mice is complicated by the ubiquitous expression pattern of this inhibitory receptor. On B cells and FDCs, RIIB is the only Fc receptor for IgG expressed. RIIB is expressed at all stages of B cell development including pre-, pro-, and mature populations. Expression levels of RIIB are modulated on different B cell populations, displaying higher levels on peripheral B cells and reduced expression on germinal center B cells [11]. IL-4 further downregulates RIIB expression on germinal center B cells [12]. RIIB expression is induced on FDCs upon antigen stimulation [11]. In addition, RIIB is expressed on immature dendritic cells, where it accounts for >75% of surface expression of FcRs on those cells [13]. RIIB deficiency does not appear to perturb the development of any of these lineages, suggesting that RIIB functions specifically in response to immune complex engagement during the active immune response. RIIB is not expressed on NK cells, T cells, or stromal cells.
ITIM Pathways The inhibitory motif, embedded in the cytoplasmic domain of the single chain FcgRIIB molecule, was defined
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FIGURE 18.1 Deficiency in complement receptors CD21 and CD35 or complement proteins C3 and C4 results in an impaired humoral response to herpes simplex virus Type I (HSV-1). Mice were infected intradermally with 2 ¥ 106 pfu of either replication-deficient strain HD-2 (panels a–c) or -competent strain KOS 1.1 (panel d) at day 0 and challenged on week 3 with a similar dose of live virus. Results indicate a normal secondary response as expected among WT controls but an impaired response in mice deficient in complement receptors or proteins.
as a 13 amino acid sequence AENTITYSLLKHP, shown to be both necessary and sufficient to mediate the inhibition of BCR-generated calcium mobilization and cellular proliferation [14, 15]. Significantly, phosphorylation of the tyrosine of this motif was shown to occur upon BCR co-ligation and was required for its inhibitory activity. This modification
generated an SH2 recognition domain that is the binding site for the inhibitory signaling molecule SHIP [16, 17]. In addition to its expression on B cells, where it is the only IgG Fc receptor, FcgRIIB is widely expressed on macrophages, neutrophils, mast cells, dendritic cells, and FDCs and is absent only from T and NK cells. Studies on FcgRIIB provided the
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impetus to identify similar sequences in other surface molecules that mediated cellular inhibition and resulted in the description of the ITIM, a general feature of inhibitory receptors. FcgRIIB displays three separable inhibitory activities, two of which are dependent on the ITIM motif and one that is independent of this motif (Figures 18.2 and 18.3). Coengagement of FcgRIIB to an ITAM-containing receptor leads to tyrosine phosphorylation of the ITIM by the lyn kinase, recruitment of SHIP, and the inhibition of ITAMtriggered calcium mobilization and cellular proliferation [16, 18]. These two activities result from different signaling pathways, with calcium inhibition requiring the phosphatase activity of SHIP to hydrolyse PIP3 and the ensuing dissociation of PH domain containing proteins like Btk and PLCg [19] (Figure 18.2). The net effect is to block calcium influx and prevent sustained calcium signaling. Calciumdependent processes such as degranulation, phagocytosis, ADCC, cytokine release, and pro-inflammatory activation are all blocked. The arrest of proliferation in B cells is also dependent upon the ITIM pathway, through the activation of the adaptor protein dok and subsequent inactivation of MAP kinases [20, 21]. The role of SHIP in this process has not
been fully defined, since it can affect proliferation in several ways. SHIP, through its catalytic phosphatase domain, can prevent activation of the PH domain survival factor Akt by hydrolysis of PIP3 [22, 23]. SHIP also contains PTB domains that could act to recruit dok to the membrane and provide access to the lyn kinase that is involved in its activation. Dok-deficient B cells are unable to mediate the FcgRIIB-triggered arrest of BCR-induced proliferation, while retaining their ability to inhibit a calcium influx, thus demonstrating the dissociation of these two ITIM-dependent pathways. The third inhibitory activity displayed by FcgRIIB is independent of the ITIM sequence and is displayed upon homoaggregation of the receptor. Under these conditions of FcgRIIB clustering, a pro-apoptotic signal is generated through the transmembrane sequence (Figure 18.3). This pro-apoptotic signal is blocked by the recruitment of SHIP, which occurs upon co-ligation of FcgRIIB to the BCR, due to the Btk requirement for this apoptotic pathway [24]. This novel activity has only been reported in B cells and has been proposed to act as a means of maintaining peripheral tolerance for B cells that have undergone somatic hypermutation. Support for this model comes from the in vivo studies of
FIGURE 18.2 Signaling pathways triggered by BCR–CD19/21–FcgRIIB co-ligation. Cellular activation is inhibited by the recruitment of the inositol phosphatase SHIP to the FcgRIIB phosphorylated ITIM sequence. See color insert.
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FIGURE 18.3 A model for the role of FcgRIIB in affinity maturation of germinal center B cells. Higher affinity BCRs rescue somatically hypermutated B cells from FcgRIIB-triggered apoptosis and negative selection. See color insert.
RIIB-deficient mice in induced and spontaneous models of autoimmunity. Recent studies have provided evidence that RIIB is involved in affinity maturation, Ig enhancement and suppression, and generation of the memory response. These various roles contribute to the overall role of RIIB in the maintenance of peripheral tolerance since perturbation in one or more of these pathways can contribute to the emergence and amplification of the autoimmune response.
Affinity Maturation Affinity maturation occurs within the germinal center, where somatically mutated BCRs undergo selection on antigen retained on FDCs [25, 26]. Antigen is retained in the form of immune complexes and involves the interaction of both complement receptors and FcRIIB with these immune complexes on FDCs. B cells also express both complement and FcRIIB. An analysis of the role of RIIB in affinity maturation thus represents the contribution of both FDCs and RIIB to this process. The role of FDC-expressed RIIB has been clarified by the development of chimeric mice in which only the FDC RIIB is deficient and the B-cell RIIB is retained [27, 28]. In these mice, potentiation of affinity maturation is seen. This potentiation likely results from the increased stringency of selection that occurs in the GC, which, in turn, results from unopposed access of FDC immune complexes to B-cell FcRIIB, as has been proposed [29]. Co-ligation of the BCR and RIIB attenuates the BCR
signal by the negative signaling role of SHIP. This negative signal would impose a requirement for a higher threshold for effective BCR stimulation and thus favor higher affinity BCRs. In addition, under those conditions where co-ligation to BCR is ineffective, ligation of RIIB alone results in an apoptotic signal, resulting in the elimination of those somatically mutated B cells with low affinity for antigen. The situation is reversed when B cells that lack RIIB expression are transferred. In that case decreased apoptosis is observed and the resulting B cells display a reduced affinity for antigen (Kalergis and Ravetch, unpublished observations). This is perhaps due to the persistence of low-affinity B cells that arise as a consequence of somatic mutation and are normally eliminated by a negative selection mechanism involving RIIB crosslinking and induce apoptosis. Since RIIB is regulated on both FDCs and B cells, the potential is provided for fine-tuning the survival and selection of B cells in the GC by the interaction with ICs on FDCs. This role of RIIB on affinity maturation could provide part of the explanation for the loss of tolerance in RIIBdeficient animals. Since RIIB would provide one mechanism for the elimination of low-affinity and potentially autoreactive BCRs that may arise through somatic mutation during the GC reaction, decreasing its expression on GC B cells could favor the persistence and eventual positive selection of these cells. This property could dominate over loss of RIIB on FDCs, since its contribution to the retention of antigen is minimal when compared to complement receptors. Enhancement of this autoimmune phenotype would be
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provided by the observation that RIIB expression on immature dendritic cells prevents IC-mediated DC maturation and the resulting T cell activation. The analysis of conditional deficiency of RIIB on B cells, FDCs, and dendritic cells will clarify these activities.
Ig Enhancement and Suppression The ability of IgG to mediate the suppression of antibody responses is perhaps the best known clinical application of feedback suppression. The administration of IgG antierythrocyte–specific antibody completely prevents the emergence of anti-erythrocyte antibody response [30, 31] and prevents the emergence of hemolytic disease of the newborn [32]. The responsible mechanism is still not defined, although experimental support for both epitope masking and RIIB-mediated inhibition has been presented. In experimental mouse models, monoclonal IgG-anti TNP suppressed the primary response to TNP-SRBCs in mice that lacked RIIB, suggesting an RIIB-independent mechanism [33]. Other studies demonstrated decreased suppression for IgG antibodies with reduced FcR binding ability or through blocking FcR engagement by antibody blockade [34]. These differences may reflect the selective effects of decreased RIIB binding on either B cells, FDCs, or APCs where different effects can be elicited depending upon the pathway affected. Here, too, discrimination between these pathways is necessary to define the principal mechanism underlying IgG suppression. The situation for enhancement by IgG is less complex. In this case, the ability of an immune complex, in the absence of adjuvant, to augment antigen presentation by APCs has been well-documented in several systems [35]. Two activities are involved in IgG mediated enhancement—the ability of IC to be more efficiently internalized by APCs through Fc-mediated pathways leading to enhanced presentation [36, 37], and the ability of ICs to induce the maturation of immature DCs through activation of FcRIII ITAM pathways [13]. This latter activity is restricted by the preferential expression of RIIB on immature DCs. This pathway provides an explanation for the observation that IgG enhancement is augmented up to several hundred-fold in RIIB-deficient animals. In the presence of adjuvants, IgG enhancement is significantly reduced, since alternative maturation signals are provided [38], and the enhanced uptake of antigen is the primary activity being followed.
ASCs and memory cells in the spleen are reduced, although the magnitude of the antibody response remained unchanged and the affinity actually increased [27]. Notably the longterm recall response was impaired. Here, too, the likely explanation points to the role of RIIB on FDCs as providing a means of limiting the ability of retained ICs from interacting with RIIB on B cells. The unrestricted interaction of RIIB on B cells with ICs could account for the perturbation in the generation of high-affinity AFC and memory cells, as discussed above. Further studies using a conditional deficiency of RIIB on these two cell populations will be necessary to resolve these pathways. The relative importance of the RIIB pathway on FDCs, as compared to the complement pathway, is discussed below.
COMPLEMENT RECEPTORS The discovery of antibody as important in recognition of bacteria directly led to the identification of a heat-sensitive component of serum that was also required for the elimination of pathogens. The component was termed “complement” for its complementary role in leading to lysis of antibody-bound bacteria. Thus, the study of the complement system has paralleled that of antibody for more than a century. It is now appreciated that the complement system represents more than twenty serum and cell surface proteins that act in concert to activate in a regulated fashion the central component C3 and the later membrane attack complex C5-C9 [39, 40]. Whereas much of the focus of early work was on proteolytic events and assembly of the membrane attack complex, work during the past several decades has uncovered an important role of the complement system in the regulation of B cells. This portion of the chapter focuses on our current understanding of the complement receptors involved in B cell regulation in vivo. Importantly, complement receptors act at multiple steps of B cell activation and help to shape not only the antibody repertoire but influence the formation of a memory response to both T-dependent and T-independent antigens [41]. Two major affects on B cells are 1) localization of antigen to lymphoid compartment; and 2) enhancement of B cell signaling via the co-receptor. Finally, we discuss recent studies that identify a role for complement receptors in the negative selection of self-reactive B-lymphocytes.
Memory Response
Early Components of Complement Influence the Humoral Response
RIIB has been postulated to play a role in both the recall response and in the switch from antibody secreting cells to memory cells in the spleen. Experimental support for a role in these processes comes from chimeric animals in which the FDC RIIB is absent. In those cases, the frequency of both
The finding of C3 activation products bound to the surface of B lymphocytes led to the speculation that complement receptors were important in regulating the immune response [42]. Indeed, transient depletion of C3 revealed a critical role for complement in humoral immunity to both
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thymus-dependent (T-D) and -independent (T-I) antigens [43]. These observations were confirmed and extended in humans and animals bearing natural deficiencies in C3 or the classical pathway (C1q, C2, C4) and later in genetargeted mice [44]. We now appreciate that the activation of complement C3 and covalent attachment to antigen “marks” the antigen as foreign and greatly enhances its immunogenicity. The classical pathway is most important for activation and binding to T-D antigens, whereas C3 binds to carbohydrate antigens probably via the lectin or alternative pathways.
Regulation of Complement Receptors: Given the central importance of complement C3 in inflammation, it is not surprising that it is tightly regulated. Most cells express regulators of complement such as membrane cofactor (MCP), decay activating factor (DAF), and complement receptor Crry that function either to displace C3 convertase or act as co-factors in its proteolytic inactivation. CD35 also participates as a regulator of complement but is more limited in its distribution. In addition to its co-factor activity, CD35 binds activated C4 (C4b) and C3 (C3b) and degradation products of C3—iC3b and C3d. In humans, it is the major immune adhesion receptor expressed on red blood cells and functions in the clearance of immune complexes. In addition, it is found on myeloid cells (dendritic cells and macrophages), follicular dendritic cells (FDC), and B cells. CD21 is a homolog of human CD35 that binds iC3b and C3d but lacks co-factor activity. It is expressed on B cells and some T cells. In the mouse, CD35 and CD21 are expressed at a single locus (Cr2), because CD21 represents a splice product of the former [45, 46]. They appear to be co-expressed primarily on B cells and FDC; however, low levels are also found on some myeloid cells, such as mast cells and dendritic cells. Like the regulatory receptors, CD21 and CD35 share a common structural motif termed complement control protein (CCP), and they are assembled from these repeating units. For example, murine CD21 and CD35 are assembled from 15 and 21 CCP repeats, respectively. In summary, most mammalian cells bear complement receptors that function to limit the activation of C3 on autologous cells. A subset of C3 receptors, CD21 and CD35, are more limited in expression and function both in binding of i.c. and adaptive immunity.
CD21 Forms a Co-Receptor on B Cells The B cell co-receptor best described for human B cells represents a complex of three receptors—CD21, CD19, and CD81. CD21 provides the major recognition domain and binds C3d-coated antigens that lead to cross-linking of CD19 and CD81. CD19 is the primary signaling receptor
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and becomes rapidly phosphorylated on cross-linking. Co-cross linking of the co-receptor and B cell receptor (BCR) results in downstream events that amplify the signal so that less antigen is required for activation and provides a unique signal-enhancing B-cell activation [47]. Thus, the coreceptor functions to lower the threshold for activation of B cells. Although CD19 has intrinsic signaling properties separate from CD21, the binding of C3d-coated antigens appears to be specifically required for the enhancement of B-cell responses. For example, co-cross linking of CD21 and BCR leads to preferential targeting of the complex into lipid rafts that enhance signaling. In mice, CD35 also forms a co-receptor with CD19 and CD81; therefore, the two receptors are treated as one (CD21/CD35) for remaining discussion in this chapter. Thus, the CD21/CD19/CD81 coreceptor provides a critical link between the innate immune system and adaptive immunity. In summary, CD21 with CD19 and CD81 forms a coreceptor that is expressed by all mature B cell subsets. An encounter with C3d-coated antigen co-ligates the coreceptor and BCR on cognate B cells and results in a reduction in the antigen signal required for a B-cell response.
CD21/CD35 Regulate B-Cell Responses As predicted by Nussensweig and colleagues two decades earlier, complement receptors are important in regulating B-cell responses. This was first demonstrated in vivo by pre-treating mice with antibodies specific for CD21/CD35 or a soluble inhibitor of CD21 (sCR2) [48]. Mice administered blocking antibody or sCR2 prior to immunization with T-D antigens failed to respond normally, and their antibody response was impaired. The defect was in the B cell compartment, because the T cells from treated and immunized mice were primed normally to the RBC antigen [49]. These studies confirmed the in vivo importance of CD21/CD35 in enhancement of B-cell immunity. However, they did not distinguish between complement receptor expression on B cells versus FDC. Thus, uptake of C3d-coated antigen by CD21/CD35 not only cross-links the B cell co-receptor but is important for localization of antigens within the lymphoid compartment.
CO-RECEPTOR SIGNALING VERSUS ANTIGEN LOCALIZATION TO FDC The examination of B cells in vitro provided important insights into how co-cross linking of the BCR and CD21/CD19/CD81 co-receptor could lower the threshold of B-cell activation. These observations raised the general question of the relative importance of CD21/CD35 expression on B cells and FDC in vivo. Thus, does the B-cell response depend equally on both co-receptor signaling and
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antigen localization? This question was addressed by the development of chimeric mice in which the B cells expressed CD21/CD35 and FDC were deficient or vice versa. Such chimeric mice were made possible by the finding that FDCs are radio resistant and apparently not restored by engraftment of adult bone marrow (BM) into irradiated recipients. For example, chimeric mice bearing CD21/CD35 on B cells, but deficient on FDC, could be prepared by the reconstitution of lethally irradiated CD21def mice with WT (Cr2+) BM. Alternatively, reconstitution of irradiated WT mice with BM prepared from Cr2-def mice yielded mice bearing Cr2+ FDC but Cr2-def B cells. In one line of chimeric mice, expression of CD21/CD35 on B cells was determined as essential for a detectable antibody response irrespective of FDC expression [50]. However, in another line of Cr2-def mice, full humoral immunity and optimal GC response required the expression of CD21/CD35 on both B cells and FDC [51]. The differences observed are probably due to variations in conditions of immunization and degree of chimerism.
B Cells Require Complement Receptors at Five Different Stages 1. Repertoire of Natural Antibody Is Altered in Cr2-def Mice Natural antibody is produced primarily by a subset of B cells termed B-1. They are distinguished from conventional B cells (B-2) by anatomical localization, repertoire, cell surface phenotype, activation, and lifespan (this subset of B cells is discussed more thoroughly in another chapter). B-1 cells are IgM-int, IgD-lo, and CD21-int, CD23-, and localize primarily in the peritoneal tissues, although they are also found at low levels in the spleen and lymph nodes (LN). Two subsets have been identified based on expression of CD5: CD5+, termed B-1a, and CD5-, termed B-1b. Unlike conventional B cells, both subsets express CD43 and CD11b. Although B-1 cells can undergo isotype switch, they are not thought to enter GC or acquire extensive somatic mutation; however, they are long lived and appear to undergo selfrenewal. Their repertoire appears to be biased towards highly conserved antigens such as phosphoryl choline, phosphatidyl choline, and nuclear antigens such as DNA and nuclear proteins. This bias might reflect their development, because they are selected during the early neonatal period in which TdT is not expressed and V-to-DJ gene rearrangement is biased towards more proximal gene expression. Their repertoire is also influenced by the positive selection by self- and enteric bacterial antigens [52, 53]. The development of B-1 cells requires an intact BCR, as defects in proteins involved in BCR signaling such as vav, PI-3 kinase, and CD19 result in a more profound loss of B-1 relative to conventional B-2 cells [54]. Alternatively,
mutations leading to hyperresponsive BCR signaling can result in an increased frequency of B-1 cells. Whether this reflects a general requirement for intrinsic signaling of BCR or encounter with cognate antigen is not clear. However, recent studies identifying the requirement of cognate antigen for B-1 cell development suggests that interaction with self-antigens or enteric bacteria is important for the initial positive selection or expansion and maintenance. This is illustrated in the ischemia/reperfusion model (I/R). I/R represents an acute inflammatory response against self following reperfusion of ischemic tissues. It is mediated by natural IgM (and IgG) and classical pathway complement. Not only are Ig-deficient animals protected from injury in the I/R model, but Cr2-def mice are also protected from full injury [55, 56]. Thus, despite apparent normal levels of serum IgM Cr2-def mice are protected in an intestinal model of I/R. Injury can be restored by reconstitution with pooled IgM prepared from WT mice. Alternatively, reconstitution of Cr2-def mice with WT peritoneal B cells also restores their susceptibility to I/R injury. Since engrafted Cr2+ B-1 cells are maintained in Cr2-def mice in the absence of stromal expression of CD21/CD35 it seems most likely that co-receptor signaling rather than FDC binding is important in the expansion and maintenance of the B-1 subset of cells. In summary, B-1 cells are a major source of natural antibody and are positively selected during early development by cognate antigen. The interaction requires complement receptors with at least some antigens to enhance antigen receptor signaling. 2. Activation of Naïve B Cells B cells, like T cells, must be tightly regulated to circumvent the nonspecific activation of bystander cells during an ongoing infection. Antigen specificity is ensured in large part by the requirement for two signals—BCR and CD40— to promote activation and expansion against specific pathogens. B cell encounters with antigen in the absence of T cell help (CD40L co-stimulation) or vice versa can result in the induction of anergy or cell death. A wellcharacterized pathway for regulation of peripheral lymphocytes is CD95 or Fas. Trimerization of the Fas receptor on peripheral B cells stimulated by CD40 alone leads to the assembly of the caspase death pathway and B-cell apoptosis. Cross-linking of BCR induces the expression of cFLIP (Fas ligand inhibitor protein) that blocks the caspase pathway and results in cell survival and expansion. Antigen affinity is important in this regulatory step, because co-ligation of CD21 and BCR can protect, whereas cross-linking of BCR alone by moderate affinity antigen can result in cell death in vivo [57]. Thus, as predicted by Carter and Fearon, the co-receptor is important in lowering the threshold for B cell activation (Figure 18.4).
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FIGURE 18.4 Complement and Fc receptors are important in the regulation of B lymphocyte responses at multiple stages of the peripheral response. Activation of naïve B cells by engagement of antigen coupled to complement C3d results in co-ligation of BCR and co-receptor and, in the presence of T cell help, leads to activation and expansion in absence of specific IgG. Activated cells initiate a germinal center within the splenic follicles that is organized by follicular dendritic cells (FDC). Complement and Fc receptors (FcRIIB) expressed on GC B cells have opposing roles in that CD21/CD35 acts to enhance BCR signaling whereas FcRIIB participates in dampening the BCR signal in the presence of specific IgG immune complexes. By contrast, both complement and Fc R expressed on FDC act to promote the antigen selection of high-affinity GC B cells. Whereas CD21/CD35 are critical in the retention of antigen complexes, FcRIIB seems to promote B cell selection by competing for Ig Fc ligand. Post GC B cells continue to require both complement and FcRIIB in continued antigen selection, at least within the spleen, as maintenance of long-term memory B cells is dependent on both receptor systems. See color insert.
In summary, the Fas pathway regulates naïve peripheral B cells to limit bystander activation during an ongoing infection. Regulation is dependent on antigen affinity and complement receptors, because low-moderate affinity antigens require co-receptor cross-linking to prevent Fas-dependent apoptosis. Thus, engagement of the BCR and co-receptor is important for the survival and expansion of naïve B cells following an encounter with many T-dependent antigens. 3. Germinal Center Survival Germinal centers (GC) represent a specialized microenvironment within the B-cell follicles of lymphoid tissue [58]. They are transient in duration and disappear within 15 to 21 days following immunization (see chapter by Kelsoe for a more detailed discussion). Within GC, activated B cells (termed centrocytes) undergo rapid expansion, isotype switch, and somatic hypermutation followed by antigen selection. As mutated centrocytes emerge from the dark zone, they encounter C3-coated antigen retained on FDC primarily via CD21/CD35 but also FcR. Survival of GC B cells within GC is dependent on T cell help (CD40 ligand), antigen, and interaction with FDC (Figure 18.4).
Co-receptor expression is also required for the survival of B cells within GC, based on several lines of evidence. Treatment of immune mice with soluble CD21 receptor (sCR2) results in rapid loss of the GC [59]. Thus, contact between C3d-coated antigens localized on FDC is essential for the survival of GC B cells. Whether the B cells are eliminated in a Fas-or FcRIIB-dependent mechanism is not known. Further support comes from studies comparing GC survival of Cr2+ and Cr2-def immunoglobulin (Ig) transgeneic (Tg) B cells (specific for hen lysozyme). In this study, the adoptive transfer of high affinity Ig-Tg B cells into mice immunized with specific antigen identified the participation of the Cr2+ but not Cr2-def Tg B cells within the GC. Thus, expression of a high-affinity BCR is not sufficient to mediate the survival of B cells in the absence of coreceptor expression. In summary, GC represents a specialized environment within the lymphoid compartment for expansion, isotype switch, somatic hypermutation, and antigen-dependent selection of high-affinity B cells. Survival requires the presence of antigen, T cell help, and contact with FDC. Co-receptor signaling independent of antigen affinity is also required and explains at least part of the need for FDC.
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4. Memory B Cells and Persistence of Antibody Secretion A hallmark of the adaptive immune response is the formation of memory B cells. These post-GC cells are antigen selected and of relatively high affinity. A subpopulation of memory cells (Bmem) differentiate into antibody forming cells (AFC) or plasma cells that persist over the long-term, primarily in the bone marrow (BM) but also in the secondary lymphoid compartment. The role of antigen in the maintenance of AFC and Bmem is controversial. Earlier studies by Gray and colleagues demonstrated a critical role for antigen in long-term antibody secretion and recall [60]. These results, combined with the observations that antibody affinity continues to increase long after GC wanes, suggests that at least some post-GC AFC precursors are continually selected by antigen such that the higher affinity clones are preferentially maintained. More recent adoptive transfer studies using chimeric mice in which FDC are deficient in CD21/CD35 or FcRIIB provide further support that antigen is important in the maintenance of Bmem cells (these results are discussed further below) (Figure 18.4). An alternative view is that Bmem, like memory T cells, are long-lived in the absence of antigen. This view is supported by results that suggest that Bmem cells are nondividing over long periods and recent elegant genetic experiments demonstrating that switching of BCR specificity after formation of Bmem did not appear to alter their survival as functional memory cells. A current view is that antigen is important for the affinity maturation and maintenance of memory B cells. However, there are long-lived plasma cells and Bmem cells that exist in the absence of antigen. In summary, antigen is retained over long periods on FDC primarily via CD21/CD35, but FcRIIB also appears to be essential for effective recall responses. The presence of antigen is important for both the affinity maturation and efficient maintenance of memory B cells.
5. Negative Selection of Self-Reactive B Cells Systemic lupus erythematosus (SLE) is an autoimmune disease characterized by antibodies specific for nuclear antigens, such as dsDNA. Although full development of disease is due to multiple gene defects, dysregulation of B cells is a major factor. Defects resulting in excess BCR signaling, such as loss of negative regulators like FcRIIB, often lead to production of anti-dsDNA or -nuclear antibodies (ANA). For example, deficiency in the regulatory Fc receptor FcgRIIB predisposes to lupus autoantibodies, as discussed above. One interpretation of these results is that the breakdown in normal regulation of self-reactive B cells within the BM and periphery results in survival of self-reactive cells that can become activated and secrete autoantibodies in the presence of T cell help.
Multiple checkpoints have evolved to limit the development and activation of self-reactive B cells. Within the BM (central tolerance), B cells that rearrange and express selfreactive receptors at the immature stage are either edited (rearrangement of additional upstream VL genes) or eliminated by apoptosis (this topic is covered in greater detail in Chapter 16). How immature B cells encounter self-antigen is not clear. However, it is most probable that mechanisms similar to those utilized in the localization of environmental antigens within the periphery are involved. Given the importance of complement receptors CD21/CD35 in the localization and retention of antigen within the lymphoid follicles, it is likely they play a similar role in the trapping of self-antigen within the BM. Thus, complement receptors could protect from lupus by enhancing the presentation of lupus antigens to self-reactive B cells at the immature stage. Defects in the retention of self-antigen could result in a loss of negative selection and escape of self-reactive B cells to the periphery. In addition, Cr2 could participate in peripheral tolerance at the transitional stage. CD21/CD35 are first expressed at the T-1-T-2 transitional stage, in which a threshold BCR signal can result in cell death rather than activation. Co-cross-linking of BCR and co-receptor would increase the sensitivity of the transitional B cell to cognate self-antigens. The humans, CD35 receptor binds both C1q and activated C4, whereas the mouse receptor only appears to bind the latter. If CD21/CD35 participates in the uptake of C4bound lupus antigens, Cr2-def mice might be expected also to display increased production of lupus autoantibodies. Additional support for a role for CD21/CD35 receptors comes from the genetic mapping of susceptibility loci in a lupus-prone strain of mice (NZM 2410/NZW) [61]. One of the susceptibility loci, Sle-1c, includes the Cr2 locus, suggesting that CD21 or CD35 might be involved in the lupus phenotype. Structural analysis of the CD21 allele (NZM2410/NZW) identified several nucleotide differences, one of which results in the addition of a carbohydrate attachment site within the region of C3d binding. Functional studies of B cells that express the mutant allele confirmed that binding of C3d is reduced and that co-receptor activity is diminished. The findings from this study suggest that the Cr2-locus encodes the Sle-1c susceptibility gene. Whether the mutation affects C4b as well as C3d binding is not known. Much of our understanding of the negative selection of B cells comes from studies with immunoglobulin transgenic mice in which the majority of B cells express a known receptor for self-antigen. One model that is particularly well characterized is the hen lysozyme (HEL) model, in which the B cells express a conventional Ig-Heavy (Hc) and Light (Lc) chain Tg that encodes a relatively high-affinity BCR. In the absence of the antigen, the B cells appear to mature normally and are responsive to antigen both in vivo and in vitro.
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However, when the mice are crossed with a strain that expresses a soluble form of the antigen (sHEL), the B cells undergo negative selection. In the BM, immature HEL-Tg B cells undergo receptor editing and limited apoptosis in response to threshold levels of self-antigen. Self-reactive Tg B cells that escape become anergized and remain unresponsive when treated with antigen in vitro. Moreover, the anergic B cells have a reduced half-life that is reflected by a reduced frequency of CD23+ mature B cells within the lymph nodes. The role of complement and its receptors was tested in this model by crossing mice deficient in C1q, C4, C3, or CD21/CD35 with double Tg mice, that is, HEL-Ig Tg ¥ sHEL Tg. Although no effect on anergy was observed in C1q- or C3-def HEL double Tg mice [62], mice deficient in either C4 or CD21/C35 failed to develop the full anergic phenotype [63]. Characterization of splenic B cells isolated from double Tg mice that were deficient either in C4- or Cr2 revealed a phenotype similar to that of single Tg mice. Thus, the B cells remained responsive to antigen in vitro, and the frequency of mature cells within LNs was similar to that of single Tg mice. By contrast, complement-sufficient controls and C3-def double Tg mice expressed the normal anergic phenotype. The finding that C3-def mice were fully anergic was not unexpected and is consistent with observations discussed above. The results suggest that C4 functions via CD21/CD35, and they both are involved in induction of anergy in the Ig Tg model. It is not clear at what stage complement receptors are involved in triggering tolerance in the lysozyme-double Tg model. This model of autoimmunity is considered one of peripheral tolerance, since the majority of the B cells escape to the periphery where they remain unresponsive. Although, as noted above, a fraction of self-reactive B cells undergo receptor editing and apoptosis within the BM, which indicates that the self-reactive B cells encounter antigen prior to the mature stage. It is not known if CD21/CD35 deficiency alters editing or apoptosis. Since B cells do not express CD21/CD35 until the late transitional stage (splenic compartment), their role in central tolerance is probably limited to binding antigen on stromal cells. Thus, the retention of C4-coated sHEL antigen on BM stroma via CD35 could enhance encounters by developing B cells and increase the efficiency of threshold signaling by an aggregation of antigen. An alternative (but not mutually exclusive) explanation is that complement affects the HEL–self-reactive B cells at the transitional stage once they express CD21/CD35. As discussed above, this is an important checkpoint, and it could involve CD21/CD35. B cells at this stage also require BAFF ligand for survival; it is possible that BAFF receptor signaling is affected by BCR cross-linking. One possibility is that co-receptor signaling at this stage via C4-coated selfantigens enhances a negative BCR signal and results in cell death or anergy.
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FRONTIERS: COMPLEMENT VERSUS Fc RECEPTORS Complement (CR) and Fc receptors are both expressed on mature B cells but play dramatically different roles in activation: CRs enhance, whereas FcRs tune-down the BCR signal. Both receptors recognize immune complexes (IC) but the isotype of the Ig greatly affects the outcome. B cell regulation via IC formed with IgM or IgG3 seems to require complement and CD21/CD35; whereas IgG1-, 2a-, and 2bcontaining IC seem to influence B cells via RIIB. For example, the administration of T-D antigen plus specific IgM or IgG3 enhances B cell response and is CD21/CD35dependent. By contrast, the presence of IgG1 IC can suppress the response of cognate B cells. Although these two receptor systems are often activated during different stages of the humoral response, they have intrinsic competing roles in regulating the germinal center reaction. An imbalance between these positive and negative regulators of BCR can have a dramatic effect on the formation of B memory and affinity. For example, B cell deficiency in CD21/CD35 results in limited survival but higher affinity, probably due to increased selection; whereas deficiency in RIIB results in reduced selection and an expansion of lower affinity B cells. Like B cells, FDC express both CD21/CD35 and RIIB. However, in contrast to B cells, both receptor systems have a positive role in formation of Bmem cells (Figure 18.4). Commensurate with a role in antigen retention, CD21/CD35 are constitutively expressed at relatively high levels on FDC, whereas RIIB appears to be regulated and expressed during an ongoing GC reaction. How RIIB is regulated is not known, but one possibility is that expression is induced in the presence of IC. Alternatively, expression could be induced by CD21/CD35 that signals independently of CD19. Absence of RIIB on FDC, but its presence on lymphoid or myeloid cells, can have a dramatic effect on the humoral response to T-D antigens. For example, chimeric mice (deficient in FDC RIIB) immunized with T-D antigen develop increased affinity of antibody but have an impaired long-term recall response [27]. One explanation is that lack of RIIB on FDC favors Fc ligand interaction with RIIB on the B cell and increased negative selection. Thus, in the absence of FDC expression of RIIB, antigen selection would become more intense and limit the frequency of surviving B cells such that only those having higher affinity survive. The presence of CD21/CD35 on FDC contributes to B cell survival by providing ligand (C3d-coated antigen) for co-ligation of BCR and co-receptor signaling. This would be important in countering the negative signal via B cell RIIB. Saturation of the lymphoid compartment during an ongoing immune response with either specific antigen or sCR2 induces a rapid elimination of the GC, as discussed
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earlier. One explanation for the elimination of B cells is RIIB signaling. Thus, a disruption of positive signaling via co-receptor and/or BCR would favor negative signaling by RIIB and elimination of the cognate B cells. It will be important to test this hypothesis in mice deficient in FcgRIIB. It is notable that both CR and FcR are protective against SLE despite their opposite roles in B-cell regulation. As discussed earlier, mice deficient in RIIB or CD21/CD35 have an increased propensity to develop lupus autoantibodies. Although both receptor systems bind IC (or IC-C3d) and participate in clearance, a more likely role for protection against lupus is via B-cell regulation. As discussed above, deficiency in either receptor can have a dramatic affect on the fate of the B cell in response to environmental antigens. Therefore, it is not unlikely that both sets of receptors participate in the regulation of self-reactive B cells. However, given the positive and negative effects of the receptors on mature B cells it seems likely that they act at different stages of B-cell development such that they both have a protective role. For example, CD21/CD35 could limit the activation of self-reactive B cells at the immature stage, where antigen encounters result in editing, cell death, or anergy. Several possible roles discussed earlier are the retention of self-antigens within the BM or co-receptor signaling in transitional B cells in the periphery. Further study in both these areas is needed. By contrast, RIIB is more likely involved in the regulation of mature B cells in the periphery. Given the nature of its ligand and the phenotype of RIIB-deficient mice, its protective role would be most prominent in the GC. An important checkpoint occurs within the GC to prevent the activation of B cells that become self-reactive as a result of somatic hypermutation. Deficiency in RIIB lowers antigen selection pressure and could allow self-reactive B cells to expand and escape the GC. In summary, in vivo studies for mice deficient in CRs and FcRs has greatly extended our understanding of B-cell regulation and how ligands for these receptor systems can modulate responses to environmental and self-antigens. However, these studies also highlight how much is left to learn. In this chapter, we have attempted to outline some of the important questions that have emerged over the past 5 years. It is clear that a better understanding of how CRs and FcRs participate in the humoral response to environmental antigens will be useful not only in future vaccine research but in helping to dissect the events leading to dysregulation of self-reactive B cells and the development of autoimmune disorders such as SLE.
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3. Takai, T., et al. (1996). Augmented humoral and anaphylactic responses in Fc-gamma-RII-deficient mice. Nature 379(6563), 346–349. 4. Bolland, S., and Ravetch, J. V. (2000). Spontaneous autoimmune disease in Fc(gamma)RIIB-deficient mice results from strain-specific epistasis. Immunity 13(2), 277–285. 5. Nakamura, A., et al. (2000). Fcg receptor IIB-deficient mice develop Goodpasture’s syndrome upon immunization with type IV collagen: A novel murine model for autoimmune glomerular basement membrane disease. J Exp Med 191(5), 899–906. 6. Yuasa, T., et al. (1999). Deletion of Fcgamma receptor IIB renders H-2(b) mice susceptible to collagen-induced arthritis. J Exp Med 189(1), 187–194. 7. Pritchard, N. R., et al. (2000). Autoimmune-prone mice share a promoter haplotype associated with reduced expression and function of the Fc receptor FcgammaRII. Curr Biol 10(4), 227–230. 8. Jiang, Y., et al. (2000). Polymorphisms in IgG Fc receptor IIB regulatory regions associated with autoimmune susceptibility. Immunogenetics 51(6), 429–435. 9. Wakeland, E. K., et al. (2001). Delineating the genetic basis of systemic lupus erythematosus. Immunity 15(3), 397–408. 10. Carter, R. H., and Fearon, D. T. (1992). CD19: Lowering the threshold for antigen receptor stimulation of B lymphocytes. Science 256(5053), 105–107. 11. Rao, S. P., Vora, K. A., and Manser, T. (2002). Differential expression of the inhibitory IgG Fc receptor FcgammaRIIB on germinal center cells: Implications for selection of high-affinity B cells. J Immunol 169(4), 1859–1868. 12. de Andrés, B., et al. (1997). FcgRII (CD32) is linked to apoptotic pathways in murine granulocyte precursors and mature eosinophils. Blood 90(3), 1267–1274. 13. Kalergis, A. M., and Ravetch, J. V. (2002). Inducing tumor immunity through the selective engagement of activating Fcgamma receptors on dendritic cells. J Exp Med 195(12), 1653–1659. 14. Amigorena, S., et al. (1992). Cytoplasmic domain heterogeneity and functions of IgG Fc receptors in B lymphocytes. Science 256(5065), 1808–1812. 15. Muta, T., et al. (1994). A 13-amino-acid motif in the cytoplasmic domain of FcgRIIB modulates B-cell receptor signalling. Nature 369(6478), 340. 16. Ono, M., et al. (1996). Role of the inositol phosphatase SHIP in negative regulation of the immune system by the receptor Fc(gamma)RIIB. Nature 383(6597), 263–266. 17. Tridandapani, S., et al. (1999). Protein interactions of Src homology 2 (SH2) domain-containing inositol phosphatase (SHIP): Association with Shc displaces SHIP from FcgRIIb in B cells. J Immunol 162(3), 1408–1414. 18. Daeron, M., et al. (1995). The same tyrosine-based inhibition motif, in the intracytoplasmic domain of Fc gamma RIIB, regulates negatively BCR-, TCR-, and FcR-dependent cell activation. Immunity 3(5), 635–646. 19. Bolland, S., et al. (1998). SHIP modulates immune receptor responses by regulating membrane association of Btk. Immunity 8(4), 509–516. 20. Tamir, I., et al. (2000). The RasGAP-binding protein p62dok is a mediator of inhibitory FcgRIIB signals in B cells. Immunity 12(3), 347–358. 21. Yamanashi, Y., et al. (2000). Role of the rasGAP-associated docking protein p62(dok) in negative regulation of B cell receptor-mediated signaling. Genes Dev 14(1), 11–16. 22. Aman, M. J., et al. (1998). The inositol phosphatase SHIP inhibits Akt/PKB activation in B cells. J Biol Chem 273(51), 33922–33928. 23. Liu, Q., et al. (1999). SHIP is a negative regulator of growth factor receptor-mediated PKB/Akt activation and myeloid cell survival. Genes Dev 13(7), 786–791. 24. Pearse, R. N., et al. (1999). SHIP recruitment attenuates FcgRIIBinduced B cell apoptosis. Immunity 10(6), 753–760.
18. Fc and Complement Receptors 25. MacLennan, I. C. (1994). Germinal centers. Annu Rev Immunol 12, 117–139. 26. Tew, J. G., et al. (1997). Follicular dendritic cells and presentation of antigen and costimulatory signals to B cells. Immunol Rev 156, 39–52. 27. Barrington, R. A., et al. (2002). B lymphocyte memory: Role of stromal cell complement and FcgammaRIIB receptors. J Exp Med 196(9), 1189–1199. 28. Barrington, R., et al. (2001). The role of complement in inflammation and adaptive immunity. Immunol Rev 180, 5–15. 29. Tew, J. G., et al. (2001). Follicular dendritic cells: Beyond the necessity of T-cell help. Trends Immunol 22(7), 361–367. 30. Bruggemann, M., and Rajewsky, K. (1982). Regulation of the antibody response against hapten-coupled erythrocytes by monoclonal antihapten antibodies of various isotypes. Cell Immunol 71(2), 365–373. 31. Heyman, B., and Wigzell, H. (1984). Immunoregulation by monoclonal sheep erythrocyte-specific IgG antibodies: suppression is correlated to level of antigen binding and not to isotype. J Immunol 132(3), 1136–1143. 32. Bowman, J. M. (1988). The prevention of Rh immunization. Transfus Med Rev 2(3), 129–150. 33. Karlsson, M. C. I., Getahun, A., and Heyman, B. (2001). FcgRIIB in IgG-mediated suppression of antibody responses: Different impact in vivo and in vitro. J Immunol 167(10), 5558–5564. 34. Kumpel, B. M. (2002). In vivo studies of monoclonal anti-D and the mechanism of immune suppression. Transfus Clin Biol 9(1), 9–14. 35. Heyman, B. (2000). Regulation of antibody responses via antibodies, complement and Fc-receptors. Annu Rev Immunol 18, 709–737. 36. Regnault, A., et al. (1999). Fcg receptor-mediated induction of dendritic cell maturation and major histocompatibility complex class I-restricted antigen presentation after immune complex internalization. J Exp Med 189(2), 371–380. 37. Wernersson, S., et al. (1999). IgG-mediated enhancement of Ab responses is low in FcRg chain deficient mice and increased in FcgRII deficient mice. J Immunol 163, 618–622. 38. Banchereau, J., and Steinman, R. M. (1998). Dendritic cells and the control of immunity. Nature 392(6673), 245–252. 39. Reid, K. B., and Porter, R. R. (1981). The proteolytic activation systems of complement. Annu Rev Biochem 50, 433–464. 40. Muller-Eberhard, H. J. (1998). Molecular organization and function of the complement system. Annu Rev Biochem 57, 321–347. 41. Fearon, D. T., and Carroll, M. C. (2000). Regulation of B lymphocyte responses to foreign and self-antigens by the CD19/CD21 complex. Annu Rev Immunol 18, 393–422. 42. Nussenzweig, V., et al. (1971). Receptors for C3 on B lymphocytes: Possible role in the immune response. In Progress in immunology, B. Amos, ed. (New York: Academic Press), p. 73. 43. Pepys, M. B. (1972). Role of complement in induction of the allergic response. Nat New Biol 237(74), 157–159. 44. Carroll, M. C. (2000). The role of complement in B cell activation and tolerance. Adv Immunol 74, 61–88. 45. Kurtz, C. B., et al. (1990). The murine complement receptor gene family. IV. Alternative splicing of Cr2 gene transcripts predicts two
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distinct gene products that share homologous domains with both human CR2 and CR1. J Immunol 144(9), 3581–3591. Molina, H., et al. (1990). A molecular and immunochemical characterization of mouse CR2. Evidence for a single gene model of mouse complement receptors 1 and 2. J Immunol 145(9), 2974–2983. Carter, R. H., and Fearon, D. T. (1992). CD19: Lowering the threshold for antigen receptor stimulation of B lymphocytes. Science 256(5053), 105–107. Hebell, T., Ahearn, J. M., and Fearon, D. T. (1991). Suppression of the immune response by a soluble complement receptor of B lymphocytes. Science 254(5028), 102–105. Gustavsson, S., Kinoshita, T., and Heyman, B. (1995). Antibodies to murine complement receptor 1 and 2 can inhibit the antibody response in vivo without inhibiting T helper cell induction. J Immunol 154(12), 6524–6528. Ahearn, J. M., et al. (1996). Disruption of the Cr2 locus results in a reduction in B-1a cells and in an impaired B cell response to T-dependent antigen. Immunity 4(3), 251–262. Fang, Y., et al. (1998). Expression of complement receptors 1 and 2 on follicular dendritic cells is necessary for the generation of a strong antigen-specific IgG response. J Immunol 160(11), 5273–5279. Hardy, R. R., and Hayakawa, K. (2001). B cell development pathways. Annu Rev Immunol 19, 595–621. Herzenberg, L. A. (2000). B-1 cells: the lineage question revisited. Immunol Rev 175, 9–22. Cariappa, A., and Pillai, S. (2002). Antigen-dependent B-cell development. Curr Opin Immunol 14(2), 241–249. Fleming, S. D., et al. (2002). Mice deficient in complement receptors 1 and 2 lack a tissue injury-inducing subset of the natural antibody repertoire. J Immunol 169(4), 2126–2133. Reid, R., et al. (2002). Functional activity of natural antibody is altered in Cr2null mice. J Immunol 169(10), 5433–5440. Fischer, M. B., et al. (1998). Dependence of germinal center B cells on expression of CD21/CD35 for survival. Science 280(5363), 582–585. MacLennan, I. C. (1994). Germinal centers. Annu Rev Immunol 12, 117–139. Fischer, M. B., et al. (1998). Local synthesis of C3 within the splenic lymphoid compartment can reconstitute the impaired immune response in C3-deficient mice. J Immunol 160(6), 2619–2625. Gray, D. (2002). A role for antigen in the maintenance of immunological memory. Nat Rev Immunol 2(1), 60–65. Boackle, S. A., et al. (2001). Cr2, a candidate gene in the murine Sle1c lupus susceptibility locus, encodes a dysfunctional protein. Immunity 15(5), 775–785. Cutler, A. J., et al. (2001). Intact B cell tolerance in the absence of the first component of the classical complement pathway. Eur J Immunol 31(7), 2087–2093. Prodeus, A. P., et al. (1998). A critical role for complement in maintenance of self-tolerance. Immunity 9(5), 721–731.
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19 Regulation of Class Switch Recombination MICHEL COGNÉ
BARBARA K. BIRSHTEIN
Laboratoire Immunologie, Faculté de Medecine, Limoges, France
Department of Cell Biology, Albert Einstein College of Medicine, Bronx, New York, USA
B lineage cells undergo two main waves of remodeling of their immunoglobulin loci. The first, V(D)J joining, occurs early in B-cell maturation in the bone marrow and results in the expression of an unmutated IgM class BCR on naïve B cells. These cells subsequently migrate to the periphery, where they later express IgM plus IgD through the alternative splicing of a single primary transcript. The second wave of remodeling, class-switch recombination (CSR), is mostly restricted to mature B cells and is triggered by specific antigen binding. This allows expression of antigen-specific heavy chains with new effector functions. Low-level CSR has been reported in fetal liver B cell progenitors (Millili et al., 1991) and may account for the generation of IgA expressing cells in knockout mice that cannot express the IgM-class BCR (Macpherson et al., 2001). CSR also occurs to some extent during responses to T cell–independent (TI) antigens that take place in the splenic marginal zone outside germinal centers (GC) (Snapper et al., 1993). However, most frequently, T cell–dependent CSR occurs concomitantly with clonal expansion in the specialized B-cell proliferation areas of peripheral lymphoid tissues that define GC. CSR appears to be regulated by the nature of the BCR-ligand, the type of T cell help, and the cytokine milieu (Stavnezer et al., 1996; Snapper et al., 1997). Some antigens and some anatomical locations promote isotype-restricted responses (e.g., mucosalassociated lymphoid tissues for IgA in mouse and human) (Garraud et al., 1995; Cavanagh et al., 2001). Germline transcription (GT) always precedes CSR to a given CH gene in the same activated cells, and the induction of CSR by specific stimuli correlates directly with their ability to induce or suppress specific GT before CSR occurs (Stavnezer, 2000). GT is detectable 4 days after primary challenge with a T-dependent antigen and increases in parallel to GC formation. As early as 12 hours after a secondary
antigen challenge, GT is detectable in B cells located in the T cell zone of lymphoid tissues and may precede the first cell division and the entry of B cells into GC or extrafollicular foci (Toellner et al., 1996). Gene-targeting studies definitively demonstrated that transcription from I promoters is a necessary prerequisite to CSR (Jung et al., 1993; Zhang et al., 1993; Bottaro et al., 1994; Harriman et al., 1996). Subsequent to the GT of IgH constant genes, there is imprecise recombination between repetitive switch (S) regions. As a targeted process, CSR tends to involve the same constant gene on both the productive and nonproductive alleles of a given B cell (Rothman et al., 1989). CSR often occurs in parallel with the generation of higher affinity antigen receptors through somatic hypermutation (SHM). Although both CSR and SHM rely on the newly discovered activation-induced deaminase (AID) and depend upon transcription of Ig genes, these are independent processes. Regulation of CSR is clearly effected at two levels: cis-acting elements that make S regions accessible to recombination in a process tightly dependent on GT, and trans-acting recombinase activities that are specifically induced in activated B cells and require AID. This review focuses on the regulation of GT and the accessibility of S regions to recombination.
Molecular Biology of B Cells
CSR REQUIRES SPECIFIC STIMULI OCCURRING IN A DEFINED GERMINAL CENTER (GC) MICROENVIRONMENT BCR Signaling to Trigger CSR Various activation pathways are connected to the transcription factors that modulate GT and CSR in B cells (see
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Figure 19.1). For example, B-cell activators, including Bcell antigen receptor cross-linkers, LPS through its interaction with CD14 and the Toll-like receptors, and CD40 agonists, induce the release and nuclear translocation of NFkB/Rel proteins. BCR signaling (extensively reviewed elsewhere by Kurosaki, 2002; and in Michael Reth’s chapter in this book), drives the entry of B lymphocytes into the cell cycle through multiple signaling pathways, resulting in the activation of IKKa, NF-kB, and AP-1. Triggering of the BCR also putatively promotes CSR through transient association of the BCR with SWAP-70, which translocates to the nucleus upon stimulation (Masat el al., 2000). In contrast, BCR-driven cell proliferation may delay the induction of CSR by T-cell dependent stimuli (Rush et al., 2002).
Cognate Interactions with T Cells CSR is especially influenced by the interaction of membrane CD40 with T cells. CD40 is a member of the TNF receptor family and is connected to the activation of NF-kB, Elf-1, and AP-1 through TRAF and MAPK/JNK pathways. In the absence of BCR cross-linking, another member of the TNFR family, CD30, is upregulated and provides a negative feedback to counteract the CD40-mediated activation of CSR in bystander B cells (Cerutti et al., 1998). Genetic defects that result in the absence of CD40L, CD40, or downstream elements of the CD40 signaling pathways, such as NEMO, have been implicated in the etiology of immunodeficiencies, such as hyper-IgM syndrome (reviewed in Durandy et al., 2001). These defects are characterized by the absence of GC and by major defects in CSR and antibody affinity maturation. The development of GC and the splenic marginal zone— environments that are conducive to CSR—is also dependent on other lymphokines of the TNF and lymphotoxin families and/or of their receptors. LTNalpha or LTNbeta knockout mice lack follicular dendritic cells and CD11c+ dendritic cells (Wang et al., 2001). In addition to lymphokines, several chemokines are required for the formation of GC. These include stromal cell-derived factor (SDF)-1a, macrophage inflammatory protein (MIP)-3a, MIP-3b, and B-cell– attracting chemokine (BCA)-1, which have been shown to sequentially direct the migration of B cells to the T cell zone and to GC after BCR ligation (Casamayor et al., 2002). B7 family co-stimulatory molecules are also involved at various levels in the control of CSR. Mice deficient in both B7–1 and B7–2, which are ligands of CD28, lacked GC and failed to generate antigen-specific IgG1 and IgG2a (Borriello et al., 1997). Another member of the B7 family, B7RP-1, specifically interacts with the inducible co-stimulator ICOS, which has homology to CD28 and is expressed on activated T cells. ICOS is essential for normal antibody responses to TD antigens, and its absence impairs both GC formation and CSR (Tafuri et al., 2001). In addition to its direct effect, CD40
activation indirectly stimulates CSR by upregulating the expression of B7h, another ICOS ligand expressed on B cells (Liang et al., 2002). The knockout of OCA-B/OBF-1 results in a severe deficiency in the production of all secondary immunoglobulin isotypes (IgG1, IgG2a, IgG2b, IgG3, IgA, and IgE). This is not due to a failure of CSR, but rather to a defect in the formation of GC and to reduced levels of transcription from switched immunoglobulin genes (Kim et al., 1996). Similarly, mice lacking the Ets protein Spi-B show a defect in GC formation and consequently display impaired IgG responses to T-dependent antigens (Su et al., 1999).
Additional Cellular Interactions with NK and Dendritic Cells CSR can occur independently of T cell interactions. Upon interaction with NK cells, either in the presence or absence of IFN-g, B cells induce GT of Cg2a together with AID transcription and are thus committed to IgG2a switching (Gao et al., 2001). CSR can also be induced in B cells in a CD40independent manner through interactions with activated dendritic cells that express high levels of the TNF family members BLyS and APRIL. CSR to Cg, Ce, and Ca genes can then occur in the presence of IL-10, TGF-b, or IL-4 (Litinskiy et al., 2002). Further secretion of class-switched antibodies requires additional BCR cross-linking and IL-15. CSR responses to TI antigens may be associated with specific signal transduction pathways. For example, the adapter molecule Gab1, which couples BCR signals to the PI-3 kinase/Akt pathway, has been recently shown to be an inhibitor of IgM and IgG1 responses to TI-2 antigens (Itoh et al., 2002). Another negative regulatory protein, the Src homology 2 domain–containing inositol-5-phosphatase (SHIP), negatively controls IgG1, IgG2a, and IgG3 responses to TI-2 antigens (Helgason et al., 2000).
PROXIMAL CIS REGULATORY ELEMENTS FOR GT Role of I Exon Transcription in CSR Various data indicate that CSR needs transcription of the S regions. All germline CH genes, except Cd, are organized such that the target S region supporting recombination is included within the first large intron of a germline transcription unit. The S region is preceded by a single noncoding exon (the I exon), and GT initiates (often heterogeneously) from a promoter 5¢ of the I exon. Deletion of the I promoter abolishes switching to the downstream S region. In some instances, constitutively transcribed drugresistance cassettes (e.g., the pgk-neor cassette) proved capable of replacing I-region promoters in directing CSR to
Feuil1
DNA-binding proteins and co-factors regulating transcription of the rearranged IgH locus
VDJ
Cm Cd
Cg3
Cg1
Cg2b
Cg2a
Ce
Ca
NF-mNR
MAR
SAT B1
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m E1
NF-mE1 (YY-1) xY
cttcctta
TFIID, TBP
TATA box
G-rich binding protein
G-rich sequence
Pax5/BSAP/SaBP
Pax5 site
Oct1/Oct2A/Oct2B
octamer / heptamer
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kB
NF-kB/p50, p52, cRel, Rel A, Rel B
=> CBP/P300
=> CBP/P300
mB, PU-box, NF-aP
PU.1, NF-aP
p (mA)
Ets-1 Elf-1 + C-fos + junB
Ets-AP1
NFE (Ets family) mE3/kE3
TFE3, USF, TFEB (bHLH-ZIP)
mE2 ,mE4, m E5/kE2
E2A products (+) / ZEB (-)/ Id control STAT1, IRF-9, IRF-1, IRF-8, IRF-4 (Pip) NF-IL6 (C/EBPb) (+) / IgEBP (-)
=> CBP/P300
O
ISRE E; C/EBP site
NF-Sm
Sm repeat
LR1 (LPS inducible)
GNCNAGGCTGAR
O
LSF ?
O
=> HDAC
=> HDAC
CACCC box
Smad3, Smad4 (+) / Smad7 (-)
TGF-b inducible element
RUNX (AML, PEBP2a)
TGF-b inducible element
ATF2
O => CBP/P300
ATF/CREB consensus
STAT6 (+) / bcl6 (-)
IL4-RE
c-fos containing complex
TGF-b inducible element
c-fos/c-jun
=> HAT => CBP/P300
AP-1 AP-2 AP-3 AP-4
Maf-p45 (NF-E2)(+) / MafK-Bach2 (-)
=>AF
MARE
: stimulatory effect
O
: inhibitory effect : positive or negative effect (modulated by additional factors) : putative site or effect non demonstrated
FIGURE 19.1 A map of the IgH locus together with the most relevant cis- and trans-acting elements. Factors potentially behaving as architectural factors (AF), or interacting with chromatin remodeling enzymes, histone acetyl transferase (HAT) and histone deacetylase (HDAC), or with the integrating factor CBP/P300, are indicated. Vertical bars at the right of binding sites indicate factors potentially forming ternary complexes. (References mentioned in the figure: 1, Scheuermann and Chen 1989; 2, Dickinson et al., 1992; 3, Herrscher et al., 1995; 4, Park and Atchison, 1991; 5, Atchison et al., 1990; 6, Mason et al., 1985; 7, Sign et al., 1996; 8, Michaelson et al., 1996; 9, Neurath et al., 1993; 10, Neurath et al., 1994; 11, Liao et al. 1994; 12, Max et al., 1995; 13, Qiu et al., 1998; 14, Gerondakis et al., 1991; 15, Stutz et al., 1999; 16, Wuerffel et al., 2001; 17, Nelsen et al., 1993; 18, Eisenbeis et al., 1995; 19, Linderson et al., 2001; 20, Rivera et al., 1993; 21, Grant et al., 1995; 22, Linderson et al., 1997; 23, Rao et al., 1997; 24, Goldfarb et al., 1996; 25, Ernst and Smale, 1995; 26, Xu and Rothman, 1994; 27, Ezernieks et al., 28, 1996; Brass et al., 1996; 29, Mao et al., 2001; 30, Wuerffel et al., 1990; 31, Williams and Maizels, 1991; 32, Drouin et al., 2002; 33, Lin et al., 1992; 34, Hanai et al., 1999; 35, Kawasaki et al., 2000; 36, Harris et al., 1999; 37, Xu and Stavnezer, 1992; 38, Zhang and Derynck, 2000; 39, Madisen and Groudine, 1994; 40, Michaelson et al., 1995; 41, Muto et al., 1998.)
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hs4
hs3B
hs1,2
3'a-88
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Se
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5' Ig2b
5' Ie
Ia Sa hs3A hs1,2 hs3B hs4
5' Ig2a
Ig2b Sg2b Ig2a Sg2a Ie Se
Sg2b
Ig1 Sg1
Sg1
5' Ig3
Sg3
Ig3 Sg3
Sm
Em enh / MAR
motif
VH promoter
PVH Em Sm
5' Ig1
MAR
1 2 3 4 5 6 8 8-13 6 8, 14-16 17-19 20 21 22 23 16, 24, 25 18, 26-28 15, 29 30 31 32 33 33 34 29, 33, 35 36 37 29,38 39, 40 37 39, 40 =>AF 41
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the downstream S region, provided their transcriptional orientation was the same as that of the I exon. However, replacement of the Ie promoter by a strong constitutively active Em-VH promoter cassette resulted only in a modest increase of constitutive switching together with a loss of cytokine-inducible Ce switching (Bottaro et al., 1994). Hence, although transcription through the S region plays a primary role in targeting CSR, it alone is not sufficient. Several studies have suggested a direct mechanistic role of S region RNA in the process of CSR through the formation of triplex structures and the destabilization of doublestranded DNA. Such RNA/DNA hybrids could lead to the formation of R loops (Tian and Alt, 2000) and/or of secondary structures that could further behave as recombination substrates. S region transcripts have also been postulated to serve as a matrix for a reverse transcription process primed onto free DNA 3¢ ends after single-strand breaks (Muller et al., 1998). Using CSR artificial substrates, the orientation of the S region with regard to transcription has been found important in one model but did not influence recombination in another case (Daniels and Lieber, 1995; Kinoshita et al., 1998). Finally, the role of S regions themselves in GT and CSR was recently challenged by the finding that deletion of most of the Sm region only partially affected CSR and promoted recombination events immediately upstream of the deletion (Luby et al., 2001). I exons by themselves are unlikely to play a specific structural role in CSR because they are weakly homologous and are not conserved between species. That they can be replaced with non-Ig sequences without altering CSR makes it unlikely that their short open reading frames encode any functional peptide. The only significant structural feature of the I exon may be its splice donor site, which allows the excision of the first intron as a lariat including S sequences when germline transcripts are processed and polyadenylated. Indeed, replacements or deletions of the I exon splice site upstream of a Sg region profoundly inhibited CSR (Lorenz et al., 1996; Heins et al., 1998). In other instances where replacement of the complete I exon with a drug-resistance cassette allowed recombination to occur, the resistance gene appeared to be spliced to the downstream CH gene. Alternatively, the cleavage of transcripts through polyadenylation could have replaced cleavage by the spliceosome (Qiu et al., 1999; Bottaro et al., 1994; Harriman et al., 1996).
Switch Sequences as Transcriptional Stimulatory Elements The Sm region has been postulated to include some transcriptional stimulatory activity in transgenes (Sigurdardottir et al., 1995). The Sg1 sequence has also been identified as a transcriptional enhancer for Ig1 transcripts (Cunningham et al., 1998). Among the various DNA-binding proteins that have been characterized with an affinity for S regions, some
potentially have regulatory activities with regard to transcription or chromatin remodeling. LR1 is a B cell–specific DNA binding complex, made up of nucleolin and hnRNP D, which binds the minor groove of DNA and has multiple sites in all switch regions and within Em. LR1 is overexpressed in activated B cells and may act both as a transcriptional activator and as a DNA bending factor, altering DNA structure and favoring recombination (Hanahaki and Maizels N, NR 2000). The DNA-binding protein, late SV40 factor (LSF), binds both Sm and Sa sequences and the histone deacetylase (HDAC) machinery, thus potentially limiting accessibility of switch regions and thereby inhibiting CSR (Drouin et al., 2002). In addition to a transcriptional stimulatory role, factors binding S regions may directly promote recombination through the formation of DNA–multiprotein complexes. For example, proteins binding the Sg3 region, including homodimers of NF-kB p50, proved necessary for CSR to Cg3 but not for GT (Wuerffel, 2001). Such protein complexes may be induced in activated cells, which then compete with factors such as Ikaros that normally recruit HDAC and limit the accessibility of Sg3 to recombination in resting cells.
Im GT and the Role of the Intronic Em Promoter/Enhancer Efficient in vivo germline CH transcription requires interactions between germline promoters and various transcriptional enhancers of the IgH locus. The B cell–specific enhancer Em, located between JH and the Cm gene, appears as a primary candidate for such interactions since it virtually includes the Im promoter region. Beside its major role in the induction of complete VDJ rearrangement, its role in CSR accessibility is restricted to the Cm gene. In the absence of Em, CSR-related deletions of downstream S regions are preserved. However, CSR occurs at a reduced efficiency, due to a decreased accessibility of the Sm region (Gu et al., 1993; Bottaro et al., 1998). Data concerning the role of the Im exon splicing are still incomplete, since deletion of the 3¢ donor site resulted in the activation of downstream cryptic splice sites that conserved the structure of the Im-Cm transcript and intron (Kuzin et al., 2000). That Em solely controls accessibility of the Sm region to CSR suggests that other regulatory elements regulate accessibility of the various downstream S regions.
Regulatory Elements Carried by the Various Non-Im Promoters and Their Inducibility by Cytokines Studies of reporter genes controlled by germline CH promoters have shown their transcriptional regulation by membrane receptors and/or lymphokines. Signal-transduc-
19. Regulation of Class Switch Recombination
tion pathways connected to surface cytokine receptors, to the BCR, and to co-stimulatory receptors proved directly responsible for the binding of specific transcription factors to the I-region promoters (reviewed in Stavnezer, 2000). Models used for the in vitro induction of class switching have also helped define conditions by which polyclonal B cell activators may direct GT and CSR to a restricted set of CH genes. For example, interactions with the T cell–bound CD40 ligand can be mimicked with anti-CD40 antibodies or CD40L-transfected fibroblasts. In addition, B cells can be activated with bacterial lipopolysaccharide (LPS) in the presence of defined mixtures of cytokines that may promote the GT of given CH genes and reciprocally inhibit the GT of other CH genes. Regulation of Cg3 and Cg2b GT The activation of murine B cells with LPS induces GT and CSR of both Cg3 and Cg2b. The Ig3 promoter carries a binding site for NF-kB together with a PU-box that may account for its LPS inducibility and for the production of IgG3 during humoral responses to TI antigens (Gerondakis et al., 1991). Like the Ig3 element, the Ig2b promoter is also responsive to BCR cross-linking and to CD40 ligation. Various motifs are found within the Ig2b sequence, including four Ets-1 sites, three C/EBP sites, and two AP-1 sites (Laurencikiene et al., 2001). During in vitro B-cell stimulation by LPS, the addition of IL-4 suppresses the Cg2b / Cg3transcription while promoting CSR to Cg1 and Ce. The molecular mechanisms that inhibit Cg3 GT in the presence of IL-4 are still unclear and may involve either the binding of repressive factors or a promoter competition mechanism with other GT promoters (namely Ie and Ig1) that are stimulated in such conditions. In contrast to Cg3, CSR to Cg2b does occur following CD40 stimulated GT (Strom et al., 1999). Regulation of Cg1 and Ce GT The stimulation of B cells in vitro with LPS plus IL-4 specifically promotes transcription and CSR to Cg1 and Ce. Both Th2 cytokines IL-4 and IL-13 bind to a membrane receptor, which includes the common g chain (gc), and activate Janus kinases JAK1 and JAK3. These kinases, in turn, mediate phosphorylation of the insulin receptor substrate proteins (IRS1 and IRS2) and the signal transducer and activator of transcription factor (STAT6). IRS proteins activate the phosphoinositide 3-kinase (PI 3-kinase) pathway and regulate cell proliferation in response to IL-4, while the phosphorylation of STAT6 allows its translocation to the nucleus (reviewed in Nelms et al., 1999). Activation by IL4 is also controlled by inhibitory pathways. For example, CD45 triggering stimulates its phosphatase activity on substrates including JAK1, JAK3, and STAT6, thus resulting in
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inhibition of the IL-4/CD40L-induced Ie GT and of subsequent CSR to Ce (Yamada et al., 2002). The mouse Ie promoter is induced by the Th2-cytokines IL-4 and IL-13. Both human and mouse Ie promoters have been shown to carry sites for PU-1 and for NF-kB, whose simultaneous mutation abolishes IL4 induction. Indeed, either PU-1 or NF-kB displays synergistic interactions with the STAT6 factor when B cells are stimulated by IL-4 (Stütz et al., 1999). Ie induction by STAT6 also needs the binding of a nearby site by AP-1, which is transiently expressed in cells activated by CD40L, LPS, or BCR cross-linking, and which may lead to chromatin remodeling through recruitment of p300/CBP (Mao and Stavnezer, 2001; Zhang and Derynck, 2000). BCL-6 appears to negatively regulate STAT6-dependent IL-4 responses, including GT and CSR to Ce, by competing with STAT6 for binding to the STAT6 site. CSR to Ce appears to be stimulated in BCL6-/- mice both in vivo and in vitro upon IL4 stimulation and is dependent on STAT6 signaling (Harris et al., 1999). The function of C/EBPb (NF-IL6) in either amplifying or inhibiting Ie GT is controversial (Stutz et al., 1999; Mao and Stavnezer, 2001). Similar to Ie, the mouse Ig1 promoter includes binding sites for STAT6, C/EBP, and PU-1 plus four CACCC boxes, a TGF b inhibitory element (TIE), an interferon response element, and an AP-3 site (Xu and Stvanezer, 1992). The binding affinity of STAT6 was ten-fold lower to the g1 promoter than to the e promoter, making Ig1 both less inducible and less dependent on IL-4 stimulation than Ie (Mao and Stavezer, 2001). Instead of an AP-1 site in the Ie element, the Ig1 STAT6 site is flanked with a site for activation transcription factor ATF2, which features direct HAT activity (Mao and Stavezer, 2001; Kawasaki et al., 2000). C/EBP may not directly bind the C/EBP site in the murine Ig1 and Ie promoter, but rather inhibits GT through negative interactions with NF-kB (Mao and Stavnezer, 2001). Strikingly, mice with a targeted deletion of C/EBP b had an increased number of surface IgG1 positive B cells (Screpanti et al., 1995). Regulation of Cg2a GT Most I promoters depend on regulation by interferons. IFNg specifically induces the Ig2a promoter (and to a lesser extent, the Ig3 promoter). In contrast, IFN-g and IFN-a inhibit GT of the Ie promoter (Xu et al., 1994; Ezernieks et al., 1996). Interferon responsive elements within the Ig2a promoter may account for the induction of Cg2a GT by IFNg both in vitro and in vivo. The IFN-g induction of GT involves JAK1 and STAT1 and a transcription factor specific for Th1 commitment, T-bet (Szabo et al., 2000). T-bet is also active in B cells and behaves as a selective transducer of IFN-g-mediated IgG2a class switching (Peng et al., 2002). Among the various transcription factors that regulate inter-
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feron-induced genes, some members of the IRF family are especially important in lymphoid cells, such as IRF-1, IRF8 (ICSBP), and IRF-4 (Pip). IRF4/Pip is involved in synergistic interactions with PU.1 and E47 (Brass et al., 1999; Nagulapalli and Atchison, 1998). IRF4/Pip also upregulates CD23 expression in activated B cells, and this trans-activation is known to be inhibited in GC by BCL6 and in plasma cells by Blimp-1. These findings support a model whereby IRF-4 function is modulated in a stage-specific manner by its interactions with developmentally restricted sets of transcription factors (Gupta et al., 2001).
complex has been demonstrated in the human Ig3 promoter upon CD40L/IL-4 induction (Shaffer et al., 1999). STAT6 and c-Rel also display a synergistic activation of the human Ig4 promoter (Agresti and Vercelli, 2002). In contrast to the mouse, the human Ie promoter is directly bound by C/EBP, and both the human Ie and Ig3 are synergistically activated by C/EBP and STAT6 (Mikita et al., 1996; Pan et al., 2000).
A DISTANT REGULATORY REGION FOR GT AND CSR: THE 3¢ IGH ENHANCERS
Regulation of Ca GT TGF-b stimulates switching to IgA and, to a lesser extent, to IgG2b. The stimulation pathway involves transmembrane receptors with serine/threonine kinase activity, connected with intracellular transducers of the Smad family. Phosphorylation of Smad proteins allows them to form heterodimers that translocate to the nucleus and regulate transcription. Other cytokines or soluble factors, such as IL-5 or the vasointestinal peptide VIP, stimulate the secretion of IgA rather than stimulating CSR by itself. The Ia promoter carries two binding sites for Smad 3 and Smad 4, which are upregulated upon TGF-b induction; in contrast, Smad 7 has an inhibitory role that may account for the observed IFN-g inhibition of IgA expression (Shockett et al., 1991; Ulloa et al., 1999). The TGF-b responsive element also includes a binding site for a transcription factor of the acute myeloid leukemia AML(RUNX)/PEBP2a family (Hanai et al., 1999). In addition, a CREB/ATF site is responsible for the constitutive basal activity of the promoter. Smad, CREB, and AML factors bind independently to the promoter but further interact with each other and cooperatively stimulate transcription, so that their functional synergy is required for maximal TGF-b–induced GT (Zhang and Derynck, 2000). The effects of the three types of transcription factors may be integrated through their association with the co-activator CBP/p300, known to form multiprotein complexes and to recruit basal transcription factors and the histone acetylation machinery (Zhang and Derynck, 2000). The Ia promoter binds the Ets proteins Ets-1 and PU.1, which may account for its LPS inducibility; an NFkB site was identified that, while playing a minimal role in GT, may independently stimulate CSR to Ca (Shi et al., 2001). Specificities of Human I Promoters Promoters located upstream of human I exons are also responsive to the cytokine environment, as assayed by transient transfection. IL-10 induces the switching of human B cells to IgG1, IgG3, and IgA (Brière et al., 1994a 1994b). Cooperative binding of STAT6 and of a p50/p65/c-Rel
Organization of the 3¢ IgH Regulatory Region Various observations have prompted a search for regulatory elements other than Em within the IgH locus. Deletions downstream of the IgH a gene were correlated with a reduction in IgH transcription, whereas the association of the cmyc oncogene with Ca downstream sequences as a result of chromosomal translocations close to the Ca gene upregulated expression of c-myc (Gregor and Morrison, 1986; Khamlichi et al., 2001). Indeed, a major 3¢ regulatory region spanning more than 30 kb has been characterized downstream of the IgH locus. This has been shown to include four lymphoid-specific transcriptional enhancers: hs3A, hs1,2, hs3B, and hs4, and likely plays a role during the antigendriven maturation of B cells (Petersson et al., 1990; Dariavach et al., 1991; Lieberson et al., 1991; Matthias and Baltimore, 1993; Michaelson et al., 1995). The hs1,2 element stands as a central element, responsive to mitogens and to cross-linking of the BCR or of CD40 (reviewed in Khamlichi et al., 2000). It is flanked by ~10 to 12 kb long inverted repeats, including two copies of a weak enhancer (hs3A and hs3B) (Chauveau et al., 1996; Matthias and Baltimore, 1993; Giannini et al., 1993; Madisen and Groudine, 1994; Michaelson et al., 1995; Saleque et al., 1997). Finally, the distal hs4 enhancer lies downstream of this palindromic structure, at about 30 kb downstream from the mouse a gene (Madisen and Groudine, 1994). In humans also, inverted repeats symmetrically flank hs1,2 while the whole region is duplicated and lies downstream of each a gene (Mills et al., 1997; Chen and Birshtein, 1997; Pinaud et al., 1997). In contrast, the rabbit IgH locus includes a single regulatory region downstream of its multiple a gene copies (Volgina et al., 2000). The region downstream of the murine 3¢ Igh regulatory region has now been sequenced (Accession #AF450245) (Zhou et al., 2002). The nearest non-IgH gene lies ~35 kb downstream of hs4. Hs1,2 and hs3 elements become demethylated and behave as transcriptional enhancers in activated or terminally differentiated B cells (Dariavach et al., 1991; Lieberson et al., 1991; Giannini et al., 1993; Fulton and Van Ness,
19. Regulation of Class Switch Recombination
1994; Matthias and Baltimore, 1993), whereas hs4 may be active throughout B-cell development (Madisen and Groudine, 1994; Michaelson et al., 1995). Although individually weak, these elements display strong synergies when associated (Madisen and Groudine, 1994; Chauveau et al., 1998). Hs1,2 is especially boosted when flanked by copies of hs3 in inverted orientation (mimicking the endogenous arrangement) (Chauveau et al., 1998). When combined, 3¢ elements both stimulate transcription in B cells and silence IgH expression in non-B cells. In addition, the combination displays some of the properties of a locus control region in that it supports position-independent expression. However, direct copy number dependence has not been observed (Madisen and Groudine, 1994; Chauveau et al., 1999; Khamlichi et al., 2000). The addition of Em to any individual 3¢ element does not show significant synergy, but a strong effect is obtained at all B-cell differentiation stages when Em is added to a combination of hs3A, hs1,2, hs3B, and hs4 (Mocikat et al., 1993; Mocikat et al., 1995; Ong et al., 1998; Chauveau et al., 1998). It thus seems that the 3¢ “weak enhancers” act as powerful co-enhancers when optimally combined (Chauveau et al., 1998).
The 3¢ IgH Regulatory Region Controls Expression of Several Non-IgM Immunoglobulins Analysis of several spontaneous or targeted truncations of the 3¢ regulatory region has clearly indicated its role in the transcription of switched heavy chain genes. In a differentiated B cell line, spontaneous deletion of the entire region was associated with a seven-fold decrease in transcription of an IgH a gene (Gregor and Morrison, 1986; Michaelson et al., 1995). Targeted replacement of 3¢aE(hs1,2) by the neoR gene in a mature B cell line also strongly affected transcription of a rearranged g2a gene lacking Em (Lieberson et al., 1995). This finding is consistent with the notion that hs1,2 controls transcription at a late B-cell developmental stage (Lieberson et al., 1995). However, it is difficult to ascertain whether suppression of g2a transcription resulted from the lack of hs1,2 per se—likely to be more critical in the absence of Em—or from disruption of the putative polarized effect of the 3¢ IgH regulatory region (through the so-called “neo effect,” discussed in detail below). A combined deletion of hs3A and hs1,2 was characterized in the 70Z/3 pre-B cell line (Saleque et al., 1999). In line with the activity pattern of hs3A and hs1,2, no apparent decrease in m expression was detected. When terminal differentiation was mimicked through fusion with myeloma cells, hs3A and hs1,2 did not seem to be required for the plasma cell–specific upregulation of m expression on the mutated allele (Saleque et al., 1999). Together, data from transgenics and cell lines suggest that the role of 3¢ IgH elements may be restricted to the reg-
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ulation of expression of class-switched Ig genes but give no firm conclusion about their role in CSR.
The 3¢ IgH Regulatory Region Controls GT of Most Constant Region Genes Analysis of B-cell maturation in mice carrying deletions or replacement of the 3¢ IgH enhancers has confirmed that IgM expression is only marginally dependent on these elements. Transcription of the m gene was unmodified after deletion of hs3A and/or hs1,2, whereas it was slightly reduced in resting B cells only, as a result of a combined deletion of hs3B and hs4 (Cogné et al., 1994; Manis et al., 1998; Pinaud et al., 2001). Both mature VDJ-Cm transcripts and Im transcripts were preserved after such deletions. In contrast to Cm, a wide-range of GT alterations was observed for several other constant IgH genes. Targeted neo replacement of the endogenous hs1,2 enhancer first suggested its involvement in CSR to downstream IgH genes (Cogné et al., 1994). The homozygous replacement suppressed the ability of cultured B cells to induce GT of Cg3 (which lies more than 120-kb upstream of hs1,2), Cg2b, Cg2a, and Ce. In agreement with the accessibility model (reviewed in Sleckman et al., 1996), in vitro switching to the corresponding Ig classes was consequently altered. Similar defects were observed for in vivo Ig production but were more severe for IgG3 and IgG2a. In vitro as well as in vivo, expression of the Ca gene was minimally affected, and normal levels of IgG1 were found (Cogné et al., 1994). Essentially, the same phenotype resulted from replacement of hs3A with a neor gene (Manis et al., 1998). Cre/lox experiments allowed the assay of isolated deletions of hs3A or hs1,2 in the absence of an inserted neor gene. Strikingly, CSR appeared normal, either eliminating any role of these enhancers in CSR or assigning them redundant or second-rate functions while relating CSR defects to “neo effects” (Manis et al., 1998). However, analysis of a joint deletion of the hs3B/hs4 enhancers revealed a severe CSR defect even in the absence of an inserted neor gene (Pinaud et al., 2001). The mutation again strongly reduced IgG2b and IgG3 production, both in vivo and in vitro. IgG2a, IgE, and IgA classes were also reduced but to a lesser extent. Although the serum concentration of IgA was slightly affected, germline transcription of the a gene and IgA secretion were decreased upon in vitro stimulation by LPS plus TGFb. In addition, studies of heterozygous mutant animals confirmed that a transcripts mostly originated from the unmutated allele. In contrast, IgG1 was minimally affected. Interestingly, a more drastic phenotype resulted from the hs3B/hs4 replacement by a neo gene than from the clean deletion of hs3B/hs4 (with a more severe reduction of serum IgE and a combined in vitro and in vivo defect of IgA) (Pinaud et al., 2001). Replacement of hs3B/hs4 with neor even affected IgM and IgG1, whereas neo replacements
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of the upstream enhancers hs3A and hs1,2 did not (Cogné et al., 1994; Manis et al., 1998).
“Neo Effects” Indicate That GT Relies on a Polarized Effect of the 3¢ IgH Regulatory Region Additional insights into the molecular basis for the polarized effect of the 3¢ IgH regulatory region came from comparisons of targeted neor insertions at various locations amid CH genes (Seidl et al., 1999). Such insertions most often resulted in a blockade of GT for all upstream but not downstream I exon promoters and have been interpreted as the blockade of a long-range and polarized function of the 3¢ regulatory region (Seidl et al., 1999). Again, the notable exceptions of Cm, Cg1, and to a lesser extent Ca, may indicate their inclusion in independent functional units. As for other insertions of neo within the IgH locus, explanations for “neo effects” in the 3¢ regulatory region may invoke competition between GT promoters and the neor promoter (Seidl et al., 1999). The neo gene would then constitute a decoy site for transcription factors normally controlling GT; in that sense, it is noteworthy that the neo gene becomes LPS-inducible upon insertion in the IgH locus (Manis et al., 1998; Seidl et al., 1998). Alternatively, the constitutively active neo promoter may behave as a boundary element and result in a functional deletion of any regulatory element located downstream of the insertion. The aggravated phenotype of mice with a neor insertion compared to the hs3B/hs4 deletion would then suggest the existence of additional regulatory elements lying downstream of hs4, a hypothesis that is not supported by the normal IgH locus expression in mice simply carrying a neo insertion downstream of hs4 (Manis et al., 2003).
MECHANISMS FOR 3¢ IGH REGULATORY REGION-MEDIATED REGULATION OF GT Altogether, mutational studies point to elements downstream of hs1,2 (i.e., to hs3B and hs4, which have not been deleted individually in the mouse) as major regulators of LPS-induced GT and CSR. Since a “natural knockout” of hs3B is featured by the human IgH locus with no associated CSR defect, and since hs3B is 97% identical to hs3A (which is itself dispensable for CSR), hs4 thus stands as the main candidate for a master GT/CSR regulator. However, deletion experiments may obviously mask functional redundancies and, on the basis of their transcriptional synergies in transgenes, it is tempting to speculate that the four 3¢ IgH enhancers may cooperate in the control of GT and CSR. The primary effect of mutations affecting the 3¢ IgH regulatory region could thus be a decreased accessibility of the
FIGURE 19.2 Modulation of holocomplexes in response to external stimulation. A hypothetical representation of loops that may allow interactions between cis-acting regulatory elements of the IgH locus and specific promoters of the locus. Both maturation stages and external stimuli likely control accessibility of the various promoters to such stimulatory interactions. In vitro stimulations stand as paradigms of targeted GT and CSR to either Cg2b/Cg3 (LPS) or Cg1/Ce (LPS + IL4).
affected CH genes through inhibition of GT, changes in chromatin structure of affected CH genes, or both (Cogné et al., 1994; Manis et al., 1998; Pinaud et al., 2001). While the long-range activity of the 3¢ regulatory region would require presence of the hs4 element, neor insertions within the IgH genes would block the propagation of this activity along the locus (Pinaud et al., 2001; Seidl et al., 1998). The 3¢ regulatory region may shift from one accessible I promoter to another following appropriate stimuli (Arulampalam et al., 1997). Bringing such remote promoters into proximity of a 3¢ IgH “enhanceasome” could involve architectural transcription factors and result in the formation of loops by the intervening regions (Figure 19.2).
Specific Interactions of I Promoters with 3¢ IgH Enhancers In addition to their specific interactions with transcription factors, I promoters may specifically interact with cis-acting elements shown to boost their transcriptional activity in reporter constructs. It is, however, unclear whether such interactions contribute to the cytokine induction of transcription. For example, in the case of the mouse Ig2a promoter, IFN-g regulation is solely dependent on the promoter itself. A strong increase of transcription not responsive to IFN-g results from the addition of either the Em enhancer or a combination of 3¢ elements (Collins and Dunnick, 1999). Experiments on transfected reporter genes also indicate that the human Ig3 and Ia promoters may be stimulated by a
19. Regulation of Class Switch Recombination
linked 3¢ IgH regulatory element, although it is not clear that these interactions contribute to induction (Pan et al., 2000; Hu et al., 2000). The murine hs1,2 enhancer has also been shown to positively interact with several I promoters, including Ig2b and Ia (Laurencikiene et al., 2001). In the absence of motifs binding cytokine-dependent transducers, the 3¢ regulatory region may solely be induced by signals resulting from BCR and/or CD40 ligation, whereas the cytokine modulation of GT would rely on each I promoter element. Accordingly, knockin neo cassettes inserted within the IgH 3¢ regulatory region were similarly inducible by LPS or LPS plus IL-4 (Cogné et al., 1994; and unpublished observations). Large transgenes, including the Cm and the Cg1 genes, are able to undergo switching in the absence of any 3¢ IgH regulatory element (Cunningham et al., 1998). Regulatory elements associated with the Cg1 gene include two DNA hypersensitive sites, one located next to the I promoter (site I) and a second immediately upstream of the Sg1 region (site II), which bind NF-kB/Rel and STAT6 (Adams et al., 2000). The Sg1 sequence by itself may act as a transcriptional enhancer for Ig1 transcripts (Cunningham et al., 1998), and Site II 5¢ of Sg1 may contribute to an LCR activity within such transgenes (Adams et al., 2000).
Transcription Factors Controlling 3¢ IgH Regulatory Elements The 3¢ regulatory elements bind multiple transcription factors, some lymphoid-specific, like those that bind to OCT, mE5, kB, mA, and mB-like motifs, and others that are ubiquitous, such as ATF/CREB family factors or the TFE3/USF factors. A given maturation stage or a given activation status may correspond to unique combinations of ubiquitous and tissue-specific factors able to display either antagonistic or synergistic interactions. NFKB Family and Transcription Factors Connected to CD40 Triggering CD40 signaling triggers cascades involving TRAF6, IKK, and NIK (NF-kB-inducing kinase), resulting in the activation of NF-kB (Brady et al., 2000). A CD40stimulated pathway resulted in the recruitment of the NFAB activating complex (composed of Elf-1, JunB, and a Fosfamily related partner) to the ETS-AP-1 motif of hs1,2 (Grant et al., 1996). CD40 is putatively associated with Ku70, a component of the NHEJ machinery involved in the junctions between S regions, which translocates to the nucleus upon CD40 ligation (Morio et al., 1999). Triggering of CD40 also results in nuclear translocation of the SWAP-70 protein, which may be involved in CSR to the Ce gene (Borggrefe et al., 2001).
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NF-kB binding sites located in 3¢ IgH enhancers likely play a major role in their regulation. Human hs1,2 alleles carrying duplicated NF-kB sites have been correlated with higher serum IgA1 levels in patients (Aupetit et al., 2000; Denizot et al., 2001). An NF-kB site in the murine hs1,2 binds a complex including p50, p65, and c-Rel; and in B lymphocyte cell lines, mutation of this site led to an increased activity of a reporter gene (Michaelson et al., 1996b). The cooperative binding demonstrated for the NFkB complex and for NFE—mutation of either of which leads to decreased reporter gene expression in plasma cell lines— suggests that NF-kB may be an activator at the plasma cell stage and a repressor before (Michaelson et al., 1996b; Linderson et al., 1997). In contrast, in hs4, a single kB site binds a complex with a stimulatory effect at all B-cell stages, in agreement with activity of this enhancer throughout B-cell development. Of note, CSR defects from knockout experiments affecting the 3¢ regulatory region are reminiscent of those found in mice lacking NF-kB components or lacking the Ik kinase (IKK) activity necessary for nuclear translocation of NF-kB (reviewed in Ghosh et al., 1998). Targeted disruption of RelA (p65) was first suggested to affect switching to IgG1 and IgA because of decreased serum levels of these isotypes (Doi et al., 1997). When studied using in vitro activated B cells, it, in fact, mostly inhibited GT of and CSR to Cg3 (Horwitz et al., 1999). Lack of p50 resulted in a substantial decrease of IgE, IgG1, and IgA serum levels (Sha et al., 1995). Upon appropriate stimulation in vitro, p50-/- resting B cells underwent substantial switching to IgG1 but markedly less to IgG3, IgE, or IgA. Interestingly, both GT and CSR were affected for all isotypes, except for IgA where normal GT occurred (Snapper et al., 1996). Whereas B cells from RelB knockout mice underwent normal CSR (Snapper et al., 1996), mice deficient in c-Rel had a severe deficiency in IgG1 and IgG2a (Köntgen et al., 1995). In vitro, stimulated B cells from mice deficient in the C-terminal transactivation domain of c-Rel failed to switch to IgG3, IgG1, and IgE. Here again, the failure to switch to IgE occurred despite normal GT of Ce, raising the possibility that NF-kB/Rel factors may have roles in the regulation of CSR in addition to regulation of GT (Zelazowski et al., 1997; Snapper et al., 1997). Thus, deficiencies in several members of the NF-kB family appear to be associated with specific CSR defects, some of them associated with GT defects. It is tempting to speculate that deficiencies in NFkB family members alter CSR through an impaired activation of the 3¢ IgH LCR and of I promoters themselves, leading to a defect in GT and/or CSR. However, additional indirect effects on CSR must also be considered, since these factors regulate multiple genes, including those of cytokines and accessory membrane receptors.
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Lymphoid-Specific Factors of the Oct, E2A, and Ets Families Oct Family Factors Octamer sites likely contribute to the B cell specificity of 3¢ IgH regulatory elements. Two imperfect octamer sites within hs3A bind Oct-1 and Oct-2 with low affinity and no cooperative binding (Matthias and Baltimore, 1993). A unique Oct site in hs1,2 is mostly regulated by a phosphorylated form of Oct-2 interacting with the co-activator OCAB/Obf1 (Tang and Sharp, 1999). As discussed below, its activity is modulated by Pax5/BSAP and may yield stimulation at the plasma cell stage and repression before (Singh and Birshtein, 1996). Finally, two Oct sites were defined within hs4, where the Oct-binding activity is mainly attributable to Oct-1 and is mostly stimulatory (Michaelson, 1996b). E2A Products Among lymphoid specific factors, E-box binding bHLH proteins encoded by the E2A gene may be especially important. The hs1,2 mE5 site binds bHLH proteins E12 and E47, whose activity is central in the commitment to B-cell differentiation (reviewed in O’Riordan and Grosschedl, 1999). Binding of these factors is controlled by dominant negative regulatory factors of the Id family (reviewed in Kadesh, 1992). Although Id1 and Id2 are responsible for the inhibition of Ig expression in non-B and in pro-B cells, Id3 is expressed in pre-B and B but not in plasma cell lines and is able to downregulate hs1,2-driven reporter genes (Sun, 1994; Meyer et al., 1995). Id3 thus likely participates in the silencing of hs1,2 in resting B lymphocytes. Strikingly, multiple E-box sites (mE2 and mE5), potentially binding E12 and E47, are also found within the hs3A, hs3B, and hs4 enhancers. ETS Family Several Ets family factors bind hs1,2, notably at the mA and mB sites. In addition, footprinting experiments allowed identification of an ETS-AP-1 motif, which was protected following stimulation of splenic B cells. Trimerization of this motif upstream of a reporter gene stably transfected in a surface IgM-expressing cell line conferred strong expression following cross-linking of surface IgM, treatment with TPA, or stimulation through CD40. In response to these stimuli, the ETS-AP-1 site was shown to bind a complex, named NFAB, and composed of the Elf-1 protein (an Etsrelated factor), together with Jun-B, c-Fos (in the case of signaling through IgM), and a c-Fos-related family member (for CD40 stimulation) (Grant et al., 1995, 1996). Another Ets family transcriptional activator, first named NF-aP and later identified as PU.1, binds hs1,2 at the plasma cell stage in close vicinity to Pax5 sites (Neurath et al., 1995; Linderson et al., 2001). Its interactions with Pax5 are discussed
below. NF-aP/PU.1 binding results in an increased synthesis of g2b, g3, and a mRNAs but had little effect on m (Neurath et al., 1995). Finally, the Ets-family protein, NFE (nuclear factor Ets-like) displays cooperative binding of hs1,2 with NF-kB (Linderson et al., 1997). Implication of Pax5/BSAP as a Regulator of Both I Promoters and 3¢ IgH Enhancers Pax5 belongs to a family of transcription factors containing a paired DNA-binding domain; it is expressed in B lymphocytes, the developing central nervous system, and adult testis (Adams et al., 1992). In the absence of Pax5, Bcell progenitors are blocked at an early stage, show a dramatic reduction in V-to-DJ recombination, and loosen their commitment to the B lymphoid lineage (Urbánek et al., 1994; Nutt et al., 1996; Nutt et al., 1999). Pax-5 binds multiple sites upstream of and within several switch regions in the IgH locus (reviewed in Busslinger and Urbánek, 1995; Michaelson et al., 1996a; Stavnezer, 1996). Important with regards to CSR, although Pax5 represses Ia transcripts and CSR to the a gene in the I.29 cell line, it binds and activates the Ie and Ig2a promoters and seems necessary for the LPS/IL-4 induction of Ie GT, as well as for the in vitro switching to IgG1 (Max et al., 1995; Liao et al., 1994; Qiu et al., 1998). Pax5 also binds to multiple sites within the murine 3¢ IgH regulatory region. Two sites within hs1,2 are occupied in pro-B, pre-B, and B-cell lines but not in plasma cells. Upon binding, Pax5 behaves as a repressor of hs1,2 (Singh and Birshtein, 1993; Neurath et al., 1994). This effect of Pax5 likely involves its ability to form ternary complexes and then to modulate the activity of factors, including Oct, NF-kB, and a G-rich motif binding protein, all shown to contribute to the concerted repression of hs1,2 in immature B cells (Singh and Birshtein, 1996; Michaelson et al., 1996b). Upon differentiation to the plasma-cell stage and Pax5 downmodulation, NF-kB and Oct factors could switch to a positive function, thanks to their respective interactions with Ets factors and with OCA-B (Singh and Birshtein, 1996). An indirect repressive function of Pax5 on hs1,2 may also be mediated through steric hindrance with the Ets family transcriptional activator NF-aP/PU.1, whose neighboring site may only be occupied in plasma cells when Pax5 is not expressed (Neurath et al., 1995). Similarly, antagonistic activities of Pax5 and PU.1 have been documented for the 3¢Ek enhancer, with Pax5 restricting the full enhancer activity to activated or terminally differentiated B cells (Maitra and Atchison, 2000). Again, within the hs4 enhancer, multiple Pax5 sites bind repressive complexes that can be detected in pre-B and B cells but not in plasma cells (Michaelson et al., 1996a). As in other tissues, Pax5 thus clearly displays dual functions in B lymphocytes: positively regulating the transcription of genes like mb-1 or
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CD19 and the commitment to the B cell lineage, while repressing immunoglobulin J chain gene expression or full activity of Ig 3¢ elements (Rinkenberger et al., 1996; Nutt et al., 1998). Pax5/BSAP expression is downregulated by Blimp-1, a key regulator of plasma cell differentiation whose overexpression also inhibits class switching (Knodel et al., 2001; Lin et al., 2002). Pax5 is additionally regulated through alternate splicing: The main active isoform, Pax5a may be under control of a dominant negative isoform, Pax5d and a stimulating isoform, Pax5e, whereas the Pax5b isoform of unknown function may somehow persist at the plasma cell stage (Zwollo et al., 1998; Lowen et al., 2001). In response to reactive oxygen species, such as those generated during immune responses through the action of type I cytokines, the Ref-1 enzyme is translocated to the nucleus in B lymphocytes. Ref-1 reduces the cysteine residues of various transcription factors that are crucial to B-cell activation during TI responses, including AP-1, NF-kB, and Pax5, and enhances their activity (Tell et al., 2000; Xanthoudakis et al., 1992a, 1992b). What is finally the role of Pax5 with regard to CSR? Pax5 levels do not constantly decrease in B cells activated through different extracellular stimuli. In LPS- or CD40Lstimulated B cells, a strong induction of an hs1,2-dependent transgene was observed while Pax5 expression levels remained unchanged (Andersson et al., 1996). The same was true in a B cell line transfected with the same reporter gene after cross-linking of surface IgM. Likely explanations may be that Pax5-dependent inhibition of hs1,2 is alleviated by these external signals. Cross-linking of OX40L (a member of the TNF/NGF-receptor family known to negatively regulate CSR) on CD40L-stimulated splenic B cells led to a 60 to 80% decrease in Pax5 levels, the reduction being detected at both the protein and the messenger levels. In vivo footprinting experiments on hs1,2 showed a loss of the Pax5 footprint and the appearance of a footprint at the aP site with an occupancy pattern similar to that observed within hs1,2 in plasma cells (Stüber et al., 1995). Pax5 has dual effects through its direct binding to I promoters themselves, repressing the Ia and activating the Ig1, Ie, and Ig2a promoters (Max et al., 1995; Liao et al., 1994; Qiu et al., 1999). These data suggest a complex pattern of Pax5-mediated repression or stimulation of the cis-acting elements that control GT and CSR. The effects of a complete Pax5 defect with regard to CSR have not been evaluated due to the resulting block of B-cell differentiation at an early stage. Finally, Pax5 may play a positive role in the induction of GT and CSR to several CH genes in activated B cells; its downmodulation occurring later on during terminal differentiation in plasma cells may then allow the high-level transcription of class-switched antibody genes. Conditional inactivation in differentiated cells (Mikkola et al., 2002) will hopefully provide information about such issues.
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3¢ IgH Regulatory Region May Promote Chromatin Remodeling Another factor that could participate together with BSAP in the repression of 3¢ IgH enhancers in immature and resting B cells is Bach2. Putative Maf recognition elements (MAREs) have been identified within hs3A and hs3B (Muto et al., 1998). These motifs, also found in the b-globin LCR, are bound by heterodimers of basic-region leucine zipper factors that include proteins of the Maf family associated either with p45 (forming the transcription factor NF-E2) or with the negative regulators Bach1 or Bach2. Bach/Maf heterodimers bind with each other, generating a multimeric and multivalent DNA binding complex, and likely play an architectural role for the assembly of the theoretical beta-globin LCR “holocomplex ” (Igarashi et al., 1998). Among these factors, Bach2 has a restricted expression in the brain and the B cell lineage, where its expression and binding to the hs3 MARE progressively decreases during maturation and is switched off in plasma cells (Muto et al., 1998). The MafK/Bach2 complex apparently represses reporter genes driven by the 3¢ IgH LCR (Muto et al., 1998). By analogy with the b-globin LCR, these findings suggest that MAREbinding heterodimers of varying composition may control the architecture of the 3¢ IgH regulatory region throughout B cell maturation. It is now accepted that the chromatin structure is an essential component of the transcriptional regulation machinery. Although the exact molecular mechanisms controlling the accessibility to promoters and enhancers of trans-acting factors are still debatable, it is clear that changes in chromatin structure near transcriptionally active genes require interactions between transcription factors, histones, and other co-factors in order to remodel and displace nucleosomes (reviewed in Kadonaga, 1998). Enhancers may recruit and/or direct histone acetyltransferase (HAT)– containing molecules (such as transcriptional co-factors) to critical regulatory regions. Indeed, transcriptionally active genes are associated with acetylated core histones (reviewed in Majumder and DePamphilis, 1995). Therefore, it was interesting to look at the effect of the 3¢ IgH regulatory region on the chromatin structure of linked genes. A combination of hs1,2, hs3B, and hs4 (hs123B4) was suggested to act as an LCR (Madisen and Groudine, 1994). This cassette deregulated the transcription of linked c-myc genes, with a shift from P2 to P1 promoter usage (Madisen and Groudine, 1994). In addition, chromatin immunoprecipitation assays revealed an increase in histone acetylation (Madisen et al., 1998). Treatment of the transfectants with an inhibitor of histone deacetylases, leading to general histone acetylation, inhibited the hs123B4-mediated highlevel expression of P1 but not that of the P2 promoter. Thus, increased acetylation may be one mechanism by which the 3¢ IgH regulatory region establishes and maintains a tran-
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scriptionally active state along linked genes over entire chromatin domains. This could be achieved by recruiting HATcontaining co-factors to first induce local histone acetylation and then facilitate its propagation throughout the chromatin domain under the influence of the 3¢ regulatory region (Madisen et al., 1998). Thus, 3¢ IgH enhancers might recruit HAT activity to counteract a repressive chromatin structure generated by HDACs recruited by proteins such as LSF. Similar to observations in NF-kB-deficient mice, in which CSR was suppressed while GT of certain constant genes was maintained, alterations of the 3¢ enhancers have been reported to affect CSR more significantly than GT (this was specially true for the Ce gene) (Pinaud et al., 2001). As with NF-kB transactivation, it is thus conceivable that the 3¢ cisregulatory elements not only stimulate GT but also promote remodeling of the S regions prior to CSR. The observation that mice carrying large human IgH transgenes can undergo some level of CSR in the absence of 3¢ regulatory elements is also noteworthy but does not allow any quantitative comparison with a normal endogenous locus (Wagner et al., 1994; Wagner et al., 1996). However, these data may suggest that elements upstream of the 3¢ IgH regulatory region are sufficient for CSR to occur. It should also be stressed that up to now, all targeting experiments within the 3¢ IgH regulatory region have only resulted in partial in vivo CSR defects and that some Ig classes (IgG1, and to a lesser extent IgA) were less affected. Compensatory mechanisms may be at work at the cell selection level in order to allow some compartments of switched memory cells to be filled up, also indicating that alternate CSR pathways may not rely on 3¢ IgH elements.
COORDINATED REGULATION OF TRANSCRIPTION, RECOMBINATION, AND REPLICATION V(D)J recombination mostly occurs in the G0 and/or G1 stage of the cell cycle, whereas cell proliferation is accompanied by downregulation of RAG activities (Hesslein and Schatz, 2001). In contrast, CH germline transcripts are mainly expressed in G1 and S phase, and CSR seems to require DNA replication (Lundgren et al., 1995; Stavnezer, 2000). How the three processes of transcription, replication, and CSR may be coordinated is still a matter of debate. The intronic enhancer, Em, may play a crucial role since it is both involved in the accessibility of Sm to CSR and associated with a putative origin of replication (Arizumi et al., 1993). In contrast, the 3¢ IgH regulatory region, as it is currently defined, is unlikely to play a role in replication since its deletion in a plasmacytoma cell line did not change the pattern of replication of the locus (Michaelson et al., 1997). The IgH locus replicates apparently in two different patterns that
change with B-cell development. In non-B cells, plasma cells, and LPS-stimulated splenic B cells, Ca replicates early in S phase and upstream CH genes replicate progressively 3¢ to 5¢ at later intervals. In contrast, all CH genes seem to replicate early in S phase in pro- and pre-B cells (Zhou et al., 2002a). A replication origin has been localized ~70-kb downstream of hs4 in a non-B cell line and may demarcate the 3¢ end of an IgH replicative domain (Brown et al., 1987; Ermakova et al., 1999; Zhou et al., 2002b). There are, however, no data on whether developmental changes in IgH replication accompany CSR.
CONCLUSION This review has focused on CSR to all non-m isotypes, except for delta. The expression of IgD results from alternative splicing that regularly occurs as part of B-cell maturation, and only rarely involves Cm deletion (Owens et al., 1991; Arpin et al., 1998). In contrast, DNA recombination is constantly needed for the other isotypes, and there are a number of elements that are now known to be essential. For example, a critical first step in CSR is GT, whose production for several isotypes is dependent on at least two of the 3¢ IgH enhancers. GT of individual isotypes requires cell–cell interaction involving CD40 and CD40L; various B cell transcription factors, including NFkB; and specific T cell cytokines. A predilection to switch to particular classes, especially IgE in allergic individuals, can be harmful for human health, and most likely is fostered by the T cell cytokine profile associated with particular Th subsets. AID appears to act only after GT formation. Mistakes in CSR have been associated with chromosomal translocations involving c-myc and are regularly detected in murine plasmacytomas, human myeloma, and Burkitt’s lymphoma. Current challenges are to identify the signals that trigger 3¢ enhancer activity during B-cell development and the mechanisms that engage these enhancers with I promoters for GT. The formation of GT clearly involves both positive and negative regulation, providing an additional arena for critical investigation.
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Xu, L., and Rothman, P. (1994). IFN-gamma represses epsilon germline transcription and subsequently down-regulates switch recombination to epsilon. Int Immunol 6, 515–521. Xu, M. Z., and Stavnezer, J. (1992). Regulation of transcription of immunoglobulin germ-line gamma 1 RNA: Analysis of the promoter/enhancer. EMBO J 11, 145–155. Zelazowski, P., Carrasco, D., Rosas, F. R., Moorman, M. A., Bravo, R., and Snapper, C. M. (1997). B cells genetically deficient in the c-rel transactivation domain have selective defects in germline CH transcription and Ig class switching. J Immunol 159, 3133–3139. Zhang, J., Bottaro, A., Li, S., Stewart, V., and Alt, F. W. (1993). A selective defect in IgG2b switching as a result of targeted mutation of the I gamma 2b promoter and exon. EMBO J 12, 3529– 3237. Zhang, Y., and Derynck, R. (2000). Transcriptional regulation of the TGFb-inducible mouse germ-line Ig a constant region gene by functional cooperation of Smad, CREB and AML family members. J Biol Chem 275, 16979–16985. Zhou, J., Ashouian, N., Delepine, M., Matsuda, F., Chevillard, C., Riblet, R., Schildkraut, C. L., and Birshtein, B. K. (2002a). The origin of a developmentally regulated Igh replicon is located near the border of regulatory domains for Igh replication and expression. Proc Natl Acad Sci U S A 99, 13693–13698. Zhou, J., Ermakova, O. V., Riblet, R., Birshtein, B. K., and Schildkraut, C. L. (2002b). Replication and subnuclear location dynamics of the immunoglobulin heavy-chain locus in B-lineage cells. Mol Cell Biol 22, 4876–4889. Zwollo, P., Arrieta, H., Ede, K., Molinder, K., Desiderio, S., and Pollock, R. (1997). The Pax-5 gene is alternatively spliced during B-cell development. J Biol Chem 272, 10160–10168.
Note Added in Proof Additional transcriptional regulators of 3¢ Igh enhancers have been described. YY1 has been shown to acquire binding to murine hs3 enhancers after LPS stimulation (Gordon et al., 2003). In addition, synergistic activation of human hs4 by NFkB and Oct-2 has recently been reported (Sepulveda et al., in press). Furthermore, several papers (Chaudhuri et al., 2003; Shinkura et al., 2003; Yu et al., 2003) have shown that I region-driven transcription through S regions in a single (physiological) direction can generate R loops, which coordinately exposes a single non-templated DNA strand that is a substrate for AID. The ability to form R loops appears to be an essential structural feature of GT prior to CSR. Chaudhuri, J., Tian, M., Khuong, C., Chua, K., Pinaud, E., and Alt, F. W. (2003). Transcription-targeted DNA deamination by the AID antibody diversification enzyme. Nature 422, 726–730. Gordon, S. J., Saleque, S., and Birshtein, B. K. (2003). Yin Yang 1 is a lipopolysaccharide-Inducible activator of the murine 3¢ Igh enhancer, hs3. J Immunol 170, 5549–5557. Sepulveda, M. A., Emelyanov, A. V., and Birshtein, B. K. (2003). NFkB and Oct-2 synergize to activate the human 3¢ Ighhs4 enhancer in B cells. J Immunol. In press. Shinkura, R., Tian, M., Smith, M., Chgua, K., Fujiwara, Y., and Alt, F. W. (2003). The influence of transcriptional orientation on endogenous switch region function. Nature Immunol 4, 435–441. Yu, K., Chedin, F., Hsieh, C-L., Wilson, T. E., and Lieber, M. R. (2003). R-loops at immunoglobulin class switch regions in the chromosomes of stimulated B cells. Nature Immunol 4, 442–451.
C
H
A
P
T
E
R
20 Molecular Mechanism of Class Switch Recombination JANET STAVNEZER,1 KAZUO KINOSHITA,2 MASAMICHI MURAMATSU,2 AND TASUKU HONJO2 1
Department of Molecular Genetics and Microbiology, University of Massachusetts Medical School, Worcester, Massachusetts, USA, 2 Department of Medical Chemistry, Graduate School of Medicine, Kyoto University, Kyoto, Japan
al., 1981; Nikaido et al., 1982; Dunnick et al., 1993). Although it is well established that CSR results in an intrachromosomal deletion of CH genes located between the rearranged VH gene and a CH gene to be expressed, its precise mechanism is still unknown. The basic questions common to all recombination reactions are selection and recognition of target DNA, mode of the cleavage, and mechanism of repair and ligation. Until very recently, there were no known specific trans-acting factors involved in CSR. Muramatsu et al. (1999, 2000) identified the novel gene AID, which is essential for CSR as well as SHM. Subsequently, AID was shown to induce CSR and SHM in non-B lymphocytes, including fibroblasts and T cells, thus demonstrating that AID is the only trans-acting factor that is required for CSR and SHM, and is specific to B cells (Okazaki et al., 2002). In this chapter, we describe the latest knowledge about the molecular mechanism of CSR, focusing on the three basic questions mentioned above and also on the functional role of AID in CSR.
Activated B lymphocytes undergo two genetic alterations in the immunoglobulin (Ig) gene locus, class switch recombination (CSR) and somatic hypermutation (SHM), responsible for isotype switching and affinity maturation, respectively (Diaz and Casali, 2002; Honjo et al., 2002; Manis et al., 2002b; Storb and Stavnezer, 2002). These two reactions appear to be very different because CSR results in the deletion of a large segment of DNA from the Ig heavy chain constant region (CH) locus, whereas SHM introduces point mutations in the variable region (V) of the light and heavy chain loci. In spite of the clear dissimilarity of the products, CSR and SHM share several characteristics in contrast to the other genetic alteration in the Ig locus—that is, V(D)J recombination, a site-specific recombination that is precisely programmed and regulated during lymphocyte development prior to antigen stimulation. CSR and SHM take place only when B lymphocytes encounter antigens or polyclonal stimulators; the targets of these genetic alterations are not programmed, but instead depend on environmental factors such as cytokines. Most important, CSR and SHM require activation induced cytidine deaminase (AID) (Muramatsu et al., 2000; Revy et al., 2000), whereas V(D)J recombination is catalyzed by RAG-1, and -2. Metaphorically, we can consider V(D)J recombination as the basic education of soldiers, and CSR and SHM as the on-site training at the battlefield. Extensive analyses of DNA sequences surrounding CSR junctions revealed that CSR is a unique type of recombination. Unlike V(D)J and homologous recombinations, CSR does not appear to require specific primary sequences or homologous sequences between joining pairs. CSR is recognized as a region-specific recombination because it occurs within a unique type of repetitive sequence, the switch (S) region, that spans a few kb 5¢ to each CH gene (Kataoka et
Molecular Biology of B Cells
OUTLINE OF MECHANISM FOR CSR S-S Recombination with Looped-Out Deletion of CH Genes In 1978, a strong correlation was found between the pattern of CH gene deletion and the Ig isotype expressed in various lines of murine myeloma cells. This led to the proposal of a deletion model for CSR (Honjo and Kataoka, 1978). Subsequent studies using cloned CH genes convincingly proved the deletion model (Cory et al., 1980; Maki et al., 1980; Rabbitts et al., 1980; Yaoita and Honjo, 1980). The deletion model predicted a specific order of mouse CH genes
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to explain the CH gene deletion profile in a given isotypeexpressing myeloma (Honjo and Kataoka, 1978). The proposed order of mouse CH genes was proven directly by molecular cloning of the entire CH gene locus (Shimizu et al., 1981; Shimizu et al., 1982). Molecular cloning and analyses of recombination junctions revealed the presence of repetitive sequences, called the S regions, 5¢ to each CH gene except for Cd (Davis et al., 1980; Dunnick et al., 1980; Kataoka et al., 1980; Sakano et al., 1980). The majority of CSR takes place between Sm and one of the other S regions. Thus, CSR occurs by S-S recombination, resulting in looping out and excision of the DNA segment located between the recombining S regions. Isolation of excised DNA segments as a circular DNA further consolidated this model (Figure 20.1) (Iwasato et al., 1990; Matsuoka et al., 1990; von Schwedler et al., 1990).
Common Features of S Regions The S regions were originally defined as functional regions where junctions of CSR are located (Davis et al., 1980; Dunnick et al., 1980; Kataoka et al., 1980; Sakano et al., 1980). They consist of a tandem array of repetitive units
C
that are clustered relatively densely in the central region and sparsely scattered in the peripheral regions, without clear borders (Kataoka et al., 1981; Nikaido et al., 1981; Nikaido et al., 1982). In mouse and humans, each of the S regions spans from 1 to 9 kb within introns preceding the CH genes (Nikaido et al., 1982; Shimizu et al., 1982; Takahashi et al., 1982). The tandem repeat units of S regions are of variable lengths and contain scattered palindromes. Characteristic features of S regions in other vertebrate species, such as birds and frogs, are summarized in Table 20.1. The functional requirement of S regions for CSR has been demonstrated by deletion of the S region from mouse endogenous Ig loci or artificial CSR substrates. In one study, deletion of the Sm core region (entire block of tandem pentameric motifs) reduced CSR by 1/2 to 1/8, although it did not entirely eliminate it (Luby et al., 2001). It is possible that the thirteen remnant S region motifs in the Sm periphery account for the residual CSR activity. In another study, a 12kb segment containing the Sg1 region was deleted, resulting in complete abolition of switching to IgG1 (Shinkura et al., 2003).
Organization of S Regions and CH Genes CSR has been found in mammals, birds, and frogs. The organization of IgH genes is very similar among these species, in which several CH genes are arrayed downstream of the VH gene cluster. This so-called “translocon-type” configuration (Figure 20.2a) is a feature of Ig genes that undergo CSR. In contrast, cartilaginous fish have the cluster-type IgH gene configuration that is composed of clusters of a unit that contains a pair of VH and CH genes (Figure 20.2a). Vertebrate species with the cluster-type Ig locus are unlikely to have class switching. It is not known, however, whether all translocon-type IgH loci can switch; although fugu, a representative bony fish, has the translocon configuration, CSR in the bony fish has not been documented. Mouse
FIGURE 20.1 AID regulates CSR, SHM, and gene conversion. Mouse IgH locus after completion of VDJ recombination is shown at the top. In the periphery, B cells are stimulated with antigen and undergo additional genetic alterations. V genes are further diversified by SHM and/or by gene conversion. The CH region exons expressed in association with the VH gene are switched from Cm to one of the downstream C genes (Ce in the figure) by CSR. The intervening segment is looped out as a circular DNA. These three DNA alterations depend on transcription of the target gene segments and expression of AID, both of which are induced by cytokines and other stimulations. (Inset) shows structure of germline transcripts. All CH genes but Cd are preceded by an I exon and an S region (oval). Upstream of the I exon is a promoter (arrow) responsive to specific cytokines. The recombination target is determined by the activation of the specific germline promoter. After transcription of the germline transcript, the S region is removed by splicing from mature germline transcripts.
Mouse CH genes span approximately 200 kb at the distal region of chromosome 12. Eight CH genes are arrayed in the following order with intervals indicated in parentheses: JH(6.5 kb)-Cm-(4.5 kb)-Cd-(55 kb)-Cg3-(34 kb)-Cg1-(21 kb)Cg2b-(15 kb)-Cg2a-(14 kb)-Ce-(12 kb)-Ca (Shimizu et al., 1981; Shimizu et al., 1982). Each CH gene is composed of three or four exons for the secretory form and two additional exons for the membrane-spanning form. Alternative splicing and alternative transcription termination regulate the balance between secretory and membrane-bound forms, as well as that between Cm and Cd expression (Alt et al., 1980; Early et al., 1980; Kemp et al., 1980; Rogers et al., 1980; Dariavach et al., 1991). There is a strong enhancer (Em) 5¢
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20. Molecular Mechanism of Class Switch Recombination
TABLE 20.1 Summary of S regions Base usage (%) *2
species human
mouse
isotype
length (kb) *1
unit length (bp)
A
C
G
T
GenBank accession no. *3
Sm
3.5
80
17
21
43
20
(X54713); X56795
Sg3
1.5
79
21
23
50
6
U39935; D78345
Sg1
2.3
79
19
22
52
7
(X17676); U39737
Sa1 Sg2 Sg4
2.5 0.9 0.7
— 79 79
11 19 18
20 23 18
47 51 58
21 7 5
Se Sa2
1.5 2.5
— —
19 —
19 —
38 —
23 —
L19121 U39934 X56796; Y12547; Y12548; Y12549; Y12550; Y12551 X56797 —
Sm Sg3 Sg1
3.5 2 6.5
20 49 49
15 12 19
15 14 17
50 57 45
20 16 17
(J00440-2) D78343 D78344
Sg2b
3.8
49
21
15
45
19
D78344
Sg2a
1.7
49
18
12
40
16
D78344
Se
2.5
40
8
17
47
22
M57385
Sa
4
80
18
14
46
23
D11468
references (Takahashi et al., 1980; Mills et al., 1990) (Mills et al., 1995; Akahori and Kurosawa, 1997; Pan et al., 1997) (Milili et al., 1991; Mills et al., 1995) (Islam et al., 1994) (Mills et al., 1995) (Mills et al., 1990; Pan et al., 1998)
(Mills et al., 1990) (Nilsson et al., 1991) (Nikaido et al., 1981) (Szurek et al., 1985) (Kataoka et al., 1980; Mowatt and Dunnick, 1986; Akahori and Kurosawa, 1977) (Kataoka et al., 1981; Akahori and Kurosawa, 1997) (Nikaido et al., 1982; Akahori and Kurosawa, 1997) (Nikaido et al., 1982; Scappino et al., 1991) (Arakawa et al., 1993)
pig
Sm
3.2
—
13
18
45
23
U50149
(Sun and Butler, 1997)
chicken
Sm1 Sm2 Sg(u)
3.7 1.4 —
101 100 40
21 30 25
18 40 15
36 20 43
22 10 17
AB029075 AB029075 (AB029077)
(Kitao et al., 2000) (Kitao et al., 1996) (Kitao et al., 2000)
duck
Sm Sg(u) Sa
2.5 3.1 2.9
20 17 20
20 21 24
31 13 32
33 47 23
17 20 21
AJ314754 AJ314754 AJ314754
(Lundqvist et al., 2001) (Lundqvist et al., 2001) (Lundqvist et al., 2001)
frog
Sm Sc Su
5 — —
145 121 —
30 26 —
16 22 —
23 12 —
31 39 —
AF002166 (AF002167) —
(Mussmann et al., 1997) (Mussmann et al., 1997)
*1 Length polymorphism is reported to exist among strains and individuals. *2 Base composition was calculated based on repeat consensus motifs except human Sa1 and Se and duck S sequences. Complementary sequence of duck Sa was used based on putative transcriptional orientation. *3 Accession numbers in parentheses indicate that the entry sequence is partial. — Not available.
to the Sm region (Banerji et al., 1983; Gillies et al., 1983; Neuberger, 1983) (Figure 20.2b). Downstream of the Ca gene is a large complex transcriptional enhancer (3¢ IgH enhancer), which spans 23 kb. It is believed that this enhancer complex is a locus-control region that regulates expression of the entire IgH locus (Pettersson et al., 1990; Dariavach et al., 1991; Lieberson et al., 1991; Khamlichi et
al., 2000). It contains five DNaseI-hypersensitive sites called HS1, HS2, HS3a, HS3b, and HS4. Transfection experiments have identified independent enhancer activity in B cells for the hypersensitive sites, each of which is confined within approximately a 1-kb region, and they have a synergistic effect when combined (Madisen and Groudine, 1994; Chauveau et al., 1998).
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tion unit contained the 3¢ enhancer (Chen and Birshtein, 1997; Mills et al., 1997; Pinaud et al., 1997). Chicken, Frog, and Bony Fish
FIGURE 20.2 Organization of IgH C genes. (a) Scheme of cluster- and translocon-type organization. (b) Organization of Ig heavy chain loci of various species. V and C genes are represented by open and closed boxes, respectively. S regions and enhancers are indicated by open and hatched ovals, respectively. Transcription proceeds from left to right except for bird Ca, whose orientation in the nonrearranged genome is right to left, as indicated by an arrow. In human, two pseudogenes (ye and yg) are shown. The cluster-type of IgH locus has only been found in cartilaginous fish. The diagrams are not to scale.
The organization of the IgH loci in chicken and duck were recently described (Zhao et al., 2000; Lundqvist et al., 2001) (Figure 20.2b), and CSR in the chicken IgH locus has been reported (Kitao et al., 2000). Three CH genes in the order Cm, Ca, and Cu (or Cg) that encode the C regions of IgM, IgA, and IgY (IgG), respectively, are present in a relatively compact region (67 kb). A remarkable feature of the bird IgH loci is that the transcriptional orientation of the Ca gene is inverted with respect to that of the Cm and Cu. Consequently, switching to IgA results in inversion of a large DNA segment containing the Cm and Ca genes. Frog is the most primitive species that switches isotype by recombination (Mussmann et al., 1997). Frogs have three Ig isotypes called IgM (m), IgX (c), and IgY (u), which may be equivalent to mammalian IgM, IgA, and IgG, respectively. Like mammalian IgH loci, the Cm gene is located immediately downstream of the VH gene cluster, and the other C genes are aligned further downstream (Figure 20.2b). The order and orientation of Cc and Cu are not yet determined. S regions are located 5¢ to the Cm and Cu genes, and their sequences are AT rich, unlike the S regions of other species (Mussmann et al., 1997). Bony fishes seem to be the first species that evolved the translocon-type Ig configuration. CH genes for IgM- and IgD-like isotypes reside in tandem downstream of the VH gene cluster (Aparicio et al., 2002; Bengten et al., 2002; Ventura-Holman and Lobb, 2002) (Figure 20.2b). However, it is unknown whether CSR occurs in bony fish or whether they have S regions. Significant levels of SHM have been demonstrated in frog and both bony and cartilaginous fishes (Wilson et al., 1995; Diaz et al., 1999; Lundqvist and Pilstrom, 1999; Lee et al., 2002; Oreste and Coscia, 2002).
Human The organization of the human CH genes is similar to that of their mouse counterpart (Figure 20.2b). The CH genes are located at the telomeric end of chromosome 14, and there are no other genes between the telomeric repeats and the Cm gene except for the VH gene cluster that spans one Mbp (Matsuda et al., 1998). The CH genes are arranged in the following order: JH-(8 kb)-Cm-(5 kb)-Cd-(60 kb)-Cg3(26 kb)-Cg1-(19 kb)-Cye-(13 kb)-Ca1-(35 kb)-Cyg-(40 kb)Cg2-(18 kb)-Cg4-(23 kb)-Ce-(10 kb)-Ca2 (Hofker et al., 1989). Duplication of a unit consisting of Cg-Cg-Ce-Ca and the presence of two pseudogenes are the unique features of the human IgH locus (Takahashi et al., 1982). There are S regions upstream of each CH gene, except for Cd and Cyg. There are two 3¢ enhancers, one each downstream of the Ca1 and Ca2 genes, indicating that the primordial duplica-
Unique Properties of CSR as Compared with Other Recombination Reactions CSR is unique among DNA recombinations because it takes place between repetitive sequences without homology or specific nucleotide sequence. The recombination junctions occur at sites with little or no homology between the upstream (or donor) and downstream (or acceptor) S regions (Kataoka et al., 1981; Nikaido et al., 1982; Gritzmacher, 1989; Dunnick et al., 1993). Occasional inversions of S regions following recombination are observed (Obata et al., 1981; Greenberg et al., 1982; Yancopoulos et al., 1986; Schrader et al., 2002). Little or no junctional microhomology is typical of a recombination reaction known as nonhomologous end-joining (NHEJ) (Roth and Wilson, 1986; Merrihew et al., 1996).
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The second unique feature is the presence of frequent nucleotide mutation, surrounding the CSR junction and extending for about 200-bp from the junction (Nikaido et al., 1982; Dunnick et al., 1989; Dunnick and Stavnezer, 1990; Dunnick et al., 1993; Du et al., 1997; Lee et al., 1998). The majority of these mutations are substitutions, but occasional deletions and insertions can be found as well. Since NHEJ is often accompanied by extensive end-processing and repair of DNA synthesis around the junction (for up to 1 kb or more) (Henderson and Simons, 1997), the presence of mutations is consistent with a NHEJ-type recombination mechanism (Lehman et al., 1994). CSR is also unique because the S regions undergoing recombination must be transcribed (Stavnezer-Nordgren and Sirlin, 1986; Yancopoulos et al., 1986; Severinson et al., 1990), and the efficiency of CSR is quantitatively correlated with the rate of S region transcription (Stavnezer et al., 1988; Lee et al., 2001). This will be discussed later and also in detail in Chapter 19. Finally, CSR is a unique type of recombination in its complete dependence on AID, a member of the cytidine deaminase RNA editing enzyme family (Muramatsu et al., 2000; Revy et al., 2000). The mode of action of AID is a matter of intense debate and will be discussed later.
Artificial Substrates to Dissect Molecular Mechanism of CSR To dissect the molecular mechanism of CSR, several groups have described experimental systems using artificial substrates that recapitulate several aspects of the endogenous IgH locus (Ott et al., 1987; Leung and Maizels, 1992; Lepse et al., 1994; Daniels and Lieber, 1995b; Ballantyne et al., 1997; Kinoshita et al., 1998; Christine et al., 1999; Stavnezer et al., 1999; Okazaki et al., 2002; Zhang et al., 2002). The stably transfected switch plasmid substrates reported by Kinoshita et al. (1998) have proven to be a valuable model system to assay CSR activities (Okazaki et al., 2002). Recombination in these plasmids is dependent on cytokine stimulation and requires AID activity. The plasmid substrate contains Sm and Sa sequences transcribed from a constitutive promoter and a tetracycline-inducible promoter, respectively (Figure 20.3a). Both S sequences are removed by splicing of the transcripts, similarly to endogenous germline transcripts. The extracellular and trans-membrane domains of mouse CD8a are located upstream of Sm and downstream of Sa, respectively, and recombination between Sm and Sa will allow expression of the extracellular and transmembrane domain sequences on the same transcript. As a result, cells harboring the post-switch substrate are able to express membrane-bound CD8a, which can be quantitated by flow cytometry. This substrate has been shown to undergo AID-dependent recombination in immortalized fibroblasts, and the recombination
FIGURE 20.3 Artificial substrate of class-switch recombination. (a) A chromosomally integrated substrate of class-switch recombination (CSR) (Tashiro et al., 2001; Lee et al., 2001; and Okazaki et al., 2002). Two S regions (Sm and Sa) are transcribed by independent promoters A and B (closed and gray arrows). Extracellular (EC) and transmembrane (TM) domains of CD8 are located upstream of Sm and downstream of Sa, respectively. Green fluorescent protein (GFP) coding sequence is fused to the TM exon. Splicing removes S sequences from the transcripts. The neomycinresistance gene (NeoR) is used to select stable transfectants. Upper and lower diagrams show the substrate structure before and after CSR, respectively. (b) FACS profile of mouse B lymphoma cell line (CH12F3-2) harboring a single copy of the artificial substrate before (left) and after (right) cytokine stimulation to induce CSR. CD8-GFP–positive cells represent cells that underwent CSR on the artificial substrate. (c) Inversion-type substrate developed by Chen et al. (2001). Upper and lower diagrams show the substrate before and after CSR, respectively.
junctions in these cells have similar features (no homology, mutations, and deletions) to endogenous switch junctions (Okazaki et al., 2002). A series of experiments using this system have revealed the following: 1) the S region is essential for recombination; 2) an inverted S region sequence is active; 3) the S region can be replaced by an artificial G-rich palindromic sequence; 4) the G-rich sequences of telomeres are inactive; 5) the AT-rich frog S region is active; and 6) inversion-type CSR can take place in the plasmid (Kinoshita et al., 1998; Chen et al., 2001; Tashiro et al., 2001).
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ISOTYPE SPECIFICITY OF CSR Germline Transcription and the Role of Germline Transcripts Since CSR occurs after activation of B cells by antigen, it can be directed to the isotype that is best suited to clear the pathogen that induced the immune response. A fundamental mechanism for determination of isotype specificity is the regulated transcription of unrearranged CH genes, although the roles of these germline, or sterile, transcripts are still unknown (Stavnezer, 2000). Numerous studies have established that isotype choice is determined by the regulation of transcription from a promoter located 5¢ to each S region by cytokines such as IL-4, interferon-g, and TGF-b1, produced by helper T cells and other cell types, along with B cell activators such as CD40 ligand and LPS (Stavnezer, 2000). Germline transcription initiates upstream of the S region at the I exon and continues through the CH gene, terminating at the poly(A) sites. Germline transcripts are spliced and polyadenylated. The I exon is spliced to the normal splice acceptor for the CH gene, resulting in deletion of the S region sequences from mature germline transcripts (Figure 20.1). The regulation of germline transcripts is discussed more fully in Chapter 19. Although they were first identified more than 15 years ago, the role of germline transcripts is still a mystery. Several studies creating targeted deletions or mutations of the I exons and their promoters demonstrated that S-region transcription in cis is required for switching (Jung et al., 1993; Zhang et al., 1993; Bottaro et al., 1994; Lorenz et al., 1995). The appearance and quantity of germline transcripts correlates with the timing and amount of CSR, suggesting that the efficiency of transcription or the amount of transcripts correlates with the efficiency of CSR (Stavnezer et al., 1988; Lundgren et al., 1995; Qiu and Stavnezer, 1998; Lee et al., 2001). The primary sequence of the germline transcript is not important, as it can be replaced by a HPRT mini-gene (Harriman et al., 1996), by a viral sequence (Hein et al., 1998), or by CD8 (Okazaki et al., 2002). Most interestingly, CSR is strongly inhibited by the ablation of a splice donor site upstream of the S region (Lorenz et al., 1995; Hein et al., 1998). This inhibition of CSR is likely due to the absence of splicing, but not to a secondary effect, such as reduced transcriptional activity, since abundant unspliced transcripts from the targeted allele were detected in cells that could not undergo CSR to IgE due to a deletion of the Ie exon and splice donor site (Bottaro et al., 1994). Lee et al. (1998) found that recombination breakpoints in the 5¢ Sm flanking region do not extend upstream of the 3¢ end of the Im exon. It is possible that this boundary of CSR breakpoint distribution is associated with requirements for splicing of the germline transcript.
The requirement for a splice donor suggests the involvement of either spliced products or the splicing machinery in CSR.
Isotype Specificity and S-Region Accessibility V(D)J recombination is regulated by chromatin accessibility (Kwon et al., 2000). Although this is likely to be true for CSR as well, very little data are available on the regulation of S-region accessibility beyond the fact that transcription of the S region is required for CSR. Interestingly, transcription of the CSR target does not appear to be required simply to remodel chromatin structure; histone H3 on a chromosomally integrated mini-switch substrate is acetylated prior to transcription, yet its transcription is essential to CSR (Lee et al., 2001). The DNA segment surrounding the Sa region and the Ca gene in the I.29m B cell line, an IgM+ B lymphoma capable of undergoing CSR to IgA, was shown to be hypomethylated, whereas the Cg1 and Cg2b DNA segments, to which this cell line cannot switch, are hypermethylated (Stavnezer-Nordgren and Sirlin, 1986). In addition, splenic B cells induced to switch to IgG1 by treatment with LPS and IL-4, show DNase hypersensitive sites upstream of and within the Sg1 segment (Schmitz and Radbruch, 1989; Berton and Vitetta, 1990). A hypersensitive site within the Sa region is induced in CH12F3-2 lymphoma cells treated with LPS+IL-4+TGF-b1 to induce switching to IgA (Ono et al., 2000). A few activating transcription factors are known to bind S regions, and it was proposed that they regulate S-region accessibility in a sequence and therefore, isotype-specific manner. NF-kB and a complex containing the E2A protein E47 have been shown to bind the Sg3 consensus repeat sequence (Wuerffel et al., 1992; Kenter et al., 1993; Ma et al., 1997; Wuerffel et al., 2001). Interestingly, mice deficient in NF-kB proteins show isotype-specific CSR defects that are not explained by lack of germline transcripts (Snapper et al., 1996; Zelazowski et al., 1997). However, since little is known about isotype-specific binding of NF-kB to S regions, it is unknown if the isotype-specific CSR defects are consistent with S-region binding patterns. Regulation of CSR by E2A activity can be explained by a combination of its effects on the I promoter activity (Sugai et al., 2003) and transcription of the AID gene (Gonda et al., 2003). Although experiments using transiently transfected switch substrate plasmids with different S regions indicate that the S-region sequences themselves can contribute to isotype specificity (Shanmugam et al., 2000; Ma et al., 2002a), experiments by other investigators do not support this notion, because they found that replacement of Sa region sequences with other S regions or with artificial sequences did not affect CSR (Kinoshita et al., 1998; Tashiro et al., 2001; Shinkura et al., 2003).
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AID, THE SOLE B CELL-SPECIFIC FACTOR REQUIRED FOR CSR Isolation, Structure, and Role of AID AID was first identified by cDNA subtractive hybridization between cDNA libraries of CH12F3-2 cells, unstimulated or stimulated in vitro to undergo CSR (Muramatsu et al., 1999). Expression of AID mRNA is upregulated in CH12F3-2 cells more than fourteen-fold during stimulation to induce CSR. AID transcripts are observed only in activated B cells and in B-cell lines representing mature B cells, but not in highly differentiated antibody-secreting cells. In vivo, the strongest expression of AID mRNA is seen in germinal centers of peripheral lymphoid organs. Thus, expression of AID transcripts is closely correlated with the anatomical location and timing of CSR. The AID mRNA encodes a small protein of 198 amino acids that has 34% amino-acid identity to an RNA editing cytidine deaminase, APOBEC-1 (Smith and Sowden, 1996). The deaminase activity of APOBEC-1 converts cytosine to uracil at the position 6,666 of mRNA encoding apolipoprotein (apo) B100, a component of low density lipoprotein. This changes a glutamine codon (CAA) to a termination codon (UAA), thereby giving rise to an alternative mRNA species that encodes a truncated form of apoB100 called apoB48, a component of chylomicron. Most importantly, APOBEC-1 does not recognize its mRNA target in the absence of a partner protein. Recombinant AID can deaminate monomeric deoxycytidine in vitro to a degree similar to that of APOBEC-1. In addition, the genes for APOBEC1 and AID are located only 15 kb apart on mouse chromosome 6 and 916 kb apart on human chromosome 12p13, and their exon–intron organizations are almost indentical (Muto et al., 2000). Recently, related family members were identified on human chromosome 22 (Jarmuz et al., 2002). Because of these similarities, AID was proposed to be an RNA editing cytidine deaminase (Muramatsu et al., 2000). Evidence from gain- and loss-of-function studies indicates that AID is essential for CSR. When AID is exogenously expressed in CH12F3-2 cells, which constitutively synthesize germline transcripts through the Sa region, these cells undergo CSR without stimulation to induce CSR (Muramatsu et al., 2000). AID-deficient mice cannot produce any Ig isotype except IgM and IgD. Moreover, CSR is not detected in AID-deficient B cells stimulated in vitro, although they can proliferate normally and express germline transcripts of all isotypes in response to cytokine stimulation (Muramatsu et al., 2000). Human AID deficiency (hyper-IgM syndrome type II) has the same phenotype as AID deficiency in mouse (Revy et al., 2000). Another surprising phenotype revealed by the studies on AID deficiency in mouse and human is an almost complete loss of SHM (Muramatsu et al., 2000; Revy et al., 2000). The results
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demonstrate that AID is required for both CSR and SHM. Later, AID was also shown to be required for yet another type of genetic alteration, gene conversion, which introduces point mutations into V genes by recombination with pseudo V genes in chicken and rabbit B cells. AID-deficient DT40 chicken B cells do not undergo gene conversion, but this activity was restored by reintroduction of AID (Arakawa et al., 2002). When AID is ectopically expressed, CSR and SHM are induced in non-B cells such as fibroblasts (Martin et al., 2002; Okazaki et al., 2002; Yoshikawa et al., 2002). On the other hand, no switching was observed when a mutant AID lacking the cytidine deaminase domain was expressed in these cells. These results demonstrate that all factors required for CSR and SHM, except AID, are constitutively expressed in non-B cells. Therefore, AID is the only B cell–specific factor that is essential for CSR and SHM.
Molecular Mechanism for AID Function Although it is clear that AID has cytidine deaminase activity, it is still not yet resolved whether AID functions by editing a precursor mRNA for an essential factor such as endonuclease or by direct deamination of dC in DNA. Recently, several experiments supporting either the RNA editing or DNA editing hypothesis have been reported. Honjo’s group tested whether CSR requires de novo protein synthesis in addition to AID itself (Doi et al., 2003). They reasoned that if AID edits an unknown precursor mRNA, the edited mRNA must be translated into a new protein species that does not exist prior to activation of AID. By contrast, if AID acts to deaminate DNA directly, de novo protein synthesis should not be required. Doi et al. (2003) developed a new strategy to examine this possibility, involving the construction of a fusion protein of AID with the estrogen receptor (ER) hormone-binding domain (AID-ER). Since AID-ER has no activity in the absence of the estrogen analog tamoxifen, AID-ER can be accumulated in cells without showing any activity. After tamoxifen addition, the AID function of AID-ER is induced, and this activity can be monitored by assaying CSR. When the AID-ER construct was expressed in aid-/- spleen B cells by retroviral infection, CSR was detectable by the digestion-circularization PCR (DC-PCR) method within 1 hour after tamoxifen addition. If cycloheximide or puromycin was added 1 hour prior to tamoxifen, CSR was drastically inhibited when assayed 6 hours later by DC-PCR. Since the addition of protein synthesis inhibitors did not reduce levels of the AID-ER protein or germline transcripts, AID activity appears to depend on de novo protein synthesis, thus supporting the RNA editing hypothesis. Although one cannot exclude the possibility that CSR requires an additional protein that is rapidly degraded during cycloheximide treatment, the authors consider that this seems somewhat unlikely since CSR was only margin-
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ally inhibited by the protein synthesis inhibitors if they were added 1 hour after tamoxifen. Nonetheless, the interpretation of inhibitor experiments always requires some reservation. Neuberger’s group reported data supporting the possibility that AID deaminates dC in DNA directly and proposed a model explaining how to introduce DNA cleavage: AID deaminates dC to dU, followed by removal of the dU base by UNG uracil-DNA glycosylase and cleavage of the phosphodiester backbone by AP endonuclease, the major players in the base excision repair machinery (Di Noia and Neuberger, 2002; Petersen-Mahrt et al., 2002; Rada et al., 2002). Deamination and cleavage would occur on both DNA strands and thereby create staggered breaks that could initiate CSR. They found that expression of AID in Eschenichia coli can augment the endogenous mutation rate by 2- to 45fold, depending on the gene examined (Petersen-Mahrt et al., 2002). This mutation rate was synergistically enhanced in E. coli lacking UNG uracil-DNA glycosylase. However, deamination of dC bases in DNA is not unique to AID, as their subsequent studies showed a greater enhancement of mutations by expression of APOBEC-1 and its family members in E. coli (Harris et al., 2002). In addition, when APOBEC-1 is expressed in fibroblasts or B lymphocytes, no augmentation or induction of CSR or SHM was observed (Eto et al., 2003). These data raise the possibility that the DNA deamination activity of AID is unique to prokaryotes. Rada et al. (2002) also found that CSR activity is reduced about ten-fold in B lymphocytes lacking UNG. The fact that UNG is required for normal levels of CSR fits well with the direct DNA deamination model. Recently, a human hyperIgM patient with mutations in the UNG gene has been found to display an identical phenotype to the mice (Imai et al., 2003). To explain the residual CSR activity in UNGdeficient B cells, Rada et al. (2002) suggested two possibilities: 1) other uracil-DNA glycosylases, besides UNG, may remove dU bases and 2) DNA breaks may be created by recognition of dU/dG mismatches by mismatch repair (MMR) proteins. As will be discussed below, MMR has been shown to contribute to CSR (Ehrenstein and Neuberger, 1999; Schrader et al., 1999; Vora et al., 1999; Ehrenstein et al., 2001). It is not known, however, if Msh2/Msh6 binds dU/dG mismatches efficiently, as the available data are contradictory (Hughes and Jiricny, 1992; P. Gearhart, personal communication). Although UNG deficiency does not reduce the frequency of SHM, it does increase the ratio of transition to transversion mutations, as predicted if UNG were to remove dU bases created by AID activity (Rada et al., 2002). Taken together, there is no question about the involvement of UNG in CSR and SHM. However, it is possible, to explain the effect of UNG deficiency on CSR by other models. For example, the recruitment of other repair
enzymes to SHM and CSR reaction sites may depend on UNG. It is important to determine whether DNA cleavage per se is affected in UNG-deficient B cells. In addition, indirect effects due to loss of one repair system have to be excluded. At this time, no direct data prove or exclude either the RNA editing or DNA editing hypothesis. Additional in vivo and biochemical experiments are essential to test whether AID is an RNA or DNA deaminase and to identify its putative partner protein(s).
CLEAVAGE OF THE S REGION Evidence for Involvement of AID in Cleavage A major goal for researchers in the field of CSR is to identify the step at which AID functions in CSR. The most appealing hypothesis is that, whether AID works directly or via modification of an mRNA, its role is to create DNA breaks in order to initiate CSR. The available data, although not yet conclusive, are consistent with this hypothesis. Petersen et al. (2001) have found that the special form of histone, g-H2AX, and the DNA repair protein, Nbs-1, which are known to be associated with DNA breaks, accumulate to form foci on the CH locus of splenic B cells induced to undergo CSR. By contrast, AID-deficient splenocytes stimulated to undergo CSR failed to form such foci. Another line of evidence indicates that DNA lesions giving rise to nucleotide substitutions depend on AID activity. Mutations are introduced into the Sm segment on unrecombined alleles in B cells treated with LPS + IL-4, but are not found in similarly treated AID-deficient B cells (Petersen et al., 2001; Nagaoka et al., 2002). Likewise, Dudley et al. (2002) found that the Sm segments in IgM hybridomas from wildtype mice, but not from aid-/- mice, often show internal deletions and nucleotide substitutions. Further evidence for AID involvement in DNA cleavage, although indirect, comes from the following observation. If AID were involved in a step downstream of cleavage, the CSR substrate in fibroblasts should be cleaved prior to AID expression, because AID is the only factor missing in fibroblasts (Okazaki et al., 2002). However, the CSR substrates in transfectants are stable, thus supporting the hypothesis that AID is required for DNA cleavage.
Evidence for Staggered Nick Cleavage Since CSR occurs by an intrachromosomal deletion, it requires two double-strand breaks (DSBs), one in Sm and the other in the downstream, acceptor, S region. Three possible mechanisms generate DSBs. The simplest is the single event of double-strand cleavage by an endonuclease. Another mechanism is separate nick cleavages on each of the two
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DNA strands, that is, staggered nick cleavages, which would yield either a 5¢ or 3¢ single-strand tail at the cleaved end. The third possibility is a nick cleavage, followed by trans-esterification; the mechanism used by RAG enzymes during V(D)J recombination (McBlane et al., 1995). This last reaction produces a DSB with one blunt end and one hairpin end. Among these, only the staggered nick cleavage requires the repair process to provide a blunt end by either DNA synthesis or exonuclease–endonuclease digestion. Consequently, either duplication or deletion occurs at the recombination junction, and this can be identified by comparing two recombination products of a single CSR event. Until recently, it was impossible to compare all four cleaved ends of a single CSR event, because the excised looped-out circular DNA carrying one recombination junction is lost from the chromosome. To examine all four cleaved ends, Chen et al. (2001) developed switch plasmids that recombine by inversion, allowing retention of the DNA segment normally deleted during CSR (Figure 20.3c). Extensive sequence analyses of inversion-type CSR products revealed that a significant number of duplications occur at the junction. Among 82 junctions they identified five duplications of 9 to 266 bp. Duplication was directly tandem at the recombination junction, thus strongly indicating the involvement of staggered nick cleavage in CSR. Deletions occurred at the junction very frequently (64 out of 82 junctions), which is also consistent with staggered nick cleavage, although deletions can also be explained by frequent double-strand cleavages. Although DSBs have been detected by LM-PCR in splenic B cells induced to switch (Wuerffel et al., 1997), these experiments did not demonstrate that the blunt-end cleavage is the initial cleavage product. It is possible that blunt-ended DSBs are generated after the repair processing of staggered ends. Moreover, it is not known whether the observed DSBs are intermediates of CSR or by-products of cleavage events during activation of B cells.
Recognition of Secondary Structures Efforts to identify specific sequences for recognition by CSR recombinase have been unsuccessful. Nonetheless, S regions are required for optimal CSR. Extensive sequencing studies of switch recombination junctions have revealed that the majority of the recombination breakpoints are localized within the S region, although a number of breakpoints are also found outside the S region, in both 5¢ and 3¢ flanking regions. Because mammalian S regions have G-rich motifs with stretches of three or four guanines, it was speculated that the ability of G-rich sequences to form a G-quartet structure (Sen and Gilbert, 1988) similar to telomere sequences, may contribute to their recognition by CSR recombinase. The identification of two frog S regions, which
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contain AT-rich sequences (Mussmann et al., 1997), indicates that even if S regions do indeed form this structure, it is not required for CSR. It has been shown that RNA–DNA hybrids are formed in vitro by the active transcription of S regions, and such a structure was suggested to be recognized by CSR recombinase (Reaban and Griffin, 1990; Reaban et al., 1994; Daniels and Lieber, 1995a; Tian and Alt, 2000; Mizuta et al., 2003). However, it is still controversial whether the RNA–DNA hybrids of S regions are formed in vivo and play a critical role in CSR. Mussmann et al. (1997) noted the presence of short palindromic tetramers in Xenopus S regions and predicted they could form stem-loop structures. Interestingly, they showed that S-S recombination junctions in frog Ig genes can be mapped to the vicinity of transitions between singlestranded regions to double-stranded regions on the predicted stem-loop structure. They found a similar association upon analysis of mammalian S-S junctions. The association of CSR junctions and stem-loop structures was also observed in a study using stably transfected CSR substrates in the murine CH12F3-2 B lymphoma cell line (Tashiro et al., 2001). It was shown that a G-rich palindromic sequence consisting of tandem restriction sites, which can form secondary structures inserted in place of the Sa segment in a switch plasmid, supported recombination on the plasmid at 50% of the level of the intact Sa region. The stem-loop structure model is consistent with several other features of CSR, including the palindromic nature of S regions. The observed correlation between transcription of S regions and CSR efficiency could be due to the fact that increased numbers of RNA polymerase molecules loaded onto S regions might lead to more frequent denaturation of S regions, thus resulting in the formation of more abundant stem-loop structures. As described above, CSR is likely to be initiated by staggered cleavages with variable spacing, which suggests independent recognition and nicking of both strands of the S regions. This is consistent with predictions of stem-loop structures at different positions on the two strands. Nonetheless, no solid evidence exists for the requirement for secondary structures for CSR.
PROCESSING AND JOINING OF DNA ENDS AFTER CLEAVAGE Models for Processing and Joining of DNA Ends During CSR The end-processing, repair, and joining mechanisms for CSR are unknown, but the available data indicate that the mechanisms are similar to those used in NHEJ. As described above, it is most likely that CSR is initiated by staggered breaks, and the donor Sm and acceptor S-region DNA ends
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would need to be processed in order to become suitable for recombination by NHEJ. The single-strand DNA ends have either a 3¢ or 5¢ overhang, and the lengths of the single-strand overhangs will differ, depending on the cut sites (Figure 20.4c). These ends could then be processed by several different mechanisms shown to occur during NHEJ (Roth et al., 1985; Henderson and Simons, 1997). Although the recombination junction during NHEJ usually forms at or near the original site of DNA breakage, extensive processing and repair synthesis can occur on either one or both strands, and on both sides of the junction (Henderson and Simons, 1997). Ends with 5¢ overhangs can be filled by DNA synthesis to create blunt double-strand breaks (Figure 20.4c). The joining of blunt ends may involve the use of a few nucleotides of microhomology. End-joining requires proteins to focus the recombination to the ends and to hold the ends in an appropriate structure. The amount of microhomology normally found at switch junctions is so limited (usually 0 to 3 nucleotides) that its length might be no more than predicted from the shared sequence motifs of the S regions (Dunnick et al., 1993). The repair synthesis involved in end-processing appears to be highly error-prone, as nucleotide substitutions are commonly found in the DNA segments surrounding S-S junctions extending at least 200 nucleotides on either side. The highly error-prone trans-lesion DNA polymerases z, i, and h have been implicated in somatic hypermutation, but it is unknown if they are involved in CSR (Diaz et al., 2001; Zeng et al., 2001; Faili et al., 2002a). Ends with 3¢ overhangs can align without prior DNA synthesis and might initiate recombination by forming a few base pairs with the 3¢ end from the other S region (Figure 20.4c). Again, this process would depend on proteins creating the synapsis. After alignment, DNA synthesis could be initiated from these ends to create the recombination junction. If the 3¢ end of one S region does not share microhomology with the 3¢ end of the other S region, it is possible that an internal alignment site is used, and the resulting flap can be removed by exonuclease or endonuclease activity (Paques and Haber, 1997; Ehrenstein and Neuberger, 1999; Schrader et al., 1999; Wilson and Lieber, 1999; Wu et al., 1999) (Figure 20.4c). Nucleotides that differ from both the donor and acceptor S regions are sometimes (<5% of the time) observed at S junctions. These apparent insertions might be explained by alignment at internal sites without excision of the single-strand tail, resulting in mutations when repaired (Dunnick et al., 1993; Chen et al., 2001; Schrader et al., 2002). It is expected that a complex of proteins, including the recombinase or AID itself, may recognize the secondary structure of the S regions, create the initial cleavages, and then may remain in a complex with the
cleaved S region ends and contribute to synapsis and joining of the DNA ends.
Proteins Involved in DNA Repair, Processing, and Joining of S Regions DNA-PK DNA-dependent protein kinase (DNA-PK) is a serinethreonine protein kinase that is activated by DNA doublestrand breaks (DSBs) and is essential for the normal repair of DNA breaks induced by ionizing radiation, chemical agents, and during V(D)J recombination (Taccioli et al., 1993; Blunt et al., 1995; Taccioli et al., 1998). Although DNA-PK appears to focus recombination to DNA ends, its precise role and mechanism of action are still unknown. Recent data suggest it may be involved in end-processing (Ma et al., 2002b; Rooney et al., 2002). DNA-PK consists of three subunits, the catalytic subunit DNA-PKcs, Ku70, and Ku80, which form a heterodimer and bind to DNA ends, nicks, and transitions between single strands and double strands (Leuther et al., 1999). Mice deficient in any of these genes do not have B or T lymphocytes. To determine the roles of these proteins in CSR, recombined V(D)J gene segments have been introduced into their normal position (“knocked-in”) within the Ig heavy and light chain loci; this allows B cells to develop normally and to have the potential for CSR. The inability of B cells from these mice to undergo CSR proves that Ku70 and Ku80 are essential to CSR (Casellas et al., 1998; Manis et al., 1998). Although these defects might be explained by the fact that without these proteins the DNA breaks induced during CSR cannot be repaired and consequently cause cell death, the investigators demonstrated that viable B cells in cultures also show a great reduction in CSR (M. Nussenzweig, personal communication). Surprisingly, when CSR was examined in B cells from mice entirely lacking DNA-PKcs, it was found the B cells switched only to IgG1, albeit at reduced frequencies (30 to 50% of wildtype cells), but not to other isotypes (Manis et al., 2002a). However, scid mice, containing a kinase-dead DNA-PKcs, switch to all isotypes with 50% efficiency compared with normal mice (Bosma et al., 2002). B cells from both mutant mice proliferate normally. The finding that the phenotypes of the kinase-dead DNA-PKcs and null mutations differ suggests that DNA-PKcs has a function that does not require kinase activity. Mismatch Repair Proteins MMR proteins in eukaryotes fall into two classes: 1) the MutS homologs (Msh1-6), which recognize DNA mismatches, loops, and other distortions; and 2) the MutL homologs
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FIGURE 20.4 Models of AID action. (a) RNA-editing model. AID deaminates cytosine (C) to uracil (U) on mRNA coding for putative class-switch endonuclease (pacman). The active form of endonuclease translated from the edited mRNA will recognize transcription-induced secondary structure and generate staggered nicks on DNA. (b) DNA-editing model. AID deaminates dC in S-region DNA to generate dU, which is either removed by uracil-DNA glycosylase or converted to T after DNA replication, giving rise to C/G to T/A transitions. Abasic sites generated by uracil DNA-glycosylase are cleaved by AP endonuclease (pacman), resulting in staggered nick cleavage of DNA. (c) Two distinct DNA repair pathways leading to SHM and CSR. Repair of staggered nicks of V-region DNA may involve low-fidelity DNA synthesis, giving rise to frequent mutations (triangles). Staggered nicks on two strands of two S regions result in two staggered double-strand breaks, repair of which requires nonhomologous end joining (NHEJ) system. Gap filling by error-prone DNA polymerases (circle) and/or digestion of single-strand overhangs by exonuclease (small pacman) or endonuclease (not shown) occurs before the joining of cleaved S regions, introducing mutations near junctions (triangle). Although SHM and CSR follow distinct repair paths, some mechanisms, such as MMR, are used in common. It is likely that DNA cleaving enzymes and DNA repair enzymes (MMR, NHEJ, and base excision repair) form a complex of mutasome for SHM and of recombinasome for CSR. It is especially important to have anchor proteins for holding two cleaved ends during CSR. Possible components of “mutasome” and “recombinasome” are listed at the bottom.
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(Pms2, Mlh1, and Mlh3 in humans), which bind to MutS homologs bound to DNA and recruit the mutation repair machinery (Kolodner and Marsischky, 1999; Marti et al., 2002). MMR proteins have additional roles besides the correction of nucleotide substitutions and small insertions or deletions created by DNA synthesis errors. In fact, Msh2, Msh6, Mlh1, and Pms2 actually contribute to the process of SHM, because mice lacking one of these genes show slight reductions in mutation frequency (~twofold) and an altered pattern of mutations (Cascalho et al., 1998; Rada et al., 1998; Winter et al., 1998; Wiesendanger et al., 2000). In addition, some MMR proteins have roles in homologous DNA recombination, binding to Holliday junctions and preventing recombination between nonhomologous sequences (Baker et al., 1995; Baker et al., 1996; Edelmann et al., 1999; Marsischky et al., 1999; Nakagawa et al., 1999; Borts et al., 2000; Evans and Alani, 2000; Harfe and JinksRobertson, 2000; Kneitz et al., 2000). MMR also contributes to DSB repair in yeast, which requires the removal of nonhomologous DNA segments adjacent to the break before the break can be repaired by single-strand annealing and gene conversion (Paques and Haber, 1997; Sugawara et al., 1997; Evans et al., 2000). Msh2 and Msh3 are required for this end-processing if 30 nucleotides or more of such heterologous sequences are present, and their role is to recruit an endonuclease complex (Rad1/XPF and Rad 10/ERCC1) to excise the heterologous 3¢ single-strand tail (Paques and Haber, 1997; Sugawara et al., 1997; Evans et al., 2000). By testing the ability of splenic B cells from mice deficient in any one of three MMR proteins (Msh2, Mlh1, and Pms2) to undergo CSR in culture, it was found that MMR proteins are involved, although not essential, for switching in these cultures (Ehrenstein and Neuberger, 1999; Schrader et al., 1999; Ehrenstein et al., 2001). Depending on the particular isotype, switch recombination is reduced by 1/2 to 1/5 of wildtype levels. The proliferation of MMR-deficient B cells in these cultures was comparable to wildtype B cells. Experiments in which the effect of Msh2 deficiency was examined during in vivo immune responses showed a greater deficit in class switching, perhaps due to effects on B-cell selection as a result of reduced SHM (Ehrenstein and Neuberger, 1999; Vora et al., 1999). Sequencing of S-S junctions from Msh2-deficient B cells induced to switch in culture revealed three differences from junctions in wildtype B cells. On the Sm side, the junctions occur entirely within the tandem repeats, and they also occur at a somewhat restricted region within the acceptor Sg3 or Sa regions (Ehrenstein and Neuberger, 1999). Second, the junctions show reduced lengths of microhomology compared to wildtype junctions, as 80% occur at 0 or 1 nucleotide of overlap, compared to 60% in wildtype junctions (Schrader et al., 2002). The third difference is an increase in the numbers of untemplated insertions, or muta-
tions, at the junctions (Schrader et al., 2002). These data seem to be consistent with a role for Msh2 in processing DNA ends, as described above for yeast DSB repair. A recent result seems to further support the endprocessing role for Msh2. Mice lacking both the tandem Sm repeats and Msh2 have a profound deficiency in CSR (Min et al., 2003). The efficiency of switching by stimulated splenic B cells of these mice was 5 to 10% of wildtype B cells, and the difference is greater than expected from the additive effects of the Sm deletion and Msh2-deficiency. The authors propose to explain the results as follows: In the absence of the Sm tandem repeats, target sites for DNA cleavage are rarer, and thus the sites of breaks on the top and bottom strands are more likely to be farther apart, giving rise to ends that might be more dependent on end-processing by Msh2. Switch recombination junctions from Mlh1- and Pms2deficient B cells were also examined and found to differ from junctions in both wildtype and Msh2-deficient cells, suggesting that the MutS and MutL homologs have different functions in CSR (Ehrenstein et al., 2001; Schrader et al., 2002). About 25% of the junctions show greatly increased lengths of microhomology, up to 10 to 14 nucleotides. Because Mlh1/Pms2 heterodimers can bind DNA in the absence of the MutS homologs, and actually cause synapsis of two different molecules (Hall et al., 2001), it is possible that Mlh1 and Pms2 may be involved in providing a structure to and/or in stabilizing the recombination intermediate. In their absence, junctions with longer lengths of microhomology, perhaps due to their increased stability, are favored. Another interpretation is that without Mlh1 or Pms2, an alternate pathway for recombination is used. Since such long identities are never observed at wildtype junctions, the results cannot simply be explained by an increase of one subset of normal junctions. Exonuclease I Several proteins that interact with MMR proteins are recruited to repair mismatches or to perform end-processing. Two of these proteins, exonuclease I and ERCC1, have been shown to be involved in CSR. Exonuclease I excises the patch of DNA containing the mismatch during postreplicative repair (Genschel et al., 2002). Mice deficient in exonuclease I have reduced antigen-specific IgG1 responses with altered S-S junctions that resemble those found in Msh2-deficient cells (A. Martin, personal communication). ERCC1 ERCC1 is an endonuclease that functions as a heterodimer with xeroderma protein F (XPF), cleaves singlestrand 3¢ tails, and is recruited by Msh2/Msh3 during DSB repair in yeast. Recently, mice deficient in ERCC1 have
20. Molecular Mechanism of Class Switch Recombination
been shown to have reduced abilities to undergo CSR and to have altered S-S junctions, with an increase in untemplated insertions, consistent with a role in end-processing (C.E. Schrader, J. Vardo, L.J. Niedernhofer, J.H.J. Hoeijmakers and J.S., in preparation). The data discussed above still leave many unanswered questions about the roles of MMR in CSR. To uncover the roles of MMR in CSR, biochemical studies will be necessary. ATM and Nbs-1 Ataxia-telangiectasia mutated (ATM) is a member of the phosphatidylinositol (PI)-3-kinase gene family, which is activated upon binding to DNA DSBs. Upon activation, it phosphorylates and activates several proteins involved in DNA repair, for example, the Nijmegen breakage syndrome protein (Nbs-1) (Rotman and Shiloh, 1999). A-T and NBS patients show similar phenotypes. ATM functions to sense DNA breaks induced during the S phase and to coordinate the cellular response to cope with the damage (Petrini, 2000; Bakkenist and Kastan, 2003). Mice and humans with mutated forms of these genes have multisystem disorders and are prone to malignancies due to defective DSB repair, although ATM and Nbs-1 are not required for the development of B cells. There are contradictory reports about serum Ig levels in humans and in unimmunized mice with ATM or Nbs-1 deficiency (van Engelen et al., 2001; Pan et al., 2002; Williams et al., 2002). Since serum Ig levels can be regulated at several levels subsequent to CSR, it is possible that CSR itself is reduced. In fact, the Sm-Sm switch junctions from A-T and Nbs-1 patients are clearly abnormal, showing unusually long microhomologies, similar to those found in Mlh1- and Pms2-deficient mouse B cells (Pan et al., 2002). The simplest explanation for why mutations of Mlh1, Pms2, ATM, and Nbs-1 genes all seem to result in S-S junctions with increased lengths of microhomology is to postulate that, in their absence, the end-joining step of CSR occurs by an alternate and perhaps common pathway. g-H2AX Another protein that responds to DNA DSBs is g-H2AX, a special form of histone H2A, which is activated within minutes after induction of DNA DSBs to recruit repair factors (Paull et al., 2000). As described above, foci containing g-H2AX and Nbs-1 co-localize with CH genes in B cells induced to undergo CSR (Petersen et al., 2001). Furthermore, B cells from mice lacking g-H2AX genes have reduced abilities (1/2 to 1/10 relative to normal cells) to undergo CSR. Although these cells showed a reduction in proliferation, the switching defect is not caused by this, because switching was compared between cells that had undergone equal numbers of divisions after activation (Celeste et al., 2002). Mice deficient in g-H2AX have a
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normal ability to undergo V(D)J recombination, although the number of B and T cells is about 50% of wildtype mice, most likely due to the chromosome damage they sustain and the reduced ability of cells to proliferate. DNA Ligase IV The protein complex DNA ligase IV/XRCC4 is essential for NHEJ and for V(D)J recombination (Gellert, 2002). A recent report indicates that DNA ligase IV may also be involved in CSR, because a human patient with a mutant form of this gene was found to have abnormal Sm-Sg junctions in peripheral blood B cells (M.R. Ehrenstein, personal communication). The abnormal junctions showed increased lengths of microhomology. Since this patient has B cells, some ligase IV function must remain, and thus these data do not indicate whether DNA ligase IV is required for CSR.
COMPARISON OF CSR WITH SHM CSR and SHM were previously thought to be mediated by different mechanisms, and the requirement of AID for both events surprised many scientists in the field. However, when we carefully compare the two events, they share several features that are critical to the molecular mechanisms of SHM and CSR. The targets of CSR and SHM must be transcribed before the reaction takes place. The efficiency of these genetic alterations is correlated with the level of transcription. The targets of both events do not have specific primary sequences, yet defined DNA regions are altered. Not only SHM but also CSR is associated with mutations. Most important, they both require AID activity. CSR has important features mechanistically distinct from SHM. Not only do the products of CSR and SHM differ but also the initiation of the two events is dissimilar. CSR requires two DSBs, one each in two different S regions. By contrast, SHM is more likely to be initiated by a single nick on one strand of the target DNA (Faili et al., 2002b). During CSR, two separate DNA ends, which originally can be located 100 kb apart, have to be held in a close proximity. Without proteins that can hold the two recombining ends close together, it is probably impossible to join the DNA ends. It is therefore reasonable to assume that CSR depends on a protein complex that differs from a SHM complex. The CSR complex, which we refer to as “recombinasome,” may contain AID or the putative protein encoded by the mRNA it edits, MMR proteins, base excision repair enzymes, errorprone DNA polymerases, and NHEJ proteins. Although SHM may not require a complex of proteins involved in DNA synapsis, the “mutasome” may contain the DNA nicking enzyme, MMR, base excision repair enzymes, and error-prone DNA polymerases. Additional components in
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these complexes may be required to target them to the correct DNA segments. During evolution, SHM appeared first in cartilaginous and bony fish, whereas these fish do not appear to have CSR (Litman et al., 1999; Flajnik, 2002). Amphibians, which evolved later, were the first to carry out both CSR and SHM. Another indication of the distinct functions of AID for CSR and SHM is found in hyper-IgM type II patients with mutations in AID who have significant levels of SHM, but almost no CSR (Revy et al., 2000). The results can be explained most easily by the possibility that AID interacts with different protein molecules when inducing SHM and CSR. Perhaps, in these patients, proteins required for CSR are unable to associate with the mutated AID, whereas proteins required for SHM interact normally. In addition, CSR and SHM are independently regulated. For example, LPS stimulation of splenic B cells efficiently induces CSR but very rarely SHM. Thus, although a single protein, AID, mediates both CSR and SHM, the two processes evolved stepwise and are regulated differently, although it is possible that in germinal center cells they occur concurrently. Altogether, the data suggest that CSR and SHM are likely to have different molecular mechanisms downstream of AID.
CONCLUSION Extensive studies have revealed several critical features of the molecular mechanism of CSR. First, CSR is dependent on the highly repetitive S-region sequences, occurring anywhere within the repeats, thus being a region-specific recombination. Second, CSR takes place only when the S region is actively transcribed. Therefore, isotype specificity is determined by the regulation of transcription through the S region. Cytokines and B cell activators, such as CD40L and LPS, regulate this transcription through the I exon promoters. Third, CSR is initiated by two nicking cleavages, generating a staggered DSB. Fourth, CSR is completely dependent on AID activity. Fifth, current evidence indicates that the repair of the cleaved ends utilizes the NHEJ repair system. Sixth, other repair proteins such as mismatch repair, base excision repair, and g-H2AX are required for efficient CSR. Taking all these features together, we present a model for CSR in Figure 20.4. Several key issues must be addressed in order to obtain a complete understanding of the mechanism of CSR. It is most important to conclusively determine whether AID itself can deaminate DNA or whether AID edits a mRNA precursor for a cleaving enzyme. This will lead to the identification of the molecular nature of the cleaving enzyme. It is equally critical to identify the components of the putative recombinase complex and to understand how it is targeted to S regions. These findings will almost inevitably lead us
to understand how the molecular mechanism of CSR differs from that of SHM. At this stage, no experimental system recapitulates physiological CSR in a cell-free system, although there have been some trials (Lyon et al., 1996; Borggrefe et al., 1998; Zhang and Cheah, 2000). Once the cleaving enzyme is identified, it may not be difficult to establish an in vitro assay system for the recognition of CSR target structures. Then, additional information should be rapidly obtained by adding or deleting candidate enzymes for involvement in CSR. We expect many fascinating and intriguing observations will be generated in this decade, and these will have a strong impact not only on understanding the mechanism and regulation of CSR and SHM, but also on the mechanisms of recombination in general.
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Tashiro, J., Kinoshita, K., and Honjo, T. (2001). Palindromic but not G-rich sequences are targets of class switch recombination. Int Immunol 13, 495–505. Tian, M., and Alt, F. W. (2000). Transcription-induced cleavage of immunoglobulin switch regions by nucleotide excision repair nucleases in vitro. J Biol Chem 275, 24163–24172. van Engelen, B. G., Hiel, J. A., Gabreels, F. J., van den Heuvel, L. P., van Gent, D. C., and Weemaes, C. M. (2001). Decreased immunoglobulin class switching in Nijmegen Breakage syndrome due to the DNA repair defect. Hum Immunol 62, 1324–1327. Ventura-Holman, T., and Lobb, C. J. (2002). Structural organization of the immunoglobulin heavy chain locus in the channel catfish: The IgH locus represents a composite of two gene clusters. Mol Immunol 38, 557–564. von Schwedler, U., Jack, H. M., and Wabl, M. (1990). Circular DNA is a product of the immunoglobulin class switch rearrangement. Nature 345, 452–456. Vora, K. A., Tumas-Brundage, K. M., Lentz, V. M., Cranston, A., Fishel, R., and Manser, T. (1999). Severe attenuation of the B cell immune response in Msh2-deficient mice. J Exp Med 189, 471–482. Wiesendanger, M., Kneitz, B., Edelmann, W., and Scharff, M. D. (2000). Somatic hypermutation in MutS homologue (MSH)3-, MSH6-, and MSH3/MSH6-deficient mice reveals a role for the MSH2-MSH6 heterodimer in modulating the base substitution pattern. J Exp Med 191, 579–584. Williams, B. R., Mirzoeva, O. K., Morgan, W. F., Lin, J., Dunnick, W., and Petrini, J. H. (2002). A murine model of Nijmegen breakage syndrome. Curr Biol 12, 648–653. Wilson, M., Marcuz, A., and du Pasquier, L. (1995). Somatic mutations during an immune response in Xenopus tadpoles. Dev Immunol 4, 227–234. Wilson, T. E., and Lieber, M. R. (1999). Efficient processing of DNA ends during yeast nonhomologous end joining. Evidence for a DNA polymerase b (Pol4)-dependent pathway. J Biol Chem 274, 23599–23609. Winter, D. B., Phung, Q. H., Umar, A., Baker, S. M., Tarone, R. E., Tanaka, K., Liskay, R. M., Kunkel, T. A., Bohr, V. A., and Gearhart, P. J. (1998). Altered spectra of hypermutation in antibodies from mice deficient for the DNA mismatch repair protein PMS2. Proc Nat Acad Sci USA 95, 6953–6958. Wu, X., Wilson, T. E., and Lieber, M. R. (1999). A role for FEN-1 in nonhomologous DNA end joining: The order of strand annealing and nucleolytic processing events. Proc Natl Acad Sci USA 96, 1303– 1308. Wuerffel, R., Jamieson, C. E., Morgan, L., Merkulov, G. V., Sen, R., and Kenter, A. L. (1992). Switch recombination breakpoints are strictly correlated with DNA recognition motifs for immunoglobulin Sg3 DNAbinding proteins. J Exp Med 176, 339–349. Wuerffel, R. A., Du, J., Thompson, R. J., and Kenter, A. L. (1997). Ig Sg3 DNA-specific double strand breaks are induced in mitogen-activated B cells and are implicated in switch recombination. J Immunol 159, 4139–4144. Wuerffel, R. A., Ma, L., and Kenter, A. L. (2001). NF-kB p50-dependent in vivo footprints at Ig Sg3 DNA are correlated with m–>g3 switch recombination. J Immunol 166, 4552–4559. Yancopoulos, G. D., DePinho, R. A., Zimmerman, K. A., Lutzker, S. G., Rosenberg, N., and Alt, F. W. (1986). Secondary genomic rearrangement events in pre-B cells: VHDJH replacement by a LINE-1 sequence and directed class switching. EMBO J 5, 3259–3266. Yaoita, Y., and Honjo, T. (1980). Deletion of immunoglobulin heavy chain genes from expressed allelic chromosome. Nature 286, 850–853. Yoshikawa, K., Okazaki, I. M., Eto, T., Kinoshita, K., Muramatsu, M., Nagaoka, H., and Honjo, T. (2002). AID enzyme-induced hypermutation in an actively transcribed gene in fibroblasts. Science 296, 2033–2036.
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Zhang, K., and Cheah, H. K. (2000). Cell-free recombination of immunoglobulin switch-region DNA with nuclear extracts. Clin Immunol 94, 140–151. Zhang, K., Zhang, L., Yamada, T., Vu, M., Lee, A., and Saxon, A. (2002). Efficiency of Ie promoter-directed switch recombination in GFP expression-based switch constructs works synergistically with other promoter and/or enhancer elements but is not tightly linked to the strength of transcription. Eur J Immunol 32, 424–434. Zhao, Y., Rabbani, H., Shimizu, A., and Hammarstrom, L. (2000). Mapping of the chicken immunoglobulin heavy-chain constant region gene locus reveals an inverted a gene upstream of a condensed u gene. Immunology 101, 348–353.
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21 Molecular Mechanism of Hypermutation NANCY MAIZELS
MATTHEW D. SCHARFF
Departments of Immunology and Biochemistry University of Washington Medical School Seattle, Washington, USA
Department of Cell Biology Albert Einstein College of Medicine Bronx, New York, USA
Three dramatic molecular events alter the immunoglobulin loci of B cells to produce an antibody repertoire that is diverse, specific, and adaptable in response to the continued but varied onslaught of pathogenic microorganisms: V(D)J rearrangement creates a diverse repertoire of functional variable (V) regions capable of recognizing many antigens; class switch recombination joins a rearranged and expressed variable region to a new downstream constant (C) region, optimizing the pathway for antigen removal; and somatic hypermutation alters variable region sequence, occurring either to diversify the pre-immune repertoire or to increase antibody affinity and fine specificity, when coupled with selection. These modifications in genomic structure are required for resistance to infection, and they also play a role in generating the pathogenic antibodies responsible for autoimmune diseases. V(D)J rearrangement and class switch recombination are discussed elsewhere in this volume. This chapter focuses on somatic hypermutation, the remarkable process of targeted mutagenesis that alters variable region sequence. The mechanism of somatic hypermutation has not yet been defined in complete molecular detail. Nonetheless, a picture is emerging of how B cells use cell-type–specific factors to initiate an attack on the immunoglobulin loci, then employ ubiquitous repair and recombination pathways to modify DNA sequence. This chapter begins with an overview of how somatic hypermutation alters DNA sequence, emphasizing the characteristics of hypermutated variable regions that are relevant to understanding the mechanism, and then discuss the hypermutation pathway itself, focusing on specific factors that may participate in hypermutation.
CHARACTERISTICS OF SOMATIC HYPERMUTATION OF IMMUNOGLOBULIN VARIABLE REGIONS
Molecular Biology of B Cells
Hypermutated Ig Genes in Mice and Humans Somatic hypermutation of mammalian antibody variable region genes was discovered more than 30 years ago (Weigert et al., 1970), and has been documented in many thousands of sequences of mutated immunoglobulin (Ig) genes that have been compiled since then. The rate of V region mutation in a mammalian B cell is one base change per kb per cell generation. Hypermutation produces an excess of replacement to silent mutations, particularly in the complementarity determining regions (CDRs), which encode residues that make direct contact with antigen (Figure 21.1). Relatively little mutation is found in the framework of the variable region, which is necessary for intact structure. The Zone of Hypermutation The rate of hypermutation is nearly a million-fold higher than the mutation rate in most somatic cells. Rampant mutation at or even near this rate could have a devastating effect, impairing cell proliferation if targeted indiscriminately to all genes, or destroying antibody function if unleashed upon the C regions of the immunoglobulin molecule, which cannot tolerate sequence variation. However, hypermutation is targeted to the variable regions of rearranged and expressed
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FIGURE 21.1 The hypermutation zone is bounded by the promoter. The murine k light chain is shown, including the upstream promoter (P), the leader (L), and fused variable (V) and joining (J) segments, the four constant region exons (Ck1–4), and the intronic and 3¢ enhancers (EkI, Ek3¢). Above, the level of mutation is diagrammed. Below, the VJ region is expanded to show the three complementarity determining regions (CDRs), which encode residues that make direct contact with antigen.
immunoglobulin genes. As discussed in greater detail below, transcriptional activation is a prerequisite for hypermutation and contributes to stimulating and targeting hypermutation. The promoter itself is the upstream boundary of mutation, but mutation extends about 1.5 kb downstream of the end of the J segment, to include a portion of the J-C intron, but exclude the C region (Lebecque and Gearhart, 1990; Figure 1). Patterns of Unselected Mutation Reveal Mechanism The targeting of mutation to hotspots is an integral part of the mutation process and is not due to selection for higher affinity antibodies as originally believed. This has been shown by three different sorts of experiments: 1) Passenger transgenes, engineered to contain nonsense codons so that they could not be biased by antigen selection, were shown to hypermutate at hotspots comparable to those found in the expressed endogenous gene (Betz et al., 1993; Fukita et al., 1998). 2) A very hot spot for hypermutation was identified within the murine Vl intron, where it could not contribute to selection (Gonzalez-Fernandez et al., 1994). 3) The targeting of mutation to hot spots was observed in cultured B cells that undergo high rates of mutation, even though there is no selection for high affinity antibodies in culture (Denepoux et al., 1997; Sale and Neuberger, 1998; Zan et al., 1999). Because hotspot targeting is intrinsic to the mutational mechanism, the mutation spectra at hotspots should reveal the mechanism. In unselected regions, transitions occur more frequently than transversions, and there is a hierarchy of patterns of mutation at each base (Jolly et al., 1996; Rada et al., 1998; Rada et al., 2002b). For example, 65% of mutations at C/G pairs are transitions. The sequence RGYW, and its complement, WRCY (R = purine, A or G; Y = pyrimi-
dine, C or T; W = A/T), is a hotspot for mutation, and this hotspot is conserved among species (Rogozin et al., 1996). The sequence WA appears also to be a hotspot, contributing mainly to mutations in the nontemplate strand (Milstein et al., 1998). However, these motifs are neither necessary nor sufficient to target hypermutation: They account for a variable fraction of mutation, and not all RGYW or AT motifs are mutated. This suggests that neighboring sequences or DNA or chromatin structures are also important, a notion consistent with the observation that certain diand tri-nucleotide motifs are preferentially mutated, whereas others escape mutation (Shapiro et al., 1999).
Targeted Mutation of Ig Genes in Other Vertebrates Targeted mutagenesis of the Ig loci is not restricted to mammals, but goes back to ancestors as distant as the earliest vertebrates. Two especially striking lessons have been learned from comparing targeted mutagenesis of Ig genes among vertebrate species: 1) Mutagenesis may occur in the absence of antigen stimulation to create a diverse repertoire of variable region sequences (chicken, sheep); or in response to antigen activation to increase antibody affinity, when coupled with clonal selection (mice, human). 2) Mutation may be nontemplated to produce exclusively single-base changes (human, mice, sheep); or may be templated by a process of gene conversion to produce short tracts of sequence changes that match germline donor genes (chicken); or both (rabbit). The nontemplated and templated mutational pathways share critical aspects of mechanism. As discussed in greater detail below, both pathways depend upon AID, a cytidine deaminase that functions in the mutational mechanism to modify the sequence of V region DNA. Levels of transacting factors can shift the balance between templated and nontemplated mutagenesis (Sale et al., 2001; see Section 8B). Chicken: Pre-Immune Diversification by Templated Mutation Targeted Ig gene mutagenesis in the chicken has been formative in thinking about the hypermutation mechanism. Chickens do not rely on combinatorial diversification to achieve a useful repertoire, but instead depend on targeted mutation of the Ig loci (Reynaud et al., 1987; Thompson and Neiman, 1987). At the chicken l1 light chain locus, a single functional V region rearranges with a single J region and then undergoes programmed sequence diversification in which a family of 25 upstream pseudo-V regions serve as templates for gene conversion of the rearranged and transcriptionally activated variable region (Figure 21.2, top).
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FIGURE 21.2 Targeted mutagenesis of Ig loci can produce templated or nontemplated mutations. Above: At the chicken l light chain locus, gene conversion diversifies a rearranged and transcriptionally active VJ region, and a family of upstream pseudo-V regions (JV) serves as templates for mutation (patches). Below: Untemplated single-base changes (balls) are produced during hypermutation of murine or human immunoglobulin genes.
Gene conversion produces characteristic mutational tracts, which can be traced to the donor pseudogene. This contrasts to the untemplated single base changes produced during hypermutation of murine or human immunoglobulin genes (Figure 21.2, bottom). Sheep: Pre-Immune Diversification by Nontemplated Mutation Like chickens, sheep diversify the preimmune repertoire by targeted mutation of rearranged and expressed genes in the absence of exposure to antigen (Reynaud et al., 1995). Pre-immune diversification in the sheep is nontemplated, thus producing single base changes with no matches in other germline genes. Rabbits: Antigen-Activated Mutation by Both Templated and Nontemplated Mutation In rabbits, hypermutation occurs following antigen activation, as in mice and humans, but produces both templated and nontemplated mutations (reviewed by Knight and Winstead, 1997). This provided some of the first evidence
that the templated and untemplated mutational pathways could be closely related.
ACTIVATION AND TARGETING OF HYPERMUTATION BY TRANSCRIPTION AND CIS-ELEMENTS Transcription Is Required for Hypermutation Levels of Mutation Correlate with Transcription Levels The importance of transcription to hypermutation has been documented in experiments analyzing the effect of systematic deletion of promoter and enhancer elements in transgenic mice and lines generated by gene targeting. These experiments show that an untranscribed gene will not hypermutate, and gene expression levels correlate with the overall level of hypermutation (reviewed by Neuberger et al., 1998; Storb et al., 1998; Wiesendanger et al., 1998; Jacobs et al., 1999). However, no single transcriptional enhancer (or other cis-element) seems sufficient to activate hypermutation, and hypermutation appears to depend on contributions from multiple elements.
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Multiple Scenarios for the Connection Between Transcription and Hypermutation Hypermutation, V(D)J recombination, and class switch recombination all share a requirement for transcription. This could simply reflect a need for the DNA target to become accessible to transacting factors (Sleckman et al., 1996). Alternatively, various mechanistic possibilities have been suggested to explain the connection between transcription and hypermutation: that a transcriptional promoter, enhancer, or the transcription apparatus itself might be a loading site for factors essential to mutation; that the nascent transcript plays a role in hypermutation; that transcription might alter DNA structure; or that changes in chromatin structure might recruit specific transacting factors through the histone code. It is also possible that hypermutation does not require transcription per se, as a transcriptional terminator inserted downstream of a promoter does not inhibit hypermutation (Reynaud et al., 2001), and mutation continues when the human line BL2 cell line is cultured in actinomycin D, a potent inhibitor of pol II transcription (Faili et al., 2002).
Targeting of Hypermutation by cis-Elements Ig Loci Contain Elements Essential for Hypermutation The discovery that an ectopically integrated k light chain transgene could undergo hypermutation revealed that all the cis-acting elements required to target and regulate mutation were contained within that transgene (O’Brien et al., 1987). Many subsequent experiments have confirmed that transgenes in a variety of ectopic locations can be targeted for hypermutation at the correct stage in B-cell development. This suggested that it would be rather easy to use deletions and mutations to identify specific motifs that were required for mutation and, through them, the DNA binding proteins that were required. This has not proved to be the case. No Single cis-Element Is Identified to Supports Ig Gene Hypermutation In fact, despite extensive searches, no single cis-element has been identified to support Ig gene hypermutation. Remarkably, neither the rearranged V region nor its associated promoter is required for hypermutation: Cassettes carrying even bacterial genes can be substituted for a V region in an Ig transgene and still undergo mutation, not only at comparable frequency, but also with a similar profile of hotspots (Yelamos et al., 1995). Numerous heterologous promoters have been shown to support hypermutation, thus eliminating the possibility that the Ig promoters contain essential motifs. Curiously, a short insert of synthetic sequence may be able to activate transgene mutation, and this may provide some insights into endogenous cis-elements or
factors (Michael et al., 2002). However, in non-B cells forced to express AID, ectopically located nonimmunoglobulin transgenes driven by viral promoter-enhancers undergo high rates of mutation and display a mutation profile similar to endogenous V regions hypermutated in activated B cells (Martin and Scharff, 2002b; Yoshikawa et al., 2002). This raises the question of whether, other than AID, any Igspecific cis-acting elements or B cell specific trans-acting factors are actually required to target the hypermutation process that acts on V regions in B cells in vivo. Targeting Elements May Be Co-Opted Immediately following the discovery of hypermutation, a number of experiments asked if regions other than the Ig loci might mutate, with negative results. More recently, it has become clear that non-Ig genes, particularly proto-oncogenes, actually do undergo hypermutation in activated B cells, though at considerably lower rates than the heavy and light chain V regions. Mutations were first found in c-myc (Johnston and Carroll, 1992), which frequently undergoes translocations in B-cell tumors, and later in BCL-6 (Migliazza et al., 1995; Shen et al., 1998) and CD95 (Muschen et al., 2000). Strikingly, multiple loci, including several proto-oncogenes, are consistently mutated in diffuse large cell lymphomas (DLCL), which originate from hypermutating B cells (Pasqualucci et al., 2001). The realization that certain oncogenes actively hypermutate has intensified the search for elements that target hypermutation.
HYPERMUTATION OCCURS WITHIN A LIMITED WINDOW OF B CELL DEVELOPMENT Hypermutation Occurs in Sequestered Microenvironments Somatic hypermutation occurs in centroblasts. Centroblasts are rapidly cycling cells that, in chickens, are sequestered in the bursa of Fabricius, an organ dedicated to B-cell development. In mammals, these cells populate the dark zone of the lymphoid germinal center. Cell surface markers characteristic of this stage of B-cell differentiation allow hypermutating B cells to be sorted to produce a greatly enriched population for experimental analysis. Nonetheless, centroblasts are not abundant in vivo, and detailed molecular analysis has required hypermutating cell lines that will grow continuously in tissue culture. Hypermutation in Transformed Cell Lines Both chicken and human cell lines have been shown to carry out targeted mutagenesis of their Ig genes, and
21. Molecular Mechanism of Hypermutation
products of mutation correctly recapitulate pathways of targeted mutagenesis in each organism: Mutation is templated in chicken and nontemplated in human lines. The ongoing targeted diversification of Ig V regions occurs in the chicken cell line DT40, generated by transformation of a bursal cell with avian leukosis virus. DT40 is readily cultured, and mutation of the Ig genes can be easily measured by using FACS to quantitate loss of surface immunoglobulin. Moreover, gene targeting occurs with very high efficiency in DT40, presumably because the levels of enzymes involved in homologous recombination are elevated to carry out gene conversion. This has made DT40 a useful model for analysis of gene function. Until a few years ago, no mammalian B cell lines were known to undergo active hypermutation in culture. However, after a convincing case was made for hypermutation in one human B cell line (Sale and Neuberger, 1998), it was quickly found that hypermutation is active in a considerable number of human B cell lines that derive from germinal center tumors. Certain lines can be induced to activate mutation by crosslinking their surface IgM and treating with helper T cells, thus mimicking the normal stimuli that turns on mutation (Denepoux et al., 1997; Zan et al., 1999). At least one line, BL2, may support efficient gene targeting (Faili et al., 2002), which will enhance the utility of human B cell lines for studies of the hypermutation mechanism. Hypermutation and Class Switch Recombination Although hypermutation and class switch recombination both occur at approximately the same time in centroblasts, each can occur without the other, and these were originally thought to be quite different processes. The discovery that AID is required for both hypermutation and class switch recombination (Muramatsu et al., 2000; Revy et al., 2000; and see chapter by Birshtein) focused attention on mechanistic parallels (reviewed by Honjo et al., 2002). Hypermutation is a mutational process accompanied by some deletions; switch recombination is a deletion event accompanied by some single base changes. Both processes occur shortly after antigen activation in murine and human B cells, are dependent upon AID, and are supported by a pathway involving uracil DNA glycosylase (Rada et al., 2002b). Switch junctions and the sites of hypermutation are typically heterogenous, but this heterogeneity diminishes in the absence of MSH2, just as the spectrum of hypermutation is more limited in Msh2-deficient mice (Rada et al., 1998; Ehrenstein and Neuberger, 1999). In spite of these important similarities, there are also striking differences, including the dependence of switch recombination but not hypermutation on factors involved in nonhomologous end-joining, particularly Ku/DNA-PK (Bemark et al., 2000; Manis et al., 2002), g-H2AX (Petersen et al., 2001), and PMS2 (Ehrenstein et al., 2001). This suggests that hyper-
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mutation and class switch recombination may be initiated by the same mechanism but differently resolved.
THE AID GENE IS CRITICAL FOR HYPERMUTATION AID as B Cell–Specific Factor Required for Hypermutation In 2000, Honjo and his colleagues discovered a single gene that is required for V-region hypermutation and class switch recombination in both mice and humans (Muramatsu et al., 2000; Revy et al., 2000). This gene was first identified by subtraction screening as a cytidine deaminase restricted in expression to activated centroblast B cells (Muto et al., 2000), hence the name activation induced deaminase (AID). AID proved also to be required for the targeted mutation of Ig genes in chickens (Arakawa et al., 2002; Harris et al., 2002), and this role may extend to all vertebrates. The AID polypeptide is only 198 amino acids in length and bears a 34% amino acid identity with the cytidine deaminase APOBEC-1 (Muramatsu et al., 2000). APOBEC-1 is an enzyme that edits a specific mRNA, converting a C to a U and introducing a premature termination in the mRNA for low-density lipoprotein in the intestine. AID and APOBEC1 are separated by only about 1 megabase on human chromosome 12p13, suggesting that they arose by duplication of a common progenitor (Muto et al., 2000). A series of recent experiments support the possibility that AID may be the only B cell–specific factor required for targeted Ig gene mutation (Martin and Scharff, 2002a). First, overexpression of AID can induce V-region mutation in B cells at the plasma cell stage in differentiation (Martin and Scharff, 2001) and in non-B cells such as fibroblasts (Yoshikawa et al., 2002) and CHO cells (Martin and Scharff, 2002b). These results further suggested that AID is responsible for restricting V-region mutation and isotype switching to a short window in B-cell development. In addition, not only is AID expression required to support hypermutation in cultured cell lines (Arakawa et al., 2002; Harris et al., 2002; Martin et al., 2002), but levels of hypermutation are proportional to levels of AID expression both in cell lines (Zhang et al., 2001; Martin et al., 2002) and in mice (Muramatsu et al., 2000). These last results suggest that AID levels are normally limiting for hypermutation, most consistent with a direct effect of AID on DNA (Figure 21.3B).
AID Answers Some Questions and Raises Others Many pieces of information are still necessary to fill out our understanding of the role of AID. Almost nothing is
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known about its enzymatic properties, or substrates or localization in vivo. In fact, AID tagged with green fluorescent protein (GFP) localizes to the cytoplasm, not to the nucleus (Rada et al., 2002a), where, analogous to APOBEC-1, it could edit mRNA. The cytoplasmic localization could also be an artifact: AID-GFP may be inactive and incapable of correct localization, or AID may depend on another factor for transport to the nucleus. This factor may be titrated by AID-GFP overexpression or unable to bind AID-GFP. The possibility of such a partner protein is suggested by analogy with APOBEC-1, which interacts with another factor to recognize its nucleic acid substrate. For these reasons, it essential to know whether other proteins interact with AID and to learn what they are and what they do. In addition, the discovery of AID has not solved the mystery of how hypermutation is specifically targeted to Ig loci. The finding that AID can activate mutation of non-Ig genes in non-B cells, including the AID transgene itself (Martin and Scharff, 2002b), suggests that specific cis-acting elements are not required to target AID (Martin and Scharff, 2002a). This is a disquieting idea, since it seems dangerous for AID to target mutations to non-Ig genes. However it is a real possibility, since specific oncogenes also seem to be targeted for hypermutation (see above).
PHASE ONE OF HYPERMUTATION: CÆU DEAMINATION AND BASE EXCISION REPAIR
Uracil DNA Glycosylase Intersects Hypermutation The notion that AID deaminates DNA was further tested by asking how uracil DNA glycosylase participates in the actual mechanism of Ig gene hypermutation (Di Noia and Neuberger, 2002; Rada et al., 2002b). Inhibition of UNG activity in a DT40 XRCC2-/- derivative, in which most mutations are point mutations (Sale et al., 2001; and see below), altered the mutation spectrum so that transitions and not transversions predominated (Di Noia and Neuberger, 2002). In ung-/- mice, the spectrum of hypermutation was even more profoundly altered, so that 95% of mutations were transitions (Rada et al., 2002b).
Phase One: Mutations at C/G These results support a picture of a mutagenic mechanism that exploits damage induced by AID in two ways: to generate transition mutations upon replication that fixes the error (Phase One A, Figure 21.3B), and to generate mainly transversion mutations upon repair of an abasic site (Phase One B, Figure 21.3B). It has previously been suggested that an early phase of hypermutation is specifically targeted to C/G base pairs (Rada et al., 1998; Wiesendanger et al., 2000). The repair of sites deaminated by AID activity could account for mutations at C/G pairs. As described below, mutations at other sites appear to be produced in a distinct phase involving mismatch repair factors.
AID Directly Modifies DNA By analogy with APOBEC-1, AID could function to edit an mRNA and thereby regulate the expression of a protein critical to switching and hypermutation. However, recent results provide strong support to the hypothesis that AID directly deaminates DNA. Although purified AID protein has not yet been shown to modify DNA in vitro, it has been possible to analyze mutagenesis induced by AID expression in different cell types and genetic backgrounds by postulating that at least a fraction of the damage incurred by AID would feed into the highly conserved and specific base excision repair pathway that repairs spontaneous CÆU deamination (see Figure 21.3A). Reasoning that, if AID does modify DNA directly, this might be evident even in bacteria, Neuberger and colleagues expressed AID in Escherichia. coli and assayed hypermutation of reporter genes (Petersen-Mahrt et al., 2002). They showed that mutation increased several-fold as a result of AID expression, mainly due to transitions at C/G pairs. Most critically, AIDinduced mutation levels increased in a strain lacking uracil DNA glycosylase (ung-), a highly conserved enzyme that specifically removes uracil from DNA.
MISMATCH REPAIR FACTORS IN PHASE TWO OF HYPERMUTATION Deficiencies in MSH2 or MSH6 Alter the Level and Spectrum of Hypermutation In animal cells, the MSH2/MSH6 heterodimer (MutSa) recognizes single base mispairs as well as mismatches created by single base deletions or insertions while the MSH2/MSH3 heterodimer (MutSb) recognizes mismatches of two to four base pairs. Both complexes then recruit the MLH1/PMS2 heterodimer (MutLa), and presumably other factors, to excise the mismatch and repair it by DNA synthesis. One might therefore have predicted that the mismatch repair pathway would correct mutations introduced during hypermutation, so that hypermutation levels would increase in mismatch-repair–deficient backgrounds. Surprisingly, deficiencies in mismatch repair actually result in either decreased or unchanged mutation levels, while altering the mutation spectrum (reviewed by Wiesendanger et al., 1998). In Msh2-/- mice, the frequency of unselected mutations is reduced about five-fold relative to Msh2+/- littermates, and
FIGURE 21.3 Deamination of CÆU by the hypermutation pathway. (a) Spontaneous deamination of CÆU is repaired by a conserved and specific base excision repair pathway (reviewed by Wood et al., 2001). First, uracil DNA glycosylase, encoded by the UNG gene, cleaves the glycosidic bond to release uracil, thus creating an apurinic site within an intact phosphodiester backbone; next, AP endonuclease (APE) nicks the backbone; then a repair polymerase primes synthesis on the free 3¢ end; and finally, DNA ligase seals the nick. (b) The AID protein may initiate hypermutation by acting directly on DNA, catalyzing the deamination of CÆU (based on Rada et al., 2002b). Mutation could become fixed upon DNA replication to produce transition mutations, with no involvement of the uracil DNA repair pathway (Phase One A). Alternatively, following base excision by uracil DNA glycosylase (UNG), gapped DNA may be a substrate for error-prone repair, producing both transition and transversion mutations (Phase One B). (c) A distinct phase of hypermutation (Phase Two) depends on MutSa, the MSH2/MSH6 heterodimer. This phase produces mutation at sites other than C/G pairs.
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C/G (rather than A/T) becomes the preferred mutation site (Rada et al., 1998; Phung et al., 1998). This altered spectrum can be recapitulated in Msh6-/- but not Msh3-/- mice, and therefore reflects involvement of the MSH2/MSH6 heterodimer, MutSa (Wiesendanger et al., 2000). In contrast to the clear function of MutSa, the role of the MutL homologs remains unclear, and hypermutation may use MutSa differently from that of other repair pathways.
Phase Two of Hypermutation: MutSa Promotes Mutation at Sites Other than C/G One plausible hypothesis is that MutSa binds mismatches that are created by AID, thus targeting them for cleavage followed by error-prone repair (Figure 21.3C). MSH2 is required to initiate expansion at triplet repeats (Manley et al., 1999), which may analogously involve targeting a cleavage activity. Alternatively, MSH2 may function after cleavage, for example, by stimulating the resection of structured ends by interacting with repair and replication factors.
DNA BREAKS IN HYPERMUTATION Break-and-Repair Pathway of Hypermutation A hypermutation pathway driven by the creation and repair of DNA breaks was originally suggested by evidence that targeted mutation of Ig genes could produce either untemplated or templated mutations (Maizels, 1995). These superficially distinct outcomes were proposed to reflect a single mutational mechanism, in which the key step is initiation of hypermutation by a DNA lesion (shown as a single-strand break, Figure 21.4), which is repaired either by error-prone DNA replication to produce nontemplated mutation (Figure 21.4, left), or by gene conversion to produce templated mutation (Figure 21.4, right). In this view, the use of the nontemplated or templated repair pathway could be determined by the balance of cellular factors. That levels of specific repair factors can indeed tip this balance is supported by the discovery that, in the hypermutating chicken cell line, DT40, ablation of RAD51 paralogs XRCC2, XRCC3, or RAD51D, causes point mutations to predominate relative to gene conversion (Sale et al., 2001).
DNA Breaks Identified in Hypermutating B Cells Single-Strand Breaks in Hypermutating B Cells SSBs have been directly identified in hypermutating murine B cells, through experiments that used ligation-
FIGURE 21.4 Break and repair model for hypermutation. Nontemplated and templated mutation may be two outcomes of a single mutational mechanism. The key step is initiation of hypermutation by a DNA lesion, shown as a single-strand break. Repair by error-prone DNA synthesis will produce nontemplated mutation (left); while gene conversion will produce templated mutation (right).
mediated PCR to examine both endogenous l1 genes and an actively hypermutating l1 transgene (Kong and Maizels, 2001). There was approximately one break per 100 V l1 regions, and breaks mapped to the zone corresponding to the portion of the l1 gene that hypermutates, bounded upstream by the promoter. About half the breaksites in the dataset from hypermutating B cells were independently identified twice or more, consistent with a targeted biological process rather than random DNA damage. Double-Strand Breaks in Hypermutation Double-strand breaks (DSBs) have also been reported in V-region DNA, but their association with hypermutation is unclear. DSBs were identified in VH regions from Ramos, a human B cell line which hypermutates in culture, and DSBs were mainly evident in G2/S phase (Papavasiliou and Schatz, 2000). DSBs were also found associated with RGYW hypermutation hotspots in a VH “knock-in” in a transgenic mouse, but levels of DSBs were comparable in samples from nonhypermutating and hypermutating B cells (Bross et al., 2000). There was a flurry of speculation that DSBs might initiate hypermutation (Papavasiliou and Schatz, 2000). However, this proposal could only be reconciled with more recent studies showing that DSBs occur in AID-deficient mice (Bross et al., 2002) and cell lines (Papavasiliou and Schatz, 2002) by hypothesizing that AID functions subsequent to DNA cleavage (Papavasiliou and Schatz, 2002). This hypothesis conflicts with the evidence that AID acts directly on DNA to initiate hypermutation (see above). In addition, it has recently been reported that Vregion mutation can be induced rapidly (within 90 minutes of receptor stimulation) in BL2 Burkitt’s lymphoma B cells, but DSBs are not evident until much later; breaks can be found not only in cells that express AID, but also in AIDdeficient cells that do not undergo mutation (Faili et al., 2002). The evidence that dissociates DSBs and V-region mutation does not completely rule out a role for DSBs, but it has raised questions about whether the DSBs are an arti-
21. Molecular Mechanism of Hypermutation
fact or at least are not required for the mutational process and hence arise late in the course of repairing lesions that initiate the process (Chua et al., 2002; Martin and Scharff, 2002a). SSBs Are Consistent with the Current View of the Mutational Mechanism SSBs—but not DSBs—would be produced in the course of the AID- and UNG-dependent hypermutation pathway (Di Noia and Neuberger, 2002; Petersen-Mahrt et al., 2002; Rada et al., 2002b), discussed above. In addition, SSBs would give rise to DSBs upon replication, in which case DSBs would be enriched at G2/S, as has been reported (Papavasiliou and Schatz, 2000); only one end would be a flush duplex end, thus explaining the apparently asymmetric structure found at DSBs (Bross et al., 2000; Papavasiliou and Schatz, 2000). Moreover, genetic analysis does not support a role for DSBs in hypermutation. The mutation of DNA-PKcs, a critical factor in DSB repair, has no effect on hypermutation (Bemark et al., 2000). The phosphorylated minor histone, g-H2AX, participates in DSB repair in response to DNA damage and in the course of both V(D)J recombination (Chen et al., 2000) and switch recombination (Petersen et al., 2001), but appears unnecessary for somatic hypermutation (Celeste et al., 2002). SSBs are further consistent with the mechanism in light of the evidence that RAD51 paralogues determine the outcome of mutation in the chicken DT40 cell line (Sale et al., 2001; and see next section). The BCDX2 complex, formed by RAD51 paralogs (RAD51B/RAD51C/RAD51D and XRCC2), binds to single-stranded DNA and single-stranded gaps in duplex DNA (Masson et al., 2001).
COMPETING PATHWAYS OF REPAIR: ERROR-PRONE DNA SYNTHESIS OR STRAND TRANSFER Error-Prone Polymerases in Hypermutation The finding that hypermutation in Msh2-/- and Msh6-/mice focuses on C/G pairs or RGYW hotspots led to the proposal that there are two distinct phases to the process of V-region mutation (Rada et al., 1998). In the current view (Figure 21.3), the first phase would be AID-dependent and target mutation to C/G pairs, while the second phase would involve recruitment of MutSa, DNA excision, and resynthesis by error-prone polymerases. Over a dozen error-prone polymerases have been identified (reviewed by Goodman, 2002; Friedberg and Fischhaber, 2002), and several produce molecular signatures consistent with a role in somatic hypermutation. Pol h is the product of the XPV gene and produces a mutational spectrum that cor-
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relates well with potential function as the major mutator at A/T in somatic hypermutation. Patients with xeroderma pigmentosa variant (XPV) disease have a decrease in A/T mutations (Rogozin et al., 2001; Zeng et al., 2001; Pavlov et al., 2002). Pol z, a Rev3 homolog, is also a good candidate, and its function in hypermutation is supported by evidence that antisense-mediated partial inactivation of pol z in both mutating B-cell lines and in mice resulted in a decrease in the rate of V-region mutation (Diaz et al., 2001; Zan et al., 2001). Targeted ablation of the genes encoding pol l or pol m does not alter hypermutation (Bertocci et al., 2002); nor does pol k-deficiency affect somatic hypermutation (Schenten et al., 2002).
RAD51 Paralogs Carry Out Strand Transfer to Produce Templated Mutation Targeted mutagenesis of chicken Ig genes proceeds by a pathway involving strand transfer and DNA synthesis from a homeologous pseudo-V region template (Figure 21.4). The chicken line DT40 recapitulates this pathway, but targeted ablation of any of the Rad51 paralogs RAD51B, XRCC2, or XRCC3 alters both the level and outcome of mutagenesis (Sale et al., 2001): The frequency of hypermutation increases approximately five-fold. This increase is due to point mutations, which accumulate at or near the conserved hypermutation hot spot, RGYW, and are thought to result from attempts to repair DNA breaks in the absence of efficient strand transfer mediated by the RAD51 paralogs. That the balance between templated and nontemplated mutation can be altered by genetic manipulation supports the notion that these pathways are closely related, as does the fact that AID is essential to both pathways.
EVOLUTION AND HYPERMUTATION The more we learn about the mechanism, the clearer it becomes that B cells have usurped conserved and ancient pathways for DNA repair in order to carry out the targeted mutagenesis of Ig genes. The universality of this pathway is highlighed by recent findings discussed above: the likelihood that AID is the only B cell-specific factor; the participation of members of the uracil DNA repair pathway; and the involvement of MSH2 and MSH6. It may appear paradoxical that factors that normally repair DNA are used in this context to create mutations. Nonetheless, this apparent misappropriation of highly conserved factors probably speaks to the importance of evolving and maintaining a dynamic mechanism for targeted Ig gene mutagenesis. In B cells, evolution occurs in real time to allow us to respond dynamically to combat pathogens, which are themselves mutating.
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Zan, H., Komori, A., Li, Z., Cerutti, A., Schaffer, A., Flajnik, M. F., Diaz, M., and Casali, P. (2001). The translesion DNA polymerase zeta plays a major role in Ig and bcl-6 somatic hypermutation. Immunity 14, 643–653. Zeng, X., Winter, D. B., Kasmer, C., Kraemer, K. H., Lehmann, A. R., and Gearhart, P. J. (2001). DNA polymerase eta is an A-T mutator in somatic hypermutation of immunoglobulin variable genes. Nat Immunol 2, 537–541. Zhang, W., Bardwell, P. D., Woo, C. J., Poltoratsky, V., Scharff, M. D., and Martin, A. (2001). Clonal instability of V region hypermutation in the Ramos Burkitt’s lymphoma cell line. Int Immunol 13, 1175–1184.
NOTE ADDED IN PROOF AID deaminates DNA directly, but only attacks single-stranded DNA Since this chapter was written, the key question of whether AID acts directly on DNA has been resolved. Four different laboratories have shown that recombinant AID deaminates cytidines in DNA in vitro, but will only attack single-stranded DNA and will not modify double-stranded DNA, or an RNA : DNA hybrid (Bransteitter et al., 2003; Chaudhuri et al., 2003; Dickerson et al., 2003; Sohail et al., 2003). Strikingly, AID will deaminate cytidines in a transcribed substrate in a reaction coupled to transcription (Chaudhuri et al., 2003), and it will also attack the exposed single-stranded region within an artificial transcription bubble (Bransteitter et al., 2003). Deamination by AID is processive, introducing multiple mutations into a few molecules while leaving many molecules free of mutation (Pham et al., 2003). These results provide an immediate and unanticipated solution to the long-standing question of why transcription is required for hypermutation: transcription transiently denatures the DNA duplex, producing singlestranded regions that are targets for deamination. They explain why levels of hypermutation are roughly proportional to transcription levels, and why some genes are highly mutated and others unmutated. They also show why no single cis-acting mutator element could be found despite the considerable effort devoted to this quest. However, while hypermutation in vivo appears to produce no apparent strand bias, experiments thus far suggest that there is a clear strand bias, at least in one stage of hypermutation: deamination by AID is overwhelmingly targeted to the nontemplate strand, while the template strand is protected from mutation (Chaudhuri et al., 2003; Pham et al., 2003). Strand bias is also observed in products of mutations
produced when AID is expressed in E. coli (Ramiro et al., 2003; Sohail et al., 2003). Further experiments must resolve this apparent paradox. Beyond deamination: AID may recruit factors specific to switch recombination AID is required for both somatic hypermutation and class switch recombination, but these two processes produce very different genomic outcomes. Very recent studies suggests that AID itself plays a role in recruiting factors that distinguish these pathways. Deletion of 10 residues from the Cterminus of AID does not affect initiation of somatic hypermutation, abolishes class switch recombination (Barreto et al., 2003; Ta et al., 2003). The identity of these factors will of great interest. Barreto, V., Reina-San-Martin, B., Ramiro, A. R., McBride, K. M., and Nussenzweig, M. C. (2003). C-terminal deletion of AID uncouples class switch recombination from somatic hypermutation and gene conversion. Mol Cell 12, 501–508. Bransteitter, R., Pham, P., Scharff, M. D., and Goodman, M. F. (2003). Activation-induced cytidine deaminase deaminates deoxycytidine on single-stranded DNA but requires the action of RNase. Proc Natl Acad Sci U S A 100, 4102–4107. Chaudhuri, J., Tian, M., Khuong, C., Chua, K., Pinaud, E., and Alt, F. W. (2003). Transcription-targeted DNA deamination by the AID antibody diversification enzyme. Nature 422, 726–730. Dickerson, S. K., Market, E., Besmer, E., and Papavasiliou, F. N. (2003). AID mediates hypermutation by deaminating single stranded DNA. J Exp Med 197, 1291–1296. Pham, P., Bransteitter, R., Petruska, J., and Goodman, M. F. (2003). Processive AID-catalysed cytosine deamination on single-stranded DNA simulates somatic hypermutation. Nature 424, 103–107. Ramiro, A. R., Stavropoulos, P., Jankovic, M., and Nussenzweig, M. C. (2003). Transcription enhances AID-mediated cytidine deamination by exposing single-stranded DNA on the nontemplate strand. Nat Immunol 4, 452–456. Sohail, A., Klapacz, J., Samaranayake, M., Ullah, A., and Bhagwat, A. S. (2003). Human activation-induced cytidine deaminase causes transcription-dependent, strand-biased C to U deaminations. Nucleic Acids Res. 31, 2990–2994. Ta, V. T., Nagaoka, H., Catalan, N., Durandy, A., Fischer, A., Imai, K., Nonoyama, S., Tashiro, J., Ikegawa, M., Ito, S., Kinoshita, K., Muramatsu, M., and Honjo, T. (2003). AID mutant analyses indicate requirement for class-switch-specific cofactors. Nat Immunol 4, 843–848.
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22 Selection During Antigen-Driven B Cell Immune Responses: The Basis for High Affinity Antibody MARK J. SHLOMCHIK, M.D., PH.D Yale University School of Medicine, New Haven, Connecticut, USA
The phenomenon of affinity maturation—the progressive increase in antibody affinity during the course of an immune response—has been known at least since the 1960s (Siskind et al., 1968). Early experiments showed that antigen (Ag) dose and competition played important roles in the elicitation of high affinity antibody (Ab). However, it took the introduction of hybridoma technology to clearly demonstrate an important mechanism behind this phenomenon— somatic hypermutation. In a series of studies in the mid-1980s, a number of workers showed that hybridomas generated during the course of primary and secondary immune responses contained somatic mutations in V regions and also secreted higher affinity Abs (Allen et al., 1987; Berek and Milstein, 1987; Clarke et al., 1985; Griffiths et al., 1984; Kaartinen et al., 1983; McKean et al., 1984). In the case of some hapten systems, notably anti-nitrophenyl (NP) and anti-oxazolone, the particular mutations that controlled higher affinity were identified (Allen et al., 1988; Berek and Milstein, 1987). Their progressive enrichment during the evolution of the immune response provided a molecular mechanism for affinity maturation. When the germinal center (GC) was identified as a major site for memory cell generation (Coico et al., 1983) and somatic hypermutation (Jacob et al., 1991b), a link was made with affinity maturation as well. This fit well with the evidence that the GC was a site of dynamic B-cell turnover, with a high rate of division as well as death, making it quite suitable for selection of higher affinity mutants (Liu et al., 1991b; MacLennan and Gray, 1986; Zhang et al., 1988). This was directly demonstrated in anti-NP response by microdissecting GC cells and sequencing their V regions; that the V regions recovered from small numbers of cells at the same site contained hierarchies of shared and unique mutations demonstrated that the GC was indeed a site of ongoing mutation and not just a location where already-
Molecular Biology of B Cells
mutated cells congregated (Jacob et al., 1991b; Jacob and Kelsoe, 1992; Jacob et al., 1993). The mutations in these sequences allowed the construction of genelogical trees based on patterns of shared mutations, which have come to be the hallmark of ongoing mutation in clonally expanding B cells. In addition, in the NP-specific GCs, a key mutation in the VH that increases affinity (Trp->Leu) (Allen et al., 1988) and is enriched in the memory response was also shown to be enriched, albeit to a lesser degree, during the ongoing GC response (Jacob et al., 1991b; Jacob and Kelsoe, 1992; Jacob et al., 1993). In spite of this progress in our understanding of affinity maturation, a number of key questions have yet to be answered. Is the GC the only place in which affinity maturation occurs? Many studies suggest that affinity maturation also occurs at other sites and times during the immune response, and these will be discussed. How are high affinity mutants selected and what is the role of competition? What is the meaning of selective advantage: increased division or decreased death? These issues will also be discussed in the context of selection at various stages. The discussion will be structured around the phases of the B-cell response and the types and mechanisms of selection at each phase. It will culminate in an integrated view of how the B-cell immune response is designed to ensure the selection of high affinity B cells and the secretion of high affinity Ab.
OVERVIEW OF THE B-CELL IMMUNE RESPONSE B cell responses initiate with BCR crosslinking by antigen. For certain antigens that provide either a high degree of crosslinking and/or an endogenous co-stimulatory signal (like LPS), a T cell–independent response can ensue.
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In this case, AFCs are formed, but neither true GCs nor somatic hypermutation and affinity maturation occur. Nor is there immunologic memory in T cell–independent responses (de Vinuesa et al., 2000; Garcia de Vinuesa et al., 1999; Liu et al., 1991b). Here we restrict our discussion to T cell–dependent responses. For these to proceed to the next stage, the initially activated B cell must encounter an activated T cell responding to a cognate Ag, that is, a peptide that the B cell is presenting, typically as a result of specific uptake of a protein Ag by the BCR (de Vinuesa et al., 2000; Lentz and Manser, 2001; Miller et al., 1995). This happens after initially activated T and B cells modulate expression of chemokine receptors and their responsiveness to chemokines so that they move to the border of the T cell and B cell zone (Ansel et al., 1999; Gulbranson-Judge and MacLennan, 1996; Luther and Cyster, 2001; MacLennan et al., 1997; Randolph et al., 1999; Tang and Cyster, 1999). There, they interact transiently and stimulate each other to propagate the T-dependent B-cell response (Liu et al., 1991b). However, this response quickly migrates to the border of the T zone and the red pulp, in the marginal sinus bridging channels (Jacob et al., 1991a; Liu et al., 1991b). The proliferative response continues there for several days and most likely remains dependent on T cells or at least is influenced by them. A substantial proportion of the B cells formed at this site differentiate to plasmablasts (Jacob et al., 1991a; Liu et al., 1991b). Plasmablasts have properties of both activated B cells and plasma cells. Like activated B cells, they continue to proliferate, are surface Ig+, and express co-stimulatory molecules and MHCII (to a variable extent), but like plasma cells they secrete Ab at high rate, express syndecan, and have reduced CD22 and B220 levels (Wehrli et al., 2001; J. William, S. Anderson, and M. Shlomchik, unpublished observations). These plasmablasts are short lived yet most likely represent most or all of the early AFCs in the splenic response. Recent evidence suggests that these plasmablasts require signals from nearby dendritic cells, possibly including via BLyS, for their differentiation and/or survival. A few of these plasmablasts most likely differentiate into long-lived plasma cells that reside in the red pulp of the spleen; these plasmablasts may also contribute to the pool of long-lived plasma cells in the bone marrow (see below). Simultaneously with the waning of the response in the marginal sinus bridging channels, the GC response in the follicles becomes more prominent. Though it is clear that the same clone of B cells can contribute to both the marginal sinus proliferative focus as well as to the GC, it is less clear whether they are independently seeded by descendents of the original T–B zone border reaction or, alternatively, whether some of the progeny generated at the marginal sinus migrate to seed the GC (Jacob and Kelsoe, 1992; Vora et al., 1998; Vora et al., 1999). In any case, in the GC, B cells pro-
liferate rapidly and this proliferation requires signals from T cells. The GC B cells take on another unique phenotype or phenotypes that is in some ways distinct from the activated B cells that are found at other sites. Unique characteristics of GC B cells have recently been defined by microarray-based gene expression analysis. Among the prominent differences at the protein level are decreased expression of surface Ig (but not surface Ig-negative) and bcl-2, and increased expression of Fas and CD86. These GC B cells undergo somatic hypermutation of their Ig V regions. In the GC, mutant B cells are subject to a competitive process in which cells with higher affinity for Ag have a selective advantage. The result is that, in general, highaffinity mutant B cells survive the GC reaction. Also, mutations create B cells with nonfunctional receptors. These too are lost from the GC population, a process termed “negative selection.” During the course of the GC reaction, particularly at the later stages, B cells can differentiate to either plasmablasts or memory cells. The signals that control this differentiation are unclear, but in some way involve CD40 and cytokines such as IL4 (Allman et al., 1996; Casamayor-Palleja et al., 1996; Liu et al., 1989). These plasmablasts and memory cells carry mutations, some of which confer higher affinity for antigen (Cumano and Rajewsky, 1986). The plasmablasts that are generated in the GC are responsible for the first wave of higher affinity Ab, which can be seen within a couple weeks of immunization. The kinetics of AFC appearance at various sites suggests that plasmablasts can migrate either to the red pulp or bone marrow, and possibly to inflamed tissues as well (Cassese et al., 2001; Hauser et al., 2002; Manz et al., 1997; Manz et al., 1998). The bone marrow provides a reservoir or niche for long-lived plasma cells. Their longevity could be determined at the time they are generated in the GC, or they could acquire a long lifespan under the influence of signals from the bone marrow milieu. In any case, it is possible that the AFCs that first arrive at the bone marrow are in the process of differentiating to long-lived plasma cells and may still retain features of plasmablasts, including expression of surface Ig (Hauser et al., 2002). As will be discussed, this could provide the substrate for further selection on the basis of affinity for antigen. Indeed, these plasmablasts/cells secrete the antibody that maintains long-term Ab titers, the affinity of which can be seen to increase for many weeks after the GC reaction is terminated (Takahashi et al., 1998). Finally, memory B cells derived from the GC reaction will recirculate through the B-cell areas of secondary lymphoid tissue, much as naïve B cells (Liu et al., 1988; S. Anderson, L. Hannum, A. Haberman and M. Shlomchik, manuscript in preparation). It was long postulated that access to Ag, presumably deposited on FDC, was required to sustain memory B cells (Gray and Skarvall, 1988; Gray and Leanderson, 1990; MacLennan
22. Selection during Antigen-Driven B Cell Immune Responses: The Basis for High Affinity Antibody
et al., 1990). However, the weight of evidence now suggests that memory B cells can be maintained without seeing the nominal Ag on which they were selected (Maruyama et al., 2000; S. Anderson, L. Hannum, A. Haberman and M. Shlomchik, manuscript in preparation). In our view, though, it remains possible that, after the GC reaction, the repertoire of retained memory B cells could still be selected by Ag, if it is present, as will be discussed. Memory B cells will, naturally, be the source of the high-affinity Ab that is secreted very early after secondary immunization. Thus, as outlined above, there a number of stages during an Ag-driven, T-dependent B-cell immune response at which selection for higher affinity can occur. These can roughly be divided into the early stages prior to the GC reaction, the GC reaction itself, and then the later stages involving postGC B cells. The details of selection in each of these stages will be covered in turn below.
AFFINITY MATURATION IN THE EARLY STAGES OF THE B-CELL IMMUNE RESPONSE Important insights into selection at the start of the proliferative B-cell immune response came from the work of Jacob et al. (1991a, 1993). Studying the response to the hapten NP, they microdissected the foci of proliferating B lineage cells in the marginal sinus bridging channels. The mature response to NP had been known from even earlier work by Reth, Bothwell, Rajewsky and colleagues (Bothwell et al., 1981; Reth et al., 1978; Reth et al., 1979) to be dominated by B cells expressing a single VH, VH186.2, in combination with a restricted CDR3 sequence and the l1 light chain. This was determined by sequencing V regions of hybridomas generated during a late primary or early secondary immunization (Bothwell et al., 1981; Cumano and Rajewsky, 1986; Siekevitz et al., 1987). Jacob et al. (1991a) first showed that the extrafollicular foci were indeed comprised of l light chain–expressing B cells. They designed PCR primers that would amplify VH186.2 genes along with a number of related VH genes of the Sm7 family. When they microdissected B cells from these sites at early time points post-immunization, amplified their V region DNA, and cloned and sequenced the products, they mainly found VH genes that were related to but were not identical to VH186.2 (Jacob et al., 1993). At later time points, they still saw these so-called “analog” VH genes, along with an increased frequency of VH186.2 genes. These results indicated that selection was already occurring in the early stages of the extrafollicular proliferative response. They also suggested that the initial response was diverse and possibly of low affinity. As there were few if any mutations in these V-region sequences, selection must have been occurring on V gene
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rearrangements per se, allowing only certain founder clones to survive. To further test the idea that the initial response was comprised of a diversity of low-affinity clones, Dal Porto et al. (1998) cloned representative “analog” VH sequences recovered in the earlier studies into IgG1 expression vectors. These were transfected into a Vl expressing myeloma cell line, yielding an antibody that had the same V regions as the cell from which the original VH region had been cloned. They then measured the affinities of these Abs using fluorescence quenching. Remarkably, these were very low, and in some cases unmeasurable—the range was <5* ¥ 104 to 2* ¥ 105 molar. Antibodies of such low affinity had not been thought to be recruited into the immune response in the first place. These results raised the question of the subsequent fate of these low-affinity B cells. Further sequencing studies showed that B cells carrying “analog” VH genes were present, but rare in early GCs, and were even more rare in later GCs (Dal Porto et al., 1998; Jacob et al., 1993). This suggested that these B cells could reach the GC, albeit inefficiently, but were rapidly lost. However, it did not indicate why this was the case. Were low-affinity cells inherently less capable of undergoing GC differentiation or, alternatively, were they simply excluded by virtue of competition from higher affinity cells? To test this idea, Dal Porto et al., in further experiments (2002), created a series of IgM Tg mice expressing a range of low-affinity receptors for NP, derived again from the V-gene sequences recovered from extrafollicular foci. It was theorized that in the Tg mice, the lowaffinity B cells would be dominant in the pre-immune repertoire, with little likelihood of higher affinity competitors. Thus, in these Tg mice the inherent potential of very low affinity B cells could be revealed. Both low (1.2 * 105 M-1) and extremely low (<5 * 104 M-1) Tg B cells generated serum Ab and GC responses. In the very low affinity mice, GC responses were small, most likely due to inefficient early clonal selection of these very low affinity cells (S. Anderson, A. Khalil, and M. Shlomchik, manuscript in preparation). Interestingly, the GCs of both the low and very low affinity Tg began to show substantial numbers of B cells expressing V genes derived from the endogenous heavy chain loci (as determined using reagents specific for the endogenous IgHb loci of the CB.17 background). The prevalence of these B cells increased over time; microdissection and sequencing showed that they included VH186.2 genes and thereby represented high-affinity competitors that must have been very rare at the start of the response yet emerged to outcompete the low-affinity B cells in the GC. Nonetheless, these experiments showed that low-affinity B cells are not blocked from GC entry per se. This further suggests that the early enrichment of VH186.2 genes that is seen as the extrafollicular response progresses is due to competition.
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However, the relatively weak GC responses of the low and especially the very low affinity B cells suggested that there could be intrinsic defects of low-affinity B cells participating in GCs, even in the absence of competition. Subsequently, Nussenzweig and colleagues made a series of similar V-gene Tg mice, using V genes that confer low, medium, and high affinity for NP in the context of the l light chain (Shih et al., 2002). These workers used a “knock in” versus Tg approach, using homologous recombination in ES cells to insert the Tg in the normal Jh locus. Thus, their V genes could undergo isotype switch and somatic hypermutation. They obtained essentially similar results to Dal Porto et al. (2002), but were able to additionally do a very elegant experiment in which they visualized the competition process directly. Using B cells of differing affinity that were allotype marked, they showed in a transfer immunization-type study that the high-affinity cells rapidly overtook and caused the elimination of the low-affinity cells; yet, when the lowaffinity cells were transferred without competitors, they were able to expand and populate GCs.
THE GC IS A SECOND MAJOR SITE FOR AFFINITY MATURATION From the previous discussion, it is clear that GCs can initially be seeded by a number of low-affinity B cells that have at least undergone partial selection during the extrafollicular reaction. It is likely that there will also be some higher affinity GC precursors, but that low-affinity cells will generally be much more numerous than high-affinity for any given antigen. In this section, we focus more on the unique diversification and selection processes of the GC, mediated via intraclonal diversification and consequent intraclonal selection. All of these depend on somatic hypermutation to create diversity. During the course of a normal immune response to a protein Ag or a transient viral infection, it seems that somatic hypermutation is restricted to the GC (Apel and Berek, 1990; Jacob et al., 1991b; Jacob and Kelsoe, 1992). Yet, although it has recently been discovered that the protein AID is required for somatic hypermutation (Muramatsu et al., 2000), the signals that induce hypermutation are poorly understood. It is not the main point of this chapter to discuss these signals, yet it is important for our purposes to recognize that T cells are required at least for the induction and most likely the maintenance of hypermutation (Dal Porto et al., 2002; de Vinuesa et al., 2000). There has been very limited success in reconstructing hypermutation in vitro using primary normal B cells (Bergthorsdottir et al., 2001; Dahlenborg et al., 2000; Kallberg et al., 1996). In those cases where some success has been reported, ligation of CD40 has seemed critical, as have certain cytokines. Yet it is difficult to separate out which signals are needed to support B-cell
proliferation and survival versus those that specifically induce hypermutation (Pound et al., 1999). Indeed, these could be even completely overlapping in the GC. In this regard, CD40 ligation is critical to an ongoing GC response per se (Bergthorsdottir et al., 2001; Foy et al., 1994; Gray et al., 1994; Han et al., 1995; van Essen et al., 1995), so it will be difficult to tease out its role, if any, in the induction of somatic hypermutation in vivo. The capacity to undergo mutation does not appear to be completely induced at the start of the GC response. Sequences recovered by Jacob et al. from GCs at day 8 postimmunization had very few mutations (1993). By days 12 to 16, there is a measurably higher rate of mutation accumulation (Jacob et al., 1993). Yet it must be said that we have relatively little specific and precise information about the kinetics of hypermutation onset. Also consistent with the idea that mutation is gradually induced in GCs is the finding that the B-cell memory compartment can contain unmutated cells (Takahashi et al., 2001). The significance of a slow onset of mutation is that founder B cells can undergo several rounds of division without the (mainly detrimental) effects of mutation, thus establishing a critical clonal mass. Indeed, a number of simulation studies have reached the conclusion that initial clonal expansion in the GC without mutation is required to prevent a high failure rate due to the deleterious effects of most mutations (Shlomchik et al., 1998); we return to this concept below. Indeed, mutations can have a variety of consequences for the B cells that harbor them. About 1/4 of all mutations are silent (S) and have no consequences. Among the replacement R mutations, it is useful to consider where in the V gene they occur to understand their effect. Mutations in the framework regions (FRs), areas that are relatively invariant and are mainly contributing to the overall immunoglobulinfold structure of the protein, are most likely to be deleterious unless they are conservative. Otherwise, they might disrupt overall Ig structure or alter the folding in a global way, such that the resulting mutant Ab is very unlikely to bind the original eliciting Ag. Shlomchik et al. have calculated that approximately half of all R mutations in FRs are nonconservative (1990). Consistent with this, in a wide variety of mature B-cell immune responses, represented either by hybridomas from secondary immunization or sequences recovered by microdissection and PCR from late GCs, the ratio of R:S mutations is about half of what would be predicted from completely random mutation, indicating that approximately half of all the R mutations that occurred eventually did not survive in the population of successful B cells (M. Shlomchik, unpublished). Conversely, in the complementarity determining regions (CDRs), R mutations could improve the affinity of the Ab, or destroy it (or have a neutral effect). The frequency of each type of R mutation depends on the particular V region as well as the nature of the Ag. Nonetheless, it is worth dis-
22. Selection during Antigen-Driven B Cell Immune Responses: The Basis for High Affinity Antibody
cussing some general considerations. Crystal structures of Abs binding to protein Ags have revealed surprisingly large combining site surfaces, with between ten and twenty residues of CDRs making direct contact with Ag (Braden et al., 1994; Braden et al., 1996; Braden et al., 1998; Padlan et al., 1989). In addition, it is clear from mutation analysis that even CDR residues that do not make direct contact with Ag can affect the affinity of the Ab indirectly (Mizutani et al., 1995; Satow et al., 1986). Thus, the potential target of amino acids that could be improved is relatively large—in the range of half of the total CDR size. Abs are not selected over evolutionary time to bind to a particular ligand, unlike the case for typical receptor–ligand pairs. Indeed, the typical affinities of Abs are much lower. Moreover, as discussed, many GC precursors will be of extremely low affinity. While many of these may not be improvable, it generally follows that there will be a relatively high frequency of R mutations that will be actually more suitable than the germline encoded residues (Furukawa et al., 1999). Again, the details of this will depend on the particular Ag–Ab pair. As long as there is both a reasonable target size for residues that can be improved and a reasonable chance to make these mutations, then the prediction is that the R : S ratio in CDRs of mature Ab responses should be greater than that predicted for random mutations (Shlomchik et al., 1990). These beneficial mutations will be fixed in the population, because the cells that carry them will have a selective advantage (we consider what that “advantage” is below). However, S mutations cannot be enriched. Indeed, high R : S ratios in CDRs are often but not always found. The reasons for this include: random fluctuation, high expected values of R : S in CDR making it hard to achieve statistical significance in further elevations, the small target size of CDR, a typically low number of mutations, and mutational hotspots that confound the analysis. Furthermore, high R : S ratios will not be found, despite the presence of antigen selection, if only a few R mutations can improve antibody; this will depend on the particular Ag and Ab system and should be more likely in the case of small, haptenic antigens. In our computer simulations, we have estimated that the target size for improvable mutations must be in the 10 to 50% range to yield consistent increases in the R:S ratio in CDRs (Shlomchik et al., 1998). This agrees with the size of combining sites in antiprotein Abs. Many investigators have used the presence of statistically significant increases in the frequency of R mutations in CDRs as evidence that selection has taken place. As long as this is done carefully, this is a reasonable inference from such data. For the reasons stated above, the absence of high R:S ratios does not mean that selection was not taking place, a conclusion that has sometimes erroneously been reached in the literature. Given that there is selection, both negative and positive, on R mutations, what is the mechanism for this selection in vivo and what does “selective advantage” mean for the
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cells? At one level, B cells with higher affinity [(which could largely stem from a slower kinetic “off” rate (Foote and Milstein, 1991)] could achieve better or longer BCR crosslinking, which may be an essential signal for the continued proliferation and survival in the GC. In addition, a higher affinity B cell would also have an advantage in capturing and processing Ag, presenting it to CD4 T cells, and thus gaining key proliferative and survival signals (Batista and Neuberger, 1998). Both of these could operate and probably do, though there is no direct evidence for one or the other. Then there is the question of what these signals mean for the cells. In other words, does a high-affinity B cell (with “selective advantage”) have a higher proliferative rate, a lower death rate, or both? Vigorous proliferation and extensive cell death are both hallmarks of the GC reaction. It is known that CD40 ligation, BCR signaling, and IL4 can all rescue GC B cells from apoptosis in vitro (Liu et al., 1989; Liu et al., 1991a). Yet these are all well-known mitogenic signals for B cells as well. At the moment, this remains a very interesting but unanswered question that should be further studied. Linked to this question is the one of how competition between cells works. Clearly, this is a local phenomenon, a concept confirmed by the fact that individual GCs can generate different outcomes simultaneously in the same spleen (Dal Porto et al., 2002; Vora et al., 1999). It seems most reasonable that B cells are competing for some scarce resources in the GC; as implied above, this could be Ag itself and/or T cell help. Competition for Ag could occur via quantitative capture and uptake by high-affinity B cells per se, such that there is literally less Ag left for low-affinity B cells if highaffinity B cells are around (Batista and Neuberger, 1998; Batista and Neuberger, 2000). A variation on this idea is that small amounts of Ab that are secreted in the GC may do the same thing; as noted before, plasmablasts are generated in the GC sporadically, and they will secrete Ab. This Ab may remain local by virtue of its capture by FcRs on FDC. Similarly, space for direct contact with a T cell should be limiting in the GC, particularly because T cells themselves are fairly sparse (Brachtel et al., 1996; Zheng et al., 1996). Higher affinity B cells that can present more peptides to T cells are more likely to form a better synapse with T cells and thus could exclude lower affinity cells. In spite of these attractive ideas, it should be said that, like many aspects of GC development, there is little direct evidence for one or the other model. One idea that has had currency is that B cells would need to bind Ag that is captured as immune complexes on FDCs and that this would drive affinity maturation (reviewed in MacLennan, 1994). Careful thought about how B cells would have access to this Ag bound up in immune complexes raises some a priori doubts about this model. Recent experiments by Hannum et al. (2000) have cast further doubt. These investigators used mice with a Tg that could not encode secreted Ig to create a system in which
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B-cell immune responses could take place in the absence of Ab to promote the capture of Ag on FDC. In these mice, normal GC reactions occurred, along with high-rate somatic hypermutation. Most notably for this discussion, very high R : S regions were seen in CDRs, comparable to the control situation in which Ab could and did complex Ag and bind it to FDC. Furthermore, particular R mutations in the CDRs of the l light chain that have been seen in normal and control anti-NP responses were also selected repeatedly in these GCs. All of these are strong indicators that effective selection was occurring in the GCs in spite of the lack of Ag on FDCs, arguing that this is not a major mechanism to drive high affinity in the GC. A final factor that must be considered in shaping selection in the GC is differentiation (Vora et al., 1999). At some frequency, dividing GC-phenotype B cells will differentiate to either memory or plasmablast cells. The signals that control this are unclear (Casamayor-Palleja et al., 1996). Yet, if affinity plays a role in these signals—for example, if high affinity cells are selected out to become memory—then this will clearly shape the outcome by “rescuing” high-affinity cells. Indeed, this may explain why the memory population in some settings appears to contain a higher fraction of cells with affinity-improving mutations than does the active GC population (Lu et al., 2001; Miller et al., 1995; Vora et al., 1999).
AFFINITY-BASED SELECTION CONTINUES AFTER THE GC REACTION HAS ENDED Up until a few years ago, the affinity maturation of the Ab response was considered to be entirely due to events in the GC. This was in spite of older data that had indicated that serologic affinity maturation could continue for months after the primary immune response, apparently well after the GC reaction had ended (Siskind et al., 1968). A potential explanation for this conundrum came from two discoveries. First, several groups reported that plasma cells could survive for long periods in the bone marrow (Manz et al., 1997; Manz et al., 1998; Slifka et al., 1998). These cells begin to accumulate 1 or 2 weeks after the onset of a primary immune response, with their numbers peaking around 4 weeks later. Evidently, these cells are very long lived, as there was little cell loss or turnover during the ensuing year. These results implicated a long-lived plasma cell population as the source of sustained serum antibody titers. Moreover, they implied that selection for higher affinity in this population might underlie the continued increase in Ab affinity observed during the first few months after immunization. Takahashi et al. (1998) directly addressed this issue by measuring the affinity of the antibody secreted by these cells in the bone marrow at various times after immunization,
including times well after the histologic GC reaction had ended. These workers noted a striking increase in the average affinity of the AFCs in bone marrow at these late times. To examine the contribution to the plasma cell population in the bone marrow from the GC, they used anti-CD40L treatments to dissolve the GC reaction at various times. This approach is effective at preventing the further development of memory cells and, when used early, blocked the accumulation of high-affinity AFCs in the bone marrow, thus indicating a GC origin for these. However, when anti-CD40L was given late, after the histologic GC reaction had ended, it had no effect on the further increase in affinity of the bone marrow AFCs. This indicated that the affinity increase was not being fed by undetected GCs and instead was based on selection among the established population, in a CD40-independent way. An analysis of memory versus AFC populations in the NP response by Smith et al. (1997) came to similar conclusions. Selection in this population was somewhat surprising, since classically plasma cells were thought to lack surface Ig and therefore should be unable to sense the presence of Ag and be selected on that basis. It is possible that the early AFCs appearing in bone marrow are plasmablasts, rather than plasma cells (Hauser et al., 2002). Plasmablasts are certainly surface Ig positive and could potentially undergo affinity-based selection. It also seems likely that there are cells intermediate between surface Ig-positive, dividing plasmablasts, and surface Ig-negative nondividing plasma cells that could undergo selection, including a possibly novel “precursor” population described by Noelle and colleagues (O’Connor et al., 2002). This aspect of the model has yet to be fully clarified. Nonetheless, it seems clear that the early phase of AFC accumulation in the bone marrow is yet another stage at which affinity maturation takes place.
AN INTEGRATED VIEW OF THE STRATEGIC DESIGN OF THE B-CELL IMMUNE RESPONSE: FUTURE DIRECTIONS An understanding of how affinity maturation occurs at multiple stages of the antigen-driven B-cell immune response provides insights into the overall design of B-cell immune responses. It also focuses attention on some of the important unresolved questions. The fact that very low affinity B cells are recruited promiscuously into B-cell immune responses (Dal Porto et al., 1998; Dal Porto et al., 2002) suggests a strategy in which, initially, a very diverse repertoire of B cells contributes. This creates a broad substrate for subsequent selection, which begins as early as the extrafollicular proliferative response in the first few days after exposure. One result of this strategy is the maximal secretion of antibody early on, albeit of low affinity. Nonetheless, since this
22. Selection during Antigen-Driven B Cell Immune Responses: The Basis for High Affinity Antibody
is IgM, it can be biologically relevant, as demonstrated, for example, in the case of an anti-phosphorylcholine Ab that is protective against Streptococcus, yet is of low affinity (Briles et al., 1982). A second consequence of promiscuous initial recruitment is the seeding of GCs with a diverse and probably large number of early precursor cells. Although most of these will not be successful, as indicated by the oligoclonal nature of the mature GC (Jacob et al., 1991a; Kroese et al., 1987; Liu et al., 1991b), it provides many possible pathways for success. Because of the random nature of somatic mutation and the “rugged affinity landscapes” of Ig V-gene sequences with respect to affinity for a given Ag (Furukawa et al., 1999; Kauffman et al., 1988), a single mutation could convert a low-affinity into a high-affinity B cell (Dal Porto et al., 1998). Thus, it seems likely that some low-affinity B cells entering the GC will be precursors for later highaffinity memory or plasma cells. These cells may even be difficult to detect in the mature GC, because they may be eclipsed by a more “garden variety” B cell with greater inherent affinity for the Ag encoded by the germline genes. Yet, it is true that secondary immune responses are often dominated by high affinity, highly mutated B cells that use V genes that are rarely if ever seen during the primary response (Berek and Milstein, 1987; Clarke et al., 1990a; Clarke et al., 1990b; Furukawa et al., 1999; Kavaler et al., 1991; Lu et al., 2001). Of course, these cells had to be present and selected during the primary response, and their seemingly sudden appearance in the memory response simply reflects the methodological inability of studies to detect them in primary GCs. Therefore, clonal selection is not happening only at the start of the immune response, but continues on, with perhaps the major filtering out of low affinity clones occurring during the GC reaction. Competition drives this selection in the GC (Shih et al., 2002), so that clonal selection at this phase occurs in a local context and is not simply attributable to the inherent affinity of a particular B cell. How this competition works is still not clear and is a major issue yet to be worked out. Related to this is the question of how selection occurs: by death, division, or differentiation? The relative contributions of these selective mechanisms are not clear. Memory cells are selected throughout the GC response, including the early phases. Memory cells can even lack somatic mutations, presumably as a result of an early exit from the GC reaction (to become a long-lived, antigen experienced cell) before hypermutation had an effect on the cell (Takahashi et al., 2001). Thus, the memory pool can be fairly heterogeneous with regard to affinity (Dal Porto et al., 1998; Furukawa et al., 1999). When selection is impaired, as in the case of a low-affinity Tg, the memory response can even be of lower affinity than the late primary Ab response, again showing that low-affinity cells can become memory cells when competition is weak (Dal Porto et al., 2002). Although
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memory B cells are thought to remain in interphase for long periods, not much precise information is available about their fate and whether there is some turnover (Schittek and Rajewsky, 1990). In particular, it is not clear whether there is a selective survival of higher affinity memory B cells over time. This is another area requiring further study. The other possible outcome of the GC response is the generation of AFCs, some of which will become long-lived. Cellular immunology experiments have shown that these can be selected, at least in the early stages of their life (Takahashi et al., 1998), but again the mechanism by which this occurs is not clear and how antigen plays a role (and what the source of antigen is) is not clear. Thus, although we know the broad outlines and outcomes of the B-cell immune response, many factors that control the process have yet to be completely understood. Among the cardinal questions that need resolution are how B-cell differentiation, whether to GC cell, memory cell, or plasmablast, is controlled during clonal expansion. What are the cellular signals, what is the role of the B-cell receptor in particular in guiding high affinity, and what is the cellular basis of selection (death and/or proliferation)? Also, the pathways of migration that B cells can take are still uncertain: Are GC founders derived from a marginal sinus precursor, or are both of these derived instead from a precursor that proliferated at the T–B border? Finally, what factors maintain longlived memory B cells, and plasma cells, and how are these related to each other? Progress in these areas will be of basic interest to immunologists and should form the science behind the future design of needed vaccines.
Acknowledgments The author thanks Drs. Garnett Kelsoe and Ann Haberman for useful discussions and critical reading of the manuscript, and Dr. Ann Haberman, Dr. Joe Dal Porto, Dr. Lynn Hannum, and Shannon Anderson for carrying out critical studies that contributed to this chapter. Supported by NIH grant R01-AI43603.
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23 Chromosomal Translocations in B-Cell Leukemias and Lymphomas A. THOMAS LOOK AND ADOLFO FERRANDO Department of Pediatrics, Children’s Hospital, Dana-Farber Cancer Institute, Boston, Massachusetts, USA
TRANSLOCATIONS ASSOCIATED WITH A BLOCK IN LYMPHOID DIFFERENTIATION
B-lineage leukemias and lymphomas are a heterogeneous group of malignancies that occur in the B-cell lymphoid compartment and have diverse cellular and histological features and different clinical characteristics. Given this pathological and clinical heterogeneity, it is not surprising that a number of different oncogenes are involved in the pathogenesis of these tumors, which involves diverse mechanisms of transformation and numerous oncogenic pathways. Among the most prominent mechanisms of B-cell oncogene activation are chromosomal rearrangements that place a proto-oncogene adjacent to an immunoglobulin (Ig) locus, resulting in the dysregulated expression of genes involved in the control of cellular proliferation, differentiation, or survival. A second mechanism of oncogene activation involves chromosomal rearrangements that fuse the coding sequences of two different genes. These fusion genes encode chimeric oncoproteins that work as constitutively active tyrosine kinases or as novel transcription factors to activate oncogenic transcriptional programs (Look, 1995). Thus, most oncogenic events that mediate the transformation of Blineage malignancies can be traced to a failure in one or more molecular pathways that control cell growth, differentiation, or survival during different stages of B-cell development (Rabbitts, 1991). In this chapter, we review the most well-characterized oncogenic events that result from translocations in leukemias and lymphomas, organized according to their effects in these fundamental regulatory processes during B lymphopoiesis (Figures 23.1 and 23.2).
Molecular Biology of B Cells
t(12;21) (p13;q22) and the TEL-AML1 (ETV6-RUNX1) Fusion Gene in Pediatric Early B-Lineage Acute Lymphoblastic Leukemias The t(12;21) is the most common translocation found in pediatric early B-lineage acute lymphoblastic leukemia (BALL), and can be detected in up to 30% of patients with this disease (Borkhardt et al., 1997; Liang et al., 1996; McLean et al., 1996; Nakao et al., 1996; Romana et al., 1995; Rubnitz et al., 1997a; Rubnitz et al., 1997b; Rubnitz et al., 1997c; Radich et al., 1997; Raimondi, 1993; Raimondi et al., 1990; Raimondi et al., 1991; Raimondi et al., 1995). Molecular assays are required for its detection because the rearranged chromosomal fragments have similar morphology by conventional cytogenetic analysis. This rearrangement results in the fusion the oligomerization domain of TEL (ETV6) on chromosome 12 and the entire coding region of AML1 (RUNX1/CBFA2) on chromosome 21 (Golub et al., 1995). TEL is a member of the ETS family of sequence-specific transcriptional repressors (Lopez et al., 1999) and is fused with different partners in other leukemic fusion oncogenes, such as TEL-PDGFRb in chronic myelomonocytic leukemia (CMML); TEL-MN1, TEL-ABL, and TEL-EVI1 in acute myeloid leukemia (AML); and TEL-JAK2 in T-cell acute lymphoblastic leukemia (T-ALL) (Golub et al., 1997). AML1 (RUNX1) encodes a transcription factor essential for definitive hematopoiesis in mammals and is also involved in the pathogenesis of myeloid leukemias through inherited or acquired mutational inactivation (Song et al., 1999; Roumier et al., 2003) or through its fusion with ETO in AML cases with the t(8;21) (Downing, 1999).
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FIGURE 23.2 Incidence of major B-cell neoplasias and characteristic translocations associated with them. CLL: chronic lymphocityc leukemia. DLCL: diffuse large cell lymphoma.
FIGURE 23.1 Frequencies of recurrent translocations and major oncogenes activated in acute lymphoblastic leukemias.
The exact role of the TEL-AML1 oncoprotein in cell transformation remains unclear, but TEL-AML1 appears to dimerize with both itself and with normal TEL protein (McLean et al., 1996). Some evidence suggests that the primary effect of the oncoprotein is its ability to compromise the transcriptional activity of AML1 (Hiebert et al., 1996). However, the observation that loss of the normal TEL allele frequently accompanies the TEL-AML1 fusion gene in ALL cases suggests that the leukemogenic effect of TELAML1 could be mediated, at least in part, through loss of function of the normal TEL protein (Cave et al., 1997; Jousset et al., 1997; McLean et al., 1996; Takeuchi et al., 1997).
11q23 Abnormalities Resulting in MLL Rearrangements in Infants and Secondary B-lineage Acute Lymphoblastic Leukemias Chromosomal translocations, deletions, and inversions involving the 11q23 chromosomal region are found in de novo acute leukemias and even more frequently in secondary leukemias that arise after chemotherapy with topoisomerase II inhibitors (Pui et al., 1989; Super et al., 1993; Ohshima et al., 1996). Especially notable is their identification in 60 to 70% of infants with acute leukemia (Chen et al., 1993; Janssen et al., 1994; Kaneko et al., 1988). More than thirty different translocations involving 11q23 have been described (DiMartino and Cleary, 1999; Look, 1997). The most common of these rearrangements in early Blineage ALL is the t(4;11)(q21;q23), present in 60% of infant, 2% of childhood cases, and 3 to 6% of adult cases (Bloomfield et al., 1989; Chen et al., 1993). These translocations join the mixed-lineage leukemia gene (MLL) a very large gene that spans 100 kb of genomic DNA (Rubnitz et
al., 1996; Cimino et al., 1998) to genes from numerous other chromosomes. Patients whose leukemic blast cells harbor MLL fusion genes have a poor prognosis. MLL is the mammalian homologe of the Drosophila trithorax (trx) gene, a master homeotic gene regulator during fly development. Trx positively controls the actions of a wide spectrum of homeotic (Hom) genes in the Antennapedia and Bithorax complexes of the fly and is required throughout embryogenesis for normal development of the insect head, thorax, and abdomen (Yu et al., 1995; Tkachuk et al., 1992; Djabali et al., 1992; Domer et al., 1993; Mazo et al., 1990). Inactivation of the murine Mll gene by homologous recombination suggests that Mll has a similar function during normal mammalian development (Yu et al., 1995; Yu et al., 1998). Mice lacking function of one Mll allele showed bidirectional homeotic transformations of the axial skeleton. These skeletal abnormalities in haplo-insufficient Mll animals are thought to result from shifts in the normal pattern of major Hox gene expression due to inadequate Mll gene dosage. Mice with an inactivation of one Mll allele also showed a number of hematopoietic abnormalities that included anemia, thrombopenia, and reduced numbers of B cells. Studies implicating mammalian Hox genes in blood cell development suggest that the phenotypes in hematopoietic cell populations of Mll hemizygous mice may also result from a mechanism involving Hox gene dysregulation (Look, 1997; Sauvageau et al., 1994; Lawrence and Largman, 1992; Lawrence et al., 1995; Sauvageau et al., 1997; Sauvageau et al., 1995; Thorsteinsdottir et al., 1997). 11q23 translocations result in the generation of chimeric RNAs that encode proteins containing the amino-terminal portion of MLL, which include MLLs, three A-T binding domains, a DNA methyl transferase homology domain, and a transcription repression domain (Tkachuk et al., 1992; Gu et al., 1992; Tkachuk et al., 1992). However, leukemogenic MLL fusion proteins do not include the two zinc-finger
23. Chromosomal Translocations in B-Cell Leukemias and Lymphomas
regions and the SET domain, which are located in the carboxyl-terminal part of the protein. To date, more than thirty different translocation partners of MLL have been cloned (Taki et al., 1999; DiMartino and Cleary, 1999). Although the truncation or loss of function of MLL may play a role in transformation (Dobson et al., 2000), a substantial number of MLL partners are involved in transcriptional regulation or show homology with transcription factors, thus supporting a specific role for at least some of these fusion proteins in the aberrant gene regulation leading to leukemogenesis (Nakamura et al., 1993; Prasad et al., 1995; Rubnitz et al., 1994). Gene expression profiles with oligonucleotide microarrays have recently shown that MLL-rearranged B-precursor ALLs (MLL B-ALL) have a characteristic gene expression signature that includes the specific upregulation of major HOX factors and the expression of numerous myeloid genes (Armstrong et al., 2002). These findings have been independently confirmed in a large group of patients studied at St. Jude Children’s Research Hospital (Yeoh et al., 2002) and support the hypothesis that MLL-BALLs are a distinct entity from ALL and AML, and that it has both lymphoid and myeloid features. Recent comparison of gene expression signatures of early B- and T-cell ALLs expressing MLL fusion genes has shown that MLL-rearranged T-cell leukemias (MLL T-ALL) are arrested at early stages of thymocyte development (Ferrando et al., 2003). In this study, both B precursor and T-cell leukemias harboring MLL rearrangements had decreased levels of gene expression involved in cell growth and proliferation. These types of MLL cases also had upregulation of specific HOX genes including HOXA9, HOXA10, HOXC6, and the HOX gene regulator MEIS1, thus suggesting that dysregulation of HOX genes plays a critical role in the pathogenesis of MLL leukemias (Ferrando et al., 2003).
t(1;19)(q23;p13) and the E2A-PBX1 Fusion Gene in Pre-B Cell Lymphoblastic Leukemias The E2A-PBX1 oncogene resulting from the t(1;19)(q23;p13) chromosomal translocation is one of the most common fusion genes found in children with ALL, occurring in 20 to 25% of ALL cases with a pre-B immunophenotype (defined by cytoplasmic but not cell surface expression of Ig heavy chain) (Mellentin et al., 1989; Kamps et al., 1990; Nourse et al., 1990; Privitera et al., 1992; Crist et al., 1990b; Raimondi et al., 1990). Patients with pre-B ALL and the t(1;19) tend to have elevated leukocyte counts and central nervous system leukemia at diagnosis (Raimondi et al., 1990; Pui, 1995; Pui, 1995; Crist et al., 1990b; Raimondi et al., 1990). Aside from the adverse impact of these features, the E2A-PBX1 fusion gene was shown to be independently associated with a poor prognosis (Crist et al., 1990b), although in recent years, intensive
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chemotherapy has significantly improved the clinical outcome in these patients (Raimondi et al., 1990). The structure of the E2A-PBX1 chimeric factor invariably contains the two transcriptional activation domains encoded by the E2A gene located in chromosome band 19p13, but not its bHLH DNA-binding/protein interaction domain (Kamps et al., 1990; Nourse et al., 1990; Mellentin et al., 1990). The homeodomain, DNA-binding domain motif of PBX1 is present in the chimeric oncoprotein and enables E2A-PBX1 to function as a transcription regulator (McGinnis et al., 1984; Scott and Weiner, 1984; Van Dijk et al., 1993; LeBrun and Cleary, 1994; Lu et al., 1994; Kamps et al., 1990; Nourse et al., 1990). PBX factors are the mammalian homologs of the Drosophila protein, Extradenticle (exd) (Rauskolb et al., 1993; Rauskolb et al., 1993). Exd binds to and works as a co-factor for the products of homeotic genes responsible for body patterning during fly development (Chan et al., 1994; Rauskolb and Wieschaus, 1994; Van Dijk and Murre, 1994; Johnson et al., 1995; Rauskolb and Wieschaus, 1994; Van Dijk and Murre, 1994; Johnson et al., 1995). Similarly, human PBX factors interact with specific HOX proteins (Chan et al., 1994; Van Dijk and Murre, 1994; Chang et al., 1995; Lu and Kamps, 1996; Neuteboom et al., 1995; Van Dijk et al., 1995; Lu et al., 1995; Lu and Kamps, 1997; Knoepfler et al., 1996; Chang et al., 1996), and this interaction determines the target genes regulated by HOX transcriptional complexes. As previously described, HOX genes play a critical role not only in embryonic development, but also in the regulation of hematopoiesis. Dysregulation of HOX gene expression is implicated in the pathogenesis of human leukemias (Look, 1997). The fusion of the transcriptional activation domains of E2A to the homeodomain of PBX1 in the E2A-PBX1 chimeric factor results in the dysregulation of transcriptional networks regulated by HOX genes during normal lymphoid development (Privitera et al., 1992; Borowitz et al., 1993). This hypothesis is also supported by studies in animal models that show that the leukemogenic effect of E2A-PBX1 is mediated, at least in part, through its effects on the differentiation of lymphoid progenitors at particular stages of development (Kamps and Baltimore, 1993; Dedera et al., 1993).
t(3;14)(q27;q32) and BCL6 Activation in Diffuse Large Cell Lymphoma The t(3;14)(q27;q32) and related translocations in B-cell lineage diffuse large cell lymphomas (DLCL) led to the discovery of the BCL6 proto-oncogene (Kerckaert et al., 1993; Ye et al., 1993; Miki et al., 1994). Rearrangements of BCL6 are found in 30 to 40% of DLCL, 4 to 14% of follicular lymphomas, and 20% of acquired immunodeficiency syndrome–associated DLCL (Willis and Dyer, 2000). BCL6 is a sequence-specific transcriptional repressor that contains
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six Kruppel-type zinc-finger motifs in its carboxyl terminal region and an amino-terminal POZ/BTP motif involved in homodimerization and heterodimerization (Onizuka et al., 1995; Bardwell and Treisman, 1994; Zollman et al., 1994). BCL6 represses transcription at least in part through the recruitment of both the SMRT co-repressor complex and a SMRT/mSIN3A/HDAC complex (Dhordain et al., 1997). BCL6 is normally expressed during the development of B lymphocytes (Cattoretti et al., 1995; Flenghi et al., 1995) and plays a critical role in the activation and proliferation of B cells within the germinal center and in the generation of proper Th2-mediated immune responses (Fukuda et al., 1997; Ye et al., 1997). BCL6 targets include numerous genes involved in B-cell activation, B-cell differentiation, inflammation, and cell cycle control (Shaffer et al., 2000). A variety of chromosomal rearrangements involving the BCL6 locus place this gene under the control of heterologous promoters (Ye et al., 1995), including the three Ig genes as well as numerous non-Ig partners, resulting in the dysregulation of expression of a structurally intact BCL6 protein (Ye et al., 1995; Ueda et al., 2002). The constitutive expression of BCL6 in lymphocyte precursors interferes with normal developmental programs that control cell survival and proliferation and contributes to the pathogenesis of diffuse large-cell lymphomas (Dalla-Favera et al., 1996).
and maintenance of B-cell fate during lymphopoiesis (Busslinger and Urbanek, 1995; Neurath et al., 1995). PAX5 mediates B-cell development both by repressing the transcription of non-B lymphoid genes and by activating the expression of B-lineage–specific genes. This ability to function as either a repressor or an activator is mediated by its interactions with the groucho family of corepressors (repression), or with positive transcriptional regulators such as the TATA binding protein (activation) (reviewed in Nutt et al., 2001). PAX5 target genes include the positively regulated BLK and BLNK tyrosine kinases (Schebesta et al., 2002), the B-cell surface marker CD19, the B-cell receptor component Ig alpha (mb-1), and the N-MYC and LEF-1 transcription factors (Libermann et al., 1999; Nutt et al., 1998). By contrast, the cell surface protein PD-1 and the p53 tumor suppressor are downregulated by PAX5 (Nutt et al., 1998; Stuart et al., 1995). Given the critical role of PAX5 in developing lymphocytes, the deregulated expression of this transcription factor may contribute to the pathogenesis of LPL by interfering with normal transcriptional programs that control B-cell development.
t(9;14)(p13;q32) and PAX5 Activation in Marginal Cell Lymphomas
t(14;18) and Activation of BCL2 in Follicular Lymphoma
The t(9;14)(p13;q32) is a rare but recurrent translocation present in 50% of lymphoplasmacytoid lymphomas (LPL), an atypical subtype of lymphoma that progresses slowly and transforms infrequently (Offit et al., 1992; Busslinger et al., 1996; Iida et al., 1996). This translocation places the PAX5 locus from chromosome band 9p13 next to the enhancers of the IGH gene located on chromosome band 14q32, thus leading to the constitutive expression of the PAX5 gene in lymphoid cells of the B-cell lineage (Ohno et al., 1990; Busslinger et al., 1996; Iida et al., 1996). The importance of PAX5 in the pathogenesis of human B-cell malignancies is highlighted by the finding of potentially activating PAX5 mutations in 57% of diffuse large cell lymphomas, generated by a mechanism involving aberrant somatic hypermutation in the absence of translocations (Pasqualucci et al., 2001). The PAX5 gene encodes the transcription factor BSAP (B-cell–specific activator protein), a member of the paired homeobox family of transcription factors. Members of this family are involved in organogenesis and embryonic development and are characterized by the presence of two DNA-binding domains: the paired box and the paired-type homeodomain. PAX5 is expressed by all stages of B-cell lineage precursors, but not in terminally differentiated plasma cells, and is a key element in the specification
The BCL2 proto-oncogene was discovered as a target gene overexpressed due to its translocation next to the IGH locus by the t(14;18), the most common chromosomal translocation among human lymphoid malignancies (Tsujimoto et al., 1985a; Bakhshi et al., 1985; Cleary and Sklar, 1985; Cleary et al., 1986). The t(14;18) is found in more than 80% of follicular center cell lymphomas, a common and generally indolent type of B-cell lymphoma that occurs almost exclusively in adults and in approximately 20% of diffuse B-cell lymphomas in adults (Fukuhara et al., 1979; Yunis et al., 1987). BCL2 is the founding member of a large family of highly conserved proteins that either inhibit or promote apoptosis (Yang and Korsmeyer, 1996). The ratio between these two subsets determines the susceptibility of cells to undergo programmed cell death (Oltvai et al., 1993; Bakhshi et al., 1985; Cleary and Sklar, 1985; Cleary et al., 1986). Functional studies showed that BCL2 promotes the transformation of lymphoid cells by inhibiting programmed cell death, rather than through more typical effects on cell differentiation or proliferation (Vaux et al., 1988; Nunez et al., 1990; Korsmeyer, 1992). BCL-2 family members are recognized by the presence of up to four conserved BCL-2 homology (BH) domains designated BH1, BH2, BH3, and BH4 (Adams and Cory,
TRANSLOCATIONS ASSOCIATED WITH SUPPRESSION OF APOPTOSIS DURING LYMPHOID DEVELOPMENT
23. Chromosomal Translocations in B-Cell Leukemias and Lymphomas
1998; Kelekar and Thompson, 1998). Many BCL2 family members form homo- or heterodimers, and many also contain a carboxy-terminal hydrophobic domain that enables them to become integral membrane proteins. BCL2 is normally located in the outer mitochondrial membrane (Nguyen et al., 1993) where it plays a major anti-apoptotic role by protecting mitochondrial integrity. In response to a death signal, BH3-only members, such as BIM and BAD, are activated. These pro-apoptotic BH3-only proteins interact with the pro-apoptotic members BAX and BAK and induce a conformation change that targets them to integrate into the mitochondrial outer membrane. Ultimately, the disruption of mitochndrial integrity by BAK and BAX results in the release of molecules, such as cytochrome C and AIF, which are responsible for the activation of the apoptotic effector machinery. The presence of constitutively high levels of BCL2 in lymphoma cells with the t(14;18) contributes to the transformation of B-cell precursors by impairing the effect of cell death signals that normally regulate the development of B cells.
t(11;18)(q21;q21) and the API2-MLT/MALT1 Fusion Gene in Marginal Cell Lymphomas The t(11;18)(q21;q21) is a recurrent abnormality present in up to 50% of extranodal, low-grade lymphomas of the mucosa-associated lymphoid tissue (MALT). This translocation results in the fusion of the inhibitor of the API2 (apoptosis protein inhibitor-2), at 11q21 with a novel gene named MLT/MALT1 at 18q21 (Dierlamm et al., 1999; Akagi et al., 1999; Suzuki et al., 1999). MALT tumors are a subtype of marginal cell lymphoma, accounting for 5 to 10% of all non-Hodgkin’s lymphomas and represent the most common subtype of lymphoma arising in extranodal sites. MALT lymphomas originate in the setting of chronic inflammation triggered by infection or autoimmune disorders and frequently involve extranodal sites including the gastrointestinal tract and lung, and the thyroid, mammary, salivary, and lacrymal glands. API2 is one of the five human members of the IAP family of inhibitors of apoptosis proteins. IAPs are structurally characterized by the presence of a BIR (baculovirus inhibitor of apoptosis repeat) motif in one to three copies, a caspase recruitment domain (CARD), and a C-terminal zincbinding RING finger domain (Salvesen and Duckett, 2002). API2 is highly expressed in lymphoid cells in the spleen and thymus and suppresses apoptosis by directly inhibiting caspases 3 and 7, as well as by blocking the activation of caspase-9 by cytochrome C (Deveraux and Reed, 1999). The API2-MLT fusion protein results in the truncation of the normal API2 C-terminal to its BIR domains, which are sufficient to mediate caspase inhibition and the suppression of apoptosis (Roy et al., 1997). In the fusion protein, this truncation results in the loss of the C-terminal RING domain and
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(in most cases) of the CARD domain, possibly accelerating the ability of the BIR domain to inhibit caspases and suppress apoptosis (Roy et al., 1997). The MLT/MALT1 gene is highly expressed in hematopoietic tissues and peripheral blood and, at high levels, in lymphoid and myeloid hematopoietic cell lines. The normal function of MLT/MALT1 is unknown, and the sequence of the fusion protein varies significantly from case to case. A consistent feature of all variants of the fusion protein identified to date is that they are fused in frame, supporting a specific function of the encoded chimeric protein.
t(17;19)(q21-q22;13) and the E2A-HLF Fusion Gene in Pro-B Cell Lymphoblastic Leukemias in Adolescents A second E2A fusion gene is created by the t(17;19)(q21q22;13) rearrangement, (Raimondi et al., 1991), which joins E2A to the HLF gene within chromosome band 17q21–22 (Hunger et al., 1992a; Inaba et al., 1992). Early B-lineage ALL cases with this translocation are infrequent, but they are easily recognized at diagnosis because of their frequent association with hypercalcemia and intravascular disseminated coagulation (DIC) (Devaraj et al., 1994; Hunger et al., 1994; Inaba et al., 1992; Ohyashiki et al., 1991; Raimondi et al., 1991). The t(17;19) is identified in 0.5 to 1% of ALL cases with a pro-B immunophenotype (Raimondi et al., 1991). This translocation results in the fusion of the E2A gene on chromosome band 19p13 with the bZIP transcription factor gene HLF in 17q (Hunger et al., 1992b; Inaba et al., 1992). The chimeric E2A-HLF oncogene encodes a protein containing the transactivation domains of E2A and the DNA binding/protein interaction domain of HLF. Analysis of the effects of E2A-HLF on cell survival has provided important insight into how E2A-HLF might take control of immature lymphoid cells. The introduction of a dominant-negative form of E2A-HLF in t(17;19)-positive cells induces apoptosis of the leukemic cells, thus suggesting that the inhibition of programmed cell death contributes to the leukemogenic effect of this fusion oncogene (Inaba et al., 1996). HLF is the mammalian homolog of ces-2, a gene involved in the control of apoptosis in a specific neuronal cell during development in the C. elegans nematode (Metzstein et al., 1996). Ces-2 activation mediates the death of the sister cell of a specific pair of serotoninergic NSM neurons in the worm by blocking the expression of the prosurvival gene ces-1, which normally prevents apoptosis (Metzstein and Horvitz, 1999). Similarly, E2A-HLF regulates the expression of SLUG, a ces-1 homolog, normally responsible for the protection of hematopoietic progenitors from DNA damage–induced cell death in mammals (Inukai et al., 1999; Metzstein and Horvitz, 1999; Inoue et al., 2002). Thus, in the same way that ces-2 induces apoptosis by
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repressing ces-1 in the worm, E2A-HLF may contribute to leukemogenesis by inducing the aberrant expression of SLUG, ultimately leading to the suppression of programmed cell death in lymphoid progenitors.
TRANSLOCATIONS INVOLVING THE NFKB PATHWAY The NFKB family of transcription factors is activated by a wide variety of stimuli and regulate cell survival in the context of numerous physiologic settings. There are five mammalian NFKB family members (NFKB1, NFKB2, RelA, Rel-B, and c-Rel) that can bind to DNA as homo- or heterodimers. Under resting conditions, these the NFKB complexes are inhibited by I-Kappa-B proteins (NFKBIA or NFKBIB) that trap NFKB factors in the cytoplasm. Phosphorylation of serine residues in the I-kappa-B proteins by IKBKA or IKBKB kinases inactivates these proteins, targeting them for ubiquitinitation-mediated degradation in the proteasome and leading to the activation of the NFKB complex. The activated NFKB complex moves to the nucleus and activates the expression of numerous genes, including cytokines, cytokine receptors, survival factors, and adhesion molecules (reviewed in Siebenlist et al., 1994). The t(10;14) (q24;q32) results in the constitutive expression of NFKB2 in -5% of low-grade B-NHL and in occasional intermediate- to high-grade B-cell lymphomas (Offit and Chaganti, 1991). The activation of NFKB2 contributes to the malignant transformation by inducing the expression of the pro-survival genes normally activated by cytokines or other activators of the immune system. The complex and pleiotrophic effects mediated by NFKB, which can act as either an activator or suppressor of apotosis, are highlighted by the t(14;19)(q32;q13), a rare recurrent event in CLL that results in the constitutive expression of the NFKB2 inhibitor BCL3 (Wulczyn et al., 1992; Franzoso et al., 1992). Despite its role as an inhibitor of NFKB, physiologic induction of BCL3 during normal T-cell stimulation results in the promotion of cell survival through the activation of genes inhibited by other NFKB members (Mitchell et al., 2001). Aberrant activation of this pathway in B cells due to BCL3 translocations may contribute in this way to the pathogenesis of B-cell lymphomas. A third translocation that alters genes involved in the NFKB pathway in human B-lineage tumors is the t(1;14)(p22;q32), which is an uncommon but recurrent event in low-grade MALT B-cell lymphomas. This rearrangement results in the aberrant expression of the BCL10 gene due to its translocation into the vicinity of the IGH locus (Willis et al., 1999; Zhang et al., 1999). BCL10 is a cellular homolog of the equine herpesvirus-2 gene E10, and contains an N terminal caspase recruitment domain (CARD) present in several pro-apoptotic and anti-apoptotic molecules. In normal tissues, BCL10 is ubiquitously expressed, with
highest levels of expression present in lymph node, spleen, and testis. The analysis of Bcl10-deficient mice has shown that Bcl10 is a regulator of lymphocyte proliferation that mediates receptor signaling by B and T lymphocytes. Activation of NFKB by wildtype BCL10 results in the induction of apoptosis in most cell types. This paradoxical pro-apoptotic and lymphomagenic effect may be explained by the presence of mutations that result in truncations of this factor. These abnormalities produce two main mutant forms of BCL10: 1) proteins truncated within the N-terminal CARD domain, and 2) proteins with truncations distal to the CARD domain. CARD-truncation mutants are unable to induce death or activate NFKB, whereas C-terminal truncation mutants have lost pro-apoptotic properties but still induce NFKB activation. These data suggested that BCL10 might normally function as a pro-apoptotic tumor suppressor, whereas CARD-truncation mutants may be nonfunctional, and C-terminal truncation mutants may work as bona fide oncogenes that mediate both anti-apoptotic and proproliferative signals mediated by NFKB transcriptional targets (Wang et al., 1998a; Sonenshein, 1997). However, the initially reported high frequency of BCL10 mutations in a variety of hematopoietic and solid tumors has not been confirmed in numerous studies. Thus, the role of BCL10 mutations in MALT tumorigenesis is not resolved and requires further study.
TRANSLOCATIONS ASSOCIATED WITH INCREASED PROLIFERATION IN LYMPHOID PRECURSORS t(8;14)(q24;q23) and MYC Activation in Burkitt Lymphoma and B-Cell Leukemia Burkitt lymphomas and surface Ig-positive B-cell acute lymphoblastic leukemias regularly harbor chromosomal translocations that dysregulate the expression of the MYC proto-oncogene (Dalla-Favera et al., 1982a; Taub et al., 1982). In 80% of these cases, MYC is translocated into the vicinity of the IgH locus, leading to the formation of a t(8;14)(q24;q23) (Taub et al., 1982; Adams et al., 1983; Dalla-Favera et al., 1982b; Taub et al., 1982; Adams et al., 1983). In 15% of the cases, the translocation involves the kappa locus at chromosome 2p11, whereas in the remaining 5% of cases, MYC is translocated next to the lambda locus in 22q11 (Emanuel et al., 1984; Erikson et al., 1983; Hollis et al., 1984; Rappold et al., 1984; Croce et al., 1983; Taub et al., 1984; Hecht and Aster, 2000). MYC translocations also can be found in many other subtypes of primary B-cell malignancy and are acquired with the transition of follicular B-cell non-Hodgkin’s Lymphoma (B-NHL) to highgrade disease (de Jong et al., 1988; Yano et al., 1992). MYC is a prototypic bHLH/leucine zipper transcription factor that binds to a canonical hexameric E-box DNA
23. Chromosomal Translocations in B-Cell Leukemias and Lymphomas
sequence (5’-CACGTG-3’) through its C-terminal domain. Its N-terminal transcriptional activator domain interacts with components of the RNA polymerase transcriptional complex (Amati et al., 1992), and the bHLH-leucine zipper (LZIP) region serves as a dimerization domain (Kato et al., 1992; Amati et al., 1993; Blackwell et al., 1993). An important mechanism of MYC transcriptional regulation is its heterodimerization with the bHLH-LZIP protein MAX (Gu et al., 1993). MYC/MAX heterodimers bind to DNA and activate transcription (Grandori et al., 1996). MAX, however, also heterodimerizes with members of an extended family of other bHLH-LZIP proteins, including MAD (Ayer et al., 1993), MXI-1 (MAD2) (Foley and Eisenman, 1999; Hurlin et al., 1997), and MNT. These heterodimers bind E-box elements and repress transcription, thus opposing MYC/MAX heterodimer activity. Since MYC has a short life, the levels of MYC protein are the limiting factor in the regulation of MYC/MAX complex activity in the cell. MYC dysregulation induced by Burkitt’s lymphoma translocations presumably increases the concentration of the activating MYC/MAX complexes and promotes the expression of genes linked to cell proliferation. MYC targets include proteins that regulate cell growth, division, death, metabolism, adhesion, and motility, some of which may play important roles in cellular transformation. Most of these target genes are activated by MYC. However, MYC can also work as a transcriptional repressor by mechanisms that are yet poorly understood. The effect of MYC on the cell cycle is mediated in part by the negative regulation of cell cycle inhibitors p27 and p21 and in part by the transcriptional activation of CDC25A, a protein phosphatase that activates cyclin-associated kinases CDK2 and CDK4 (Mateyak et al., 1999; Muller et al., 1997; Galaktionov et al., 1996). Other relevant MYC target genes are the ARF tumor suppressor (Zindy et al., 1998), the catalytic subunit of human telomerase (Wang et al., 1998b; Wu et al., 1999), and genes involved in diverse metabolic pathways, including nucleotide (Bello-Fernandez et al., 1993; Mai and Jalava, 1994; Miltenberger et al., 1995; Boyd and Farnham, 1997; Pusch et al., 1997; Boyd et al., 1998; Bush et al., 1998) and protein biosynthesis (Rosenwald et al., 1993; Schuldiner et al., 1996; Jones et al., 1996).
Activation Cell Cycle Regulation in Mantle Cell Lymphoma and Myeloma The t(11;14)(q13;q34) joins the IG locus in chromosome 14 with the so-called BCL1 region on the long arm of chromosome 11 (Erikson et al., 1984; Tsujimoto et al., 1984). This translocation is characteristic of mantle cell lymphoma, but can also be detected in 15 to 20% of multiple myeloma cases (Fonseca et al., 1998) and occasionally in other B-cell malignancies, including splenic lymphoma with villious lymphocytes and B-cell prolymphocytic leukemia (Raynaud et al., 1993). The t(11;14) is now known to place the cyclin
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D1 locus in the vicinity of the IGH enhancers in chromosome 14, inducing the constitutive expression of otherwise structurally normal cyclin D1 protein in lymphoid cells bearing this translocation (Withers et al., 1991; Rosenberg et al., 1993; Rimokh et al., 1994). The identification of a proto-oncogene in the BCL1 region in chromosome 11 was initially difficult, and significant effort was necessary to identify the cyclin D1 gene (CCND1) located 110 kb distal to the original BCL1 breakpoint (Tsujimoto et al., 1985b; Louie et al., 1987; Rabbitts et al., 1988; Meeker et al., 1989; Lammie et al., 1991; Rosenberg et al., 1991; Withers et al., 1991; Brookes et al., 1992; Sander et al., 1993). D-type cyclins (cyclin D1, D2, and D3) are positive regulators of cell cycle progression that act in concert with their catalytic partners, cyclin-dependent kinases 4 and 6 (CDK4 and CDK6), to form catalytically active cyclin D/CDK holoenzymes. Cyclin D/CDK complexes phosphorylate the retinoblastoma protein (Rb) and lead to the activation of E2F transcription factors and ultimately to the expression of genes involved in cell growth and proliferation (reviewed in Fields and Jang, 1990). Several different classes of mitogenic signals control the expression of cyclin D2 and cyclin D3 in B cells. Constitutive activation of cyclin D1 expression uncouples cyclin D activity from mitogenic signals, driving a proliferative stimulus that contributes to the transformation of B-cell progenitors (Lukas et al., 1994; Banks et al., 1992). Also in support of this hypothesis, recent data from a gene expression profiling in a large series of mantle cell lymphomas has shown that the levels of expression of cyclin D1 in this disease are associated with the upregulation of genes involved in cell proliferation (Rosenwald et al., 2003). The few cyclin D1 negative mantle cell lymphoma cases detected in this series showed high levels of expression of cyclin D2 and cyclin D3. Prime examples of cell cycle machinery dysregulation in human malignant B-cell lymphomas are cases of splenic lymphoma with villious lymphocytes, in which CDK6 is frequently activated due to a t(2;7)(p12;q21) (Corcoran et al., 1999). In addition, two other translocations involving cyclin D loci have been associated with B-lineage malignancies. A t(12;22)(p13;q11), inducing the aberrant expression of the cyclin D2 gene, has been described in one case of chronic lymphocytic leukemia (CLL) with progression to high-grade lymphoma (Ritchter’s syndrome) (Qian et al., 1999). In occasional cases of multiple myeloma, the malignant plasma cells harbor a t(6;14), which causes overexpression of the cyclin D3 gene (Shaughnessy, Jr. et al., 2001).
t(9;22)(q34;q11) and the BCR-ABL Fusion Gene in Adults with Early B-Lineage Acute Lymphoblastic Leukemia The t(9;22), which creates the der(22) Philadelphia chromosome (Ph), is the cytogenetic hallmark of chronic
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myeloid leukemia. However, it is also found in about 3 to 5% of children and about 30% of adults with B-precursor ALL (Devaraj et al., 1995; Westbrook et al., 1992; Tuszynski et al., 1993; Saglio et al., 1991). The t(9;22) has ominous prognostic implications in both adult and pediatric ALL cases (Arico et al., 2000; Chessells et al., 1997; Crist et al., 1990a; Preudhomme et al., 1997; Schlieben et al., 1996; Secker-Walker et al., 1997; Uckun et al., 1998). Despite the use of intensified chemotherapy, relapse rates are extremely high, and allogenic bone marrow transplantation early in first complete remission is currently the best therapy to secure long-term control of this disease (Arico et al., 2000; Kroger et al., 1998; Marks et al., 1998; Uckun et al., 1998). The t(9;22) transposes the 3’ portion of the ABL gene on chromosome 9 to the 5’ region of BCR on chromosome 22, thus generating a fusion mRNA that encodes a chimeric tyrosine kinase oncoprotein (Gordon, 1999). Although routine karyotyping does not distinguish between the t(9;22) in CML and ALL, molecular analysis of the BCR and ABL proto-oncogenes, which are rearranged in both diseases, has revealed that the breakpoints of the chromosome 22 BCR gene differ in CML and ALL. This difference results in a larger transcript encoding a 210-Kd protein in CML cases and a shorter transcript encoding a 190-Kd protein in ALL cases (Chan et al., 1987; Clark et al., 1987; Kurzrock et al., 1987). Both the P190 and P210 BCR-ABL proteins can induce a myeloproliferative syndrome in vivo in mice when they are expressed in hematopoietic progenitors (Daley and Baltimore, 1988; Elefanty et al., 1990). Mechanistic studies of these fusion proteins have documented multiple signaling pathways that are activated and contribute to leukemic transformation by BCR-ABL, including the cell cycle– regulated genes MYC and cyclin D1 (Afar et al., 1994; Afar et al., 1995), the transcriptional signal transducer STAT5 (Shuai et al., 1996), and the RAS signaling pathway, which is coupled to BCR-ABL by adapters such as GRB2, SHC, and CRKL (Goga et al., 1995; Senechal et al., 1996; Raitano et al., 1995). The development of STI-571, a synthetic tyrosine kinase inhibitor of the BCR-ABL oncoprotein, with specific antileukemic effects in leukemias expressing BCR-ABL, has broadened the therapeutic options in the management CML and best exemplifies how the discovery of the molecular basis of leukemia subtypes can result in the development of novel therapies with selective antitumor activity (Sausville, 1999; O’Dwyer and Druker, 2000).
t(4;14)(p16;q34) and Activation of FGFR3 and MMSET in Multiple Myeloma The t(4;14)(p16;q34) is a translocation only detected by molecular approaches. It is present in 15% of MM cases (Avet-Loiseau et al., 1998; Chesi et al., 1998). This
rearrangement places two different loci on chromosome 4 band 4p16, under the control of Ig heavy chain regulatory elements in chromosome 14. These oncogenes are the fibroblast growth factor receptor 3 gene (FGFR3) and the five exons of MMSET/WSCH1 gene. FGFR3 is a member of the high-affinity fibroblast growth factor receptor family of tyrosine kinases, which is not normally expressed in plasma cells. Constitutive expression of this receptor as a consequence of the t(4;14) is frequently associated with point mutations that constitutively activate the kinase activity of this receptor in MM cell lines. Activation of FGFR3 signaling in MM cells results in the phosphorylation of the STAT3 and MAPK signal transducers. It also synergizes with the action of IL6, a key cytokine for the growth and survival of MM cells. In fact, the activated products of this translocation can ultimately induce the proliferation and survival of MM cells even after IL6 withdrawal (Plowright et al., 2000). MMSET encodes a nuclear factor with an HMG domain, four PHD-type zinc fingers, and a SET domain. The presence of a PHD and a SET domain relates MMSET to the MLL gene on chromosome band 11q23, which is translocated with multiple partners, especially in infant and secondary leukemias. Although the role of MMSET in the transformation of MM cells is not fully understood, the absence of variant translocations and the simultaneous dysregulation of FGFR3 and MMSET in t(4;14) cases suggests that both genes may play a role in the pathogenesis of MM cases with this translocation.
CONCLUSION What has been learned from chromosomal translocations about the pathogenesis of B-cell lineage leukemias and lymphomas? First, the existence of two different mechanisms of proto-oncogene activation: 1) an intact proto-oncogene is placed in the vicinity of an Ig gene that induces its aberrant expression in B-cell precursors, and 2) a fused chimeric gene forms a novel oncoprotein, often with novel transcriptional regulatory properties or constitutively active tyrosine kinase activity. Second, we have learned a great deal about the pathways that cause the transformation of B-cell precursors, many of which are also involved in the control of normal B-cell proliferation and survival. Finally, the arrival of the BCR-ABL tyrosine kinase inhibitor STI571 on the clinical scene demonstrates how the identification of the genes altered by chromosomal translocations, together with the analysis of their biochemical properties and their effects on cell proliferation and survival, have opened the field for the development of novel, highly specific anti-leukemia and anti-lymphoma drugs. Ultimately, ample reasons predict that new treatments that specifically target oncogenic signal transduction pathways
23. Chromosomal Translocations in B-Cell Leukemias and Lymphomas
will change the therapeutic management and the outcome of patients affected with these diseases.
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24 Classification and Characteristics of Mouse B Cell–Lineage Lymphomas HERBERT C. MORSE III Laboratory of Immunopathology National Institute of Allergy and Infectious Diseases National Institutes of Health Rockville, Maryland, USA
The history of mouse B cell–lineage neoplasms most likely begins in 1878, when Eberth described a mouse with massive splenomegaly associated with a heavy lymphocytic infiltrate in the liver but no lymphadenopathy or thymic enlargement (Eberth, 1878)—possibly a splenic marginal zone lymphoma in current terminology. In the 1950s, Thelma Dunn developed the first rigorous classification of mouse lymphomas that distinguished lymphocytic leukemias arising at an early age in the thymus from “reticulum cell neoplasm, Type B” lymphomas (Dunn, 1954). These “sarcomas” arose in older mice in the spleen, internal lymph nodes (including the mesenteric), and Peyer’s patches, findings we now recognize as characteristic of many B cell–lineage lymphomas. Plasmacytomas (PCT) comprised another category of lymphoid disease in her classification, without clear ties to lymphocytic or reticular neoplasms. Studies by Potter and others demonstrated that PCT, the neoplastic counterpart of Ig-secreting plasma cells, could be readily induced in certain strains of mice through the injection of peritoneal irritants (Potter and Boyce, 1962). This provided the first insights into relations between histologically defined diagnoses from the Dunn classification and tumor types later associated with the B cell lineage. The finding that serum paraproteins were associated with SJL “reticular neoplasms” (McIntire and Law, 1967) extended this connection to spontaneously occurring lymphomas. It was not until normal B cells were critically defined by the presence of surface immunoglobulin—a breakthrough attributable to the efforts of Moller, Pernis, Sell, Gell, Coombs, Raff, and others—that mouse B cell–lineage lymphomas could be identified with certainty (Shevach et al., 1972) and related developmentally to plasma cells and plasmacytomas.
Molecular Biology of B Cells
Over the ensuing decade, well documented B cell–lineage lymphomas were reported as occurring either spontaneously (Kim et al., 1979), following administration of carcinogens, or subsequent to immunization and cell transfer. The monumental discovery of the molecular mechanisms involved in the generation of functional Ig opened the floodgates to understanding B-cell development from D–J recombination in pro-B cells, to mutation and classswitch recombination in the germinal center (GC), to the post-GC evolution of memory and plasma cells. As a result, it could be said with certainty that mice infected with Abelson murine leukemia virus (MuLV) (Abelson and Rabstein, 1970), known later to harbor the v-abl oncogene, developed pre-B cell lymphomas or, under other conditions, plasmacytoma (Potter et al., 1973). Cell lines developed during these years have been invaluable for studies of B-cell differentiation, responses to ligation of surface receptors including the B cell Ig receptor (BCR), and targets for altering gene expression patterns by transfection or infection. Multiple examples of PCT, Abelson virus–transformed pre-B cell lines, WEHI 231 (immature B), 70Z (late pre-B), and BCL1 (mature B) represent only a portion of a remarkably rich repertoire. Studies of other lines have revealed the existence of previously unappreciated subsets of normal B cells, such as Ly1/CD5+ B cells (Lanier et al., 1981; Davidson et al., 1984) and lineage flexibility between B celland myeloid-lineage cells (Holmes et al., 1986; Davidson et al., 1988; Klinken et al., 1988; Principato et al., 1990). In humans, the combined powers of clinical history, histopathologic studies, and the molecular and phenotypic assessments of leukemias and lymphomas set the stage for progressively refined understandings of disease heterogeneity and certainty of diagnosis. This culminated in the recent
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366 adoption of an internationally recognized, consensus World Health Organization classification of hematopoietic neoplasms (Jaffe et al., 2001). For B-cell neoplasms, the weight given to different parameters in making a diagnosis varies from one disorder to another. However, the understanding that specific chromosomal alterations—mostly balanced translocations in B cell–lineage lymphomas—are associated with particular histologic features has provided remarkable synergy in defining tumor types. These translocations, which are often readily seen on routine karyotyping, serve to activate the transforming activity of latent proto-oncogenes (see Dalla-Favera, this volume). For B cell–lineage lymphomas, this is exemplified by the “BCL” series of translocations and oncogenes for specific lymphoma types. Examples include the activation of cyclin D1 in mantle cell lymphoma, as a consequence of translocations at the BCL1 site, and BCL6 in diffuse large B cell lymphoma (DLBCL). Most notably, the diagnosis of Burkitt lymphoma (BL) cannot be conclusively made without the demonstration of an Ig/MYC translocation (Dalla-Favera, this volume), and the same translocation is probably present in all cases of Burkitt-like lymphoma (BLL) (Jaffe et al., 2001). Characterizations of less commonly occurring translocations have provided a continuing rich source of information related to the pathogenesis of distinct B cell–lineage neoplasms. Phenotypic markers are also powerful components of the diagnostic armamentarium for human B-cell lymphomas. The expression of CD5, for example, is almost completely restricted to cases of chronic lymphocytic leukemia and mantle cell lymphoma (Jaffe et al., 2001). Efforts to define mouse B cell–lineage lymphomas using a similar compilation of criteria date to the 1980s (Pattengale and Taylor, 1983; Fredrickson et al., 1985, 1994). These efforts revealed several obvious, and other less apparent, obstacles to achieving the goal of a classification system that successfully combines histologic, phenotypic, and molecular features to establish diagnoses. At the level of karyotyping, for example, mouse chromosomes are acrocentric and vary little in size, making it difficult to detect changes other than complete chromosomal or segmental gain or loss. The only true success story in karyotyping mouse lymphomas was the identification of recurring translocations involving the IgH or IgL loci and MYC in mouse pristane-induced plasmacytomas (refigured in Potter and Wiener, 1992). Phenotypic studies of mouse lymphomas using flow cytometry as one facet of a classification system have proved to be of less value than in analyses of human lymphomas. For example, studies of an extensive series of mouse B cell–lineage lymphomas have shown that nearly 85% are CD5+, irrespective of histologic type (H.C. Morse, III, unpublished observations). This said, new opportunities to identify balanced translocations and other chromosomal anomalies using spectral karyotyping (SKY) or to define regions of chromosomal loss
Morse
or gain by comparative genomic hybridization (CGH) may bring our understanding of chromosomal alterations to a more prominent place in unraveling the pathogenesis of mouse B cell–lineage neoplasms. An ever-increasing list of antibodies for use in flow cytometry and immunohistochemistry should provide additional opportunities for crossspecies comparisons.
COMPARATIVE CLASSIFICATIONS OF MOUSE AND HUMAN B CELL–LINEAGE NEOPLASMS The many mechanisms now available for enforcing unscheduled expression or abrogating the expression of desired genes have resulted in ever-increasing numbers of new strains of genetically engineered mice. Substantial numbers of these were produced to model human neoplasms as a way to reveal previously unappreciated mechanisms involved in transformation, progression, and metastasis. Of these, many developed lymphoid neoplasms as a desired outcome. However, other transgenic or knockout strains produced with no eye to cancer biology developed hematopoietic neoplasms as unexpected by-products. The terminology used in describing these “intentional” and “accidental” neoplasms has varied tremendously from publication to publication. For example, a tumor defined as a diffuse large B-cell lymphoma in one report might be classified simply as a mature B-cell lymphoma in another. As a result, it can be difficult if not impossible to determine what lymphoma type was seen, how it might relate to lymphomas uncovered in other reported settings, and to relate these lesions to human disorders. This type of inconsistency reflected in large part the absence of a consensus nomenclature that would permit the classification of specific lesions using accepted criteria and terminology that were understandable and useful. Earlier efforts to do this have usually been the work of individuals or small groups, with results that were often idiosyncratic and sometimes difficult to relate to contemporaneous schemes for the classification of human neoplasms. To enhance the value of the literature to the community of scientists involved in studies of hematopoietic neoplasms, an international committee was convened under the sponsorship of the Mouse Models of Human Cancers Consortium. Representatives included experts in human and mouse hematopathology and nonpathologists involved in generating disease models. The challenge was to develop a consensus nomenclature for diseases seen to occur spontaneously, in genetically engineered mice and in bone marrow transduction/ transplantation systems, which would provide a community standard for use in publications describing hematopoietic neoplasms in mice. To facilitate opportunities for comparing and contrasting mouse to human disorders,
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24. Classification and Characteristics of Mouse B Cell-Lineage Lymphomas
the WHO classification of human disorders (Jaffe et al., 2001) was used as a model for developing nomenclatures for mouse nonlymphoid (Kogan et al., 2002) and lymphoid (Morse et al., 2002) neoplasms. The terminology used for the classification of mouse B cell–lineage neoplasms (Table 24.1) often reflects that used for human diseases where it was felt that neoplasms in the two species were sufficiently similar, but differs where parallels between seemingly related diseases are less certain. Both classification schemes make use of the clinical, histologic, immunophenotypic, and genetic characteristics of specific disorders, with all modalities contributing to an establishment of the human categories. For the mouse, however, some categories are known only from their clinical occurrence and histologic features. As a result, the foundation for the proposed classification for mouse lymphomas is considerably less strong, and changes can be expected as more data become available. The types of data that are likely to result in changes of both classification systems are rapidly becoming apparent. First, the application of cDNA microarray technology to studies of human B cell–lineage lymphomas has opened new avenues to understanding the heterogeneity of lymphomas within a single histologic type. Studies of DLBCL revealed two subsets with distinct gene expression signatures associated with different clinical outcomes (Alizadeh et al., 2000). Second, analyses of Ig variable (V) region sequences in cases of sporadic CLL revealed two subsets. One has germline, unmutated V regions, and the second has mutated sequences (Cai et al., 1992; Schroeder and Dighiero, 1994; Maloum et al., 1995; Oscier et al., 1997; Fais et al., 1998), suggesting origins from pre-GC naïve B cells and GCexposed B cells, respectively. These distinctions are also of clinical significance, because the unmutated cases have a less favorable overall survival than the mutated cases. Similar observations have been made in studies of familial cases of CLL (Sakai et al., 2000). Remarkably, gene expression profiling of mutated and unmutated cases revealed a homogenous profile similar to that of normal memory B cells (Klein et al., 2001; Rosenwald et al., 2001). The application of microarrays and V-region sequencing in studies of mouse lymphomas is in its infancy. A quick overview of the lists of human and mouse B cell–lineage neoplasms indicates that, although substantial numbers of human lymphoma types have no parallels among the known mouse lymphomas, only two lymphoma types may be mouse specific. One of the simplest explanations for this disparity is that the lymphoma/leukemia types that are seemingly human specific occur at low frequencies in older individuals (median ages of 55 to 70) or in the setting of HIV infection. The seeming deficit in mouse diseases might also reflect the fact that only a limited number of strains have been studied intensively for lymphomas throughout their lifespan. Inbred, retrovirus-congenic NFS.V+ mice and the
TABLE 24.1 Comparative classifications of mouse and human B cell-lineage lymphomas and leukemias Mouse
Human
Precursor B cell neoplasms Precursor B cell lymphoblastic lymphoma/leukemia (pre-B LBL)
Pre-B LBL
Mature B cell neoplasms Small B cell lymphoma (SBL)
Chronic lymphocytic leukemia (CLL)/ small lymphocyte lymphoma (SLL) B cell prolymphocytic leukemia Lymphoplasmacytic lymphoma/ Waldenstrom macroglobulinemia Mantle cell lymphoma Hairy cell leukemia
Splenic marginal zone B cell lymphoma (SMZL)
SMZL
Follicular B cell lymphoma (FBL)
Follicular lymphoma
Diffuse large B cell lymphomas (DLBCL) Morphologic variants Centroblastic (CB) Immunoblastic (IB) Histiocyte-associated (HA) Subtypes Primary mediastinal (thymic) (PM)
DLBCL Morphologic variants (CB) (IB) Histiocyte/T cell rich Subtypes PM Intravascular Primary effusion
Plasma cell neoplasms Plasmacytoma (PCT) Extraosseus PCT (PCT-E) Anaplastic plasmacytoma (PCT-A)
Plasma cell neoplasms PCT-E
Burkitt lymphoma (BL)
BL
Burkittlike lymphoma (BLL) B-natural killer cell lymphoma (BNKL)
BLL
Plasma cell myeloma Solitary PCT of bone Primary amyloidosis Heavy chain diseases MGUS Extranodal MZL–MALT-type Nodal MZL
AKXD recombinant inbred (RI) strains are prominent examples (Fredrickson et al., 1985, 1999; Mucenski et al., 1986, 1987; Gilbert et al., 1988; Hartley et al., 2000; Morse et al., 2001a). This contrasts with the vast number of lymphomas submitted for diagnosis in the outbred human population. There are also significant differences in the development, phenotype, and characteristics of intracellular signaling that distinguish mouse and human B cells (Morse et al., 2003). These could mediate species-specific differences in the sus-
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ceptibility of B cells to transformation. For instance, IL-7 is absolutely required for the development of B cells in mice but not in humans (Peschon et al., 1994; Puel et al., 1998). In addition, humans deficient in BTK have no B cells in blood and are profoundly hypogammaglobulinemic, whereas deficient mice have modestly reduced numbers of B cells and levels of Ig (Khan et al., 1995; Kerner et al., 1995). The dominant sites of origin for B-cell lymphomas also differ for mice and humans. Most mouse B-cell lymphomas develop in the spleen and then spread to lymph nodes and other tissues. In contrast, lymph nodes are the most common source of B-cell lymphomas in humans. Finally, most mouse strains that develop B-cell lymphomas at high frequencies express high levels of endogenous MuLV. These viruses contribute to the pathogenesis of lymphomas by acting as insertional mutagens and may well contribute to B-cell lymphomas as surrogates for translocations. Of interest, the one mouse disease regularly associated with translocations that activate the MYC gene—pristane-induced plasmacytoma—develops with similar latency and frequency in the presence or absence of endogenous ecotropic MuLV (Potter et al., 1984).
FIGURE 24.1 Time course of lymphoma development in NFS.V-congenic mice.
TABLE 24.2 Frequencies of lymphoma types in selected strains with a high incidence of B cell-lineage lymphomas1 Percent of B lymphoma type Lymphoma type
CHARACTERISTICS OF MOUSE B CELL–LINEAGE LYMPHOMAS The most common spontaneous or induced hematopoietic neoplasms of mice are lymphomas. In strains other than AKR, C58, and some of the AKXD RI strains that die with precursor T-cell lymphoblastic lymphomas of the thymus before 1 year of age, malignancies of mature B cells dominate the field. The frequency of these neoplasms is highly strain dependent, and they develop with latencies usually between 1 and 2 years. An understanding of “background” lymphomas is critical for understanding whether tumor incidence, latency, or type is altered in transgenic or knockout mice. Detailed aging studies of mice such as B6;129 (Haines et al., 2001) provide critical baseline information. In strains such as NFS.V+, having a high incidence of B cell–lineage lymphomas, the latencies for different types of lymphoma are quite similar (Figure 24.1). However, even in lymphoma-prone strains, the distribution of lymphomas among the different tumor types may vary greatly (Table 24.2) (Hartley et al., 2000; Taddesse-Heath et al., 2000; Morse et al., 2001a). The histologic diagnoses of all cases presented in this table were made by a single pathologist, Dr. T. N. Fredrickson, and are thus internally consistent. In comparison with the tumors of NFS.V+ mice, the AKXD series stands out for the absence of DLBCL(CB) or DLBCL(IB), but a high proportion of DLBCL(HA). Although the number of cases is small, the absence of MZL from the CFW panel is equally striking. It is well worth noting that the category of MZL was not appreciated as a
+
NFS.V <12
AKXD
CFW
8.0
<5
SBL
12.5
1.5
<5
MZL
39.4
34.7
<5
Pre-B LBL
FBL
6.5
8.0
35
BLL
19.8
27.6
30
DLBCL CB IB HA
13.9 7.6 <1
<1 <1 20.1
17 4 4
Number of cases
368
199
23
1 Data are exclusive of composite cases comprising two different B cell lymphoma types or a T cell and B cell lymphoma. For NFS.V+ mice, cases with oligoclonal expansions of B cells were also excluded. 2 Because rearrangements of Ig light chain were not regularly evaluated, the true frequency may be higher.
common lymphoma type until Fredrickson and Lennert described it as a distinct entity (Fredrickson et al., 1999). Similarly, DLBCL(HA) was not recognized until a panel of lymphomas collected in the mid-1980s and carefully dissected for molecular features was re-evaluated for histologic features (Morse et al., 2001a). These two developments are salient examples of the synergy that can be achieved through the combined powers of histopathology and molecular biology. The following section covers some features of the histopathology of mouse lymphomas along with current understandings of molecular mechanisms and their relationship to human pathology. For readers interested in develop-
24. Classification and Characteristics of Mouse B Cell-Lineage Lymphomas
ing a better understanding of the histologic features of these lymphomas, useful images are available in several excellent references (Fredrickson and Harris, 2000; Taddesse-Heath and Morse, 2000).
A Precursor B-Cell Lymphoblastic Lymphoma/Leukemia (Pre-B LBL) These lymphomas and leukemias are composed of smallto medium-sized blast cells with scant cytoplasm, an oval nucleus, condensed chromatin, and often a central nucleolus. They exhibit a high mitotic index and extensive apoptosis, and frequently sport macrophages filled with apoptotic bodies (tingible body macrophages) that generate a “starry sky” appearance. This presentation is indistinguishable from that of precursor T-cell LBL or the more mature sIg+ BLL. However, molecular and phenotypic features of pre-B LBL are those of normal pre-B cells. They exhibit rearrangements of IgH but not IgL and express early B cell–lineage surface markers. In contrast to most B cell–lineage tumors, they present with lymphadenopathy rather than splenomegaly. Spontaneous tumors of this type are well documented in SL/Kh mice (Yamada et al., 1994) and the AKXD RI strains (Morse et al., 2001a), although the latter have not been characterized for lack of surface Ig and other phenotypic features of pre-B LBL. Pre-B LBL also develops rapidly in strains with oncogene transgenes such as Em-myc (Adams et al., 1985) or MT-BCR/ABL (Heisterkamp et al., 1990); at low frequency in TEL/AML1 bone marrow transduction/ transplantation (BMT/T) studies (Bernardin et al., 2002); in BMT/T analyses of the transforming activity of BCR/ABL (Li et al., 2001); or in mice infected with any of a series of acutely transforming retroviruses (reviewed in Rosenberg and Jolicoeur, 1997). Mice bearing both Em-myc and Empim-1 transgenes develop the disease in utero (Verbeek et al., 1991). Human pre-B LBL is associated with a series of fusion genes including BCR/ABL, AF4/MLL, PBX/E2A, and TEL/AML1 that individually are found in 2 to 20% of cases. Histologic, phenotypic, and molecular characteristics shared with the human disease, and the ease with which “phenocopy” mouse models can be generated with relevant constructs, indicate that these are truly homologous diseases.
Small B-Cell Lymphoma (SBL) Spontaneous lymphomas of this type are seen at varying frequencies in old mice of different strains (e.g., Table 24.2) and can be associated with secondary leukemias in perhaps a third of cases. The leukemic phase of the disease is characteristic for all mature B-cell lymphomas in that it appears to represent a spillover into the blood of a neoplasm that originates in spleen or node. This contrasts with human CLL/SLL, a similar disease, in which leukemia precedes lymphoma. The cells are small, about the size of normal
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cells of the mantle zone, with scant cytoplasm and condensed nuclear chromatin. Mitoses and apoptotic bodies are rare. The disease involves the spleen and lymph nodes and often the marrow, even in nonleukemic cases. In some cases, progression to immunoblastic lymphoma can be seen in discrete areas, with many immunoblasts and residual small lymphocytes, a phenomenon know as Richter’s transformation in human CLL/SLL. Recently, Bichi and collaborators described mice bearing an Em-TCL1 transgene with a VH promoter that targets expression to early B cells. Mice bearing this transgene developed CD5+ small-cell leukemias that appeared to arise in the peritoneum and spread to the spleen before appearing in the blood (Bichi et al., 2002). The fact that the cells are CD5+ is not particularly telling because, as mentioned previously, the great majority of mouse B cell–lineage lymphomas regardless of type are CD5+. Some cases of Em-TCL1 transgene-driven lymphomas closely resemble spontaneous SBL, whereas others have a substantial contribution from prolymphocyte-like cells (H.C. Morse III and J.W. Ward, unpublished observations). TCL1 is normally expressed in B cells until shortly after they enter the GC. These observations suggest that TCL1 or other genes in the TCL1 pathway may contribute to B cell–lineage lymphomas. This suggestion is reinforced by contemporaneous studies from the Teitell laboratory that demonstrated that another TCL1 transgene drives the development of several types of mature B-cell lymphoma (Hoyer et al., 2002). The AKT kinase pathway may well contribute to B-cell transformation in these systems, because TCL1 functions as a coactivator of AKT (Laine et al., 2000). Human CLL/SLL, the disease most similar to mouse SBL, has no established genetic cause although over 80% of cases have abnormal karyotypes (Jaffe et al., 2001). The terminology of SBL was adopted for this mouse disorder instead of SLL because similarities to the human disease are less consistent than for preB LBL (Morse et al., 2002).
Splenic Marginal Zone Lymphoma (MZL) This lymphoma occurs spontaneously at relatively high frequency in NFS.V+ mice and AKXD RI strains (Table 24.2) but was initially appreciated in the autoimmune strain, NZB (Yumoto et al., 1980). The tie of splenic MZL to autoimmunity is strengthened by unpublished observations that two strains of mice bearing three lupus-susceptibility genes (Morel et al., 2000) develop MZL (B.P. Croker, J.M. Ward, T.N. Fredrickson, H.C. Morse III, and L. Morel, manuscript in preparation). Studies of large numbers of mice showed that MZL begins in and is usually confined to the spleen. Restriction of disease to the spleen is remarkably similar to human splenic MZL, which can often be cured by splenectomy. In contrast to the cells of SBL and pre-B LBL, cells of MZL have moderately to highly abundant
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eosinophilic cytoplasm in association with an oval nucleus and nucleoli that are often hard to define. When the disease is confined to the marginal zone, mitoses are rare, and there is no apoptosis. Studies of mice that involved biopsy of the spleen when there was only modest enlargement—followed by autopsy when mice presented with advanced disease— showed that the disease progresses from low- to high-grade (Fredrickson et al., 1999). With progression, the cells increase in size and take on features of centroblasts while invading the red pulp and compressing the white pulp. Mice deficient in p53 develop an accelerated form of the disease (Ward et al., 1999). Notably, mutations and/or allelic loss of p53 were detected in nearly 20% of human SMZL (Gruszka-Westwood et al., 2001). Mutations of FAS are found in a substantial proportion of human nonsplenic MZL cases (Seeberger et al., 2001), but their potential role in the mouse disease has not been investigated. It is clear, however, that the FAS signaling pathway plays a tumor-suppressor role for B cell–lineage malignancies, as evidenced by the development of these neoplasms in mice and humans with germline mutations of FAS (Davidson et al., 1998; Lim et al., 1998; van den Berg et al., 2002) or FASL (Wu et al., 1996). The cell of origin of mouse MZL is unequivocally the splenic MZ B cell, whereas for the human disease it is most unlikely that this is the case (Du et al., 1997; Morse et al., 2001). Thus splenic MZL was a fully appropriate terminology for a mouse disease that may not have a true counterpart in humans, even though the names are the same.
Follicular B-Cell Lymphoma (FBL) This is the most common type of lymphoma found in old mice of many strains as well as in Em-Pim-1 transgenic mice (Repacholi et al., 1997). The disease initially develops in the spleen or lymph nodes but can disseminate widely with time. At the microscopic level, the lymphoma can be seen to occupy the white pulp but lack the eponymous follicular pattern featured by almost all human follicular lymphomas. The expanded white pulp is composed of a mixture of centrocytes and centroblasts, the normal components of the light zone of the GC (follicle center). The centrocytes, which typically predominate, are smaller and possess cleaved, angulated, elongated, or noncleaved nuclei with clumped chromatin and inconspicuous nucleoli. The larger centroblasts have round vesicular nuclei, often with two nucleoli appended to the nuclear membrane. Over 80% of cases of human follicular lymphoma have the t(14;18)(q32;q21) translocation that activates BCL2. Unpublished studies from two laboratories have failed to identify any changes in the genomic organization of the Bcl2 locus by Southern hybridization (P.K. Pattengale and P. Leder, unpublished observations; H.C. Morse III, unpublished observations). Efforts to model this disease in
mice by the transgenic approach produced high-grade lymphomas and PCT in one transgenic strain but not in others (McDonnell and Korsmeyer, 1991; Strasser et al., 1993). The lymphoma phenotype of the first transgenic did not persist after transfer to an inbred C57BL/6 background (H.C. Morse III, unpublished observations). The differing presentations and molecular features of human follicular lymphoma from the spontaneously occurring mouse disease led to the designation of FBL.
Diffuse Large B-Cell Lymphoma: Centroblastic and Immunoblastic Morphologic Variants [DLBCL(CB) and DLBCL(IB)] DLBCL, like FBL, is a spontaneous disease of old mice but also can be seen in Em-Pim-1 transgenic mice. DLBCL can develop by transformation from FBL or may appear de novo with a similar presentation of splenomegaly and lymphadenopathy. The lymphomas are composed of large transformed B cells. Specific cytologic profiles permit the distinction of morphologic variants, including centroblastic (CB) and immunoblastic (IB). Cells of the CB subset are cytologically similar to those described above for centroblasts in FBL. According to the thinking of Fredrickson and Harris (2000), three variations on this theme relate to cell of origin. These are designated follicular, marginal zone, and diffuse, the two former categories defining distinct anatomic compartments that yield cells with indistinguishable cytology. The cytologic consistency extends to diffuse CB lymphoma that has overgrown all normal landmarks, thereby preventing assignment of origin. DLBCL(IB)s are distinguished by the presence of substantial numbers of immunoblasts—large cells with intensely staining abundant cytoplasm and nuclei with prominent, sometimes bar-shaped, eosinophilic nucleoli. Mitotic activity and apoptosis can be prominent features. Histiocyte-associated (HA) DLBCL is a recently described subset of large cell lymphomas. Its existence was uncovered by reviewing the histologic features of AKXD RI lymphomas that have been a rich source of information on mutagenic proviral integrations and were previously well characterized for clonal rearrangements of IgH, IgL, and Tcr. By histologic criteria, nearly 10% of the AKXD hematopoietic neoplasms had features of histiocytic sarcomas, malignant proliferations of macrophages (Morse et al., 2001a). When the diagnoses were aligned with molecular features of the neoplasms, it showed that almost all tumors of this type were clonal for rearrangements of both IgH and IgL, indicating that a B-cell lymphoma comprised a substantial proportion of the cell population that, under the microscope, appeared to be dominated by macrophages. This tumor
24. Classification and Characteristics of Mouse B Cell-Lineage Lymphomas
group was not monomorphic, as a review of lymphomas of NFS.V+ mice with the same diagnosis revealed the presence of clonal TCR as well as Ig rearrangements (Morse et al., 2001a), features of a composite B cell/T cell macrophagerich tumor type. The DLBCL(HA) lymphoma type in the mouse may correspond to the histiocyte-rich/T cell-rich DLBCL in the WHO classification (Jaffe et al., 2001). Much work needs to be done before any firm conclusions regarding relatedness can be drawn. Finally, we have found a unique disease in mice infected helper-free with the replication defective virus (Aziz et al., 1989; Chattopadhyay et al., 1989) that causes a retrovirusinduced immunodeficiency syndrome in mice termed MAIDS (Knoetig et al., 2001). After a long latency, mice presented with dyspnea due to thymic enlargement resulting from clonal proliferations of immunoblastic B cells (H.C. Morse III and S.K. Chattopadhyay, unpublished observations). The lymphomas were transplantable, indicative of their malignancy and origin from GC-experienced B cells. These lymphomas have provisionally been termed primary mediastinal (thymic) DLBCL to suggest similarity to the human disease of the same name (Jaffe et al., 2001). Considerable work will be required to characterize this tumor type and relate it to other DLBCL. The identification of morphologic variants among human DLBCL is characterized by poor intra- as well as interobserver reproducibility. This, combined with the fact that there were no discernible molecular or clinically important associations with these subtypes, led to the suggestion that these may not be helpful distinctions. The analysis of DLBCL by gene expression profiling, however, has led to the identification of two subsets with clear clinical implications (Alizadeh et al., 2000). One subset, termed GC-like, has a significantly better prognosis than that termed activated B cell–like. How these distinctions relate to the histologically defined subsets of mouse DLBCL remains to be determined. Genetic determinants of human DLBCL include translocations of BCL2 in about 30% of cases as a probable marker for their previous existence as follicular lymphomas and transformation to higher grade. Another 30% have translocations involving BCL6, in partnership with a series of other genes. The vast majority of human DLBCL also have mutations in the 5¢ noncoding region of the gene regardless of whether the tumor has a BCL6 translocation (Migliazza et al., 1995). The characteristics of the mutations parallel those found in Ig V-region sequences for strand preference and other features, but occur at a rate ten-fold reduced from that of Ig genes. This contrasts with mouse DLBCL and PCT, which exhibited no mutations in homologous 5¢ noncoding sequences (Hori et al., 2002). However, mutation frequency for Ig sequences was not known for most mouse lymphomas examined for Bcl6 mutations. If the level of Ig mutation was very low, it is possible that a log reduction in rate at the Bcl6
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locus would not be detectable. As mentioned above, efforts to model BCL2 translocation effects in the mouse have not been successful. Generating a model of BCL6 translocations has proven equally elusive, with a series of transgene and retroviral constructs littering the landscape, perhaps soon to be joined by a knockin.
Burkitt Lymphoma (BL) The first attempts to model a human B-cell disease in mice were based on the expectation that a MYC gene deregulated by the presence of Ig regulatory sequences would reproduce human BL or mouse PCT, species-specific tumor types that nonetheless share a mechanism for driving aberrant expression of the same proto-oncogene. The resulting Em-myc mouse developed instead mostly pre-B LBL, with some that were sIg+ (Adams et al., 1985). Mice bearing a human IgH-containing YAC and a knocked-in MYC developed IgM+ IgD+ CD25- CD43+ B-cell lymphomas, consistent with the transformation of immature B cells (Butzler et al., 1997; Palomo et al., 1999). Deletion of the intronic heavy chain enhancer from the construct had no effect on tumor development. Recently, transgenic mice were described that carry a mutated human MYC gene regulated by sequences from the 3¢ lambda enhancer region (Ï-MYC mice) (Kovalchuk et al., 2000). The mice developed lymphomas diffusely involving the spleen and nodes. The cytology of the cells was distinct from that seen in any other mice, and the sections featured a striking starry sky appearance. The lymphomas were uniformly IgM+ IgD-/dull CD5- CD23-. IgH variable regions were basically germline. In spite of the last finding, it was felt that the disease might be a model for human BL. Since then, a number of approaches, detailed below, were employed to validate the model. Together, they established that these lymphomas were not GC experienced and therefore could not be a valid model of human BL. The end result is that the classification of Burkitt lymphoma will likely be dropped when the scheme shown in Table 24.1 is re-evaluated.
Burkitt-Like Lymphoma (BLL) These lymphomas, histologically indistinguishable from pre-B cell LBL or pre-T cell LBL, are common neoplasms of mice over 1 year of age (Table 24.2) (Fredrickson and Harris, 2000; Morse et al., 2002). They are distinguished from pre-B LBL by the presence of surface Ig. Previous studies demonstrated that some of these lymphomas express BCL6 protein and that 15% had changes in the genomic structure of Bcl6 (Qi et al., 2000). A translocation into the locus at a site homologous to the major translocation cluster in human DLBCL was mapped in one cell line (Qi et al., 2000). This suggested that these lymphomas might be a
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subset of DLBCL, resulting in their designation in earlier publications as DLBCL(LL), for lymphoblastic lymphomalike (Taddesse-Heath and Morse, 2000; Hori et al., 2001). The mature B-cell phenotype, the morphologic similarities to BL, and the distinctness from FBL or DLCL, however, led to the adoption of the term Burkitt-like. It is more than likely that this term will be also be dropped when the nomenclature is next revised. Based on early studies of Ig V-region sequences, the tumors appear to be GC-passaged, in the case of TCL1-induced tumors (Hoyer et al., 2002), but germline for those of NFS.V+ mice (Z. Du, F. Stevenson, T.A. Torrey, and H.C. Morse III, unpublished observations). This suggests a situation analogous to that seen in human CLL/SLL, in which cases have either germline or mutated Ig V regions.
Plasma Cell Neoplasms Plasma cell tumors of mice are neoplasms composed of Ig-secreting cells. They cover a broad morphologic spectrum from small, well-differentiated plasma cells to less mature plasmablasts and immunoblasts to high-grade anaplastic tumors. Some cases present as homogeneous populations of one morphologic type and others as a mixture of cells in various stages of differentiation. Three subsets are recognized in the nomenclature for plasma cell tumors: plasmacytoma (PCT), extraosseous PCT (PCT-E), and anaplastic plasmacytoma (PCT-A) (Table 24.1). Spontaneous cases of mature PCT are uncommon, occurring in spleen and lymph nodes of C3H, C57BL/6, and a few other strains of mice 1 year of age or older, thus making them difficult to study. The finding that PCT could be regularly induced at high frequency with much shorter latencies opened this tumor type to intensive study. Best understood are PCT that develop on the mesentery of BALB/c, NZB, or F1 mice following intraperitoneal injections of pristane (Anderson and Potter, 1969; Warner, 1975; Morse et al., 1978). The latency in pristane-primed mice is shortened and penetrance increased by infection with any of a series of acutely transforming retroviruses, a raf/myc virus being but one example (Troppmair et al., 1989). In the Bethesda proposals, the classification of PCT applies to these cases occurring in the peritoneum. Mature PCTs are also seen in the spleens and/or lymph nodes of mice bearing Em-v-abl (Rosenbaum et al., 1990), Il6 (Kovalchuk et al., 2002), and Bcl-2 (Strasser et al., 1993) and Bcl-XL transgenes (M. Potter, personal communication). These cases garnered the designation of extraosseous PCT (PCT-E). Finally, anaplastic PCT (PCT-A) have been observed as spontaneous neoplasms in the spleens and lymph nodes of NFS.V+, C57BL/6, and Em-v-abl mice. Indeed, the full spectrum of PCT types is seen in Em-v-abl transgenic mice (Rosenbaum et al., 1990; Fredrickson and Harris, 2000). Of importance, an intramedullary plasma cell
disease that is more like human multiple myeloma (MM) has been described in aging animals of one strain of mice, C57BL/Ka, but then only at a very low frequency (Radl, 1981). The difficulties inherent in this system have resulted in its not being widely adopted. Much remains to be learned before it could be accepted as a model for MM. These pristane-induced PCTs have been remarkably important tumors in biology, generating homogenous immunoglobulins that permitted structural characterization of antibody bound to antigen, providing fusion partners for hybridomas, and uncovering the regular association of MYC-activating chromosomal translocations with a unique disease in a manipulable model system (Potter and Wiener, 1992). They were instrumental in defining genetic determinants of PCT induction (Mock et al., 1993; Potter et al., 1994), the role of cytokines on growth regulation (Nordan and Potter, 1986), the effects of environment (Byrd et al., 1991), and the consequences of pharmacologic intervention to reduce inflammation (Potter et al., 1985). Shen-Ong, Cole, and their collaborators were the first to uncover the genetic determinants of PCT transformation (1982). They demonstrated that the Myc gene on chromosome 15 is usually translocated into the IgH locus on chromosome 12 by illegitimate class switch recombination, resulting in deregulation of Myc expression. Janz and collaborators have demonstrated that similar translocations can be found in pre-neoplastic conditions and that, once formed, they may undergo extensive remodeling (Muller et al., 1994; Kovalchuk et al., 1997). Mechanistically similar events are responsible for Ig translocations that activate CCND1, CCND3, MAF, and FGFR3/MMSET in MM as the probable initiating events in the transformation process (Bergsagel and Kuehl, 2001). Translocations involving the MYC locus are seen in 15% of primary MM, but they partner infrequently with the IgH locus and are currently considered to be secondary rather than initiating events (Shou et al., 2000; Bergsagel and Kuehl, 2001; Avet-Loiseau et al., 2001). The unusual genetic susceptibility of BALB/c mice has been dissected in increasingly fine detail with crosses to PCT-resistant DBA/2 (Mock et al., 1993, 1997; Potter et al., 1994), thus demonstrating the contributions to susceptibility of an efficiency allele at the Pctr1 modifier locus (Zhang et al., 2001). Similar investigation of susceptibility to PCT induced by an Em-v-abl transgene is at an earlier stage, but shows multiple tumor modifier loci differences between high-susceptibility BALB/c and lowsusceptibility C57BL/6 mice (Symons et al., 2002). Remarkably, BALB/c mice raised specific-pathogen free (SPF) are resistant to induction with pristane (Byrd et al., 1991). Similar but less powerful effects of environment on tumor development have been observed with NFS.V+ mice. Mice raised SPF develop the same spectrum of lymphomas as mice raised in conventional conditions,
24. Classification and Characteristics of Mouse B Cell-Lineage Lymphomas
but with a time course offset by nearly 5 months (J.W. Hartley, T.N. Fredrickson, and H.C. Morse III, unpublished observations).
B-Natural Killer Cell Lymphoma This lymphoma, which has no known counterpart in human disease, occurs only in thymectomized (SL/KhxAKR/Ms)F1 mice (Lu and Hiai, 1999). The lymphomas present with splenomegaly and lymphadenopathy in mice averaging around 1 year of age. Affected mice could live several months after first exhibiting splenomegaly. Histologic studies show diffuse involvement of lymphoid tissue and infiltration of the liver with lymphoblasts, centroblasts, and immunoblasts that, by electron microscopy, were found to contain large granules. The granules are positive for lysozyme and acid phosphatase. By flow cytometry, the tumor cells are shown not only to express IgM, B220, CD5, and CD11b but also to express NK1.1. This unique combination of attributes suggests their designation as B-natural killer cell lymphoma.
PATHOGENESIS Proviral Insertional Mutagenesis The molecular mechanisms involved in the pathogenesis of spontaneous lymphomas occurring in low-incidence strains have received little attention. This contrasts with our understandings of pathogenesis in strains that develop leukemias or lymphomas at high frequencies. Almost all these strains have one or more germline copies of full-length, replication-competent ecotropic MuLV that are expressed at high levels and contribute to disease through their reinsertion in somatic cells in the vicinity of protooncogenes or, less commonly, tumor suppressor genes. This process, known as proviral insertional mutagenesis, was first described in studies of avian bursal lymphomas where it was found that MYC was activated by recurring insertions near the locus (Hayward et al., 1981). Traditionally, the integration sites were cloned in phage, and unique sequence probes were developed from the virus flanks that could be used to probe DNA from other tumors to detect structural changes. Studies of Moloney MuLV-induced rat T-cell lymphomas by Tsichlis and his collaborators uncovered recurring integrations at several sites, one of which was later shown to be MYC (Tsichlis et al., 1983, 1985). The newly acquired integrations were clonal and involved a high proportion of cells, lending credence to the proposition that the mutations were causally linked to lymphoma development or progression. Recently, the techniques of inverse PCR (IPCR) and PCR-based splinkerette amplification have been used to analyze sequences at proviral integrations in large-scale
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screens of mouse leukemias and lymphomas (Li et al., 1999; Hansen and Justice, 1999; Suzuki et al., 2002). This approach has also been used to identify genes that can collaborate with preset loss- or gain-of-function mutations to induce transformation (Hwang et al., 2002; Lund et al., 2002; Mikkers et al., 2002). The studies of Suzuki et al. were focused primarily on B-cell lymphomas of AKXD mice that were well characterized histologically (Morse et al., 2001a). The combined studies identified 152 loci that were targeted in more than one tumor, thereby defining a common retroviral integration site (CIS). The linked genomic sequences provided tags for the discovery of genes involved in cancer (Suzuki et al., 2002). Nineteen of the CIS localized to genes previously known to be mutated in human lymphoid and nonlymphoid hematopoietic neoplasms including PAX5, MYC, and CCND1. Genes at other CIS are frequently mutated in nonhematopoietic tumors; ZFP217 (breast) and NMYC1 (neuroblastoma) are two such examples. More than 50 of the CIS were restricted to B-cell lymphomas, with eight being specific for BLL and one specific for MZL (Suzuki et al., 2002). These data demonstrate that proviral tagging can be an extremely powerful approach to identifying genes involved in cancer. An important distinction between the traditional cloning of integration sites and the IPCR approach is that the latter provides no information on the proportion of cells in a primary tumor that contain the integration. Indeed, Southern analyses of tumors using probes cloned from IPCR virus flanks sometimes fail to demonstrate any change in the genomic structure of the site in the very tumors from which the virus was cloned. One interpretation is that these CIS could be progression genes present in subclones that have yet to become dominant. Alternatively, they could be sites of little or no functional import that are passively carried along in a small clone. It will be important in future studies to relate integration sites in primary tumors to gene expression determined by cDNA and tissue microarrays to see how frequently genes at specific CIS are altered in their expression for a particular type of lymphoma. The power this approach brings for understanding genes that can contribute to B-cell neoplasia is substantial, but how does this information relate to the pathogenesis of human non-Hodgkin lymphoma? Previous studies demonstrated that Pim family genes were activated in a high proportion of lymphomas occurring in Em-myc mice infected with Moloney MuLV (van Lohuizen et al., 1991). The cooperativeness of these genes for transformation was proven by studies of Em Myc/Em Pim1 double transgenic mice that died of lymphomas perinatally (Verbeek et al., 1991). Altered expression of PIM1 may also contribute to the pathogenesis of human DLBCL as a consequence of point mutations in the 5¢ noncoding region that could affect transcription (Pasqualucci et al., 2001). Notably, similar mutations of PIM1 were not found in human BL, so PIM may
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not cooperate with MYC for transformation in this setting. It is possible that mutations of genes that act either downstream or parallel to PIM may serve to cooperate with MYC in human BL, much as activation of Pim1 functions in the tumors of Em-myc mice. Species-specific differences in genes that complement dominant genes for transformation would clearly complicate efforts to fully model human lymphomas in mice using transgenic or other approaches.
Gene Expression Profiling The analyses of human lymphomas for gene expression patterns through the use of microarrays has uncovered previously unappreciated subsets of specific tumor types with clear clinical implications (Alizadeh et al., 2000; Klein et al., 2001; Rosenwald et al., 2002). Recently, using oligonucleotide arrays we studied a large number of primary mouse lymphomas and cell lines for the expression of 7,400 genes. The primary tumors included cases of MZL, SBL, DLBCL(IB), DLBCL(CB), FBL, BL, and BLL. All the diagnoses were made and reviewed by a single pathologist, Dr. T. N. Fredrickson, obviating interobserver bias. We also studied PCT cell lines. For controls, we tested the spleens of nude mice as representative of resting B cells in vivo and LPS-stimulated nude spleen as an activated B-cell population. These studies have yielded some interesting results. First, using a hierarchic clustering algorithm to view the data (Figure 24.2), we found a remarkable concordance for diagnoses made almost exclusively on histologic grounds and
molecular profiling. All cases of primary BL and BLL, as well as the PCT cell lines, fell into three exclusive branches of the family tree. The groupings of DLBCL(CB) and SBL were only slightly less tight. All the MZL fell into a single third-order branch of the tree, but may comprise two subsets. Most FBL localized to the same branch that contains the MZL, although some cases appeared to be only distantly related. The category of DLBCL(IB) was the most widely dispersed. The sets of “resting” B cells clustered perfectly and lay with the low-grade MZL and FBL. Activated B-cell preparations also grouped nicely, lying next to PCT and BL. The same data parsed by a multidimensional scaling algorithm showed that PCT and BL lay away from a massing of other lymphoma types. The distinction of BL and PCT from the other lymphoma types does not seem to be attributable to a MYC signature. If PCT and BL are removed from the picture, SBL, MZL, DLBCL(CB), and BL form their own distinct clusters (Figure 24.3). Thus, two independent, unsupervised modes of analysis demonstrated that histologically defined lymphomas often grouped similarly when examined for their patterns of gene expression. These results appear to provide a firm foundation against which to test other B cell–lineage lymphomas that occur spontaneously or after manipulation of the mouse genome. This brings us back to the starting point of this chapter and the rationale for studying mouse neoplasms. If specific mouse B lymphoid neoplasms can now be identified by their histologic and molecular fingerprints, do these have any resemblance to features of purportedly similar human diseases and, if so, can we use these models to better under-
FIGURE 24.2 Hierarchic clustering of mouse B cell-lineage lymphomas.
24. Classification and Characteristics of Mouse B Cell-Lineage Lymphomas
FIGURE 24.3 Multidimensional scaling analyses of gene expression patterns for mouse B cell-lineage lymphomas.
stand pathogenesis and develop effective treatments? Can these mouse diseases provide us with novel understandings of B-cell transformation, irrespective of their relations to human neoplasms? The answer to the latter question appears to be an unequivocal yes, given the numerous important understandings of proto-oncogenes and tumor suppressors that derive from analyses of insertional mutations. The answer to the first will be known only after extensive efforts to validate each mouse disease in relationship to its apparent human counterpart. The first such test has been performed with mice bearing a Ï-MYC transgene that were presented as a model of BL (Kovalchuk et al., 2000). This proposition was based on the demonstration of marked histologic and phenotypic similarities and a quite distinct presentation from that of Em-myc mice that exhibited an early, marked expansion of pre-B cells in bone marrow, blood, and spleen (Adams et al., 1985; Langdon et al., 1986). Ï-MYC transgenic mice had normal splenic and nodal architecture and a quite normal bone marrow population of B-cell precursors prior to tumor development. In studies designed to examine the types and frequencies of mutations occurring in normal tissues and lymphomas of Ï-MYC mice using a lacZ reporter gene, it was found that translocations, deletions, and inversions were much more common than point mutations (Rockwood et al., 2002). SKY analyses of cell lines derived from the tumors revealed that translocations, mostly nonreciprocal, involved almost all autosomes as well as the X-chromosome. The fact that chromosomal instability is not a feature of human BL suggested that at least some elements of this model were at odds with the clinical disease in humans. Further studies clearly indicated that, whereas human BL is a neoplasm of cells that have experienced the GC, the lymphomas developing in the mouse model had not. Two exper-
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iments were particularly telling. First, when the Ï-MYC transgene was crossed onto a Bcl6 null background, yielding mice incapable of generating GC (Ye et al., 1997; Dent et al., 1997), lymphomas developed with comparable frequency and latency in -/- and +/- mice (G. Cattoretti, and R. Dalla-Favera, unpublished observations). Second, analyses of Ig V-regions from a number of tumors showed them to be all germline (F. Stevenson, R. Dalla-Favera, unpublished observations). In addition, both microarray analyses of gene expression and flow cytometric studies indicated that the lymphomas had features of cells that were recent migrants from the bone marrow (T. McCarty. T. A. Torrey, and H. C. Morse III, unpublished observations), possibly mirroring transitional B cells (Carsetti et al., 1995; Allman et al., 2001). The conclusion to be drawn from this work is that the Ï-MYC transgenic mouse does not provide a model for human BL. PCT is the only other well-studied mouse lymphoma type with some similarities to a human disease. We have used array data to determine those genes that most distinguish PCT from other types of B cell-lineage lymphomas. The distinguishing genes included a series previously shown to be differentially expressed in PCT (Pet2, Xlr3a, Xlr3b, Tacstd1) (Bergsagel et al., 1992a, 1992b, 1994), others recently shown to be highly expressed in human MM (Tra1, Xbp1, Irf4, Sdc1, Grp58) (Zhan et al., 2002; Claudio et al., 2002), and some regulated by BLIMP (Vegf, Icsbp) (Shaffer et al., 2002). Other tantalizing clues to pathogenesis include the upregulation of Snrpn, an imprinted gene (Leff et al., 1992). Loss of imprinting (LOI) is very common in solid tumors, but has not been thoroughly evaluated in lymphoid neoplasms. This minor entrée into the world of epigenetics and cancer will rapidly expand in studies of mouse as well as human lymphoma and leukemia.
CONCLUSION Many of the tools that have been used so effectively to characterize and understand human B cell–lineage lymphomas have not gained full purchase with investigators studying spontaneous mouse disorders or trying to model human diseases. CGH, SKY, testing for LOI and changes in CpG methylation, tissue arrays, and the expanded use of DNA microarrays will contribute much to our understanding. Having a system of standardized nomenclature for lymphomas and a view into how molecular profiling can augment histologic studies provides an important foundation for progress.
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25 B Cells Producing Pathogenic Autoantibodies CONSTANTIN A. BONA
FREDA K. STEVENSON
Department of Microbiology, The Mount Sinai Medical Center, New York, New York, USA
Molecular Immunology Group, Tenovus Laboratory, Southampton University Hospitals Trust, Southampton, United Kingdom
During evolution, vertebrates developed an immune system able to cope with a broad universe of foreign antigens, with limited reactivity to self-antigens. It is to the host’s advantage to maintain macromolecular homeostasis, which can be perturbed by foreign molecules, and to prevent tissue damage due to microbes or their toxic products. The germline genes of lymphocytes have been selected by evolutionary forces to encode proteins able to function in the innate and adaptive immune responses. However, B cells also undergo somatic mutation in their variable region genes during the process of affinity maturation of antibody responses. This generates new potential binding sites and, since autoreactivity could arise, requires control in the periphery. It was clear to Ehrlich that the immune system must avoid the horror autotoxicus of self reactivity (1); since then, the ability of the immune system to distinguish self from nonself has been revealed as an intrinsic property. Burnet (2) postulated that unresponsivness to self-antigen results from the process of deletion of self-reactive clones during ontogeny (clonal deletion) leading to central tolerance. Central tolerance occurs during lymphocyte ontogeny as a result of negative selection in central lymphoid organs: in bone marrow where B cells are generated, and in the thymus where T cells develop. The concept of negative selection of B cells is well supported by studies carried out in transgenic mice. In mice expressing VH and VL genes encoding antibodies specific for allogeneic MHC class I molecules (3), hen egg laysozyme (4), or an allelic form of the CD8 self antigen (5), B cells specific for these antigens are deleted. Deletion occurs in the bone marrow during the differentiation of pre-B (sIg-) to immature (sIgM+) B cell stages (6). Deletion of self-reactive
clones may also occur in peripheral lymphoid organs, particularly in immature HSA+ B cells exported from bone marrow to enter into the T-cell zone of the white pulp of spleen (7). Negative selection operates at the cellular level through the alteration of signaling pathways or by apoptosis initiated by the interaction of B-cell receptor with autoantigen (reviewed in 8). Ehrlich’s concept of horror autotoxicus prevailed for several decades in spite of the fact that, as early as 1903, Donath and Landsteiner (9) described paroxysmal cold hemoglobinuria, the first identified autoimmune disease. Ehrlich himself showed that in patients afflicted with disease, hemoglobin is present in the blood of a ligatured finger after immersion in cold water and then warming it to 37°C (10). Burnet (2) postulated that, in spite of the deletion of self-reactive clones during ontogeny, autoreactive clones could arise later in life because of receptor mutation during affinity maturation. This concept was supported by data showing that antibody specific for phosphorylcholine acquired the ability to bind to DNA after the introduction of mutations in the VH gene (11). Ensuing years have seen numerous evidences of the imperfection of the immune system in distinguishing self from nonself, and the consequent development of autoimmune diseases. In the late 1940s, the breaking of central tolerance was shown by injecting brain or testicular tissue in adjuvant, thus inducing rapid acute disseminated encephalomyelitis in Rhesus monkeys (12) or aspermatogenesis in guinea pigs (13) respectively. In 1956, Rose and Witebsky (14) induced autoimmune disease, manifested by the enlargement of thyroid gland, following injection of rabbits with rabbit thyroid extract.
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These findings led Nossal (15) to propose a new concept—clonal anergy—a tolerant state in which selfreactive clones are rendered unresponsive by low doses of antigen, but are not deleted. This concept is supported by observations in transgenic mice expressing the HEL gene encoding for secreted or membrane-expressed molecules. When HEL was produced in soluble form at 1 nM in blood, low-affinity B-cell clones escaped from tolerance and were anergized. In contrast, high-affinity, immature B cells were deleted in transgenic mice expressing the cellular form of HEL, since the receptor occupancy by self antigen exceeded a critical threshold (16). More recently, studies in transgenic mice expressing V genes encoding autoantibodies showed that self-reactive lymphocytes can escape deletion by receptor editing, a process in which the transgenic VL gene is replaced with an endogenous VL gene (17). However, receptor editing may not be restricted to autoreactive B cells, since recent findings showed that nonautoreactive B cells may also be targeted for receptor editing at the pre-B cell stage (18). (Current concepts of receptor editing and its role in B-cell development are described by Nussenzweig and Weigert in this volume.) Both genetic and environmental factors are involved in the pathogenesis of autoimmune diseases. Excepting a few animal models in which the autoimmune disease is due to a single gene defect (SLE in MRL (19), gld mice (20) or BxSB mice (21), scleroderma in tight skin mice (22,23), inherited interstitial nephritis in kd/kd mice (24), and the polyautoimmune syndrome in motheaten mice (25), the majority of animal models of autoimmune diseases, and human disease, are multifactorial. Mutations in genes encoding autoantigens, MHC genes, and polymorphism in the promoter or structural genes encoding cytokines, chemokines, or costimulatory molecules may contribute. Furthermore, other factors such as molecular mimicry, defects in tolerance, the engagement of ignorant, nontolerant lymphocytes, hormonal factors, infections, and environmental factors have been considered as possible contributors to autoimmune disease. Based on tissue injury, the autoimmune diseases have been classified into two major categories: organ specific and systemic. Intensive research during the past decades aimed at understanding the effector mechanisms involved has led to another classification: 1) autoimmune disease mediated by pathogenic T cells such as multiple sclerosis, IDDM, uveitis, dermatomyositis and Crohn’s disease; and 2) autoimmune diseases mediated by pathogenic autoantibodies. This chapter is limited to describing autoimmune diseases mediated by pathogenic autoantibodies (Table 25.1)
TABLE 25.1 Autoimmune diseases mediated by pathogenic autoantibodies Disease
Target autoantigens
1. Organ- or cell-specific autoimmune disease Idiopathic thrombocytopenic purpura
Platelet antigen (GPIb, IIb, IIIa, IA/IIA)
Autoimmune hemolytic anemia: cold warm
Red blood cell antigens Ii Rh(e), Kell, LW. Vel
Paroxysmal hemoglobinuria
P antigen
Primary autoimmune neutropenia
Neutrophil-associated antigens (NA1, NA2, NB1, NB2, CD11B/CD18)
Myasthenia gravis
Acetylcholine receptor
Graves’ disease
Thyrotrophin receptor
Hashimoto disease
Thyroglobulin and thyroid peroxidase
Insulin-resistant diabetes (Kahn syndrome)
Mutated and uncleaved insulin receptor
Pemphigus vulgaris
Desmoglin
Bullous pemphigus
Hemidesmosomes (BP180, BP230)
Autoimmune polyendocrine syndrome
Mutation of autoimmune regulatory gene
Addison’s disease
Adrenal cell antigens and steroid 21 hydroxylase
Stiff-Man syndrome
Glutamic decarboxylase and amphiphysin
Rassmussen’s encephalitis
Glutamate receptor
2. Systemic autoimmune disease Neonatal lupus
Ro antigen
SLE
dsDNA and Sm
Cryoglobulinemia
RF specific for Fc gamma portion of IgG
Goodpasture’s disease
Type IV collagen
SUBSETS OF AUTOANTIBODIES Autoantibodies can be classified into three major subgroups.
Natural, Polyspecific Autoantibodies This subset constitutes a substantial fraction of the selfreactive repertoire. “Natural” serum antibodies are found in most species in various Ig classes (IgM, IgG, IgA). They bind with a moderate or low affinity to structurally dissimilar epitopes borne by self and foreign molecules (26). One example is a human monoclonal IgM specific for the capsular polysaccharide of group B meningococci and
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Escherichia coli K1 (poly-N-acetyl neuraminic acid), which cross-reacted with denatured DNA (27). Natural antibodies are predominant in the fetus and newborn (28) and are encoded by unmutated V germline genes (29). In humans and mice, they are produced by CD5+ B cells (30), a subset enlarged in some animal strains prone to autoimmune disease (31). Natural autoantibodies exhibit a high idiotypic connectivity (32). They represent a phylogenetically conserved system, which may function as the first barrier of defense in primitive animals (and possibly in vertebrate species) and may contribute to the elimination of metabolites or cell breakdown products (33). Polyspecific autoantibodies are able to cross-react with a range of antigens and can be found in increased concentration in the blood of patients with autoimmune diseases, without induction of injury to normal tissues (34).
Autoantibodies with Exquisite Specificity for Autoantigens These autoantibodies exhibit high binding affinity to selfantigens and can be found at low levels in healthy humans and animals. Their concentration can be increased in some autoimmune diseases and may be used as a diagnostic criterion (e.g., anti-topoisomerase I in scleroderma, anticentromere in CREST syndrome, anti-tRNA synthetase or anti-Jo1 in polymyositis, anti-U1RNP in MCTD), and in certain conditions they can be pathogenic. An example is represented by anti-thyroglobulin autoantibodies. Antithyroglobulin antibodies with similar epitope specificity are found in 50 to 70% of patients with autoimmune thyroiditis and 10 to 20% of normal subjects (35,36). Antithyroglobulin antibodies do not appear to be pathogenic, since passive transfer in animal models does not cause thyroiditis (37).
Pathogenic Autoantibodies These autoantibodies appear to be responsible for the onset of autoimmune disease that causes injury to tissue bearing specific target autoantigens. Although they do not exhibit particular immunochemical properties, they demonstrate distinct physiopathological properties.
CRITERIA TO DEFINE PATHOGENIC AUTOANTIBODIES In the original formulation, Koch proposed postulates for linking a specific disease to a given microbe. Witebsky et al. (38) adapted Koch’s postulates to define autoimmune diseases. Based on the progress of knowledge in this field, we revised Witebsky’s postulates and proposed additional criteria to define pathogenic autoantibodies (39,40).
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First Postulate: Identification of Autoantigen and Demonstration of Its Ability to Induce Autoantibodies and Autoimmune Disease in Animals Following identification of autoantigens by reactivity with serum from autoimmune patients, molecular biological techniques have allowed the cloning of genes encoding autoantigens and the production of recombinant proteins. Among these are human Sm (41), acetylcholine receptor (42), Ku antigen (43), histidyl tRNA synthetase (44), and UI-RNP (45). The immunogenicity of recombinant self proteins was demonstrated in vivo by their ability to induce autoantibodies and eventually an experimental autoimmune disesase. Examples include the induction of myasthenia gravis in mice immunized with AchR (46), of pemphigus in desmoglein 3-/- mice injected with desmoglein 3 (47), or of glomerulonephritis in sheep injected with type IV collagen (48).
Second Postulate: Induction of Damage or Disease by Passive Transfer of Autoantibodies Several examples of autoimmune diseases result from the natural transfer of pathogenic autoantibodies. Perinatal autoimmune diseases can occur subsequent to transfer of maternal antibodies. Examples include transient myasthenia gravis observed in newborns born to affected mothers (49). Anti-AchR antibodies can be transferred to the fetus via placenta or milk (50). The termination of myasthenia gravis in newborns has been suggested to occur through the production by infants of anti-idiotype antibodies against maternal anti-AchR autoantibodies (51). Neonatal transient Graves’ disease is a relatively rare condition resulting from the transfer of anti-TSR autoantibodies in normal (52,53) or premature infants (54). More rarely, neonatal hypothyroidism has been described in infants born to mothers affected by Hashimoto thyroiditis with anti-TSR autoantibodies (55) and anti-thyroid microsomal autoantibodies (56). Perinatal lupus in infants with congenital heart block is caused by the transfer of anti-Ro antibodies and, despite early treatment, these infants have a high mortality during the first year (57), thus demonstrating pathogenicity of maternal autoantibodies. Neonatal lupus can also arise due to transferred anti-Ro and anti-La antibodies (58). A rare case of neonatal pemphigus foliaceus was observed that appeared to be secondary to transplacental transfer of maternal antidesmoglein autoantibody (59). The induction of human disease has also been observed following experimental transfer of autoantibodies. In a unique experiment, Harrington injected himself with plasma from a patient with idiopathic thrombocytopenia. The
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injection caused platelet depletion and a severe bleeding disorder (60). In human-to-animal transfer experiments, it was shown that the injection of Ig from patients with pemphigus vulgaris into newborn mice caused typical blisters (61). Injection of monoclonal anti-AchR autoantibodies into mice and rats can cause muscle weakness and those alteration of myograms typical for myasthenia gravis (62,63).
encoding a cold hemagglutinin. These mice exhibited anemia and significant reticulosis after injection of antigen (CAGAS) (71). Table 25.2 illustrates human autoimmune diseases fulfilling the criteria defining pathogenicity of autoantibodies.
GENETICS OF AUTOANTIBODIES Third Postulate: Isolation of Pathogenic Autoantibodies From Affected Organs Using appropriate techniques, pathogenic autoantibodies have been isolated from the platelets of patients with idiopathic thrombocytopenia. Similarly, cationic anti-dsDNA antibodies were isolated from the kidney of lupus patients (64), anti-collagen IV autoantibodies from kidney or lung of patients with Goodpasture’s syndrome (65), or anti-type II collagen autoantibodies from the cartilage of patients with relapsing polychondritis (66). This criterion is strong but indirect evidence for pathogenicity.
Fourth Postulate: Occurrence of Autoimmune Disease in Transgenic Mice Expressing V Genes Coding Pathogenic Autoantibodies The occurrence of autoimmune diseases in transgenic mice expressing murine or human V genes encoding pathogenic autoantibodies represents direct and strong evidence for the role of autoantibodies in the induction of disease. The first transgenic murine model for autoimmune hemolytic anemia was generated by Okamoto et al. (67), who expressed the V genes encoding an anti-RBC autoantibody isolated from a hybridoma obtained from an NZB mouse that spontaneously developed hemolytic anemia. About 50% of Tg mice bred in conventional facilities exhibited anemia, with a hematocrit of <30% and accumulation of agglutinated RBC in the spleen. Interestingly, the same mice bred in germ-free conditions did not develop anemia, unless injected with LPS. This suggested that autoreactive B cells escape central tolerance, but their activation depends on environmental, possibly inflammatory, factors. Several transgenic mice expressing V genes encoding anti-DNA antibodies have been generated. In one model, Tg B cells binding to DNA were present, but no anti-DNA antibodies were secreted, thus indicating that the autoreactive B were anergized via a receptor editing mechanism (68). In contrast, in a second model in which the VH gene was combined with Cg2a, but not Cm, mice spontaneously secreted IgG anti-DNA antibodies and developed a mild nephritis (69). This observation correlates with the fact that pathogenic anti-DNA antibodies are IgG and not IgM (70). Another model is a Tg mouse expressing the human V genes
Molecular Characteristics of Murine Autoantibodies In mice, the functional VH genes are assembled from a minimum of 134 VH (variable), 12 D, and 4 JH genes in the repertoire (72). Although not yet fully sequenced, indications from the large VH J558 family in C.B-20 mice suggest that there may be only a few pseudogenes (73). VL genes derive from about 140 Vk, four Jk, and three Vl and three Jl gene segments. Among the Vk gene segments, it appears that about two-thirds are potentially functional (74) (see Chapter 2). In most cases, antibody specificity and idiotypic specificity are phenotypic markers of a VH gene with a VK or Vl gene. In contrast to other mammalian species, 97% of mouse Ig express k light chains. Interestingly, studies carried out in k-/- showed that, although there are only three functional Vl genes in mice, the antibody repertoire for antigens (75) or self antigens (76) can be rescued by pairing of different VH with Vl genes. Hybridoma technology facilitated the studies of the utilization of various VH and VK families in autoantibodies. With a few exceptions, autoantibodies utilize VH and VK genes from various families. A striking exception is represented by CD5+ B cells producing anti-bromelain–treated RBC antibodies. In this case, the majority of autoantibodies use VH11 or VH12 gene families (77–79). In contrast, an analysis of V-gene usage in hybridomas producing autoantibodies with various specificities obtained from motheaten mice, in which >90% of B cells express CD5, showed that autoantibodies exhibiting different specificities for autoantigens utilized V genes from VH X24, J606, J558, S107 and 7183, and, VK1, VK4, VK10, and VK19 families (80). It is noteworthy that the B cells from viable motheaten mice producing anti-RBC autoantibodies use the same VH11 and VK9 family genes as the peritoneal CD5+ B cells producing anti-bromelain–treated RBC (81). Study of V-gene usage in pathogenic autoantibodies specific for collagen type II showed derivation from various families (J558, X24, 7183, and 36–60). However, a biased usage of VK21E was observed in autoantibodies specific for the C1 epitope of collagen type II (82). Recently, a comprehensive analysis of V genes expressed in hybridomas producing autoantibodies was performed (8). This analysis showed that certain VH gene families such VH 7183, S107, and VH11 are overrepresented, whereas others
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TABLE 25.2 Autoimmune diseases mediated by autoantibodies fulfilling the criteria of pathogenic antibodies Criteria
Isolation of Ab from lesion
Disease
Transfer of disease Maternal Human
Experimental animal
Induction of disease in animals with the antigen reacting with pathogenic Ab
Occurrence of disease in transgenic mice expressing VH + VL encoding pathogenic autoantibodies
Myasthenia gravis
+
+
+
+
ND
Pemphigus
+
+
+
ND
ND
+
Graves’ disease
+
Hashimoto disease
+
Polychondritis
+
Warm hemolytic anemia
+
Cold hemolytic anemia
+
Idiopathic purpura thrombocytopenia
+
Goodpasture’s syndrome
+
SLE
+
+
+
+
+
+
+
+
ND
+
ND
ND
ND
ND
+
ND
+
ND
+
+
ND
ND
+
ND = not done or unknown.
FIGURE 25.1 Expected and observed frequency of the utulization of VH gene families in murine autoantibodies (Kaushik and Bona (8)).
such as J606, 36-09, and Gam3.8 were under-represented compared with that expected from genomic complexity (Figure 25.1). With respect to D-segment genes, the observed usage by autoantibodies was comparable to the expected frequency (p < 0.05). Analysis of the recombina-
tion of four VH genes (J558, 7183, VH11, and Q53) and three D genes (SP2, FL16, and Q52) showed that it occurs randomly (p < 0.05). Analysis of VK gene families showed that VK4, VK9, VK10, and VK24 gene families were overexpressed in B cells producing autoantibodies compared to
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FIGURE 25.2 Expected and observed frequency of the utilization of Vk families in murine autoantibodies (Kaushik and Bona (8)).
the expected frequency of VK gene family complexity (Figure 25.2). Recombination of VK with JK showed a preference for JK2 in B cells producing autoantibodies. Except for biased usage of a few VH and VK gene families, generally the utilization of V gene family in autoantibodies was comparable to random, utilization in B cells activated by LPS. Previously, we have shown that the pairing of VH and VK variable regions occurs stochastically in polyclonally activated murine B cells (83). Analysis of VH and VK pairing in 447 autoantibodies showed that the pairing is random, with the exception of anti-BrMRBC autoantibodies, in which a biased pairing of VH11 with VK9 and VH12 with VK4 e was observed. Overall, our analysis showed that the usage in some VH JH and VK JK respectively shows some bias in hybridomas producing autoantibodies, as compared with the expected usage based on the complexity of V gene families or their usage in polyclonally activate B cells. The biased usage may reflect the process of positive and negative selection of the precursor of B cells specific for self-antigens.
Molecular Characteristics of the Human V-Gene Repertoire A major advantage of studying human V genes is that the potentially functional germline repertoires of VH and VL have been fully mapped. Details of the organization of the repertoires are given in other chapters. The ability to compare expressed sequences with the corresponding
germline sequence has provided the basis for our understanding of V-gene usage and of the role of somatic mutation in the development of autoantibody responses. Polymorphic differences have been identified that may vary among different ethnic groups (84) and could influence disease susceptibility. They include variations in copy number, insertional or deletional polymorphisms, and the occurrence of different alleles of the same gene segment (85). Deletions may be important, since deletion of the VH hv3005 (V3-30.3) gene has been associated with rheumatoid arthritis (86), lupus, and chronic idiopathic thrombocytopenic purpura (87). Compared to insertions and deletions, sequence polymorphism appears to be low (85). However, it is evident in certain VH gene segments, such as the VH3b subfamily, which is highly associated with antibody responses against Haemophilus influenzae type b (88). In fact, it has been suggested that the known differences in susceptibility to infection may be associated with the absence of certain V genes (88). Another example of a polymorphic gene is the V169 gene segment, which is commonly used in antibody responses against hepatitis C (89) and often encodes rheumatoid factor activity (90). Interestingly, V1-69 is also preferentially used in a subset of the B-cell malignancy chronic lymphocytic leukemia, suggestive of a superantigenic drive on the cell of origin (91,92). The V1-69 gene segment shows extensive polymorphism, comprising at least thirteen alleles, with variable copy numbers (93). For the 51p1-related variants, increased copy number is clearly
25. B Cells Producing Pathogenic Autoantibodies
associated with increased expression (94). Although a high copy number has not been found to be associated with susceptibility to rheumatoid arthritis, there is an intriguing association between a sequence variant and the development of disease in the Czech population (95). Knowledge of the available repertoire, and the development of technology for analyzing V-gene usage in single B cells, has allowed analysis of the expressed repertoire in human blood IgM+ B lymphocytes (96,97). This has indicated important aspects of V-gene usage and of somatic mutational activity in normal B cells that subsequently reveals deviations in malignancies and in autoimmunity. In both CD5+ and CD5- B cells, expression of the different VH gene families largely reflected the available germline repertoire. However, within families, differential expression of certain V-gene segments occurred. Within the large VH3 family, there was frequent usage of certain gene segments, such as V3-23 (Figure 25.3). Interestingly, preferential expression was confined to productive rearrangements, indicating selective processes acting on the expressed gene, rather than preferential recombinatorial events. However, the latter mechanism appears to account for increased expression of other genes, such as V4-59 and V3-07. A surprisingly small number of VH genes accounted for the
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majority of the functional repertoire of both B-cell populations. Thus, nine VH family members (18% of the functional repertoire) were used by >50% of B cells. Similar biases were observed in expressed Vk genes and are discussed in Chapter 3. Somatically mutated VH sequences, presumed to derive from memory B cells, were detected mainly in the CD5- subset, with an incidence of ~50% in all three individuals analyzed. There were interesting differences in VH gene usage between unmutated and mutated sequences, assumed to result from selective pressure during affinity maturation (97). In B cells that had rearranged both alleles, the retained nonfunctional sequences could be analyzed. Since both alleles undergo somatic mutation, this provided an opportunity to investigate the distribution of mutations in the absence of antigen selection (98). The surprising finding was that clustering of replacement mutations in the CDR1 and CDR2 occurred naturally, prior to selection. This is likely to be due to susceptible sequence motifs or “hotspots” in these regions (99). The only distinctive feature of functional sequences appeared to be a reduced number of replacement mutations in the FRs, presumably required to maintain structure. These findings perturbed those of us who were assigning B cells to “antigen selected” or “non-antigen selected”
FIGURE 25.3 The repertoire of VH gene segments expressed by B cells from normal adult blood. Single CD19+ B cells from a normal adult donor were separated by FACS, and rearranged VH genes in seventy-five cells sequenced. The percentage of B cells with recombined individual and potentially functional VH gene segments is shown. See color insert.
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on the basis of clustering patterns. Valiant attempts are still being made to make such assignments (100) but, since one replacement amino acid can have major effects on affinity, we must be cautious in our interpretation. In summary, the mapping of the germline repertoire in human B cells has provided an invaluable resource for understanding normal B-cell development and the influences exerted on it to generate the expressed repertoire. The importance to those engaged in analyzing deviations from normal behavior in autoimmune diseases or in B-cell malignancies cannot be overestimated.
MOLECULAR AND IMMUNOCHEMICAL CHARACTERISTICS OF HUMAN PATHOGENIC AUTOANTIBODIES Molecular and genetic studies of autoantibodies associated with human autoimmune disease have revealed a number of principles that provide insight into normal B-cell behavior, and into its subversion in autoimmune disease. Clearly, autoantibody-producing B cells exist in the normal repertoire and can be generated in the periphery during somatic mutation. The drive exerted on these B cells by exogenous agents or by autoantigens is an important factor in the development of autoimmune disease. This section, provides examples where immunogenetic analysis has indicated the route taken by the B cells to autoreactivity, and the consequent interaction with autoantigen associated with disease.
Cold Agglutinins: A Consequence of a Superantigenic Drive on B Cells The first example is of a directly pathogenic autoantibody against a red-cell carbohydrate antigen. Binding to the target antigen leads to clinical symptoms associated with red-cell agglutination and complement fixation, including autoimmune hemolytic anemia and acropathy, with occasional cases of gangrene (101). It is now clear that the autoantibodies, commonly known as cold agglutinins (CAs), bind the carbohydrate autoantigen via an unconventional binding site, rather than via the CDRs (102–104). Therefore, the antigen is acting as a B-cell superantigen with a broad activity reminiscent of T-cell superantigens (105). This is not the only example of a human B-cell superautoantigen, but it is highly illustrative, since it has been possible to track the production of autoantibodies from the normal repertoire to the pathogenic setting. Pathogenicity can arise from either a monoclonal or a polyclonal B-cell response, and the molecular interaction between the autoantibodies and the target antigen has now been mapped by mutagenesis and crystallography (106,107).
In many cases, CAs are monoclonal IgM proteins and are produced by a B-cell tumor, often benign in character (101). The target antigen on the red cell is composed of N-acetyl lactosamine disaccharides, which can be linear (i antigen) or branched (I antigen). As noted for other anticarbohydrate antibodies (108), binding is stronger at low temperature, and the clinical effects are largely due to agglutination of red cells in the cooler peripheral circulation. CAs are present at low levels in normal individuals, and levels can rise transiently following infection with EpsteinBarr virus, cytomegalovirus, or Mycoplasma pneumoniae (109). An investigation of VH utilization in CAs revealed the striking finding that all the monoclonal and polyclonal IgM proteins with anti-I/i specificity are encoded by the same VH gene segment, V4-34, a member of the VH4 family, (102–104). This mandatory requirement for V4-34 sequence, which could be combined with a range of VL and CDR3 sequences, strongly suggested that the I/i antigen was binding to a distinctive FR sequence outside the conventional binding sites in CDRs. Early data using mutagenesis located the site to the FR1 (103). More recent investigations, combined with the crystallographic data (105,106), have revealed a unique hydrophobic patch in FR1, involving a tryptophan (W) residue (residue 7) on b-strand A and an AVY motif (residues 23–25) on b-strand B, likely to be involved in binding to the sugar residues. Interestingly, the carbohydrate antigen tolerates many different CDR3 sequences in the autoantibodies, but certain sequences are nonpermissive (104,110). These include sequences containing basic amino acids, which convert the anti-carbohydrate activity to an anti-DNA specificity (110). The crystal structure allows an influence of CDRH3 on binding via FR1, consistent with the mutagenesis data (Figure 25.4a and 25.4b) (106,107). Since binding of the I/i antigen is to the unmutated FR1 of V4-34-encoded Ig, it can be regarded as a superantigenic interaction. There are several known B-cell superantigens, mostly derived from pathogens, with Staphylococcal Protein A (SpA) being prototypical (111). SpA has two binding sites for IgG, one of which recognizes Fcg and the other Fab (112). Binding to Fab is superantigenic and, in common with the red-cell I/i antigen, confers an ability to interact with a range of B cells via specific germline-encoded sequences in VH. SpA enhances bacterial virulence, possibly via its stimulatory activity on B cells (113). Binding of SpA involves most members of the VH3 family, again without preference for particular VL or CDRH3 sequences. The antibody recognition sequences lie in FR1, FR3, and the adjacent residues of CDR2 (114). These have been mapped in the crystal structure derived from the Fab fragment, an IgM antibody complexed with SpA (115). From these examples, and from consideration of the structure of the V region, it is clear that FR1 and FR3 of VH are able to interact with soluble
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(a)
(b) FIGURE 25.4 Ribbon diagram of a molecular model of an Ig Fv region showing VH (darker shading) and VL (paler shading). (a) The amino acids in FR1 identified as being involved in recognition of the I carbohydrate antigen by mutagenesis (H7trp, H22cys, H23ala, H24val, H25tyr) are indicated. (b) The space-filling representation of H7trp, H23ala, and H25tyr reveals a hydrophobic patch in FR1 that is a candidate for interaction with sugar residues. See color insert.
antigens, usually with low affinity (116). Similar considerations apply to VL, where the preferential usage of certain gene segments by antibodies against pathogens such as bacterial polysaccharides has been noted (reviewed in 117). Although interactions may be of low affinity, pentameric IgM, or possibly cell surface IgM, will have a significant
avidity for such superantigens, and this may be important in the natural immune response. It is also possible for antibodies to bind both superantigens and other antigens. One example is the anti-Rh D IgM MoAbs, some of which, including the well-studied MAD-2 MoAb, are encoded by V4-34 (118). These MoAbs can bind
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to Rh D+ red cells and, in the cold, to I/i antigen (119). The interaction with Rh D is specific and is not mediated by FR1, since other V4-34-encoded IgM proteins do not bind (119). This ability of autoantibodies to recognize autoantigen via an unconventional binding site, while leaving open the ability of the CDRs to bind another antigen, has been described for an IgM rheumatoid factor, with crystallographic evidence for two available sites (120). The I/i antigen is an autoantigen expressed on the surface of red cells, B cells, granulocytes, macrophages, and platelets, but also expressed by some bacterial pathogens, such as Listeria monocytogenes and Streptococcus pneumoniae Type XIV. Cold agglutinins therefore represent another example of cross-reactivity between autoantigens and the antigens of infectious agents (121). Intriguingly, the observation that B cells expressing V4-34–encoded Ig are also stimulated by herpesviruses, including EBV and CMV, suggests that there may be viral superantigens taking advantage of the hydrophobic site (122). Although it is not known if this is a virulence factor, activation of the V4-34–expressing B cells, which comprise 5 to 10% of the normal repertoire, could be useful for the virus, possibly in establishing latency. The consequent stimulation of production of the encoded IgM accounts for the rise in serum levels and the occasional evidence for cold agglutination in infected individuals (122). It has been uniquely possible to track utilization of the V4-34 gene in Ig produced in a range of clinical situations, due to the availability of a mAb specific for the FR1 sequence of the gene product (123). The mAb, 9G4, binds FR1 close to the superantigenic binding site and inhibits cold agglutination (123). Tracking has has shown that normal levels are not increased in a range of common autoimmune diseases including rheumatoid arthritis (124), but are specifically elevated in lupus. The V4-34–encoded Ig can be detected in involved renal tissue, and fluctuations in serum levels are associated with disease status (125). It is clear that some of the IgM and IgG anti-DNA autoantibodies of lupus can be encoded by V4-34 (92), and that specificity appears to lie in the basic amino acids of the CDRH3. Interestingly, these antibodies can also recognize gangliosides expressed by B cells (126) and are capable of the complement-independent killing of B lymphocytes (127). However, the elevation of V4-34-encoded Ig in lupus is greater than can be accounted for by anti-DNA activity and reaches levels similar to those post infection with EBV. The role of this Ig in the disease process remains unclear, and the possibility that it reflects reactivation of endogenous herpesviruses due to immunosuppression must be considered.
Anti-Mitochondrial Antibodies: An Example of a Route to Autoreactivity This example illustrates a route to the development of autoreactivity in a B cell that does not initially produce
autoantibody, to one that acquires autoreactivity through somatic mutation. The potential for the acquisition of autoreactivity during the process of somatic mutation is clear and is likely to be controlled at the level of availability of T-cell help. In autoimmune disease mediated by IgG antibodies, this regulatory mechanism has apparently been bypassed. Since immunogenetics and mutagenesis can be used to retrace the steps of differentiation, it is possible to investigate at which stage of B-cell development autoreactivity is gained. The illustration is again taken from a human autoantibody that appears to be directly pathogenic, in this case being characteristic of the disease primary biliary cirrhosis (PBC). This autoimmune disease involves the liver, mainly of females, and leads to cirrhosis and death from portal hypertension or liver failure, unless a liver transplant is available. The hallmark of PBC is an anti-mitochondrial antibody (anti-M2) that gives a characteristic immunofluorescent pattern in liver sections (128). The autoantibodies are largely directed against the E2 subunit of the pyruvate dehyrogenase complex (PDC-E2) and tend to focus on the inner lipoyl domain, which encompasses the site of attachment of lipoic acid at a specific lysine at the active site of E2 (129). Much effort has been directed to determining whether the antigen is expressed at the surface of biliary epithelium and to the mechanisms by which antibodies mediate disease (129). For structural studies, it has been possible to establish IgG mAbs from patients who have the characteristic specificity of the hallmark serum antibodies and to study their interaction with the E2 antigen (130,131). Among the ten IgG mAbs, there was a striking incidence of the l light chain (8/10), well above the tendency to favor Igl in heterohybridomas (130,131). There was also a tendency to produce IgG3, reflecting the known predominance in serum anti-M2 antibodies. Immunogenetic analysis of four available IgG antibodies, all of which had classical anti-M2 reactivity, and recognized lipoylated target E2 antigen, revealed a range of VH gene usage, with 3/4 derived from VH3 and 1/4 VH4 (131). Although all were Vl, a range of gene segments was also used, and this variability in V genes does not point to a superantigenic drive on the B cells of origin. All sequences were somatically mutated, consistent with passage through a germinal center and presumed conventional antigen selection leading to the high affinity for E2 antigen (131). To address the question of the developmental process leading to the autoantibodies, we expressed one mAb as Fab on the surface of phage particles. Expressed Fab bound to E2 autoantigen by ELISA, and allowed us to assess the role of replacement somatic mutations in recognition. We found that the CDR3 sequence of VH was critical for recognition, as expected, and that the CDR3 of VL was also important. However, the removal of somatic mutations from VL did not change specificity but led to only a small decrease in avidity. The surprising finding was that removal of
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somatic mutations from VH, with retention of the parental VL sequence, led to a change in specificity from the inner lipoyl domain to a different epitope in the E2 protein (132). Although there is no evidence that V regions comprising this combination of mutational patterns existed in the patient, the change in specificity is reminiscent of epitope shifting, and does suggest that similar shifts may be part of the route to autoreactivity. Immediate biological relevance was obtained by removing all the somatic mutations from VH and VL and effectively reverting the Ig to that expressed by the original B cell prior to antigen encounter. Interestingly, this germline-encoded Fab had no specificity for any part of the E2 autoantigen. The conclusion is, therefore, that the initiating antigen was different and that autoimmunity developed during the process of somatic mutation (Figure 25.5). The abnormality in patients is likely to be either unusual availability of CD4+ T cells able to help autoantibody production, or a failure to eliminate or suppress the autoreactive B cells from the germinal center.
Anti-Thyrotropin Receptor (TSHR) in Graves’ Disease Graves’ disease is a unique human autoimmune disorder characterized by diffuse goiter, with clinical and biochemical features of hyperthyroidism. In addition, 30 to 70% of cases exhibit ophthalmopathy. The disease is mediated by autoantibodies specific for TSHR that stimulate cAMP production, iodine uptake, and increased synthesis of
thyroglobulin and thyroid peroxidase, with increased growth and proliferation of thyroid cells. The pathogenicity of antiTSHR autoantibodies was clearly demonstrated in humanto-human transfer of antibodies. Thus, neonatal Graves’ disease was observed in infants born to affected mothers due to transplacental transfer that led to a stimulation of fetal thyroid cells (52,53). In addition, as in the case of Harrington’s experiment in hemolytic anemia, Adams et al. (133) demonstrated thyroid stimulation in healthy subjects infused with serum from patients with Graves’ disease. The TSHR is a G-protein link made up of two a and b subunits that use cAMP and phosphoinositol pathways for signal transduction. Anti-TSHR autoantibodies are produced by the B cells that accumulate within hypertrophied thyroid glands and recognize both linear and conformational epitopes (134). The anti-TSHR autoantibdy response is oligo- or pauci-clonal as demonstrated by restricted light chain (135) and GM allotype (136) usage. However, analysis of V gene expression in EBV-transformed B cells from Graves’s patients showed that various VH and VK genes were used (137,138), as illustrated in Table 25.3. It is to be noted that the same combination of VH and JH segments confers different antibody specificity. Sequences showed mutations in both CDR and FR segments. The pattern of somatic mutations of V genes encoding anti-TSHR autoantibodies is consistent with stimulation by antigen and affinity maturation.
Anti-Platelet Autoantibodies in Autoimmune Idiopathic Thrombocytopenic Purpura (AITP) AITP is an autoimmune disease characterized by persistent thrombocytopenia resulting in bruising, purpura, and even life-threatening bleeding. The pathogenicity of the autoantibodies is well supported by several findings: 1) The occurrence of transient thrombocytopenia in infants born to a mother with AITP (139), 2) thrombocytopenia associated with bleeding and seizure due to human to human transfer of Ig from AITP patient to a healthy subject (60), and 3) elution of antibodies from platelets (140).
TABLE 25.3 V-gene segments expressed in anti-TSHR antibodies produced by B-cell clones isolated from patients with Graves’ disease FIGURE 25.5 Model of the route to development of autoantibodies characteristic of primary biliary cirrhosis (PBC). High-affinity somatically mutated IgG autoantibodies derived from patients with PBC bind to the inner lipoyl domain of the pyruvate dehyrogenase complex. Reversion of VH and VL sequences to the germline sequences of the naïve IgMexpressing B cell of origin leads to complete loss of reactivity to the autoantigen. This suggests that another antigen, possibly a pathogen, initiated the B-cell response. Reversion of VH to germline, with retention of mutations in VL, changes specificity from the inner lipoyl domain to a different part of the protein, the E1/E3 binding domain. Shifting of epitope specificity in vitro may reflect similar events in vivo. See color insert.
Cell clone B6B7 101-2 71-2 79-4 141-1 82-1
VH
D
JH
VL
JL
V4-59 V3-7 V2-26 V3-23 V5-51 V4-59
3–9 1–7 3–10 3–16 2–15 3–9
6 4 6 4 6 4
Vk1 02/012 Vk3 A27 Vk Vk Vk Vk
Jk2 Jk4
V-region sequences reported by Akamizu et al. (139) and Shin et al. (138).
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In AITP, the majority of autoantibodies are specific for a variety of epitopes of integrin aIIb,b3 glycoprotein (GP aIIb and bIIIa). Fujisawa et al. (141) tried to define the structure of linear epitopes by generating synthetic peptides encompassing the amino acid sequence of the entire b3 chain of integrin. They found that both murine mAbs and human autoantibodies bind to the two peptides corresponding to amino acid residues 721–744 and 742–776. A study of human mAbs showed that they bind to a peptide corresponding to 222–238 amino acid residues (142) and to a neoantigen of b3 chain (143). In addition, it was found that 15 to 30% of antibodies from AITP patients bind to GPIbIX and 10 to 20% bind to GPV glycoproteins expressed on the surface of platelets. RFLP revealed a deletion of hv3005 (V3-30.1) in a high percentage of patients with chronic AITA (87). The V genes of anti-platelet autoantibodies were studied in human hybridomas or EBV-transformed B cells from AITP patients. Autoantibodies specific for integrin IIb heavy chain expressed genes from the VH3 and VK1 families (144). Another human mAb, specific for the GPIa/IIa platelet antigen, also used these families (145). One mAb specific for aII,b3 platelet antigen utilized VH4 and Vl2. A large 30-nucleotide insertion was detected in CDR2 of the VH gene encoding this antibody (146), and somatic mutations were identified. Anti-platelet autoantibodies display a major cross-reactive idiotype termed DMId (147). Sequencing of the VH genes of six human mAbs specific for GPIb antigen showed that usage mainly of the VH4 family, with one from VH1 having evidence for somatic mutation (148). Using the approach of recombination of VH and VL to generate antibody activity, Fab fragments were generated from subjects immunized with alloantigen and having antiglycoprotein IIb-IIIa antibodies. In humans, this platelet antigen exhibits allelic polymorphism, and the immunization of a negative recipient with antigen-positive platelets induces an antibody response. The V genes of the Fab fragments showed polyclonality with derivation from various families. Thus, one Fab fragment specific for the Leucin33 of GPIIa used a VH1 gene most homologous with DP7 (V1-46) and an unmutated VL2-1-11 gene. The VH1 gene showed eighteen point mutations, with eight replacement amino acids (149). Another VH gene in recombinant Fab was derived from VH4 (150). In general, the Fab fragments obtained from AITP patients showed a large repertoire of VH and VL germline genes, with 40 to 50% of both the VH and VL genes exhibiting nonrandom patterns of somatic mutations in CDR (151,152). Although these are all synthetic autoantibodies and may therefore not reflect natural antibodies, there is a suggestion that anti-GPIIb-IIIa antibody responses in AITP patients and in subjects immunized with alloantigen may be driven by platelet antigen. The events triggering such a response are as yet unknown.
HUMAN PATHOGENIC AUTOANTIBODIES WITH MURINE COUNTERPARTS Although it is often difficult to draw parallels between human autoimmune disease and mouse models, there are some examples where antibodies of a similar specificity are present in both and where insight into human disease can be provided. Examples of these are given in this section.
Anti-Acetylcholine Receptor (AchR) Autoantibodies in Myasthenia Gravis The neuromuscular transmission defect characterizing myasthenia gravis (MG) is mediated by autoantibodies specific for postsynaptic nicotinic AchR. Anti-AchR autoantibodies are produced by B lymphocytes lodged in peripheral lymph organs and in the thymus. The role of the thymus in the secretion of autoantibodies is well documented by the presence in the medulla of a large number of B-cell clusters and germinal centers (GC), indistinguishable from the GC found in the tonsils of healthy subjects. In thymic GC, the AchR is trapped in FDC, as assessed by abundant binding of 125-I a bungarotoxin (BT). Phenotypic analysis of thymus GC shows extensive infiltration of B cells (153). In MG patients who develop thymoma, in addition to antiAchR autoantibodies, antibodies against other antigens, such as the ryanodine receptor, titin, and myosin, were identified (154). Anti-AchR autoantibodies are clearly directly pathogenic and fulfill the defining criteria based on the following findings: • Injection of AchR in normal animals but not in B-cell knockout mice (155) induces the production of autoantibodies and causes experimental disease similar to human MG • Transplacental transfer of maternal antibodies can cause the disease in humans (49), and injection of anti-AchR mAbs induces MG in mice (62). In infants, the disease is transient and its development correlates with the titer of antibodies specific for fetal AchR receptor (156). The rare yet dramatic neonatal disease produced by transfer of maternal antibodies is manifested as artrogryposis multiplex congenital syndrome, characterized by an irreversible contracture of muscles and neuropathies (157) • Anti-AchR autoantibodies can be isolated from the myasthenic muscle (158). The AchR is a pentameric glycoprotein composed of a2,b,d a chains in adult, and in fetal life of a2, b, g and d chains. More than 60% of anti-AchR antibodies are directed against the extracellular moiety a subunit mapping to amino acid residues 67–76 (159). Some antibodies associated with
25. B Cells Producing Pathogenic Autoantibodies
MG without thymoma are specific for the g subunit, whereas in MG restricted to ocular muscle, antibodies are directed against the e chain (160). The majority of anti-AchR IgG autoantibodies in myasthenic patients are of high affinity and are produced by CD5- B cells (161). In terms of V-gene usage, the autoantibody response in MG is quite heterogeneous. An early study showed expression of multiple VH and VK gene families by in situ hybridization in thymic GC of MG patients (162). This was confirmed later by the sequencing the V genes of B cells producing anti-AchR autoantibodies. The VH and VL genes utilized by autoantiAchR antibodies derived from peripheral blood B cells immortalized with EBV, in hybridomas, or in synthetic Fab fragments isolated from PBL- or thymus-derived phage combinatorial libraries from MG patients were also analyzed (153,163–166). These studies clearly showed usage of VH and VK genes from various families with no preferential usage of D-segment genes or J genes (Table 25.4). Most were somatically mutated, thus suggestive of antigen selection (153). Injection of AchR into genetically susceptible mice induces autoantibodies, muscle weakness, and alteration of the myograms typical for human disease (63). Hybridomas established from affected mice showed reactivity with at least three epitopes of AchR. One is a major immunodominant epitope overlapping with the Ach binding site. A second epitope is near the Ach and a-BT binding site D, since some mAbs were able to inhibit the binding of a-BT to AchR. The third category of antibodies recognizes overlapping epitopes on the g and d subunits (167). Analysis of the VH gene usage in this set of hybridomas demonstrated that at least six different families were used, with a significant deviation from stochastic usage, resulting in an overexpression of VH J606 and Vgam3.8 families (168). It is noteworthy that this biased
TABLE 25.4 V-gene segments expressed in human anti-AchR autoantibodies Source PBL
System EBV transformed
VH
D
JH
V5-a V5-a V2-05
6-6 6-6
6 6 5
Vk3 A27 Vk3 A27 Vk1 L
Jk2 Jk2 Jk1
—
3
Vl1b
Jl2
PBL
Hybridoma
V3-7
PBL
Fab combinatorial library
V3-23 V3-23 V3-23 V3-11
Thymus
Fab combinatorial library
V3-53 V4-30.1 V1-46 V3-11
VL
JL
Vk3 A27 Vk3 A27 Vk1 02/O12 6a 4 3 4 5
Vk3 A27 Vk1 02/O12 Vk3 A27 Vk3 L25
Jk3 Jk2 Jk3 Jk4
This information summarizes the data reported in references 163, 164, 165, and 166.
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usage was observed in hybridomas obtained from C57BL/6 mice and not from BALB/c mice, which are not susceptible to the induction of experimental disease. The usage of VH and VK genes in a panel of 19 pathogenic mAbs revealed highly homologous VH-D-JH and VL-JL (>90%) sequences (168), suggesting a pauci-clonal response. Three antibodies had VH sequences close to VH7183 germline genes. The sequence comparison of VH CDRs showed important homology among antibodies specific for the major immunodominant epitope of AchR. In summary, anti-AchR autoantibodies show a diversity of VH and VL genes, with some biased usage of two VH gene families and some highly homologous VH genes related to VH7183 in a few murine pathogenic autoantibodies.
Pathogenic Autoantibodies in Lupus Both human and animal models for SLE are associated with autoantibodies specific for a multitude of antigens (ANA, ssDNA, dsDNA, Z-DNA, Ro, nRNP, Sm, Ku, La, P1/P2, laminin, HLA class I promoter-binding proteins, nucleolin, histones, CD45 isoforms). Autoantibodies specific for these autoantigens are also associated with other autoimmune diseases such as Sjogren’s syndrome, MCTD, RA, and scleroderma, and may also be found at low levels in healthy subjects. Among these, anti-Ro autoantibodies fulfill the criteria of pathogenic autoantibodies since they are responsible for neonatal lupus with complete heart block (168). Neonatal lupus is a rare disease characterized by transient annular erythematous plaques that develop 2 to 8 weeks after birth and disappear by 6 month of age as maternal antibodies decrease in the newborn circulation (170), and by permanent atrial-ventricular block (170,171). Skin biopsies from infants with neonatal lupus demonstrate slight edema of papilliary dermis, with the dilated blood vessels and perivascular lymphocytic infiltration associated with deposition of Ig and C3 at the basement membrane zone of epidermis (157). In children with neonatal lupus with heart block, mortality was 16%; 73% of these died during first 12 months (58). Ro protein is present in two isoforms of 52- and 60-kDa. The 52-kDa protein bears two epitopes, one immunodominant corresponding to amino acid residues 169–291 containing the leucine zipper, and another more restricted subset corresponding to residues 1–78 containing zincfinger domain (173). In a study of 59 sera from mothers whose children had neonatal lupus, 95% displayed high titers of antibodies specific for 52-kDa Ro protein (173). In 10 Japanese infants with neonatal lupus with skin lesion and heart block, maternal antibodies reacted with both 52- and 60-kDa Ro proteins (58). Although there is no information on the V-gene structure of maternal anti-Ro antibodies causing neonatal lupus, a study of synthetic Fab fragments from a combinatorial library from a patient with Sjogren’s
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syndrome showed usage of VH3 and VH5 genes paired with VK3 and Vl1 (174). Usage of VH3 (V3-21 and V3-23), combined with Vl2 and Vk1 respectively, was also seen in two unmutated IgM autoantibodies, specific for Ro 52-kd antigen, isolated from a patient with primary Sjogren’s disease (175). Although combinatorial libraries and IgM antibodies may not reflect disease-associated antibodies, these data suggest that there is no preferential usage of V genes by anti-Ro antibodies. In contrast to anti-Ro antibodies, anti-dsDNA and antiSm autoantibodies do not fulfill all the criteria of pathogenic autoantibodies. Anti-ds DNA autoantibodies cannot be induced by immunization with antigen, nor can neonatal lupus be induced by transplacental transfer. However, in lupus patients, anti-DNA antibodies were eluted from kidney and skin lesions and, in some cases, infusion of cationic human anti-DNA mAbs into mice can induce lesions. The anti-dsDNA antibodies are characteristic of lupus, but IgM anti-dsDNA can be found in healthy subjects (176). In these cases they bind also to ssDNA and exhibit low affinity for DNA and other self-antigens. In contrast, anti-dsDNA antibodies in lupus patients exhibit high affinity for DNA and have switched to IgG (177). Early studies of mAbs obtained from BW mice that spontaneously develop lupus similar to human disease showed that a subset of anti-dsDNA pathogenic antibodies are cationic. These antibodies promote nephritogenicity, fix complement, and exhibit high affinity for anionic DNA (177). They also bind to antigens in glomerular basement membrane, in particular heparan sulfate, and may share cross-reactive idiotypes. Cationic anti-DNA antibodies have been eluted from the kidney of lupus mice, thus indicating that they may be causally related to the occurrence of nephritis (178). Cationic anti-dsDNA antibodies are also present in the sera of lupus patients. Although antibodies with pI = 8.5 bind to both DNA and heparan sulfate, neutral antibodies with a pI = 7 bind exclusively to DNA (179). This again suggests that the binding to heparan sulfate may contribute to the development of nephritis by triggering local inflammatory reactions. Liveneh et al. (180) showed that a subset of human cationic anti-DNA antibodies that express Vl share a cross-reactive idiotype, 8.12, indicating an oligoclonal origin of this subset of human anti-DNA antibodies. The utilization of V genes in human pathogenic anti-ds DNA antibodies was studied in EBV-transformed B cells, Bcell hybridomas, and IgG Fab libraries obtained from lupus patients. Analysis of V genes showed that they used some VH genes frequently used in the normal repertoire, such as V1-69, V3-23, V3-07, V4-34, and V4-39, and some less commonly used, such as V1-46, V3-64, V3-74, and V4-61. The CDR3 sequences showed some particular features. First, the CDR3 sequences appeared to involve D-D fusion and uncommon reading frame processes. Second, CDR3 sequences can be rich in positively charged amino acids,
such as arginine and lysine (181–184). The introduction of arginine may be associated with the acquisition of DNA binding in pathogenic anti-dsDNA antibodies (185,186). Two human V4-34–encoded IgG anti-DNA antibodies had a predominance of basic amino acids in CDR3 (110,183). Mutagenesis to remove these from one of the moAbs abrogated anti-DNA activity (187). In mice, the position of the arginine residue in CDR3 was found to be critical for binding to dsDNA (188). Combinatorial IgG Fab phage libraries from a patient with lupus also revealed prominence of basic amino acids in CDR3 with a concentration in the N-terminal region (189). The importance of this sequence for the recognition of DNA was confirmed by transplanting sequence and specificity into an unrelated antibody. However, many anti-DNA antibodies do not have a basic CDR3, and there are clearly multiple routes to this specificity (190). Both human and murine anti-dsDNA antibodies express a broad range of VL genes, with VK1-3 families, also common in normal B cells, being slightly more frequent. However, the hallmark of V-gene features consist of a high rate of replacement somatic mutation in CDRs and the introduction of arginine substitutions that may facilitate DNA binding. B cells exhibiting such mutations may results from positive selection or the lack of negative selection of precursors of anti-dsDNA antibody-forming cells.
Pathogenic Anti-Phospholipid Antibodies Anti-phospholipid syndrome (APS) is an autoimmune disease characterized by thrombosis, thrombocytopenia, and recurrent fetal loss (191) caused by pathogenic autoantibodies. Initially, APS was associated with lupus and autoimmune thrombocytopenia, but the latter can occur in the absence of other autoimmune diseases. In patients, APS is manifested by arterial, cardiac, dermatologic, and placental pathology characterized by miscarriage and fetal loss. In animals, anti-phospholipid antibodies (aPL) were found in strains prone to lupus (MRL, NZWxBSB). In addition, in MRL/lpr mice, the presence of anti-cardiolipin antibodies was associated with an increased rate of fetal resorption (192). The pathogenicity of APL antibodies was demonstrated in human–mouse transfer experiments. Passive transfer of polyclonal or monoclonal anti-cardiolipin antibodies into mice induces fetal resorption (193). Although passive transfer of anti-cardiolipin antibodies causes abortion, it does not cause the venous thrombosis observed in human disease. However, an increased size and duration of thrombus formation in the veins of mice was noted after the injection of IgG fraction from human sera (194) or monoclonal antibody (195) from APS patients. Autoantibodies associated with APS are directed against various plasma proteins, such as b-2 glycoprotein1, prothrombin, annexinV, and protein C (196). It is noteworthy that, in APS patients, the
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25. B Cells Producing Pathogenic Autoantibodies
anti-PL antibodies tend to bind cardiolipin in association with b2-glycoprotein. MAbs binding to cardiolipin (CL), b-2-glycoprotein1, and phospholipids have been generated from PBL or tonsil lymphocytes from healthy subjects and from PBL or splenic lymphocytes of patients with primary or secondary APS or lupus. From the analysis of the structure of the VH and VL genes encoding anti-phospholipid antibodies (199–202), several conclusions emerge: • There is no preferential usage of V-gene families in APL autoantibodies, with almost all VH gene families being represented in combination with various D and JH segments (Table 25.5). With regard to VL, VK or Vl genes can be involved in combination with JK or Jl segments respectively, in autoantibodies obtained from healthy subjects or patients with primary or secondary APS or lupus. • There is no biased usage of V genes expressed in autoantibodies displaying various specificities against cardiolipin, cardiolipin + ss or dsDNA, b2 GP1 glycoprotein, or phosphlipids. • The majority of anti-PL antibodies exhibit mutations, in particular replacement mutations that are clustered in CDRs. • The pathogenic anti-PL autoantibodies exhibit an accumulation of arginine in the VH CDRs compared to nonpathogenic anti-PL autoantibodies. As for anti-DNA antibodies, the accumulation of this basic amino acid may be responsible for the specific binding to phospholipids (201,202)
CONCLUSION Knowledge of the available repertoire of unrearranged variable region genes, and of the selection of these for recombination by normal B cells, allows insight into the perturbation of V-gene usage in disease. Sequence analysis also reveals whether B cells have undergone somatic mutation, which is presumed to occur in the germinal center following antigen encounter. Autoreactive antibodies can clearly arise at both stages, and in many cases may be of no pathological consequence. However, some IgM autoantibodies with low affinity but high avidity can be dangerous, and innocuous IgM antibodies can become pathogenic following somatic mutation and apparently aberrant antigen selection. Either inappropriate T-cell help, or a failure to delete or modify B cells in the periphery, allows the progression through maturation and isotype switch to high-affinity autoantibody production. We have focused on autoimmune diseases in which autoantibodies are clearly pathogenic. We have drawn together data from murine models and human diseases to illustrate the molecular features of autoantibodies, and to point to possible routes to disease. No unify-
TABLE 25.5 V-gene segments expressed in human monoclonal Abs against phospholipid antigens from healthy subjects and from patients with primary or secondary APS Origin
Specificity
VH
D
JH
VL
JL
V4-39 V4-39 V4-39 V1 V3-33
D2-8 D2-8 D3-10 D3-10
4 4 1 4 6
VK1 VK1 VK2 Vl2 Vl3
JK4 JK4 JK2 Jl2/3 Jl1
Secondary APS CL1 human b2GP1 + CL CL15 human b2GP1 + CL CL25 humanb2GP1 + CL
V1-2 V4-34 V3-23
D1-14 D7-27 D3-16
5 6 4
Vl3 Jl2/3 VK1 JK2 VK3 JK2
Lupus 18-2 1-17 C119 C471 RSP4 R149 AH2 DA3 UK4
V3-23 V3-23 V3-23 V3-64 V3-30 V1-69 V5-51 V5-51 V3-23
5 5 3 4 4 6 6 6 4
VK3 VK3 Vl1 VK2 Vl1 Vl1 Vl2
Healthy subjects Fetal liver A431 CL+ssDNA
V6-1
3
Child’s tonsil Kim13.1 CL KIM4-6 CL + dsDNA
V1-69 V3-30
5 6
VK3 JK4 Vl1 Jl3
Adult PBL B122 PL + ssDNA B6204 PL + dsDNA A10 CL + dsDNA A431 CL + dsDNA H3 CL A5 CL Bou53 CL H5 CL
V1-18 V3-23 V6-1 V6-1 V-1 V3 V3 V4-39
4 4 3 4 4 6 6 5
VK1 JK2 VK3 JK1
Primary APS IS1* bovine IS2 bovine IS3 human IS4 human BH1 PL***
b2GP1 b2GP1 b2GP1 + CL** b2GP1 + CL
CL + ssDNA CL + ssDNA PL + ssDNA PL + ssDNA CL CL PL PL PL
D6-13 FL16 D2-2 D1RL
JK4 JK2 Jl1 JK2 Jl2 Jl2 Jl2
Vl4 Jl2/3 Vl3 Jl2 VK1 JK2
* name of human MoAb. ** cardiolipin. *** phospholipid. The data presented in this table summarize those reported by Rahman et al. (190),Logtenberg et al. (197), Siminowitch et al. (200), Harner et al. (198), Hohman et al. (201), and Chukwuchocha et al. (199,202).
ing hypothesis has emerged, and it is likely that the control of autoreactive B cells can be subverted at many points. However, understanding the nature of the consequent pathogenic autoantibodies may provide opportunities for preventing their effects.
Acknowledgments The authors thank Dr. Brian Sutton for permission to reproduce his ribbon diagram (p. 389). Dr. Stevenson thanks the Leukemia Research Fund, the Arthritis Research Council, and Tenovus UK for grant support.
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26 Immunodeficiencies Caused by B-Cell Defects FRANCISCO A. BONILLA AND RAIF S. GEHA Division of Immunology, Children’s Hospital, and Department of Pediatrics, Harvard Medical School, Boston, Massachusetts, USA
clinical and laboratory phenotypes associated with these various forms of agammaglobulinemia are essentially identical. The majority of patients described clinically to date have XLA, which accounts for about 85% of all cases of agammaglobulinemia (Conley et al., 2000). This section focuses on the clinical features of this group of patients. The incidence of XLA is not known precisely, but is estimated to be approximately 0.5/100,000 live births (Sideras and Smith, 1995). Symptoms have their onset between 6 and 12 months of age in most patients (Lederman and Winkelstein, 1985). Immunoglobulin transferred across the placenta from the maternal circulation during gestation affords considerable protection during the early months of life. Diagnosis is frequently delayed until between 1 and 3 years of age. At that time, the IgG is almost always <1 g/L; IgA and IgM are <0.1 g/L. B cells are much less than 1% of circulating lymphocytes in the majority of patients. The major types of infectious complications of XLA encountered in one review of 96 patients are listed in Table 26.2 (Lederman and Winkelstein, 1985). Respiratory tract bacterial infections account for the majority, with frequent occurrences of gastrointestinal, skin, and bone and joint infections, as well. Agammaglobulinemic patients are susceptible to intestinal giardiasis (LoGalbo et al., 1982). Aseptic joint arthritis may be due to infection by mycoplasma and ureaplasma organisms (Franz et al., 1997; Furr et al., 1994). Enteropathic viruses such as ECHO (enterocytopathic human orphan) viruses, coxsackie, and poliovirus may cause an acute or chronic encephalomyelitis syndrome in agammaglobulinemic individuals (Katamura et al., 2002; Misbah et al., 1992). This complication occurred more commonly prior to the era of intravenous gamma globulin therapy, but is still encountered occasionally, even with such therapy.
Defects of human specific immunity are usually classified with respect to their effects on humoral immunity (antibody or B-cell deficiencies), cellular immunity (T-cell deficiencies), or both (combined deficiencies) (1997). In this chapter, we review the clinical and molecular aspects of human immunodeficiency disorders arising from known mutations of genes important principally for B-cell function (Table 26.1). These comprise two major phenotypes. In the agammaglobulinemias, B-cell development is generally completely blocked in the bone marrow and serum immunoglobulins are absent (Conley et al., 2000). In hyper IgM syndromes, B cells develop normally in the bone marrow and are found in normal numbers in the circulation, but they are not able to cooperate properly with T cells to generate complete antibody responses, or are intrinsically incapable of undergoing immunoglobulin class-switch recombination (Bonilla and Geha, 2001; Revy et al., 2000; Imai et al., 2003).
CLINICAL FEATURES OF THE AGAMMAGLOBULINEMIAS The classical clinical description of agammaglobulinemia was offered by Bruton in 1952, when he reported the case of a male child lacking serum gamma globulin with severe recurrent bacterial infections of the respiratory tract (Bruton, 1952). This disease subsequently became known as Bruton’s, or X-linked agammaglobulinemia (XLA). The molecular defect in this disorder affects a gene encoding a tyrosine kinase, now called Bruton’s tyrosine kinase, or BTK. The ensuing decades have brought the discoveries of additional gene defects associated with forms of agammaglobulinemia having autosomal recessive inheritance. The
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TABLE 26.1 Immunodeficiencies resulting from B cell gene defects Disease Agammaglobulinemia X-linked or Bruton’s Autosomal recessive
Hyper IgM syndrome Autosomal recessive
Protein
Gene
Disorders
Reference(s)
Map location
BTK m heavy chain l5 Iga BLNK
BTK IGHM IGLL1 CD79A BLNK
Xq21.33–22 14q32.33 22q11.23 19q13.2 10q23.2–23.33
CD40 AID UNG
TNFRSF5 AICDA UNG
20q12–13.2 12p13 12q23–24.1
TABLE 26.2 Major infectious complications of XLA Complication
TABLE 26.3 Rare infectious complications of XLA
H. influenzae conjunctivitis
(Hansel et al., 1990)
Chlamydia conjunctivitis with corneal scarring
(al Ghonaium et al., 1996)
Adenovirus pneumonia
(Siegal et al., 1981)
M. hominis pneumonia
(Roberts et al., 1989)
P. carinii pneumonia
(Alibrahim et al., 1998)
Cryptococcal thoracic empyema
(Wahab et al., 1995)
Recurrent purulent triaditis
(Wray et al., 1981)
Recurrent pneumococcal arthritis
(Peters et al., 2000)
Chronic coxsackie virus infection
(O’Neil et al., 1988)
U. urealyticum endocarditis and osteomyelitis
(Frangogiannis and Cate, 1998)
P. aeruginosa sepsis
(Urbach et al., 1983, Zenone and Souillet, 1996)
% affected* Ecthyma gangrenosum
(Kim et al., 2000)
Flexispira/Helicobacter sepsis
(Cuccherini et al., 2000, Gerrard et al., 2001)
Respiratory tract Otitis media Sinusitis Mastoiditis Bronchitis Pneumonia
59 14 4 9 56
Gastrointestinal tract Diarrhea Perirectal abscess
32 3
Disease
Skin Cellulitis
28
Hematologic disorders Neutropenia
Nervous system Meningitis Encephalitis
10 6
TABLE 26.4 Noninfectious complications of XLA
Musculoskeletal Arthritis, Septic Aseptic Osteomyelitis
8 11 3
Generalized sepsis
10
* Data from (Lederman and Winkelstein, 1985).
Patients with agammaglobulinemia are also prone to paralytic poliomyelitis caused by vaccine strains (Wright et al., 1977). However, at least one reported patient with classic XLA survived wildtype poliovirus infection (Sarpong et al., 2002). Additional rare infectious complications of XLA are listed in Table 26.3. The most common causes of death are the sequelae of acute infection or respiratory failure due to chronic infection (Lederman and Winkelstein, 1985). A variety of noninfectious complications of XLA may also occur (Table 26.4). Prominent among these is neutropenia, which may occur in 10 to 25% of patients (Farrar et al., 1996; Lederman and Winkelstein, 1985). This appears to be predominantly related to acute or chronic infection and resolves with antimicrobial and immunoglobulin replacement therapy. A few autoimmune disorders, such as Crohn’s
Hypersensitivity Drug reaction
Reference(s)
(Farrar et al., 1996, Kozlowski and Evans, 1991) (Kudva-Patel et al., 2002)
Autoimmune disease Type 1 diabetes mellitus (Martin et al., 2001) Regional enteritis/Crohn’s disease (Cellier et al., 2000) Malignancy Gastric adenocarcinoma Colorectal cancer Squamous cell lung carcinoma B cell lymphoma T cell lymphoma Other disorders Amyloidosis Monoclonal gammopathy
(Bachmeyer et al., 2000, Lavilla et al., 1993) (Lederman and Winkelstein, 1985) (Echave-Sustaeta et al., 2001) (Lederman and Winkelstein, 1985) (Kanavaros et al., 2001) (Meysman et al., 1993, Tezcan et al., 1998) (Gemke et al., 1989, Hendrickx et al., 1979)
disease (Cellier et al., 2000), and type 1 diabetes (Martin et al., 2001) have been seen in some patients with XLA. Presumably, these are mediated exclusively by cellular immune responses against target tissues. Drug hypersensitivity has also been reported, also due to a cellular response (KudvaPatel et al., 2002). Finally, a variety of malignancies have
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26. Immunodeficiencies Caused by B-Cell Defects
been reported in patients with XLA (Table 26.4). However, it is not apparent that there is an increased rate of malignancy in these patients.
Atypical XLA As described above, the majority of patients with XLA present in childhood with bacterial respiratory tract infections and other complications. Many may die without aggressive therapeutic and preventive antibiotic regimens and replacement therapy with gamma globulin. A small, but still unquantified fraction of individuals having BTK mutations have less severe clinical and immunological phenotypes. Several patients diagnosed with common variable immunodeficiency in a Japanese registry were found to have BTK mutations (Kanegane et al., 2000). Some patients have been misdiagnosed with even milder forms of antibody deficiency such as transient hypogammaglobulinemia of infancy (Kornfeld et al., 1995), or selective polysaccharide antibody deficiency (Wood et al., 2001). Several of these atypical patients have survived without specific therapy, only to be diagnosed in adulthood (Hashimoto et al., 1999; Kornfeld et al., 1997; Usui et al., 2001). Even within the same kindred, including siblings, males having the same BTK mutation may have discordant phenotypes (Bykowsky et al., 1996; Gaspar et al., 2000; Kornfeld et al., 1997).
al., 1999). As of September 2002, 461 distinct mutations had been catalogued from 758 patients in 656 separate kindreds. Table 26.5 enumerates the types of mutation identified according to the regions of the BTK gene affected. Figure 26.1 also shows the distribution of mutations across the coding regions and introns of the BTK gene. Missense mutations account for 1/3 of all genetic lesions, while deletions constitute 1/4. Nonsense, insertion, and splice-site mutations each comprise roughly 1/10 of the total. Mutations are distributed fairly evenly through the different functional domains of the molecule (Figures 26.1 and 26.2). Most are found in the kinase domain (the largest), which has 44% of the total. The SH3 domain has relatively fewer in comparison to other regions of the molecule. Figure 26.2 shows the distribution of mutations in BTK in kindreds with respect to the amino acid sequence of BTK. Arginine residues are most often affected by missense and nonsense mutations. Those at positions 13 and 28 (PH domain), 255 (SH3), 288 (SH2), and 520, 525, 562, and 641 (kinase) are frequently affected in XLA families. Additional frequent sites of mutation include M1 in the PH domain, and C506 and G594 in the kinase domain. A consistent genotype–phenotype correlation among patients with XLA has not been found with respect to either
BTK Mutations in XLA BTK mutations in patients with XLA were first described in 1993 (Tsukada et al., 1993; Vetrie et al., 1993). A database of BTK mutations in XLA patients was established in 1995. It can be accessed via the World Wide Web at http://www.uta.fi/laitokset/imt/bioinfo/BTKbase/ (Vihinen et
FIGURE 26.1 Distribution of BTK mutations. The exon boundaries are indicated by vertical lines and are numbered. The numbers of families having exon or intron mutations in each region are indicated. This figure was redrawn based on a figure at the web page url: http://protein.uta.fi/BTKbase/tables.html (reproduced with permission).
TABLE 26.5 Distribution of BTK mutations in XLA according to type and location* Mutation types
Missense Nonsense Insertion, inframe Insertion, frameshift Insertion Deletion, inframe Deletion, frameshift Deletion, gross Splice-site, inframe Splice-site, frameshift Splice-site Other Total Percent
Upstream
Pleckstrin homology
Tec homology
Src homology 3
Src homology 2
Tyrosine kinase
Downstream
Other
Total
Percent
0 0 0 0 0 0 0 0 0 0 2 3 5 1.1
28 11 1 16 0 4 27 0 2 5 10 0 104 22.6
4 5 0 13 0 2 16 0 0 0 3 0 43 9.3
0 9 0 3 0 0 11 0 1 1 2 0 27 5.9
25 9 0 2 0 1 9 0 1 3 8 0 58 12.6
93 25 2 16 1 5 36 0 1 7 17 0 203 44.0
0 0 0 0 0 0 0 0 0 0 0 0 0 0.0
5 0 0 0 2 1 0 13 0 0 0 0 21 4.6
155 59 3 50 3 13 99 13 5 16 42 3 461 100.0
33.6 12.8 0.7 10.8 0.7 2.8 21.5 2.8 1.1 3.5 9.1 0.7 100.0
* Data from BTKbase: http://www.uta.fi/laitokset/imt/bioinfo/BTKbase/.
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FIGURE 26.2 Distribution of mutations in families having XLA. Each bar represents the relative number of families having each type of mutation (missense, nonsense, insertion, deletion) according to the amino acid/region of the BTK molecule. This figure was redrawn based on a figure at the web page url: http://protein.uta.fi/BTKbase/familybars.html (reproduced with permission).
B cell number or clinical course (Holinski-Feder et al., 1998; Tao et al., 2000). Identical genetic lesions have been found in patients with or without circulating B cells, or with severe or mild clinical courses. Although R28 mutation leads to a relatively mild phenotype in the xid mouse strain, it can be associated with a classical severe XLA phenotype (de Weers et al., 1994). Conversely, even mutations that lead to great reduction in the amount of BTK protein may be associated with a relatively mild clinical course. However, splice-site mutations that permit the production of small amounts of functional BTK protein or conservative amino acid substitutions do tend to lead to milder phenotypes (Conley et al., 1998; Saffran et al., 1994).
Three patients with XLA and sensorineural deafness were found to have deletions encompassing BTK and 4 to 19 kilobases of 3¢ downstream DNA (Richter et al., 2001). These deletions eliminate the deafness-dystonia protein gene associated with the Mohr-Tranebjaerg syndrome. The biochemical consequences of BTK mutations have been studied. Several distinct pleckstrin homology domain mutations have been shown to have reduced binding to inositol 1, 3, 4, 5 tetraphosphate, indicating the importance of membrane localization of BTK for execution of its signaling function (Fukuda et al., 1996). Mutations within the BTK SH2 domain lead to significant reductions in phosphotyrosine binding as a result of altered conformation,
26. Immunodeficiencies Caused by B-Cell Defects
selective disruption of key physicochemical interactions, or both (Mattsson et al., 2000). Several kinase domain mutations have been shown to impair activity in vitro (Maniar et al., 1995). In some cases, the mutant protein was expressed at normal levels in patients’ cells. A modeling study predicted that eight distinct BTK mutations described in XLA clustered together on one face of the kinase domain, thus suggesting that they might similarly disrupt kinase activity (Vihinen et al., 1994).
XLA with Growth Hormone Deficiency In 1980, Fleisher and colleagues described kindred patients with clinical features of XLA in association with growth hormone deficiency (Fleisher et al., 1980). These individuals had recurrent sinopulmonary bacterial infections, reduced or absent circulating B cells, low immunoglobulin levels and impaired antibody responses, short stature and delayed bone age, and failed to produce growth hormone when appropriately challenged. Additional individuals with this association have been described, but it is not yet clear that it represents a single distinct disease entity. Linkage analysis in one kindred mapped the genetic lesion to the BTK locus (Conley et al., 1991). In another study, in one patient, an intronic mutation within the BTK gene (1882+5 GÆA) led to exon skipping with premature termination and loss of the 61 C-terminal amino acids (Duriez et al., 1994). In another patient, an insertion in BTK codon 157 in exon 6 (CAGÆCAAG) led to premature termination in codon 193 (exon 7) (Abo et al., 1998). However, in another kindred, the BTK cDNA sequence contained only one silent base substitution (1899 CÆT), BTK mRNA had normal size, and BTK protein was expressed at normal levels in western blots (Stewart, et al., 1995). One genetic analysis of polymorphic markers flanking the BTK gene ruled out a contiguous deletion in three patients with XLA and GHD (Vorechovsky et al., 1994). Thus far, no mutations other than in BTK have been described in these patients. Also, the basis of growth hormone insufficiency in those patients with established BTK mutations has not been elucidated.
B-Cell Development and Function in Patients with XLA BTK is expressed in all B-cell stages except plasma cells, myeloid cells, and platelets (de Weers et al., 1993; Oda et al., 2000). BTK has critical roles in signaling through pre-B and mature B cell Ig receptors (Aoki et al., 1994; de Weers et al., 1994). BTK also participates in signaling pathways triggered by the ligation of the platelet collagen (glycoprotein VI) receptor (Oda et al., 2000), the high affinity IgE receptor (Kawakami et al., 1994),
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FcgRII (CD32) (Oda et al., 2000), CD19 (Kitanaka et al., 1998), and receptors for interleukins-5 (Sato et al., 1994) and -6 (Matsuda et al., 1995). Despite the involvement of BTK in a variety of signaling pathways that do not depend on immunoglobulin, there are no apparent clinical consequences of myeloid cell or platelet dysfunction in XLA. Some early studies of B-cell development in XLA patients showed a block following D-JH heavy chain Ig gene rearrangement in EBV-transformed bone marrow B-cell lines (Ichihara et al., 1988; Schwaber 1983; Schwaber, 1992; Schwaber and Chen, 1988). However, other studies of rearranged Ig heavy chain genes in similar bone marrow–derived B-cell lines demonstrated complete VHDJH rearrangement with diverse VH gene usage, with a repertoire of VH genes, D regions, and JH genes characteristic of fetal liver B cells (Milili et al., 1993). Somatic mutations were observed, but very seldom. Studies of EBV-transformed peripheral blood B-cell lines from XLA patients also revealed completely rearranged Ig heavy chain genes (Mensink et al., 1986). Mortari et al. observed the usage of a single VH gene (4.18) in peripheral blood EBV transformed clones derived from one patient (1991). They found N-region nucleotide addition only between VH and D, and proposed that DJH recombination had occurred through “illegitimate” recombination in regions of signal sequence homology—characteristics of B cell immaturity (pre-B cell stage) (Ichihara et al., 1989). In another study, IgM-, IgD-, IgG-, and IgA-producing lines were obtained (Anker et al., 1989). Both k and l light chain genes were used. Southern blot analysis of genomic rearranged Ig heavy chain genes showed clonal unrelatedness among cell lines from individual patients. Further studies of similarly derived peripheral B-cell lines showed usage of all VH gene families, although in one report, no somatic mutation was seen (Timmers et al., 1991). The same group later reported that IgM-associated kappa light chains were derived from all four Vk families, and that some lines displayed somatic mutations (Timmers et al., 1993). IgM-surrogate light chain complexes may be found on bone marrow B-cell precursors in XLA patients, albeit with reduced frequency in comparison to controls (Genevier and Callard, 1997). A more recent study of cytoplasmic m chain and VpreB expression in XLA bone marrow cells showed relatively increased pro-B cells (m-/VpreBhi) and early preB cells (mlo/VpreBhi), with a reduction in the populations expressing cytoplasmic m chains in the absence of surrogate light chain (Nomura et al., 2000). Similar data have been reported by others, who documented normal numbers of nuclear TdT+/cytoplasmic CD22+/surface CD10+/CD19+ bone marrow cells in XLA patients with absent peripheral B cells (Campana et al., 1990). B-cell development halted with the expression of cytoplasmic m chains, which were only observed in the precursor cells of half of the patients
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studied. A similar block in the pro-B to pre-B cell transition was also reported by another group (Noordzij et al., 2002). This study also documented near normal bone marrow precursor B cell populations in a patient with low levels of normal BTK due to a splice site mutation. However, peripheral B cell numbers remained low, indicating the requirement for BTK at several points during B-cell development. One flow cytometric analysis of the small number of circulating B cells that develop in some XLA patients has shown that they express higher amounts of surface IgM and CD38, and lower amounts of surface HLA-DR and CD21, and cytoplasmic bcl-2, in comparison to controls (Conley, 1985). All express CD20, about 80% express surface IgD. Both k and l light chains are expressed, as may be expected from the genetic analyses described above. Another report documented a phenotype of CD5-/CD19+/CD20+/CD21- of XLA peripheral blood B cells (Nonoyama et al., 1998). Calcium flux is impaired upon cross-linking surface IgM on bone marrow B cells and B-cell lines from XLA patients (Genevier and Callard, 1997). EBV-transformed peripheral blood B-cell lines from XLA patients respond with Ig secretion to IL-6, whereas response to IL-4 may be impaired (Lau et al., 1989). In this study, however, response of control cell lines to IL-4 was small. Another analysis of peripheral blood B cells from XLA patients stimulated with anti-CD40 and IL-4 showed a normal induction of surface CD23 expression, and the production of IgE (Nonoyama et al., 1998). These authors further reported the induction of low amounts of specific antibody in vivo following immunization with bacteriophage X174. Other authors have also reported low primary and secondary responses to bacteriophage in XLA patients (Leickley and Buckley, 1986).
T-Cell Function in Patients with XLA Peripheral blood T cells from patients with XLA respond normally to nonspecific plant mitogens, various monoclonal antibodies, and recall antigens (Plebani et al., 1997). For this reason, some argue that XLA patients should be vaccinated with killed or inactive component vaccines. One study has shown normal HBSAg-specific T cell numbers, as well as normal HBV-induced interferon-g and interleukin-4 production in vitro by T cells from XLA patients immunized with HBV vaccine (Paroli et al., 2002). One group observed that TH1 responses dominate in XLA patients, based on preferential interferon-g production in response to phytohemagglutinin or tetanus toxoid (Amedei et al., 2001). In this study, T cells from XLA patients expressed higher amounts of LAG-3 (associated with interferon-g producing TH1 cells) and lower amounts of CD30 (associated with TH2 cells) in comparison to controls. In addition, higher plasma levels of interferoninducible protein 10, and lower levels of macrophage-
derived chemokine were interpreted as further indicating TH1 over TH2 predominance. Another study did not find any differences in the TH1/TH2 profile in XLA patients compared to controls (Paroli et al., 2002).
Diagnosis of and Carrier Detection for XLA The assessment of BTK protein level and activity have been used as confirmatory tests for XLA (Gaspar et al., 1998). BTK expression in monocytes (Kanegane et al., 2001) or platelets (Futatani et al., 2001) may be measured by flow cytometric methods, and this has been used as a diagnostic test for XLA. In a study of 106 Japanese agammaglobulinemic patients with low circulating B cells, flow cytometric analysis of monocyte BTK content identified 93 patients with XLA on the basis of low or undetectable BTK (Kanegane et al., 2001). Only two patients with near normal levels of BTK were found to have missense mutations, yielding a sensitivity for this method of 98%. In males with characteristic clinical and laboratory features, a positive family history consistent with X-linked inheritance may establish the diagnosis of XLA. Molecular diagnosis is desirable in order to continue to define the specific phenotypic and biochemical features that may be associated with particular BTK mutations. Only approximately 50% of XLA patients will have a positive family history (Conley et al., 1998; Lederman and Winkelstein, 1985), thus, additional measures (such as the flow cytometric methods noted above) must be undertaken to establish the correct diagnosis in these patients. Flow cytometric determination of cytoplasmic BTK in peripheral blood monocytes or platelets may also be used to assess XLA carrier status in females, who show two populations, BTKdim and BTKbright (Futatani et al., 1998; Futatani et al., 2001; Kanegane et al., 2001). In one Japanese study, 81% of patients’ mothers were identified as carriers based on flow cytometric analysis of monocyte BTK content (Kanegane et al., 2001). Four of 13 mothers showing normal staining carried mutant BTK alleles. Bone marrow CD34+/CD19+ pro-B cells in female XLA carriers show random X chromosome inactivation (Conley et al., 1994). Since B cells cannot develop from hemopoietic precursors that inactivate the functional BTK gene, a nonrandom pattern of X inactivation (i.e., retention of the chromosome carrying functional BTK) is observed in the mature B cells of female carriers of XLA (Conley and Puck, 1988; Fearon et al., 1987). A variety of polymorphic markers linked to the BTK locus have been described and have proven useful for chromosome linkage studies to demonstrate carrier status in females at risk, or for prenatal diagnosis (Conley, 1993; Journet et al., 1992; Kwan et al., 1994; O’Reilly et al., 1993; Parkar et al., 1994; Schuurman et al., 1988; Sweatman et al., 1993). The earliest methods used methylation-dependent restriction fragment length polymor-
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phisms to distinguish active from inactive X chromosomes. Similar methods have been applied to various linked markers such as DXS255 (Hinds et al., 1993) or the human androgen receptor gene (Allen et al., 1992; Wengler et al., 1997). PCR-based methods have also been used to analyze the androgen receptor gene (Allen et al., 1994; Alterman et al., 1993), the monoamine oxidase A gene (Hendriks et al., 1993), and the BTK locus itself (Moschese et al., 2000). Using this method, among maternal ancestors of males having BTK mutations, 84% of mothers, and 16% of maternal grandmothers are found to be carriers, indicating the high rate of new mutations (Conley et al., 1998). Germline mosaicism for BTK defects has been observed both in males (Hendriks et al., 1989), and in females (Puck et al., 1995). As a result, mothers of males with sporadic XLA, who are not demonstrated to be carriers, have a low (estimated <5%) but unquantifiable risk of having another affected son. BTK mutations have also been found in monozygotic female twins, although absent from their mother, thus suggesting germline mosaicism, or mutation in the zygote (Curtis et al., 2000).
TABLE 26.6 Mutations in autosomal recessive agammaglobulinemias Gene IGHM
The IgM heavy chain is first expressed on the surface of developing pre-B cells in a complex with the surrogate light chain (heterodimer of l5 and VpreB) and the signal transducing molecules Iga (CD79a) and Igb (CD79b). IgM heavy chain gene mutations were first described in six agammaglobulinemic patients by Yel et al. in 1996. Between ten and fifteen such patients have been identified to date (Conley et al., 2000; Milili et al., 2002). A variety of mutations leading to the inability to express IgM have been described (Table 26.6). These patients have clinical and laboratory features identical to classical XLA, perhaps with a tendency toward more severe phenotype and earlier diagnosis of agammaglobulinemia (Conley et al., 2000). Current clinical laboratory methods do not reliably distinguish between serum IgM levels of 0 and approximately 0.1 g/L. Thus, a molecular lesion involving IgM cannot be excluded in patients reported to have very low levels of serum IgM. Flow cytometric analysis of bone marrow cells from one patient showed normal numbers of TdT+/CD34+/CD19+ pro-B cells, and absence of TdT-/CD34-/CD19+ pre-B cells (Conley et al., 2000).
Agammaglobulinemia Due to Mutations Affecting l5/IGLL1 The human genes encoding the l5 and VpreB proteins, respectively, are called IGLL1 (also called 14.1) and
75 kb deletion >260 kb deletion
1831 G Æ A
1768 T Æ G
IGLL1
C Æ T in codon 22 425 C Æ T
CD79A
A Æ G exon 3 splice G Æ A intron 2
BLNK
Agammaglobulinemia Due to Mutations of the m Heavy Chain Gene
Mutation
A Æ T intron 1 splice
Consequence
Reference
Loss of D, JH and Cm genes Loss of some VH, all D and JH, and C genes Abnormal splicing, loss of membrane IgM C536G, loss of cysteine required for intrachain disulfide bond
(Yel et al., 1996) (Yel et al., 1996)
(Yel et al., 1996)
(Yel et al., 1996)
(Q22X), premature STOP codon P142L, abnormal protein folding
(Minegishi et al., 1998) (Minegishi et al., 1998)
Abnormal splicing, loss of exon 3 Presumed abnormal splicing
(Minegishi et al., 1999) (Wang et al., 2002)
Abnormal splicing, mRNA not detected
(Minegishi et al., 1999)
VPREB1. Minegishi et al. have described one male agammaglobulinemic patient who is a compound heterozygote for mutations of IGLL1. One allele contained a mutation (CÆT in codon 22) leading to a premature stop codon in exon 1 (Table 26.6). The other allele was apparently generated by gene conversion with the l5 pseudogene IGLL3 (also called 16.1), and contained three base substitutions, including one that changed an invariant proline to leucine. This led to abnormal protein folding and expression. Flow cytometry of CD19+ peripheral blood B cells (0.06% of all lymphocytes) from this patient revealed a normal IgMlo/CD38lo phenotype. Only 6% of bone marrow B cells expressed CD19; these were predominantly TdT+/CD34+/cm-, thus indicating a block at the pro-B to preB cell transition. The same group of authors described a high rate of gene conversion events leading to polymorphisms in l5 in healthy individuals (Conley and Rohrer, 1995). They studied 134 unrelated individuals and found thirteen variant alleles, nine (69%) of which led to amino acid changes in l5. Most of these were consistent with gene conversion events involving the l5 pseudogenes IGLL2 (16.2) and IGLL3 (16.1). Gene conversions with Cl genes, as well as novel single nucleotide polymorphisms, were also seen. Two individuals homozygous for a T148I polymorphism were studied. Both had B cell numbers at the low end of the normal range, and one had slightly low serum IgM.
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Agammaglobulinemia Due to Mutations Affecting Iga Iga (CD79a) is a signal transducing component of B-cell Ig receptors. It is directly associated with the Ig heavy chain, and links it via noncovalent interactions with another signal transducing component of the receptor, Igb (CD79b). Iga contributes both to pre-B cell and mature B-cell Ig receptors. The cytoplasmic portion of Iga contains an immunoreceptor tyrosine containing activation motif (ITAM) and links the receptor to Src family tyrosine kinases. To date, two patients having Iga mutations have been reported in the medical literature (Minegishi et al., 1999; Wang et al., 2002). They have a clinical phenotype indistinguishable from the classic form of XLA, and have agammaglobulinemia with very low numbers of circulating B cells. One female patient was homozygous for a splice-site mutation (Table 26.6), and <0.01% of peripheral lymphocytes were CD19+ (Minegishi et al., 1999). Analysis of bone marrow B-cell development was identical to that seen with mutations affecting IGHM, thus indicating a block at the pro-B to pre-B cell transition. Transcripts encoding TdT, RAG1, VpreB, and l5 were found in normal abundance; transcripts of fully rearranged Ig heavy chains were markedly reduced. The repertoire of rearranged VH genes was limited. Another male patient was homozygous for a splice-site mutation in intron 2 at the +1 position (Table 26.6).
Agammaglobulinemia Due to Mutations Affecting B-Cell Linker Protein The B-cell linker protein (BLNK) is a signal transduction adapter molecule whose expression is restricted to B cells in humans. This molecule links Syk-family kinases with phosphoinositide 3¢ kinase and MAP kinase pathways during B-cell receptor signaling. To date, BLNK mutation has been described in a single male patient with agammaglobulinemia and absence of B cells (Minegishi et al., 1999). He was found to be homozygous for two base substitutions—one was a silent change in codon 10, and the other was a splice mutation (Table 26.6). This patient had <0.01% circulating CD19+ lymphocytes. Only 0.3% of bone marrow cells were CD19+. These cells exhibited a block at the pro-B to pre-B cell transition, as seen in patients with mutations affecting Cm, l5, and Iga.
AUTOSOMAL RECESSIVE HYPER-IGM SYNDROME Hyper-IgM syndrome (HIGM) is an eponym given to a group of immunodeficiencies characterized most often by low levels of IgG, IgA, and IgE together with normal or elevated IgM. The X-linked form of this disorder (HIGM1)
results from a mutation in the gene encoding tumor necrosis factor superfamily member 5 (TNFSF5), also called CD154, or CD40 ligand (Bonilla and Geha, 2001). This molecule is expressed predominantly on activated T cells, and plays an important role in T-cell interactions with antigen presenting cells. It also regulates B-cell activation and isotype switching, and the induction of effector activities of mononuclear cells (Bonilla and Geha, 2001). Clinical features include bacterial respiratory tract infections as seen in agammaglobulinemia, together with opportunistic infections and neutropenia. Autosomal recessive forms of this disorder have long been recognized. Three gene defects have recently been identified in this group of patients.
Mutations of CD40 The ligand for TNFSF5 (CD154) is tumor necrosis factor receptor superfamily member 5 (TNFRSF5), also called CD40. TNFRSF5 is expressed principally on B cells, dendritic cells, and monocytes and macrophages, and may also be expressed in a variety of other cell types (van Kooten and Banchereau, 2000). Mutation of this molecule leads to a combined immunodeficiency indistinguishable from HIGM1 (Ferrari et al., 2001). Three individuals from two separate kindreds, each having consanguinity, have been described. Clinical features included bacterial respiratory tract infections, P. carinii pneumonia, and neutropenia. Circulating lymphocyte populations and responses to polyclonal T-cell mitogens were normal. Flow cytometric analysis showed that most circulating B cells were IgM+/IgD+; CD27+ cells were greatly reduced. CD40 was not detected on B cells. Serum IgG ranged from 0 to 1.8 g/L, IgA was undetectable, and IgM ranged from 0.81 to 4.0 g/L. B cells did not produce immunoglobulin in vitro in response to stimulation with FcgRII-expressing L cells, anti-CD40 monoclonal antibody, and interleukin-10. One individual was homozygous for a silent mutation in codon 136 (threonine, nucleotide 455 AÆT). This change at the fifth position in exon 5 led to complete skipping of this exon in mRNA transcripts, with a resulting frameshift in codon 135 and chain termination in codon 191. Two other patients were homozygous for a C83R mutation (nucleotide 294 TÆC), which is presumed to lead to abnormal protein folding, processing, or both. In one of these patients, CD40 with abnormal electrophoretic mobility (apparently smaller) was seen in western blots.
Mutation of the Activation Induced Cytidine Deaminase and Uracil-DNA Glycorylase Genes The activation induced cytidine deaminase enzyme (often abbreviated AID, although the gene designation is AICDA) is a DNA-editing enzyme that has a central role in the processes of Ig class-switch recombination and somatic hypermutation (Honjo et al., 2002; Muramatsu et al., 2000),
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changes in Ig light chain gene usage associated with affinity maturation (Meffre et al., 2001), and gene conversion (Arakawa et al., 2002). Revy et al. reported the occurrence of AID defects in a group of eighteen patients with autosomal recessive hyper-IgM syndrome from 12 families (2000). This form of HIGM is often referred to as HIGM2. These patients exhibited recurrent respiratory and gastrointestinal tract infections, but opportunistic infections were not seen. In addition, many (13/18 or 72%) displayed significant hypertrophy of secondary lymphoid tissues due to enlargement of germinal centers. These patients had very low levels of serum IgG (<0.06–1.3 g/L) with very elevated IgM (1– 37 g/L). This range of IgG is lower, and IgM higher, than is seen in X-linked HIGM (Bonilla and Geha, 2001). The AICDA mutations identified in these patients included missense and nonsense mutations in coding regions, as well as 9 and 19 base pair deletions, and one larger deletion spanning exon 3. Most patients were homozygous for mutations, but some compound heterozygotes were identified. Somatic mutation of the VH3 gene family member V3-23 was reduced between three- and seven-fold in comparison to controls. Germinal centers (GC) in tonsils from these patients revealed normal mantle zones, interfollicular areas, and follicular dendritic cell network. GC B cells had a high proliferation frequency, and many coexpressed sIgD, in contrast to normal GC B cells. The majority of proliferating cells were CD38+/sIgM+/sIgD+ “founder” B cells (Lebecque et al., 1997). Minegishi et al. studied 27 patients with HIGM lacking CD154 mutations and found AID defects in 18 (67%) (Minegishi et al., 2000). A group of 14 individuals of French-Canadian ancestry apparently shared a founder mutation, as they were all homozygous for the same point mutation in exon 3 leading to altered amino acid sequence (R112C). Two additional mutations were identified— another base substitution at codon 112 (R112H), and another exon 3 mutation leading to premature chain termination (W84X). These authors also identified two common polymorphisms, one intronic and one silent base substitution, in a population of 100 controls. This group of patients was also affected by bacterial respiratory tract infections, the majority had had pneumonia, and 28% had bronchiectasis. Other bacterial infections, including sepsis, were seen; there were no opportunistic infections; 50% had lymphoid hyperplasia. In vitro, peripheral B cells from four patients with AICDA mutations showed increased expression of CD23 and CD25 after stimulation with IL-4. However, there was no increased response in the presence of anti-CD40. B cells from HIGM2 patients produced only 0.1% or less of the amount of IgE secreted by control B cells when cultured with anti-CD40 and IL-4. Noguchi et al. recently described polymorphisms of the AICDA gene in a group of Japanese families with members with and without asthma (2001). These authors found that a 7888C/T polymorphism (a silent mutation in exon 4) was
significantly associated with alterations in serum IgE level in adults. Specifically, 7888C/C was associated with higher levels of serum IgE in comparison to 7888C/T or 7888T/T, with a reported significance of p = 0.02. One rare variant allele and two additional polymorphisms were described, but there was no clinical correlation. It remains to be determined how and to what extent the silent 7888C/T polymorphism might contribute to the pathogenesis of asthma or other atopic diseases. This association may also reflect the linkage of the polymorphism to another locus. Mutations of uracil-DNA glycosylase (UNG) have recently been described in three patients with an autosomal recessive form of hyper-IgM syndrome (Imai et al., 2003). These individuals have clinical and laboratory phenotypes very similar to patients with AID deficiency. Ig class-switching is severely impaired, and there is a selective defect in somatic hypermutation involving transversion at dG or dC.
TABLE 26.7 Single gene mouse knockout models with selective humoral immune defects Gene
Protein (Reference)
Models with reduced B cell development and dysgammaglobulinemia and/or impaired antibody responses BCL3* B-cell CLL/lymphoma 3 transcription co-activator (Franzoso et al., 1997) FES V-FES feline sarcoma viral/V-FPS fujinami avian sarcoma viral oncogene homolog (tyrosine kinase) (Hackenmiller et al., 2000) CD22 CD22 antigen (Otipoby et al., 1996) IL5RA Interleukin-5 receptor a (Yoshida et al., 1996) LYN Lyn tyrosine kinase (Hibbs et al., 1995) MAP3K14 Mitogen-activated protein kinase kinase kinase 14 (Yin et al., 2001) PIK3R1 Phosphatidylinositol 3-kinase, regulatory subunit, polypeptide 1 (p85 alpha) (Suzuki et al., 1999) POU2AF1 POU domain, class 2, associating factor 1 (transcription co-activator) (Schubart et al., 1996) Models with normal B cell number and dysgammaglobulinemia and/or impaired antibody responses CCR6 Chemokine (C-C motif) receptor 6 (Cook et al., 2000) CD23 CD23 antigen (Yu et al., 1994) CD28 CD28 antigen (Shahinian et al., 1993) CD81 CD81 antigen (Maecker and Levy, 1997, Tsitsikov et al., 1997) IRF4 Interferon regulatory factor 4 (Mittrucker et al., 1997) LTB Lymphotoxin-a isoforms a and b (Koni et al., 1997) NFKB1 Nuclear factor of kappa light polypeptide gene enhancer in B-cells 1 (p105) (Sha et al., 1995) RELB Nuclear factor of kappa light polypeptide gene enhancer in B-cells 3 (Weih et al., 1995) TNFRSF13B Tumor necrosis factor receptor superfamily 13B (von Bulow et al., 2001) * These are official HGNC designations for the human genes corresponding to the murine model.
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MURINE MODELS OF HUMAN B-CELL DEFICIENCY Mice with a targeted disruption of each of the genes listed in Table 26.1 have been generated. In some cases, observation of the murine phenotype prompted the search for the same mutation in humans having similar features. Table 26.7 lists knockout mouse models for relatively selective B-cell deficiencies. More details regarding the immunological phenotypes of these models may be viewed in a searchable database of mouse knockouts with immunodeficiency at http://immunology.tch.harvard.edu/. These genes may be viewed as candidates for study in humans having similar immunological characteristics.
CONCLUSION It is quite striking that all of the genetic defects of bone marrow B-cell development so far identified in humans directly involve elements that are critical for signaling via Ig receptors expressed on B-cell precursors. The implications of this with respect to the evolution of the human immune system remain to be fully appreciated. It is doubtless that study of naturally occurring human genetic mutations, together with targeted gene disruption methods in animal models, will continue to go hand-in-hand to further our understanding of immune physiology.
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27 Diverse Forms of Immunoglobulin Genes in Lower Vertebrates GARY W. LITMAN
MARTIN F. FLAJNIK
GREGORY W. WARR
Department of Molecular Genetics, All Children’s Hospital, St. Petersburg, Florida Immunology Program, H. Lee Moffitt Cancer Center and Research Institute, Tampa, Florida Department of Pediatrics, University of South Florida, Children’s Research Institute, St. Petersburg, Florida, USA
Department of Microbiology and Immunology, University of Maryland, Baltimore, Maryland, USA
Department of Biochemistry, and Center for Marine Biomedicine and Environmental Sciences, Medical University of South Carolina, Charleston, South Carolina, USA
somatic variation of Ig genes has been described (Lewis and Hsu, this volume). Despite the current inability to definitively identify Ig or Ig-like genes in species that are found below the phylogenetic level of the jawed vertebrates, recent efforts to describe candidate Ig-like molecules, as well as the more widespread use of genomics technology, hold considerable promise for elucidating both the distant phylogenetic origins of the diversified Ig-like genes and the mechanisms that have diversified complex families of humoral immune receptors.
Over the past several years, considerable progress has been made in characterizing the structure, organization, and regulation of expression of immunoglobulin (Ig) gene loci in a large number of lower vertebrate species. It presently is possible to define the major patterns of diversification of Ig structure within the most significant evolutionary radiations of the jawed vertebrates (Litman et al., 1999; Flajnik, 2002). When viewed collectively, the form and function of the Ig genes are complex and exhibit a high degree of interspecies variation, even at relatively early points in vertebrate phylogeny. In marked contrast, the organization and diversification of T-cell antigen receptor (TCR) genes has remained relatively stable in those species in which Ig and TCR genes can be compared. Neither the evolutionary basis for the extraordinary variation in Ig structure and organization nor the relationships of many of the differences in Ig structure to humoral immune function are understood (Flajnik, 2002). Although this chapter describes many of the unique Ig gene structures found in modern representatives of phylogenetically ancient species and attempts to describe systematic trends in Ig phylogeny, the primary emphasis has been placed on defining those features of structural diversification that potentially are most significant for interpreting the function and genetic regulation of the Ig genes as antigen binding receptors. Several other chapters in this text explore in greater depth some of the lower vertebrate model systems, including avians, which have factored prominently in achieving our current understanding of unique aspects of gut-associated lymphoid tissue (GALT) immunity (Knight, this volume) and cartilaginous fish, in which complex
Molecular Biology of B Cells
CARTILAGINOUS FISH: AN UNUSUAL EXAMPLE OF GENE MULTIPLICITY Both pentameric and monomeric forms of IgM can be detected at equivalent levels in the serum of a cartilaginous fish (Marchalonis and Edelman, 1965; Marchalonis and Edelman, 1966). In addition, these species possess at least two other Ig isotypes designated IgX, IgW, or IgNARC, depending on the species in which it was identified (which will be referred to as IgW in this chapter as a matter of convention), and IgNAR. IgM constitutes approximately half of the serum protein (Clem et al., 1967), with concentrations exceeding 20 mg/ml, whereas IgNAR levels range from 0.1 to 1 mg/ml. IgW has been difficult to detect in the serum, due in part to its apparent sensitivity to proteolysis (H. Dooley and M. Flajnik, unpublished observation).
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IgM genes are organized in clusters containing one VH (variable heavy chain) two DH (diversity heavy chain), one JH (joining heavy chain), and a single set of CH (constant heavy chain) exons; rearrangement generally occurs only within a cluster (Hinds and Litman, 1986; Kokubu et al., 1988b). A single VH family, which can exhibit >90% amino acid identity between the VH domains within a species (Kokubu et al., 1988a), predominates in the three species of sharks that have been characterized most extensively to date. However, extensive heterogeneity in the germline complementarity determining regions (CDRs) has been shown in Heterodontus francisci (horned shark) (Kokubu et al., 1988a), Carcharhinus plumbeus (sandbar shark) (Shen et al., 1996), Ginglymostoma cirratum (nurse shark; L. Rumfelt and M. Flajnik, unpublished observation), and Raja erinacea (little skate; F. Harding and G. Litman, unpublished observation). The number of m heavy chain genes differs between species; very large numbers of m heavy chain genes are found in horned shark (Kokubu et al., 1988b; Kokubu et al., 1988a), little skate (Harding et al., 1990a) and Raja eglanteria (clearnose skate; A. Miracle and G. Litman, unpublished observation). The horned shark has the largest genome of any elasmobranch examined thus far (Schwartz and Maddock, 2002); the relationship of the variation in gene number to genome size or to expansions and contractions of this multigene family is unclear. Single-cell PCR analyses have demonstrated that only a single potentially productive heavy chain gene transcript can be detected in the majority of peripheral blood lymphocytes in clear nose skate, thus suggesting that haplotype exclusion occurs in this species (Eason and Litman, 2002). IgNAR is an H-chain dimer and does not associate with light chains (Greenberg et al., 1995; Roux et al., 1998). The VH regions of IgNAR are not related closely to the corresponding VH regions of IgM or IgW, but rather are somewhat more similar to the V regions of TCR or Ig light chain genes (Greenberg et al., 1995; Richards and Nelson, 2000). VH domains in IgNAR are free and flexible, and have been shown to be useful for expressing soluble antigenrecognition molecules from phage-display libraries (Dooley et al., 2003; Nuttall et al., 2002; Nuttall et al., 2001). Two IgNAR types (I and II), described in nurse sharks (Roux et al., 1998), are distinguished primarily by the presence of non-canonical cysteines that are presumed to form disulfide bonds that result in structural compaction. Some of these cysteines are encoded by the D (preferred reading frames) or J segments. It is likely that the two IgNAR types exhibit different conformations based on these unusual disulfide bridges and as predicted from protein modeling studies (Diaz et al., 2002). The longer length of CDR3 in IgNAR type I sequences is consistent with the predicted conformational differences, despite the findings that both type I and type II genes contain three D regions and generally undergo four rearrangement events (Roux et al., 1998; Diaz et al., 2002).
The IgNAR genes hypermutate extensively (Greenberg et al., 1995; Diaz et al., 1999), and replacement/silent (R/S) ratios in CDR1 for the type I form and CDR2 for the type II form are consistent with positive selection. The relationship of IgW to IgNAR suggests that both genes existed before the common elasmobranch ancestor 220 million years ago (Greenberg et al., 1996; Anderson et al., 1999). The last four CH domains of IgNAR are homologous to those of IgW and have led to the suggestion that an en bloc duplication of two exons encoding CH domains apparently gave rise to the ancestral IgW and IgNAR gene from a precursor having four CH domains. Since IgM also has four CH domains, it was proposed that IgW/IgNAR was derived from an IgM-like ancestor (Anderson et al., 1999), although phylogenetic analyses cannot predict whether the IgM or IgW V regions is older (Bernstein et al., 1996). Because an IgW homolog has been identified in Protopterus aethiopicus (lungfish; see below), it stands to reason that both an IgM-like and an IgW-like gene were present in the common ancestor of all extant, jawed fish, thus confounding the identification of which represents the more primordial form. Pentameric IgM precedes the appearance of monomeric IgM in the serum (Fidler et al., 1969). Recently it was shown that the major forms of IgNAR are not expressed at high levels at birth (Diaz et al., 2002; Rumfelt et al., 2002). However, transcripts of IgW genes can be detected early in both skates and nurse sharks (Miracle et al., 2001; L. Rumfelt and M. Flajnik, unpublished observation). Recent data in nurse shark have shown that neonatal splenic white pulp is composed entirely of Ig+ B cells; by 3 months of age, white pulp containing non-Ig+ cells appears and presumably represents T-cell zones that become populated by class II+ dendritic cells (Rumfelt et al., 2002). IgNAR+ cells become detectable concurrently in both red and white pulp, and IgNAR protein appears in the plasma. The rather late appearance of monomeric IgM, IgNAR, and organized T-dependent areas in secondary lymphoid tissues suggests that acquired immunity is suboptimal in young animals. The presence of extended families of totally or partially germline-joined Ig clusters is a property of Ig genes in cartilaginous fish (Kokubu et al., 1988a). One of the nurse shark IgM heavy chain clusters (IgM1gj) is fully germline-joined (VDJ) and lacks the CH2-encoding exon, resembling the number of domains found in mammalian IgG heavy chains (Rumfelt et al., 2001). The IgM1gj V region gene is single copy, monotypic, and related more closely to the IgM VH regions of horned shark than to the predominant IgM VH of nurse shark. IgM1gj is expressed early in the ontogeny in the spleen and epigonal organ, and later in life is detectable only in the epigonal organ. Recently, it was found that a type I germline-joined light chain gene is a preferential partner of the IgM1gj heavy chain (E. Hsu and M. Flajnik, unpublished observation). Along these lines, it is notable that one of the
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four IgNAR clusters (type III) has a “germline-joined” D1/D2 segment, and also is expressed preferentially early in ontogeny (Diaz et al., 2002). Despite the presence of Nregion additions in the early type I and type II sequences and the presence of V-D1/D2 and D1/D2-J boundaries in the type III gene, the CDR3 length of the type III IgNAR is highly constrained, suggesting that a particular binding site selects for a specific ligand—that is, a partially germlinejoined gene that behaves as though it were entirely joined. Notably, N-region addition has been shown to occur very early in ontogeny in the clearnose skate (Miracle et al., 2001), and extremely low levels of TdT expression were associated with N-region additions in Ig genes, a phenomenon that also has been observed in TCRa in mouse (Cherrier et al., 2002). The structural organizations of the different immunoglobulins in cartilaginous fish are compared with those found in species representative of more recently derived points in evolution (Figure 27.1).
BONY FISH: IG HEAVY CHAIN GENES RESEMBLE IGM AND IGD Prior to the description of Ig heavy chain (IgH) genes in fleshy-finned fish (see below), the bony fish were considered the defining taxon in the transition from the multicluster form of IgH organization seen in the cartilaginous fish to a single “translocon”-IgH locus, such as found in mammals. As a group, the bony fish IgH locus typically encodes a single IgM-like heavy chain and a second class of Ig, which has been termed IgD-like (and is discussed further below). Extensive studies in different mammalian species have demonstrated the importance of RNA processing, which permits the co-expression of IgM and IgD and enables the expression of each class of Ig in either the membrane receptor or secreted form. RNA processing also plays a significant role in the regulation of Ig expression in lower vertebrate species. IgD has been identified in the mammalian orders of rodents, primates, and artiodactyls (Abney, Parkhouse, 1974; Zhao et al., 2002) as well as in bony fish (Wilson et al., 1997; Hordvik et al., 1999; Stenvik, Jorgensen, 2000). The specific function of IgD in mammals is not understood, and comprehensive efforts to detect IgD in other species, including Anas platyrhynchos (duck) (Lundqvist et al., 2001) have not been successful. The absence (presumably due to loss) of IgD from certain evolutionary lineages, as well as its extreme structural variation (two Cd domains in mouse compared with up to seven Cd domains in the fish IgD-like molecule) suggest that it may be functionally redundant, which is supported by the targeted disruption of the IgD locus in mice (Roes and Rajewsky, 1993; Nitschke et al., 1993). The Cd gene typically is found immediately 3¢ of the Cm gene; this linkage relationship permits the co-expression of m and d IgH through the alternative processing of a long
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primary transcript that includes the rearranged VDJ, Cm, and Cd genes. Thus the significance of d-type heavy chain sequences that occur 5¢ of the Cm gene in Fugu rubripes (pufferfish) and Ictalurus punctatus (channel catfish) (as discussed below) is difficult to evaluate. Despite their use of a similar mechanism of RNA processing, there are significant differences between the genes encoding IgD in bony fish and mammals. In bony fish, the Cd-encoding mRNA is chimeric and always contains the exon that encodes the Cm1 domain (Wilson et al., 1997; Hordvik et al., 1999; Stenvik and Jorgensen, 2000; Hordvik, 2002). Since the Cd1 domain is missing the Cys residue involved in H/L covalent bonding in at least one species of bony fish, the Cm1 exon between the VDJ sequence and the Cd1 exon may be required to effect disulfide bonding between the d-IgH chain and the light chain (Wilson et al., 1997). In mammals, a single d gene encodes the membrane as well as secreted forms and differential RNA processing produces mRNA with different C-terminal sequences, which specify the different forms; whereas, in bony fish the membrane and secreted forms of the d-IgH chain are encoded by different genes (Bengten et al., 2002; Aparicio et al., 2002). The gene for the membrane form of the Cd gene in catfish is downstream of the Cm gene, whereas the gene for the secreted form is upstream of the Cm gene (Bengten et al., 2002). Given the physical separation of the membrane and secreted forms of the Cd gene, it is unclear how the secreted form of IgD can be expressed as a functional antibody. The VH, DH, and JH segments that encode the VH domains of bony fish immunoglobulins show strong conservation in terms of structure and organization when compared to those of other vertebrates that also possess IgH loci in the “translocon” organization. Similarities include the sequential organization along the chromosome of VH, DH, and JH segments, with canonical recombination signal sequences, and the presence of large numbers of VH genes that fall into characteristic families. For example, there are at least seven families of VH in the catfish (Ventura-Holman et al., 1996). The evolutionary relationships of VH gene families within the vertebrates is a fascinating and complicated story that is beyond the scope of this review but is discussed in Ota et al. (2000). The organization of the IgH loci in bony fish appears to have undergone disruption. In addition to the presence of a d gene sequence upstream of the m gene (discussed above), the IgH locus of catfish contains Cm pseudogenes and VH gene–encoding segments (including germline-joined VDJ) that are distributed over distances of at least 700 kb (Ghaffari, Lobb, 1999b; Bengten et al., 2002).
Expression of Membrane Receptor and Secreted Forms of Igs The membrane receptor and secreted forms of every class of Ig in mammals and birds, with the exception of IgD, are expressed from a single gene by a highly conserved mech-
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FIGURE 27.1 Immunoglobulin classes and their phylogenetic relationships in the jawed vertebrates. (A) Schematic of the vertebrate Ig classes identified to date. IgM is a pentamer or hexamer in all tetrapods, but is a tetramer in bony fish and either a pentamer or monomer in the cartilaginous fish. IgD varies in the number of constant domains depending on the species examined, both in mammals and bony fish (see text). Note that based on amino acid sequences, there are no detectable hinge regions in the majority of non-mammalian vertebrate isotypes, despite the “Y” shape of all Igs shown in the figure. (B) Approximate times (in millions of years) for the appearance of the various Ig classes in evolution. See text for details on the emergence of IgM, IgMlgj, IgD, IgW, and IgY.
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anism, in which the hydrophobic membrane-spanning tail of the membrane receptor form is encoded separately by one or (more typically) two exons that lie 3¢ of the polyadenylation signal for the secreted form of the message. The membrane receptor–encoding mRNA is produced by splicing the membrane exon(s) into a cryptic splice donor site in the terminal exon of the secreted form, as illustrated for IgM in Figure 27.2. This mechanism also is highly conserved for Igs in amphibians, and for IgM in the cartilaginous fishes (Ross et al., 1998). The major exceptions to this type of differential processing are found in the Ig of bony fishes as well as for the IgW and IgNAR isotypes in cartilaginous fish (see below). The structure of the m heavy chain genes of teleost fish conforms to the pattern of four secretory exons and two transmembrane exons, which is highly conserved throughout higher vertebrates (Figure 27.2). Different species of teleost fish generate the membrane receptor form of IgM by
FIGURE 27.2 Alternative pathways, within the vertebrates, by which the message encoding the membrane receptor form (mm) of the IgM heavy chain is produced. A phylogenetic tree of the vertebrates is shown, along with the distribution of known pathways of RNA processing to produce the mm message. Cm, the 4 constant region exons of the m gene; TM, the 2 exons encoding the transmembrane tail of the mm heavy chain. The cleavage/ polyadenylation signal (filled black circle) for mm is shown, but the splicing pattern and cleavage/polyadenylation signal for the secretory form of the m chain are omitted for clarity. Mya, million years ago.
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alternative RNA processing, in which the transmembrane exons are spliced to the splice donor site at the 3¢ boundary of the Cm3 exon (Figure 27.2) rather than to a cryptic splice site within Cm4 (Wilson et al., 1990; Bengten et al., 1991; Lee et al., 1993). A somewhat similar process has been characterized in Notothenia coriiceps (yellowbelly rockcod) a member of the highly cold-adapted notothenioid fish from the Antarctic. The transmembrane form of the m chain is spliced to the splice donor site at the 3¢ end of the Cm2 exon in this species (Ota et al., 2002). The extracellular portion of the transmembrane sequence of the rockcod is exceptionally long and might reflect adaptation to the extreme cold of the Antarctic Ocean. The teleosts represent the most highly evolved group of the ray-finned bony fishes or actinopterygian fishes; the chondrostean (sturgeons, paddlefish) and holostean fishes (bowfin, gars) are representative of earlier evolutionary stages in this lineage (Figure 27.2). The splice patterns by which the membrane receptor forms of IgM are generated in the holostean fishes (data are not available for the chondrostean fishes) have been studied in Lepisosteus osseus (longnose gar) and Amia calva (bowfin). In both species, the “mammalian” Cm4 cryptic splice donor site and the “teleost” Cm3 splice donor site are used to generate two in-frame species of mRNA-encoding transmembrane forms of the m heavy chain (Wilson et al., 1995). The most conservative interpretation of this observation is that during the evolution of the bony fish, there was a period of instability in which multiple pathways were used for generating the mRNA encoding transmembrane m form and that the Cm3 Æ transmembrane m form became the dominant pathway. The unusual transmembrane m mRNA in rockcod (Ota et al., 2002) and a third pathway in the bowfin, in which the transmembrane exons are spliced to a cryptic site within Cm3 (Wilson et al., 1995), suggest instability in RNA processing in the actinopterygian fishes. The utilization of so many different RNA processing pathways in bony fish could relate to defects in the processing machinery. Specifically, B cells in bony fish cannot splice the primary transcript from a mammalian Ig gene, whereas mammalian B cells splice the transcript from a fish m gene using the pathway that has been defined in teleosts (Ross et al., 1998). Bony fish provide a unique view of alternative possibilities for the utilization of B-cell receptors, some of which can be examined through transgenesis approaches in mammalian systems that are amenable to the study of interactions of defined cell lineages and are more suitable for functional assays. Distinctive patterns of Ig mRNA processing also have been identified in cartilaginous fish. The secreted form of the IgW isotype has been identified in short (two CH domains) and long (six CH domains) versions in the little skate (Greenberg et al., 1996; Anderson et al., 1999; Kobayashi et al., 1984; Harding et al., 1990b; Anderson et al., 1994) and presumably represent alternatively spliced
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isoforms. The long isoform has been described in the sandbar shark (Bernstein et al., 1996) and nurse shark (Greenberg et al., 1996), and short isoforms have been identified in both shark species (L. Rumfelt and Flajnik, in preparation) and possibly in Chlamydoselachus anguineus (frill shark) (Kobayashi et al., 1992). The short isoforms have an unusual secretory tail that is not homologous to the Cterminus of any other vertebrate heavy chain isotype. Although skate, nurse shark, and horned shark share a number of cysteine residues and an N-linked glycosylation site in this region, they otherwise are highly divergent. This low level of sequence identity is uncommon among the orthologous IgH and IgL chains in cartilaginous fish. The short form of IgW, which is not expressed in all nurse sharks tested and may depend on the immune status and state of health of the animal, may be important in limiting inflammatory responses in cartilaginous fish, as has been proposed for the alternative forms of IgY in some birds (Flajnik, 2002). Like the transmembrane forms of IgM found in bony fish, the IgW transmembrane proteins in horned and nurse sharks are generated by alternative splicing. The transmembrane region is contiguous to the CH4 domain in all cDNA clones analyzed to date and thus the heavy chain of the IgW transmembrane protein is the same size as the transmembrane form of IgM heavy chains (L. Rumfelt and M. Flajnik, in preparation). PCR analyses have confirmed that this four-CH domain molecule is the principal IgW transmembrane type. However, there may be minor forms in which the transmembrane exons are spliced to exons encoding other domains. Transmembrane forms of IgNAR splice either to the CH3 or to the CH5 domain exons; to date, no unusual secretory forms of IgNAR have been identified.
LOBE-FINNED FISH: A “TRANSITIONAL” ARRANGEMENT OF RECOMBINING ELEMENTS In contrast to the broadly divergent taxa that encompass the bony fishes, the lobe-finned fishes, which include the coelacanth, are the sole extant occupants of a major radiation of the vertebrates, which were assumed to be long extinct prior to the discovery of a living species of Latimeria chalumnae (Comoran coelacanth) in 1938. The genomic locus encoding VH genes in the coelacanth possesses general features of both the cluster organization found in the cartilaginous fish and the translocon organization seen in other vertebrates. Specifically, each VH segment is separated by ~200 bp from a DH segment, a linkage feature reminiscent of IgH genes in cartilaginous fish. Genomic Southern blot and differential library screening analyses are consistent with a large family of VH genes. The failure to detect JH or CH (Cm) regions in VH-containing genomic phage clones, has been interpreted to indicate that these gene segments are not
localized to within 10 to 20 kb of VH. Bacterial artificial chromosomes (BACs) encoding VH clones were isolated recently from Latimeria menadoensis (Indonesian coelacanth), a closely related species (Holder et al., 1999). The close linkage of VH and DH segments detected previously in the Comoran coelacanth has been confirmed in the Indonesian species (Amemiya, personal communication). A BAC contig containing the Cm exons was isolated; however, efforts to identify VH elements in the contig using an ubiquitous PCR primer strategy were unsuccessful. Likewise, a Cm exon could not be detected in VH-containing BACs. Outside of the close linkage of VH and DH, the organization of the Ig locus in the coelacanth differs from that of the cluster-type loci described in cartilaginous fishes.
FLESHY-FINNED FISH: AN ANCIENT ORIGIN FOR ISOTYPE DIVERSITY The fleshy-finned fishes, which include several different genera of lungfishes, represent another major vertebrate radiation that diverged prior to the tetrapods. A m-type heavy chain, as well as two different non-m-type IgH isotypes, has been characterized recently in the African lungfish, Protopterus aethiopicus (Ota et al., 2003). Extensive sequencing of lungfish Ig cDNAs as well as RNA blot analyses suggest that both the m and non-m isotypes, which vary in length, are encoded at separate loci. The two non-Igm isotypes share overall length as well as significant sequence identity with the short and long forms of the IgW heavy chain, which heretofore were considered as being unique to cartilaginous fish (see above). The presence of an IgW-type molecule in both cartilaginous and fleshy-finned fish, and its absence in the bony fish and the tetrapod lineage is difficult to reconcile with the phylogenetic relationships of vertebrates and underscores the ancient origins of Ig isotype diversity. The most conservative interpretation is that the IgW-type genes were lost from the teleost and tetrapod lineages. As with the findings for coelacanth, it appears as if an element of the cartilaginous fish, in this case “independent” loci, occurs in a representative species that diverged more recently in phylogeny. The findings with lungfish emphasize the difficulties in reconciling genetic findings with Ig genes to the known patterns of phylogenetic divergence in vertebrates.
AMPHIBIANS AND REPTILES: THE POSSIBLE ORIGINS OF CLASS SWITCHING The amphibians demonstrate an example of yet another distinct Ig system and represent the most phylogenetically early example of Ig class switching, which dominates all
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subsequent evolutionary radiations of the vertebrates. In terms of both functional immunity and Ig structure, Xenopus laevis (African clawed frog), which possess three distinct IgH isotypes, is the most extensively characterized of the amphibians. In addition to IgM, Xenopus possess another multimeric Ig molecule designated as IgX (Schwager and Hadji-Azimi, 1984; Hsu et al., 1985), which at the sequence level is most related to IgM (Amemiya et al., 1989) but is expressed in a thymus-independent fashion, predominantly in mucosal tissue (Mussmann et al., 1996). IgY represents a third isotype of amphibian Ig, which in terms of expression is related to both IgG and IgE in mammals and can be induced in T cell–dependent secretory responses, a characteristic of mammalian IgG. Despite the distant relationship of IgY and IgG, the cytosolic tails of their transmembrane regions are related, which potentially is of regulatory consequence (Mussmann et al., 1996). Ig class switching in Xenopus is a component of the secretory immune response and is associated with AT-rich switch boxes, which exhibit sequence changes (insertions, deletions, mutations) in regions of predicted secondary structure (Mussmann et al., 1996). Despite the different nature of the switch sequences in mammals and amphibians, it is likely that they function in an equivalent manner in somatic cells. An extensive variation in germline VH sequences, which is consistent with mammalian VH family diversity, has been demonstrated in essentially all lower vertebrate species examined thus far, with the exception of the cartilaginous fish (Haire et al., 1990). In Xenopus 11 different VH gene families and multiple JH families have been described, and comparable levels of variation have been described in other species (Hsu and Lewis, this volume). There is little basis to believe that limitations in genomic complexity per se reduce the overall level of somatic variation in heavy chain genes. Although reptiles factored in the earliest descriptions of Ig gene structure in lower vertebrates, they have not been subject to comprehensive analysis at the Ig gene level. Nevertheless, it is possible to draw several general conclusions regarding the diversity of Ig genes: 1) evidence has been presented for extensively diversified VH genes in both Caiman crocodylus (caiman) and Chelydra serpentina (snapping turtle) (Litman et al., 1999); 2) the Ig genes in snapping turtle map to separate chromosomal linkage groups by in situ hybridization; and 3) the extraordinary complexity of Ig genes seen in reptiles is lost in the avians, which typically utilize single functional heavy and light chain V genes to effect extensive diversity through a gene conversion mechanism (Knight, this volume). Several aspects of Ig gene complexity in reptiles make them particularly significant models in terms of understanding the role of large families of diversified germline genes in the generation of immunological diversity. The dichotomy presented by the complexity of Ig genes in basal reptiles versus the absence of
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germline variation of functional V elements in both IgH and IgL genes in their evolutionary descendents—the avians— provides a striking example of the fluidity of Ig gene organization and diversification.
LIGHT CHAIN GENES: DIVERSE STRUCTURES AND ORGANIZATION IN LOWER VERTEBRATES The organization of light chain genes in a number of different lower vertebrate species is illustrated in Figure 27.3. Light chain genes in cartilaginous fish and teleosts are of the cluster type; those of Acipenser baeri (sturgeon), a chondrostean fish, are in a translocon arrangement. Defining which organizational type was ancestral, which was derived, and how gene organization could change so dramatically in the lineage that includes the modern representatives of bony fish underscores the complications in classifying light chain genes in lower vertebrates (Bengten et al., 2000), although there are recognizable common features of certain light chains. Three types of light chains (types I, II, and III) have been detected in sharks and skates, and possibly are present in all cartilaginous fish (Rast et al., 1994). Contraction and expansion of the genes has been noted in each group, and the relative abundance seems to correlate with the level of expression—for example, in nurse sharks, the type III genes are most abundant and predominate in the serum (Sledge et al., 1974; Greenberg et al., 1993). The three light chain types are encoded by cluster-type genes. The type II genes are entirely germline-joined in all species studied to date (two species of skate, nurse shark, sandbar shark, Hydrolagus colliei [ratfish]) (Rast et al., 1994; Hohman et al., 1993; Anderson et al., 1995; Lee et al., 2002). The limited number of type II light chain genes in nurse sharks permitted a comprehensive study of somatic mutation and led to the identification of an unusual hypermutation mechanism (Lee et al., 2002). An expansion of light chain genes appears to have occurred in skate. Exhaustive analyses have established that the type I genes are completely germline-joined (Anderson et al., 1995), whereas in horned shark, type I genes are all in the split configuration (Rast et al., 1994). In nurse sharks, most of the type I and type III genes are split, but germlinejoined forms also have been identified. Phylogenetic analysis established that one of the type III genes appears to have been germline-joined within ~10 million years, confirming the hypothesis proposed earlier that germline-joining is an evolutionarily derived characteristic, due to RAG activity in the germline (Lee et al., 1999). Although it has been difficult to assign a k- or l-like character to type I and type II light chains, phylogenetic analyses place the type III light chain V regions in the kappa lineage (Rast et al., 1994; Greenberg et al., 1993; Hsu, Steiner, 1992). Whereas the VL
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FIGURE 27.3 Schematic showing the organization of light chain genes in the various vertebrate taxon. The transcriptional organization of all gene segments is depicted in the conventional sense (L Æ R = 5¢ Æ 3¢) except where arrows below the V segments indicate that they are in opposite transcriptional orientation to the J and C segments. Genes that are in cluster organization are indicated by parentheses, ( )n. Both type I and type II genes in elasmobranchs may or may not be joined in the germline. Data are taken from Figure 27.7 of the review by (Bengten et al., 2000; Reynaud et al., 1987; Zezza et al., 1992; Ghaffari and Labb, 1993; Daggfeldt et al., 1993; Timmusk et al., 2000; Lundqvist et al., 1996; Rast et al., 1994; Shamblott, Litman, 1989).
regions in cartilaginous fish tend to cluster in phylogenetic trees with VL regions from other vertebrates (especially the type III), the CL regions form an elasmobranch “supercluster,” emphasizing the independent evolution of the VL and CL domains in these species (Rast et al., 1994; Greenberg et al., 1993; Sitnikova and Nei, 1998). Several possibilities can be invoked to explain the three different types of light chains in the elasmobranchs: 1) association of heavy chains with the different light chains permits alternative conformations that are selectively advantageous in antigen recognition; 2) light chain preferences (or exclusion) may exist for the IgX or IgM heavy chains— monoclonal antibodies to different light chain types
(presently unidentified) immunoprecipitate IgM but not IgW heavy chains (Greenberg et al., 1996); or 3) some of the fixed specificities encoded in the germline-joined (especially the type II) genes may be particularly effective in recognizing particular epitopes on pathogens. It is possible that the cluster-type organization allows for a level of functional plasticity that is not possible to achieve in the translocon configuration. Five different types of light chain genes, which can be categorized in three families, have been described in bony fish: F, G, L1, L2, and L3. Catfish F and G as well as cod and trout L1 are described as k-related (Bengten et al., 2000). Bony fish IgL loci are in the cluster-type
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organization (reviewed in Pilstrom, 2002), consisting of (VJ-C)n or (V-V-J-C)n (Ghaffari and Lobb, 1997a), in which the V segments are in opposite transcriptional orientation to the J and C segments. However, the L2 light chain type of trout also exhibits a cluster-type organization consisting of up to three V segments per cluster, in the same transcriptional orientation as J and C (Timmusk et al., 2000). A phylogenetic analysis of bony fish CL regions shows segregation into three clusters, specifically: (zebrafish 1, trout/cod 1, and catfish G), (trout and zebrafish 2), and (catfish F, zebrafish 3) (Haire et al., 2000b). One of the unusual features of light chain genes in bony fish relates to the large number of sterile transcripts found in zebrafish and in other species of bony fish (reviewed in Haire et al., 2000b). This sterile transcription may be related to unusual features of the enhancers described in bony fish IgL loci, as discussed below. Three classes of light chain genes, s, r, and l, are encoded at separate loci in Xenopus (Schwager et al., 1991; Zezza et al., 1992; Haire et al., 1996). The r locus contains at least four Vr and five Jr segments, as well as a single Cr gene, and is classified as k-like on the basis of both Vr sequences and overall organizational features of the r locus (Hiom et al., 1998; Zezza et al., 1992). The s-type light chain consists of multiple Vs1 and Vs2 as well as Cs1 and Cs2 segments (Schwager et al., 1991); both types of genes rearrange to their respective Js types. Expressed Vs sequences are highly conserved, and the diversity of s and r light chain genes in Xenopus may be markedly restricted. The l light chain type in Xenopus represents an unequivocal homolog of higher vertebrate l light chains. Six distinct polygenic and polymorphic Vl families, as well as two Jl segments and two distinct Cl exons, have been described (Haire et al., 1996). The germline variation of l genes argues against a repertoire restriction in rearrangement events involving these genes. The single light chain gene in avians can be assigned unequivocally as a l type (Reynaud et al., 1983). Whereas the systematics of vertebrate light chains is not critical to understanding either their diversity or function, it nevertheless has been suggested that the light chain genes should be grouped into three broad categories or lineages. Lineage A contains those genes with greatest similarity to the mammalian l, lineage B includes genes with a relationship to mammalian k, and lineage C genes lack a clear relationship to k or l and contain only light chain genes from poikilothermic species. It is possible that further genomic investigations may reveal conserved synteny; that is, that similar common genes will be found linked to Ig genes in different species, which could shed light on the relationships between disparate species across wide phylogenetic ranges. Such syntenic relationships exist for many gene families; however, Ig genes, which encode molecules that mediate responses to a vast range of foreign pathogens (many of which pose unique challenges to individual species), may lie
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outside standard modes of consideration in terms of the evolution of multigene families.
TRANSCRIPTIONAL CONTROL OF IG GENES IN LOWER VERTEBRATES Promoters The promoters of both IgH and IgL genes of mammals initially were observed to contain two highly conserved motifs: an octamer element (sometimes extended by 2 bp and termed a decamer motif) of consensus sequence ATG CAAAT (Parslow et al., 1984) and a TATA box. These two elements initially were thought to be sufficient to confer B cell–specific transcription in mice (Wirth et al., 1987). Subsequent functional studies have implicated additional motifs (including the pentadecamer and E-box sites) in the integrated function of mammalian Ig promoters (Artandi et al., 1994; Bemark and Leanderson, 1997; reviewed in Calame and Sen, this volume). Given the clear conservation of promoter structure in the mammals, it is of interest to determine whether this conservation extends to the promoters of Ig genes in lower vertebrates. With some notable exceptions, promoter regions of the Ig genes of lower vertebrates, including bony and cartilaginous fishes [collected in the IMGT database (LeFranc, 2001), http://imgt.cines.fr:8104/ home.html], a reptile (Litman et al., 1983), and the coelacanth (Amemiya et al., 1993) are very similar to those of mammals and birds. Furthermore, the conserved octamer and TATA box motifs in these species are often found in combination with additional sites such as E-box and k-Y motifs (Timmusk et al., 2002). Although relatively few studies have been conducted on promoter function in the transcriptional regulation of lower vertebrate Ig gene expression, the available evidence indicates that they function in a B cell–specific manner (Magor et al., 1994; Timmusk et al., 2002). Some notable exceptions can be found to the consensus structure that was discussed above for Ig promoters. Octamer sequences are not present in the IgM-type VH genes in shark or the IgW genes in the skate. The shark IgM VH promoter region contains a TCRb-type CRE element that closely resembles the promoter region found in mammalian TCRb genes. In marked contrast, the light chain genes and the hypermutating NAR genes (Greenberg et al., 1996) in cartilaginous fish possess an octamer motif in their promoter regions (Shamblott and Litman, 1989; Hohman et al., 1993; Rast et al., 1994). Defining an absolute role for octamer motifs in Ig promoters is complicated further by the consideration of Ig genes in the rainbow trout. Trout possess three classes of light chain, all of which are arranged in cluster organization. The promoter regions of L1 and L3 possess an octamer as
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well as TATA and E-box motifs. However, the L2 promoter lacks both typical octamer and TATA box motifs, but can be shown to function in a B cell–specific manner, and a function can be demonstrated for a k-Y box using deletion analyses (Timmusk et al., 2002).
Enhancers The efficient transcription of all vertebrate Ig loci that have been examined experimentally depends not only on promoters but also on B cell–specific transcriptional enhancers (Calame and Sen, this volume), which not only function in Ig gene transcription, but also factor in V(D)J recombination, class switching, and somatic hypermutation (Inlay et al., 2002; Serwe and Sablitzky, 1993; Cogne et al., 1994; Bachl et al., 1998). The mammalian IgH locus typically contains several enhancers; the most active enhancers are found in the JH to Cm intron (the Em enhancer) and 3¢ of the Cm gene cluster, downstream of the Ca exons (the hs1, 2; hs3a; hs3b; hs4 enhancers). Enhancers are found in the Jk to Ck intron, as well as downstream of the Ck gene (Inlay et al., 2002). Three enhancer regions are found downstream of the human Cl gene cluster (Asenbauer et al., 1999). Although the functional studies of enhancers in the Ig loci of lower vertebrates are restricted thus far to bony fish, some unexpected findings have been made in terms of enhancer location, structure, and function. Enhancers can function from any position within a reasonable distance of
the transcription start site. “Reasonable distance” for mammalian Ig enhancers can involve thousands (if not tens of thousands) of bp, which complicates their identification in lower vertebrate Ig genes, because consensus sequences that define transcription factor-binding motifs are short, common, and can be distributed throughout an Ig locus. The enhancer of the IgH locus of the catfish was identified in the intron separating the m and d genes, as well as overlapping the second transmembrane segment-encoding exon of the m gene (Magor et al., 1994). Not only does this enhancer (designated Em3¢) occur in an unexpected location, but it lacks a clear-cut homology with any of the known mammalian IgH enhancers. The Em3¢ enhancer is large (approximately 1.8 kb) and contains multiple octamer and E-box motifs of both consensus and variant sequence. Extensive mutational analyses have shown that the function of this enhancer depends on a single consensus mE5 site and multiple octamer motifs (Cioffi et al., 2001). The sites of transcriptional enhancers in the IgH locus of the mouse and the catfish are compared in Figure 27.4. Em, the best-studied mammalian IgH enhancer, is flanked by matrix-attachment regions and contains a tightly linked array of individual transcription factor–binding motifs, including E-boxes (mE1–5), an octamer site, and mA and mB motifs. Although these sites exhibit redundancy (Dang et al., 1998), and many of these motifs contribute significantly to function in mammals, the “core” of the Em enhancer has been defined as consisting of mA and mB sites flanking (in mouse), a mE3 site, or (in
Mouse L VH1
L VHn D1 D13 JH1 JH4 Cm Cd Cg3 Cg1 Cg2bCg2a Ce Ca
DQ52 Em Enhancer
Catfish VH(n)
D(n)
JH(9)
3’ Enhancers
Cm
Cd
Em 3’ FIGURE 27.4 Schematic comparison of the mouse and catfish immunoglobulin heavy chain gene loci indicating the positions of the enhancers (orange) in relation to the VH exons, D and JH segments, and CH exons. The mouse 3¢ enhancers are, from left to right: hs3a; hs1,2; hs3b; hs4. See color insert.
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humans) a CBF site (Tian et al., 1999; Nikolajczyk et al., 1997). The mA and mB sites bind members of the Ets family of transcription factors (Nelsen et al., 1993), whereas mE3 binds TFE3 and CBF binds core-binding factor/PEBP2. On the basis of these observations, the evolutionary relationship of the IgH enhancers of bony fish and mammals is obscure. However, it has been suggested that the enhancer in the primordial vertebrate IgH locus would have been 3¢ to the CH gene cluster, and that its translocation and duplications during vertebrate evolution have had important functional implications (Magor et al., 1999). For example, it is clear that the position of the Em3¢ enhancer of the catfish is incompatible with the evolutionary development of Ig class switching. Its location between the m and d genes would unavoidably result in the deletion of this enhancer during the chromosomal recombination involved in class switching, thus resulting in transcriptional inactivation of the classswitched locus. Additional information on enhancer activities in lower vertebrates comes from studies of the light chain genes in another bony fish, Gadus morhua (Atlantic cod) (Bengten et al., 2000). IgL in cod, like those in other bony fish, are organized in clusters. Six clusters were examined for enhancer activity, and three were found to possess regions downstream of the CL gene that would drive enhancer transcription. This high frequency of enhancers in the IgL gene clusters of a bony fish may account for the elevated frequency of noncoding light chain transcripts in the cod and other bony fish (Daggfeldt et al., 1993; Ghaffari and Lobb, 1993). Whereas the studies of enhancers in IgH and IgL loci of bony fish have yielded highly informative data concerning the diversity of the transcriptional control of Igs during the evolution of vertebrate immunity, our understanding of the evolution of enhancer function is far from complete. Studies on the transcriptional control of Ig genes in the elasmobranchs and in the poikilothermic tetrapods (amphibia and reptiles) are particularly relevant.
A UNIFYING HYPOTHESIS TO EXPLAIN THE ORIGINS OF THE ADAPTIVE IMMUNE RECEPTOR Phylogenetic analyses have suggested that a g/d TCR-like ancestor may have predated a/b TCR and IgH/IgL (Richards and Nelson, 2000), implying that direct antigen recognition, perhaps by a cell surface receptor, arose first in evolution. Hood and colleagues argue that phylogenetic analyses over such large evolutionary distances obscure true relationships among the antigen receptor genes (e.g., the relationships of the molecules in the phylogenetic trees impose multiple loss/gain of D segments in the different antigen receptor families), and suggest a model based upon extant genomic organizations (Glusman et al., 2001). An alternative phylo-
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genetic scenario has been proposed in which an ancestral chromosomal region possessed linked genes encoding both chains of an ancestral antigen receptor heterodimer, one having D segments and the other not. One en bloc duplication led to the divergence of Ig and TCR, and a second duplication gave rise to the a/b and g/d TCR gene complexes. The a and d TCR loci remain closely linked on human chromosome 14 in all vertebrates analyzed, and a pericentric inversion is suggested to have separated the TCR b and g loci, which are linked on human chromosome 7. D segments only would have emerged once, and the existence of inverted V elements in the TCR b and d loci is explained. This model does not predict whether Ig or TCR is older, but a rather simple view of receptor evolution is provided, consistent with the suggestion by Kasahara that genomewide or large en bloc duplications played a major role in the emergence of adaptive immunity (Kasahara, 1998).
IMMUNE MOLECULES IN JAWLESS VERTEBRATES The jawless vertebrates (Agnatha), which are comprised of two extant groups (hagfish and lampreys), are the sole surviving remnants of one of the two major vertebrate evolutionary radiations. Although the issue of their separate common versus single common ancestor origin is presently unresolved, they represent the most divergent forms relative to jawed vertebrates in terms of anatomical, biochemical, and physiological characteristics (Forey and Janvier, 1993; Takezaki et al., 2003). The nature of the immune mediators in the jawless vertebrates is generally acknowledged to represent one of the most significant unanswered questions in evolutionary immunology. Agnathans lack both spleen and thymus, which are found in all jawed vertebrate groups. The inducible immune response in lamprey to both bacteria and an erythrocyte antigen is associated with a highly specific molecule(s) of a labile, lectin-like character (Rast et al., 1995; Litman et al., 1993). Humoral immune reactivity to bacteriophage similarly is associated with a molecule exhibiting a labile character (Marchalonis and Edelman, 1968). Owing to the “instability” of immune mediators in lamprey, recent efforts at detection have been directed at identifying homologs of Ig, TCR, MHC I and II, and RAG genes (at the nucleotide level). To date, none of these efforts have been successful. However, the approaches that have been employed would require ~60% overall nucleotide identity between a higher vertebrate probe and target sequence for successful library screening or two short regions of near absolute nucleotide identity with higher vertebrate Ig for a PCR-based approach, such as that used to identify TCR genes in cartilaginous fish (Rast et al., 1997). The failure to identify the immune receptors and related gene products by conventional methods
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could relate to their absence or to either qualitative or quantitative complications; for example, low levels of expression (and inhomogeneous cell populations) and/or transient expression of immune effectors at different stages of development or immune stimulation. Nevertheless, the negative results have been interpreted to indicate that the adaptive immune system was acquired after the divergence of jawless and jawed vertebrate forms. An alternative approach to the problem of receptor identification has used large-scale cDNA sequencing. Recent reports describe the results of sequence analysis of ~8,000 ESTs from lymphoid-like cells, defined as protolymphocyte, from the intestine of the sea lamprey (Mayer et al., 2002; Uinuk-Ool et al., 2002). Although a number of transcripts were identified that are homologous to genes associated with higher vertebrate lymphocytes—CD45, CD9/CD3e, BCAP, and CD98—known effectors of immune function or molecules that are integral to either somatic diversity or antigen presentation were not identified. Notably, CD45 also has been detected in the hagfish (Nagata et al., 2002). The failure to identify lymphoid transcription factors and IgSF-related transcripts in the large-scale screens was surprising, given their broad distribution and relating high expression on the surfaces lymphoid cells of higher vertebrates. In other studies, a lamprey ortholog of Spi C, which is involved in lymphocyte differentiation in higher vertebrates has been described (Shintani et al., 2000; Anderson et al., 2000), as has Ikaros, another transcription factor associated with lymphocyte development (Haire et al., 2000a; Mayer et al., 2002), and a homolog of GATA2/3 (J. Cannon, R. Haire and G. Litman, unpublished observation), which could factor in lymphocyte development. A direct cloning approach that requires sequence identity over only a single short peptide region (three amino acids) has been described recently (Cannon et al., 2002). Using this approach, several V region–containing cDNAs have been recovered from lamprey (J. Cannon, R. Haire and G. Litman, unpublished observation). Whereas the identification of possible immune receptors as well as lymphoid-related transcription factors are encouraging, these findings must be interpreted carefully, bearing in mind that structurally homologous molecules may not necessarily represent functional equivalents. The broad evolutionary theme of the utilization of similar genetic material for markedly different purposes is being reinforced steadily with growing amounts of genomic sequence information. It is important to recognize that clues to the existence of alternative mechanisms of adaptive immune function also could come from the identification of molecules such as AID or other molecules that function in the somatic component of adaptive immune diversity. Immune receptors in jawless vertebrates could be membrane-bound and exhibit diverse innate specificities, such as seen in the nonrearranging diversified families of NITRs in bony fish (Figure 27.5). The ectodomains of
FIGURE 27.5 General architectural features of a TCR, a novel immunetype receptor (NITR) and inhibitory killer-cell immunoglobulin-like receptor (KIR). V = variable region, C1 = type 1 constant region, C2 = type 2 constant region, V/C2 = V-type C2 region (equivalent to I-type), TM = transmembrane, ITIM = immunotyrosine-based inhibitory motif, J = joining region motif containing FGXG, GXG = glycine-X-glycine (a core feature of J regions), small ovals represent cysteine residues and a positive intramembrane charge is indicated.
NITRs are V regions that can be classified in up to thirteen distinct families in pufferfish; further diversification is evident within families. The majority of NITRs described to date encode immunoreceptor tyrosine-based inhibitory motifs (ITIMs) in their cytoplasmic tails but putative activating forms also have been identified. Although their ligand specificities are not understood, NITRs resemble killer-cell immunoglobulin-like receptors (KIRs) in overall organization and represent the only gene family other than Ig and TCR in which diversified V regions potentially account for recognition specificity (Litman et al., 2001). Finally, it is critical to recognize that antigen binding receptor gene homologs in jawless vertebrates may take an entirely different form, thus requiring that their identification be initiated at the functional level.
PROTOCHORDATES: DIFFERENT CONTEXTS FOR DIVERSIFIED V REGIONS The possibility exists that identification of “immunerelated,” diversified V region–containing IgSF members may be achieved in basal protochordates, in which “potential precursors” might be detected more easily than in the more derived higher vertebrates. The draft genome of a urochordate, the sea squirt (Ciona intestinalis), an ascidian, was reported recently (Dehal et al., 2002). Whereas several genes involved in cell signaling and development, as well as some
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genes that are known to be involved in innate function, were identified, it was not possible to identify genes that are related to those involved in adaptive immunity. Recently, attention has been focused on Branchiostoma floridae (amphioxus), a cephalochordate, because: 1) morphologically and developmentally it closely resembles the last common ancestor of all vertebrates, 2) it has become a significant model in developmental biology, and 3) the complete sequencing of its genome has been proposed. Multiple families of novel, presumably bifunctional genes (VCBPs) consisting of two diversified V-like regions and a chitinbinding domain have been identified in this species. The patterns of diversification of the VCBPs and their tissue-specific expression in the gut have been described (Cannon et al., 2002). The presence of chitin-binding domains at the C-termini of all known VCBPs raises new questions about the broad role of bifunctional molecules in protochordates and invertebrate immunity, and how interactions with chitin (widely distributed in arthopods, fungi, bacteria, and other organisms) may relate to immune function. The overall features of VCBPs strongly suggest that they represent immune-type molecules. Specifically, they are highly polymorphic and contain regionalized substitution hotspots that may be somatically diversified. In that somatic mutation is not necessarily linked to segmental rearrangement, this mechanism of diversification may have preceded the origins of Ig and TCR-like genes in evolution (Du Pasquier et al., 1998). The discovery of VCBPs and the recognition that other IgSF members may function in innate recognition in other invertebrates has opened new avenues to investigate the origins of immune recognition. Although the genes may turn out to not represent true orthologs of IgSF genes found in jawless and jawed vertebrates, they unequivocally establish the presence of highly diversified families of protochordate V genes that likely function in some aspect of immune recognition in this species.
Acknowledgments We would like to thank Barbara Pryor for her editorial assistance. GWL, MFF, and GWW are all supported by grants from the National Institutes of Health.
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28 Immunoglobulin Genes and Generation of Antibody Repertoires in Higher Vertebrates: A Key Role for GALT DENNIS LANNING,1 BARBARA A. OSBORNE,2 AND KATHERINE L. KNIGHT1 1
Department of Microbiology and Immunology, Stritch School of Medicine, Loyola University Chicago, Maywood, Illinois, USA 2 Department of Veterinary and Animal Science, University of Massachusetts, Amherst, Massachusetts, USA
In higher vertebrates, immunoglobulin (Ig) gene rearrangements, B-cell development, and the generation of the primary antibody repertoire occur in a variety of ways. In mouse and human, they occur mostly in bone marrow. In other higher vertebrates, Ig gene rearrangements and B-cell development can occur in tissues such as spleen and yolk sac. Many of these species, including chicken, rabbit, cattle, and sheep, also use gut-associated lymphoid tissues (GALT) for development of the primary antibody repertoire. In these species, B-lineage cells migrate to GALT, where B-cell expansion and somatic diversification of Ig genes occurs. Generally in these species, B lymphopoiesis occurs early in development and does not continue throughout life. Whereas mouse and human use combinatorial joining of multiple V, D, and J gene segments to generate a large antibody repertoire, most members of the GALT-species utilize only a limited number of Ig gene segments in V(D)J gene rearrangements. Further, unlike mouse and human, which possess VH genes from all three VH groups, the VH genes of the GALT-species generally belong to only one VH group, either group B or C (Sitnikova and Su, 1998). Likewise, the Vk and Vl genes of the GALT-species generally belong to only one group. Another difference between members of the GALT-species and mouse and human is that some of the GALT-species utilize gene conversion as well as hypermutation to somatically diversify their Ig genes. We review Bcell development and generation of the antibody repertoire in higher vertebrates, excluding mouse and human, highlighting the importance of GALT in these processes.
typical GALT species to which others are compared. Of all avian species, we discuss primarily the chicken.
Organization of Chicken Ig Genes Although the chicken heavy and light chain loci contain multiple V, (D), and a single J gene segments, the loci differ from those of other species in that they contain only one functional V and J gene segment. The sole functional light chain V gene segment, Vl1, lies 1.8 kb upstream of a single Jl gene segment and about 3.6 kb upstream of a single Cl locus. Twenty-five V pseudogenes lie within 19 kb upstream of Vl1. Most of these pseudogenes lack recombination signals, and all of them lack upstream promoter and leader sequences. The pseudogenes lie in both transcriptional orientations with respect to Vl1 (Reynaud et al., 1985, 1987). The only functional heavy chain V gene segment, VH1, lies about 30 kb upstream of the Cm locus and 15 kb upstream of a single JH gene segment (Reynaud et al., 1989, 1991). Eighty to one-hundred VH pseudogenes spaced an average of 0.8 kb apart lie within a 60- to 80-kb region upstream of VH1. The VH pseudogenes lack recombination signals, as well as promoter and leader sequences, and the 3¢ end is fused to a D-like segment. As discussed below, this structural feature facilitates VDJ gene diversification. As in the Vl locus, VH pseudogenes lying in opposite transcriptional orientation with respect to VH1 are interspersed among VH pseudogenes sharing the same orientation as VH1. Although the heavy chain locus contains only a single functional VH and a single JH gene segment, it contains sixteen functional D gene segments. Fifteen of the D gene segments are, however, highly homologous, with several encoding the same amino acids (Reynaud et al., 1991). All of the D gene segments exhibit a strong bias for usage in
AVIANS Of all species that use GALT to develop the antibody repertoire, the chicken is the best studied and is the proto-
Molecular Biology of B Cells
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reading frame 1, which encodes primarily hydrophobic and aromatic amino acid residues, particularly Gly, Ser, Ala, Tyr, and Cys. A similar D reading frame bias has been reported in other species, suggesting that the amino acid composition of the D region is important for light chain pairing or selection of developing B cells. We do not yet understand why multiple, highly homologous D gene segments have been maintained in the chicken heavy chain locus. Because they are utilized with strong reading frame bias, their contribution to antibody repertoire diversity can be only minor. Reynaud et al. (1991) suggested that their major contribution to heavy chain diversity might be through the formation of D–D junctions. They found such junctions in 25% of DJ gene rearrangements and in 20% of rearranged but preselected VDJ genes in bursal B cells. The formation of D–D junctions is unexpected because chicken D gene segments are flanked at both ends by 12-bp recombination signal sequences; D–D recombination therefore appears to violate the 12/23 base pair rule. D–D junction formation expands the range of potential heavy chain CDR3 lengths to fifteen to thirty amino acids. There appears to be no requirement in the chicken for matching CDR3 lengths during pairing of the heavy chain with the light chain, in which CDR3 varies little in length (McCormack et al., 1989c; Reynaud et al., 1991).
Ig Gene Rearrangement DHJH gene rearrangements are first detectable by PCR analysis in the yolk sac at day 5 to 6 of embryogenesis and at other hematopoietic sites a few days later (Reynaud et al., 1992). The first fully rearranged V(D)J genes are detectable at embryonic day 8 to 9 in the spleen, blood, and yolk sac, and V(D)J genes are detectable in the bursa 1 to 2 days later (Reynaud et al., 1992). Ig gene rearrangement can, however, occur after B-cell precursors migrate to the bursa. This was demonstrated by the isolation of retrovirally transformed Bcell precursors containing only DJ rearrangements from the bursa at embryonic day 12 (Banatar et al., 1992). The detection of B-cell recombination excision circles (BRECs) in the embryonic bursa is also consistent with ongoing Ig gene rearrangement (McCormack et al., 1989b; Reynaud et al., 1992). The bursa, however, is not required for the induction or completion of Ig gene rearrangement, as B cells expressing surface IgM were produced in chickens from which the bursa was removed early in embryonic development (60 hours of incubation) (Jalkanen et al., 1983). The simultaneous appearance of the first detectable VDJ and VJ gene rearrangements at embryonic day 8 to 9 shows that the chicken heavy and light chain loci are not rearranged in the sequential manner characteristic of murine B cells (Reynaud et al., 1992). In murine B cells, VDJ rearrangement generally precedes VJ rearrangement by 2 to 3 days (Osmond, 1991). In contrast, analysis of Ig gene rearrange-
ments in individual retrovirally transformed chicken B cells from embryonic day 12 bursa demonstrated that, although heavy chain DJ gene rearrangement appears to occur first, it can be followed by either VJ or VDJ gene rearrangement (Banatar et al., 1992). Chicken Ig genes therefore appear to be rearranged stochastically. Consequently, rearrangement of the chicken light chain locus is not regulated by the expression of a productively rearranged heavy chain gene. Unlike human and murine B cells, chicken B cells therefore probably do not express a pre-B cell receptor (pre-BCR). Unlike murine B cells, few chicken B cells contain nonproductively rearranged Ig genes. In more than 90% of chicken B cells, the unexpressed light chain locus is in germline configuration, and the unexpressed heavy chain locus generally contains a DJ gene rearrangement. The frequency of productive Ig gene rearrangement, however, appears to be no higher in chicken B cells than in those of mice (McCormack et al., 1989b; Reynaud et al., 1991). Allelic exclusion in chicken B cells is therefore not mediated by the expression of a productively rearranged allele but by an alternative mechanism that is not currently understood. This mechanism probably evolved to limit the possibility of out-of-frame rearrangements being rendered productive by gene conversion. It has been proposed that the restriction of Ig gene rearrangement to a brief period during embryonic development imposes a time constraint that limits the probability that both alleles of the heavy and light chain loci will be rearranged. Lauster et al. (1993) demonstrated that the V-J intron contains a negative regulatory element with strong transcriptional silencing activity. Transient inactivation of this silencer after DJ gene rearrangement would provide the recombination machinery with a brief period of access to the locus, with rearrangement presumably restoring silencer activity at the other allele. Such a regulatory mechanism would make it unnecessary to coordinate downregulation of the RAG genes with the expression of a functional Ig molecule. Expression of the RAG genes, in fact, continues in the bursa, where RAG-2 is expressed at high levels and RAG-1 at much lower levels (Carlson et al., 1991; Reynaud et al., 1992). It is nonetheless not known how time-restricted regulation of Ig gene rearrangement could allow the efficient rearrangement of one heavy and one light chain locus while keeping the frequency of double rearrangements at both loci low. Limited B Lymphopoiesis Although B lymphopoiesis and Ig gene rearrangement continue throughout life in mice and humans, these processes occur only during embryonic development in the chicken. In allotype suppression experiments in which chicken embryos heterozygous for the IgM allotype were treated with anti-allotype antibodies, severe long-term deletion of IgM allotype-expressing B cells was induced, thus
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demonstrating that B lymphopoiesis does not continue after hatch (Ratcliffe and Ivanyi, 1981). Similarly, BRECs produced during V(D)J gene rearrangement were readily detectable in the spleen and bursa during embryonic development but were no longer detectable shortly before hatch (McCormack et al., 1989c; Reynaud et al., 1992). Restriction of B lymphopoiesis to an early stage of development implies that long-lived and/or self-renewing B cells maintain the adult chicken B-cell compartment.
Generation of Antibody Diversity In mice and humans, combinatorial rearrangement of multiple V, (D), and J gene segments generates an enormous number of unique Ig heavy and light chains. In contrast, all chicken B cells utilize the same Vl1, Jl, VH1, and JH gene segments during rearrangement of the light and heavy chain loci (Reynaud et al., 1985, 1989). Although several functional D gene segments are present in the chicken heavy chain locus, their contribution to VDJ gene diversity is relatively minor, for reasons discussed above. In addition, little junctional diversity is generated during joining of V, D, and J gene segments, further restricting potential heavy chain diversity (McCormack et al., 1989c). In contrast to humans and mice, therefore, little antibody diversity is generated in the chicken during Ig gene rearrangement. Instead of generating antibody diversity, Ig gene rearrangement in chickens generates substrates for gene conversion, the primary mechanism by which Ig genes are diversified. Gene conversion transfers tracts of nucleotides from upstream V pseudogenes into equivalent regions of the rearranged VJ and VDJ genes (Reynaud et al., 1987, 1989; Thompson and Neiman, 1987). This process introduces novel nucleotide sequences into the rearranged VJ and VDJ genes while leaving the donor V pseudogenes unchanged (Carlson et al., 1990). Although Vl pseudogenes share a high degree of homology with Vl1, they differ in nucleotide sequence primarily in and around the CDRs, which ultimately contribute to the antigen-binding site of the antibody molecule (Reynaud et al., 1987). A similar concentration of nucleotide differences in the CDRs occurs among VH pseudogenes. VH pseudogenes are homologous to fused VH1 and D gene segments, and gene conversion events can introduce nucleotide changes into the D region, as well as into VH1, in rearranged VDJ genes (Reynaud et al., 1989). Pseudogene usage is influenced by several factors, including sequence homology, transcriptional orientation, and proximity to the rearranged V-gene segment (McCormack and Thompson, 1990). Each rearranged VJ and VDJ gene can undergo several gene conversion events involving a variety of pseudogene donors, resulting in the generation of considerable heavy and light chain diversity. In another avian species, the Muscovy duck, rearrangement of several different VL gene segments, in addition to gene conversion,
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contribute to Ig light chain diversity. Hence, not all birds are restricted to gene conversion–mediated Ig gene diversification (McCormack et al., 1989a). The Bursa of Fabricius Diversification of chicken Ig genes occurs during B-cell development in the bursa of Fabricius, a lymphoid organ associated with the hindgut (Figure 28.1). Between days 8 and 15 of embryogenesis, the bursa is colonized by a single wave of B-cell precursors (Houssaint et al., 1976). B-cell precursors first populate mesenchymal tissue within the bursa, and 20,000 to 40,000 of these precursors subsequently migrate across the bursal epithelial basement membrane and begin proliferating in epithelial buds that ultimately develop into bursal follicles (Pink et al., 1985; Ratcliffe et al., 1986). During proliferation, these cells accumulate somatic gene conversion events at their Ig loci, resulting in the generation of a diversified primary antibody repertoire. Shortly before hatch, B cells begin emigrating from the bursa into the periphery, and emigration continues until the bursa involutes several months after hatch. An important function of the bursa during embryonic development is to selectively expand B-cell precursors that have productively rearranged their Ig genes and are thus able to express surface IgM. Although few V(D)J gene rearrangements isolated from the embryonic bursa at days 10 to 13 are productive, their incidence increases markedly over subsequent days so that more than 90% of isolated V(D)J gene rearrangements are productive by day 18 (McCormack et al., 1989b; Reynaud et al., 1991). Critical B lineage developmental checkpoints include precursor transit across the basement membrane and proliferation within epithelial buds (Reynaud et al., 1992). Both are dependent on surface IgM expression. Because B-cell precursors encounter these checkpoints before diversifying their Ig genes, which is also when they express nearly identical B cell receptors (BCR),
FIGURE 28.1 Schematic diagram of development of the primary and secondary antibody repertoires in chicken.
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it was speculated that the specificity encoded by the prediversified BCR plays a critical role in precursor selection, perhaps by recognizing endogenous ligands within the bursa (McCormack et al., 1989b; Reynaud et al., 1989; 1991). Sayegh et al. (1999a), however, demonstrated that B-cell precursors expressing a truncated BCR that lacked variable region domains colonized and rapidly proliferated within bursal epithelial buds. Furthermore, rearranged light chains in these cells underwent gene conversion-mediated diversification at levels indistinguishable from those in normal Bcell precursors (Sayegh et al., 1999b). Therefore, although colonization of, and proliferation within, epithelial buds are dependent on B-precursor surface IgM expression, the specificity encoded by the prediversified BCR plays no role in these processes. After hatch, the bursa functions as a site of contact between exogenous antigens transported from the gut and lymphocytes developing within the bursal follicles (Sorvari et al., 1975). It has therefore been suggested that exogenous antigens might influence the maturation or emigration of developing bursal B cells (Paramithiotis and Ratcliffe, 1993). In support of this idea, Sayegh and Ratcliffe (2000) found that B-cell precursors expressing a truncated BCR, although capable of colonizing and proliferating within bursal follicles, underwent apoptotic death after hatch and failed to emigrate to the periphery. This observation suggests that interaction with exogenous antigen within the bursa represents a developmental checkpoint governing the maturation and/or emigration of bursal B cells expressing diversified BCRs. After emigrating to the periphery, mature B cells undergo additional V(D)J gene diversification in secondary lymphoid tissues during antigen-specific responses (Arakawa et al., 1996). During these responses both gene conversion and somatic hypermutation contribute to V(D)J gene diversification.
of the duck heavy chain locus containing the Cm, Ca, and Cn genes and definitively established that no Cd gene or Cd remnant resides in this region. The Ig heavy chain constant region genes of both chicken and duck are organized in the order Cm, Ca, Cn (Lundqvist et al., 2001; Zhao et al., 2000). In both species, the Ca gene is in inverted transcriptional orientation with respect to the Cm and Cn genes. Class switch to IgA expression therefore requires the inversion of an 18-kb (in chicken) or 27-kb (in duck) genomic region containing both the Cm and Ca genes. Although the inversion of the Ca gene must have occurred in a common ancestor of the duck and chicken, it is not known whether all birds share this feature. The chicken and duck constant region genes are considerably longer than their mammalian homologs, primarily because they contain exceptionally long introns (Kitao et al., 1996; Lundqvist et al., 2001; Magor et al., 1994). Interestingly, however, in the chicken but not in the duck, the Cn gene lacks an intron separating the exons encoding the CH1 and CH2 domains of IgY (Zhao et al., 2000). Putative class switch repetitive sequences have been identified in the 5¢ introns of the Cm and Cn genes of the duck and chicken, and the Ca gene of the duck (Kitao et al., 2000; Lundqvist et al., 2001). Summary Although the processes underlying the generation of the primary antibody repertoire are better understood in the chicken than in other GALT species, many unanswered questions remain. The mechanisms controlling Ig gene rearrangement and allelic exclusion, for example, and the selective processes operating at different stages of B-cell development are not currently understood. It has become increasingly apparent, however, that the strategy chickens use to generate their primary antibody repertoire is surprisingly widespread among mammalian species.
CH Genes Avian species express three immunoglobulin classes, IgM, IgA, and IgY (Kincade and Cooper, 1973; Magor et al., 1998; Mansikka, 1992; Parvari et al., 1988). Although sometimes referred to as IgG, IgY shares homology with both mammalian IgG and IgE and is thought to have descended from the evolutionary precursor of these two classes (Parvari et al., 1988; Warr et al., 1995). Two isoforms of IgY, termed IgY and IgY(DFc), have been described in the duck (Magor et al., 1994; Magor et al., 1992). Both isoforms are encoded by the same IgY gene, with IgY(DFc) resulting from the use of a unique terminal exon located between the exons encoding the CH2 and CH3 domains. IgY(DFc) therefore lacks the functionally important FC region required for secondary effector functions (Peter, 1995). An avian homolog of IgD has not been identified. Lundqvist et al. (2001) recently sequenced a 48.8-kb region
LAGOMORPHS The order Lagomorpha includes two families, Leporidae and Ochotonidae. Most lagomorphs are members of the Leporidae family, which includes hares, domestic rabbits, cottontail rabbits, old world (or wild) rabbits, and jack rabbits. Most studies of lagomorph Ig genes have been performed with domestic rabbits, which is the focus of our review.
VH, D, and JH Gene Segments The VH locus of rabbit spans over 750 kb and appears to contain approximately one hundred VH, fourteen DH, and six JH gene segments (Becker et al., 1989; Chen et al., 1996; Currier et al., 1988; Gallarda et al., 1985). As in other
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mammals, the D gene segments are located within the 70kb span between the 3¢-most VH gene segment, VH1, and the JH gene segments. Both functional and nonfunctional VH genes are found within the VH locus, and all of them belong to the VH3 family, or group C, of VH genes (Sitnikova and Su, 1998; Tutter and Riblet, 1989).
because they increase the stability of the V-region domain or because they may be important for antigen-independent interaction with intestinal microbes and subsequent somatic diversification of Ig genes.
VH, D, and JH Gene Segment Usage
Kappa
The D-proximal VH gene, VH1, is used in 80 to 90% of VDJ gene rearrangements (Knight and Becker, 1990). The VH1 gene of the three heavy chain haplotypes, a1, a2, and a3, encodes the allelically inherited VHa allotypes, a1, a2, and a3. Approximately 20% of serum Ig molecules do not have the VHa allotype, and they are designated VHa-negative. These VHa-negative regions are encoded by multiple VH genes, including VHx, VHy, and VHz (Friedman et al., 1994). Although the preferential usage of VH1 is likely due to its preferential rearrangement during fetal development (Tunyaplin and Knight, 1995), the molecular basis for this is not known. The preferential rearrangement is not likely due to a preferred recombination signal sequence (RSS) because the RSS flanking VH1 is identical or highly similar to those flanking most functional germline VH genes analyzed. VH1 may be preferentially used because its close proximity to DJ gene rearrangements makes it most accessible to the recombination machinery (Yancopoulos et al., 1984). D and JH gene segments are also preferentially used in V(D)J gene rearrangements. Although both JH2 and JH4 are found in DJ gene rearrangements, over 90% of the VDJ genes utilize JH 4. Four of the eleven D gene segments, D1, D2a, D2b, and D3 together account for nearly 80% of the D regions of VDJ genes (Becker et al., 1990; Chen et al., 1993; Friedman et al., 1994). This D and JH gene segment usage is similar in both productive and nonproductive VDJ gene rearrangements, indicating that preferential usage results from preferential rearrangement rather than from selective processes.
Unlike most mammals, rabbits have two Ck germline genes, Ck1 and Ck2, which appear to be separated by 1 Mb (Benammar and Cazenave, 1982; Emorine and Max, 1983; Hole et al., 1991). Each Ck gene is associated with a separate 5¢ cluster of Jk gene segments. Of the five Jk gene segments associated with Ck1, Jk2 is the only functional one in most rabbits. It is not known whether each Ck gene is also associated with its own set of Vk gene segments (Emorine et al., 1983a; Emorine and Max, 1983; Heidmann and Rougeon, 1982; 1983). In normal rabbits, 80 to 90% of Ig molecules have k light chains and nearly all of them are encoded by Ck1. Ck2 is rarely expressed, except in rabbits with the b9 k-chain allotype, in wild rabbits, and in mutant Basilea rabbits (Kelus and Weiss, 1977; Mage et al., 1984). In Basilea rabbits, a mutation in the mRNA splice site of Ck1 likely limits expression of Ck1 light chains (Lamoyi and Mage, 1985). Although only a few Vk germline gene segments have been cloned from recombinant libraries (Ayadi et al., 1990; Heidmann and Rougeon, 1984; Lieberman et al., 1984), Sehgal et al. (1999) PCR amplified and sequenced Vk gene segments from kidney DNA and VJk genes from a cDNA library of bone marrow from 1-day-old rabbits, before the Ig genes are somatically diversified. Thirty-nine Vk gene segments were identified, and twenty-eight of them were found expressed as RNA.
Evolutionary Conservation of VHa Polymorphisms Lagomorph Ig is of particular evolutionary interest because the same a1, a2, and a3 VH allotypes are conserved in many different species, including domestic rabbits, wild rabbits, and snowshoe hares (Van der Loo et al., 1996; Su and Nei, 1999). Su and Nei (1999) estimated that these allotypic polymorphisms have been conserved for nearly 50 million years, suggesting that they provided an evolutionary advantage to lagomorphs. In rabbit, the VHa allotypes are encoded in framework region (FR)1 and FR3 of the predominantly utilized VH gene segment, VH1 (Knight and Becker, 1990). Although the molecular basis for the maintenance of these polymorphisms through evolution is not known, we suggest that they have been maintained either
Kappa and Lambda Light Chain Gene Segments
Lambda Lambda chains generally comprise less than 10% of total rabbit Ig light chains (Dray et al., 1963). Relatively little is known about either the proteins or the genes that encode them. By Southern blot analysis, the germline appears to contain approximately eight Cl genes, six of which are cloned and designated Cl1–Cl6 (Duvoisin et al., 1986; 1988). Of these, only two, Cl5 and Cl6, appear to be expressed. By Southern blot analysis, five Vll genes have been found, and at least two of them appear functional (Hayzer et al., 1987; Hayzer and Jaton, 1989a,b).
B-Cell Development and Generation of Antibody Diversity A model of B-cell development and generation of the primary and secondary antibody repertoires in rabbit is
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shown in Figure 28.2. B lymphopoiesis occurs in fetal liver and bone marrow, with DJ and VDJ gene rearrangements beginning around 14 days’ gestation (Tunyaplin and Knight, 1995). Because almost no B lymphopoiesis was detected in bone marrow in adult rabbits (Crane et al., 1996), Jasper et al., 2003) examined bone marrow of rabbits of different ages to determine the time at which B lymphopoiesis wanes. As determined by immunofluorescence and PCR, the number of precursor B cells and the number of V to DJ recombination excision circles (BRECs) steadily decreased beginning at 3 weeks of age. By 16 weeks, no precursor B cells or BRECs were found, indicating that B lymphopoiesis terminates several weeks after birth. Thus, it appears that the B cells generated early in life are those from which all antibody specificities are derived throughout life. Presumably rabbit B cells have a long lifespan and/or are self-renewing. The primary heavy chain antibody repertoire is generated through preferential rearrangement of Ig gene segments followed by somatic diversification of VDJ genes. Because VH1, JH4, and a small number of D-gene segments are preferentially rearranged, combinatorial rearrangement of IgH gene segments does not contribute widely to the B-cell repertoire. The VDJ genes in peripheral B cells remain undiversified for approximately the first 3 weeks of life. However, by 6 to 8 weeks of age, essentially all VDJ genes have undergone somatic diversification. The resulting repertoire is often referred to as the primary or pre-immune antibody heavy chain repertoire (Crane et al., 1996). In contrast to the limited VH gene usage in VDJ gene rearrangements, at least thirty Vk gene segments are rearranged early in development. This more extensive use of Vk gene segments in VJk rearrangements may in part compensate for the limited number of VDJ gene rearrangements and give rise to a moderately large primary antibody repertoire prior to somatic diversification (Sehgal et al., 2000).
FIGURE 28.2 Schematic diagram of development of the primary and secondary antibody repertoires in rabbit.
VDJ and VJk genes somatically diversify by two mechanisms—gene conversion-like recombination and somatic hypermutation (Becker and Knight, 1990; Lanning and Knight, 1997; Sehgal et al., 2000). The term “gene conversion-like” is used because definitive evidence that VDJ genes diversify by gene conversion requires that the donor VH gene be identified and analyzed in individual cells to establish that the nucleotide sequence is unchanged. Those genes have not been indentified unequivocally in rabbit. To date, chicken l genes are the only Ig genes for which gene conversion has been established. Somatic diversification of rabbit IgH genes by hypermutation was analyzed by determining the sequence of the region 3¢ of VDJ genes, a region for which no homologous donor sequences are known in the germline (Lanning and Knight, 1997). Extensive mutation was found in this region, and the mutations were typical of somatic hypermutation, including a preference for transitions over transversions, strand bias, hotspots, and decreasing frequency of mutation with increasing distance from the VDJ gene. GALT and the Intestinal Flora Are Required for Development of the Primary Antibody Repertoire GALT plays an essential role in both somatic diversification of Ig genes and B-cell expansion. Soon after birth, B cells likely migrate from bone marrow and fetal liver, the sites of B lymphopoiesis, to GALT where they proliferate and somatically diversify their Ig genes (Figure 28.2) (Crane et al., 1996; Pospisil et al., 1995; Vajdy et al., 1998). The somatic diversification of Ig genes after birth is unlike that in chicken, where most Ig gene diversification occurs in the bursa of Fabricius before hatching. Because newborn rabbits are protected by both transplacental and colostral Ig, they may have no need to expand the antibody repertoire until levels of maternal Ig decrease a few weeks after birth. Lanning et al. (2000) determined whether somatic diversification of rabbit Ig genes in GALT was developmentally programmed or was driven by the intestinal flora. For these experiments, all organized GALT except the appendix was removed at birth, and the lumen of the appendix was ligated so that it could not interact with the intestinal flora. They found that GALT, especially follicles with proliferating B cells, did not develop and that nearly 90% of the VDJ genes did not undergo somatic diversification. These data demonstrated that neither B-cell proliferation nor the somatic diversification of Ig genes is developmentally programmed; rather these processes are driven by the intestinal flora. By raising sterilely derived newborn rabbits in an environment in which they were not exposed to the microbiota of conventional rabbits but instead to the microbiota of a normal laboratory environment, Lanning et al. (2000) also showed that not all microbiota could induce somatic diversification of Ig genes. More recently, by introducing individual intesti-
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nal bacteria into the lumen of ligated (germfree) appendices, some microorganisms, such as a combination of Bacillus subtilis and Bacteroides fragilis, were found to promote the development of follicles with proliferating B cells in GALT (K. Rhee, D. Lanning and K.L. Knight, unpublished data). Presumably, the proliferation and expansion of B cells and diversification of Ig genes that occurs in GALT a few weeks after birth and in response to the microbiota, result in the primary B-cell and antibody repertoires that provide initial protection for the rabbit from invasion by pathogenic microorganisms. Although GALT development and somatic diversification of Ig genes in GALT require intestinal flora, we think these processes are not antigen driven. Lanning et al. (2000) introduced individual organisms such as Bacteroides fragilis, Clostridium difficile, and Escherichia coli into the ligated (germfree) appendix, and those bacteria did not promote GALT development, or presumably, Ig gene diversification, even though they could serve as antigens. Recently, Sehgal et al. (2002) analyzed somatic diversification of Ig genes in clonally related B cells that occurs in the appendix a few weeks after birth and compared it with diversification of Ig genes in clonally related B cells from spleen during an antigen-specific immune response. They found that during development of the primary antibody repertoire in the appendix, all three CDRs of the Ig genes diversified extensively, whereas during an antigen-specific response in spleen, only CDR1 and CDR2 diversified extensively. These observations suggest that Ig gene diversification in GALT differs from that of an antigen-specific response. We speculate that somatic diversification of Ig genes might be the result of B cells being activated through the BCR by a superantigen, through a Toll receptor, and/or through cytokines produced by local T cells or dendritic cells. Whether T cells are required for GALT development or somatic diversification of the Ig genes in GALT has yet to be determined. The Secondary Immune Responses To investigate whether the gene conversion-like recombination that contributes to development of the primary antibody repertoire also occurs during secondary immune responses, Schiaffella et al. (1999) and Winstead et al. (1999) examined hapten-specific VDJ genes from splenic germinal centers and from popliteal lymph node–derived hybridomas. They identified clonally related sequences and found that they had both single nucleotide changes, which are characteristic of somatic mutation, and clusters of mutations, including codon insertions and deletions, which are characteristic of gene conversion. Similar results were obtained for hapten-specific VJk chain genes, indicating that they also diversify by gene conversion as well as by somatic hypermutation during secondary immune responses (Sehgal et al., 2000). As described above, gene conversion is also
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used to somatically diversify Ig genes in chickens during secondary immune responses, thus indicating that gene conversion is not limited to the generation of the primary antibody repertoire but is also used during antigen-induced immune responses.
CH Genes Like other mammals, rabbits have CH genes that encode each of the isotypes, IgM, IgG, IgA, and IgE (Figure 28.3). The rabbit CH chromosomal region differs from that of most other mammals in that 1) only a single Cg gene is present; 2) thirteen nonallelic Ca genes are present, and 3) no Cd gene has been found. Recently, the nucleotide sequence of genomic DNA 15 kb 3¢ of Cm was determined, and no Cd gene or Cd remnant was identified (Lanning et al., 2003). For many years IgD had been found only in some teleosts, rodents, and primates, and it appeared that the Cd gene may have been eliminated during the evolution of other mammals. However, Zhao et al. (2002) recently identified Cd in cattle, pigs, and sheep, suggesting that IgD may be more common among mammals than previously thought (discussed below). The region of the rabbit CH chromosomal locus that corresponds to the region containing Cd in other mammals is occupied by LINE and C repeats. If Cd is present in rabbit, it must reside in a location different from that in other species. The most unusual feature of the CH chromosomal region in rabbit is the presence of thirteen tandem Ca genes. All thirteen of these genes are expressible in vitro. However, in vivo, Ca3 and Ca8 are not expressed, and the other genes are differentially expressed in various mucosal tissues. Because the expression of germline Ia-Ca transcripts precedes isotype switching, Spieker-Polet et al. (2002) used a luciferase reporter gene assay to examine the Ia promoters and determine to what extent differential expression was due to differences in the Ia promoters. They found that Ca3 and Ca8 are not expressed because the Ia3 and Ia8 promoters were defective. In contrast, they found that all other Ia promoter regions were functional and could be induced by TGFb and IL-2. However, the strength of promoter activity
FIGURE 28.3 Organization of IgCH chromosomal region in rabbit. Solid boxes = Ca genes; ovals = switch regions.
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did not correlate with levels of Ca gene expression in vivo, and consequently, the molecular basis for the differential expression of IgA isotypes is not known. We suggest that the 3¢a enhancers may contribute to this regulation. The expansion of Ca genes in rabbits occurred in an ancestor of lagomorphs. Burnett et al. (1989) showed by genomic Southern blot analysis that members of both families of lagomorphs, Leporidae and Ochotonidae, have multiple Ca genes. Because the location of lagomorphs in the mammalian phylogenetic tree is still in question, it is not known in which ancestor the duplication of Ca genes occurred. It is also not known whether the expansion of Ca genes conferred a survival advantage to lagomorphs. Functionally, all the Ca genes appear similar in that all IgA isotypes bind poly-immunoglobulin receptor and can be transported across the epithelium into the lumen (Schneiderman et al., 1989). Further, all IgA isotypes activate complement by the alternative pathway (Schneiderman et al., 1990). The reason for the expansion of Ca genes remains a mystery.
Enhancer Regions IgH and k intronic enhancers as well as 3¢aE enhancers, but not 3¢Ek enhancers, have been identified in rabbit Ig chromosomal regions. The intronic enhancer, Em, is located 4.5 kb 5¢ of Cm, and its sequence is highly similar to that of mouse and human (Mage et al., 1989). Transgenic rabbits, in which Em was used to drive the tissue-specific expression of c-myc, developed B-cell leukemia soon after birth (Knight et al., 1988). In contrast, transgenic rabbits with cmyc driven by the intronic Ek enhancer (Emorine et al., 1983b) developed one of several different types of tumors, including B-cell lymphoma, basal cell carcinoma, embryonic carcinoma, hepatoma, and ovarian carcinoma (Sethupathi et al., 1994). These findings indicate that intronic Ek functions in cells other than B-lineage cells. A 3¢aE hs1,2 enhancer was identified approximately 8 kb downstream of the distal Ca gene, Ca13 (Volgina et al., 2000). More recently, a negative regulatory element, designated Ca-NRE, which negatively regulates the hs1,2 enhancer activity on Ia promoters has been found associated with eight of the thirteen Ca genes (Volgina and Knight, unpublished data). Ca-NRE may contribute to the differential expression of the Ca genes in various tissues. Summary Rabbits share some features of the human and mouse B lymphoid systems and other features of the avian B lymphoid system. For example, rabbits, mice, and humans share a similar genomic organization of Ig genes and, in all three species, B-cell development and Ig gene rearrangement take
place in the bone marrow. In contrast, like chickens, rabbits preferentially use one VH gene segment during VDJ gene rearrangement, require GALT for somatic diversification of Ig genes and development of the primary antibody repertoire, and undergo waning of B lymphopoiesis with age. Yet, rabbits differ from all these species in requiring the intestinal microflora for somatic diversification of Ig genes during the generation of the primary antibody repertoire. Thus, rabbit B-cell development, incorporating characteristics of both the chicken and the mouse and human B-cell developmental pathways, might provide insight into the evolutionary relationship between these two fundamentally different strategies for generating a primary antibody repertoire.
ARTIODACTYLS The order Artiodactyla encompasses cloven-hoofed animals with an even number of toes and includes such diverse animals as pigs, sheep, cattle, bison, deer, camel, and hippopotamus. Immunoglobulin genes of artiodactyls such as sheep, cattle, and swine have been studied much less extensively than their counterparts in human, mouse, rabbit, chicken, and perhaps even fish. However, enough information is available to provide a partial view of the organization of Ig genes, mechanisms of generation of diversity, and B-cell development in these species. Although differences exist among the expressed IgG subclasses, antibody structure in Artiodactyla is the same as in other mammals with one outstanding exception. Antibodies of the camelids (camels, llamas, and dromedaries) are strikingly different from those of most other vertebrates. Instead of the usual two heavy- and two light-chain structure, approximately 50% of camelid antibodies are comprised of two heavy chains and no light chains (Hamers-Casterman et al., 1993; Nguyen et al., 2001; van der Linden et al., 2000). These antibodies are referred to as heavy-chain antibodies (HCAb), and the VH regions are designated VHH. Various antigens appear to differentially elict HCAb or conventional IgG. In a study by van der Linden and colleagues (2000), llamas were immunized with a variety of antigens, and some antigens elicited primarily a conventional antibody response whereas others induced predominantly HCAbs. In HCAb, the light chain–binding domain, CH1, of g heavy chains, is absent due to a nonfunctional splice signal at the 3¢ end of the CH1 exon (Figure 28.4). HCAbs are easily secreted from cells because the conserved amino acids in conventional VH regions that interact with VL are absent, and without the CH1 domain, chaperones will not bind and retain the heavy chain in the endoplasmic reticulum. Several crystal structures of HCAb associated with antigen have been determined, and these molecules have the classic Ig fold with conserved intradomain disulfide bonds and the
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FIGURE 28.4 Schematic representation of conventional antibodies compared with HCAb in camelid. Note the lack of light chain and the absence of a CH1 domain in camelid HCAb. Also note the extended hinge region in HCAb. Interchain disulfide bonds are indicated by -S-.
CDR clustered at one end of the molecule (Nguyen et al., 2001). In many instances, CDR1 and CDR3 of HCAbs are considerably longer than in conventional VH molecules and are stabilized by an intradomain disulfide bond between these two regions.
VH, D, and JH Gene Segments The genomic size and content of the heavy chain loci of any of the artiodactyls is not known, probably because of the lack of resources applied to genome projects of agriculturally relevant animals. However, the data that do exist provide clues about the organization of heavy chain gene segments in these animals. An estimated fifteen VH genes have been identified in sheep, and all of these genes belong to a single VH family, VH1, within group B of VH genes, as defined by Sitnikova and Su (1998). This family is most related to VH4 in human (Dufour et al., 1996). Within the JH locus, six gene segments have been identified, but only two are functional; the other four are pseudogenes (Dufour and Nau, 1997). Thus far, no DH gene segments have been isolated, but cDNA sequences predict that these gene segments exist and have yet to be identified. In cattle, approximately ten to twenty germline VH gene segments have been identified (Kaushil et al., 2002; Jackson, 2002). All belong to a single family (Bo VH1) that, like sheep, is analogous to human VH4 or group B (Sinclair et al., 1997; Saini et al., 1997; Jackson, 2002). Only one JH gene segment currently is known, and this sequence is most similar to human JH4 and human JH2 (Jackson, 2002). Similar to sheep, no DH gene segments have been isolated. It is possible that DH gene segments may be longer in cattle than sheep because CDR3 lengths vary a great deal, with some as long as twenty-one amino acids compared to the seven to twelve amino acids observed in mouse or human CDR3 (Berenes et al., 1997; Lopez et al., 1998). This longer
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length of CDR3 could be due to longer D gene segments in cattle, the use of multiple D gene segments, nontemplated addition of bases during VDJ recombination, or even a combination of some or all of these mechanisms (Kaushik et al., 2002; Jackson, 2002). However, these explanations remain speculative, and determination of the germline sequences of the D-gene segment cluster will resolve this issue. The VH repertoire in pigs is similar to that in cattle and sheep and also appears restricted, with only five VH gene segments known to be expressed (Butler et al., 1996b; 2000; Butler, 1998; Sun et al., 1998). These VH genes are members of group C. Pigs also have a single identified JH gene segment and two DH gene segments. Again it must be stressed that the exact number of genes and their precise organization remains to be elucidated, and progress in genome sequencing of domestic artiodactyls will help characterize the Ig repertoire of these animals. Both VH and VHH gene segments are found in the germline of camelids, and both have normal promoter sequences as well as rearrangement signal sequences. Forty different VHH and fifty VH germline gene segments have been identified by PCR (Nguyen et al., 2000), but the total number of VH and VHH gene segments is not known. All known VHH gene segments are members of seven subfamilies within VH family III, group C. Similar to other vertebrates, camelids have DH and JH gene segments. Although it is not known whether the VH and VHH gene segments are clustered or are on separate chromosomes, it seems likely that they are clustered because the same DH and JH gene segments are associated with both VH and VHH gene segments (Nguyen et al., 2000). The genomic organization of camelid Ig heavy chain gene segments is complicated by the observation that HCAbs are exclusively IgG. If VH and VHH gene segments are clustered and utilize the same DH and JH gene segments, it is difficult to understand how a primary transcript encoding VHH/DH/JH in association with the IgG constant region is produced. A detailed map of the camelid heavy chain locus is required to better understand the observed protein data.
Kappa and Lambda Light Chain Genes Sheep and cattle express primarily l light chains. However, in both species, there is evidence of a functional k chain locus. In sheep, there are at least six Vl families (Reynaud et al., 1991) and probably more than fifty germline Vl gene segments. Among the expressed genes, only four of the six families appear to be represented (Charleton et al., 2000). Two Jl gene segments have been identified in sheep (Jeong et al., 2001). Cattle, like sheep, have many germline Vl gene segments; however, many of them appear to be pseudogenes. The functional Vl gene segments likely belong to two distinct families, one of which is used pre-
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dominantly in VJl gene rearrangements (Parng et al., 1996). Similar to sheep, cattle have one Jl gene segment, thereby limiting the amount of diversity that can be generated by V-Jl gene rearrangement (Parng et al., 1996; Meyer et al., 1997). Although cattle and sheep do not express high levels of k light chain protein, both species possess functional k chain gene loci. The k chain locus is poorly characterized in either species, but cDNA sequences from sheep indicate that at least six separate gene families may exist (Charleton et al., 2000). Southern blot data also suggest that the k locus in sheep might have been duplicated (Hein and Dudler, 1998). Cattle have several functional Vk gene segments and a functional Jk gene segment (S. Pyarajan, K. Curnick, and B.A. Osborne, unpublished data). It is not apparent from published studies why l light chain expression predominates in cattle and sheep, because the k loci appear capable of expressing functional k light chain protein. Pigs, unlike their barnyard compatriots, cattle and sheep, express approximately 60% k and 40% l light chains (Butler and Brown, 1994). Both the number and organization of germline light chain genes are unknown. Characterization of k and l loci in sheep, cattle, and pigs should provide valuable insights into the mechanisms that regulate vertebrate light chain gene expression. In camelids, genes encoding k and l light chains used to produce conventional Ig molecules have been cloned from cDNA of peripheral blood cells and found to be similar to those of conventional light chain genes (Nguyen et al., 2001). These findings, taken together with the presence of conventional VH, DH, and JH gene segments, suggest that the conventional Ig molecules and genes are similar to those of other vertebrates.
Generation of Antibody Diversity Because the germline Ig gene loci of artiodactyls are poorly characterized, it is difficult to comment unambiguously on the mechanisms used by these animals to generate a diverse antibody response. Clearly, the limited number of
V-gene segments used to generate the primary repertoire, as well as very few J-gene segments, suggests that artiodactyls must use mechanisms other than combinatorial diversification to generate a primary antibody repertoire. In fact, early studies by Weill and Reynaud and coworkers over a decade ago suggested that sheep diversify the Ig repertoire primarily through extensive somatic hypermutation of a few rearranged VJ sequences (Table 28.1) (Reynaud et al., 1991). The data supporting somatic mutation as the primary mechanism of Ig gene diversification is based on a comparison between sequences of several germline genes and Ig cDNA sequences. Many of the differences between these sequences are single point mutations, suggesting that these genes undergo somatic hypermutation in human or mouse. Because all of the germline Ig genes have not been isolated in sheep, another interpretation is that the potentially mutated genes may merely be new, closely related germline genes. Indeed, recent evidence from Charlton and colleagues using a bacteriophage display library suggests that the germline heavy chain repertoire may be more diverse than previously reported (2000). Until the entire VH and VL loci are sequenced, it is difficult to unequivocally attribute the differences between known germline genes and expressed genes to somatic hypermutation. More recent data from cattle and pigs indicate that, in addition to somatic mutation, templated recombinational processes, such as gene conversion, might contribute to the diversification of the primary repertoire. Several years ago, when sequencing l light chain genes from cattle, Parng et al. (1996) observed that sequence differences between a germline gene and an expressed gene could be attributed to blocks of sequences found in other germline genes. In many instances, these could be found in nonfunctional or pseudogenes, suggesting that, as in chicken, pseudogenes could act as sequence reservoirs and contribute to the diversification of the functional repertoire by donating sequences through gene conversion. Although this occurred in the diversification of the l light chain repertoire, heavy chain diversification appeared to happen primarily through somatic hypermutation rather than gene conversion (Jackson, 2002).
TABLE 28.1 Generation of primary antibody repertoire in GALT-species of mammals Mechanism of Ig gene diversification Organism Chicken Rabbit Sheep Cattle
Site of Blymphopoiesis
Site of primary repertoire diversification
Gene conversion
Somatic mutation
Requirement for intestinal flora
Spleen, yolk sac Bone marrow, fetal liver Spleen Spleen
Bursa of Fabricius GALT/Appendix Ileal Peyer’s Patch Ileal Peyer’s Patch
+++ +++ -? +?
+ +++ +++ +++
No Yes No ?
GALT = gut-associated lymphoid tissue.
28. Immunoglobulin Genes and Generation of Antibody Repertoires in Higher Vertebrates
Again these conclusions require tempering and await the complete sequencing of both heavy and light chain loci in order to assess the contribution of germline sequence to the expressed repertoire. Similar to cattle, sequence data from pigs indicated that at least part of the diversity-generating machinery could utilize gene conversion (Butler et al., 1996). However these data were complicated by the use of PCR to amplify expressed Ig cDNAs for sequencing. For example, data from the same group showed that gene chimeras resembling gene conversion could be generated by PCR (Sinkora et al., 2000). Once again, without a complete or near complete compilation of germline sequences, it is difficult to determine which mechanisms generate antibody diversity. Diversification of the primary repertoire in camelids is poorly understood, but in the absence of the combinatorial association of VH and VL chains in HCAbs, it is not clear how a large repertoire of HCAbs is generated. Perhaps this repertoire is, in part, a reflection of a large CDR3. Nguyen et al. (2000) generated hypervariability plots of germline and rearranged VHH genes and found considerable variation around the antigen-binding loops. They proposed that mutations in this region occur through an antigen-independent process to increase the diversity of the repertoire. An ileal Peyer’s patch has been described in camelids (Alluwaimi et al., 1998), indicating that they may be members of the GALT-species. If so, it is possible that this antigenindependent somatic diversification of Ig genes occurs in GALT either directly or indirectly in response to the intestinal flora. A practical advantage to camelid homodimeric Ig molecules, HCAbs, is that such molecules can be readily synthesized and purified by standard molecular genetic methods. The VHH antibodies can be expressed as soluble proteins in bacteria and yeast (Nyugen et al., 2001), and such antibodies may be considerably easier to produce in vitro than the conventional antibodies that require both VH and VL. This finding, coupled with the observation that VHH preferentially interacts with the active site of many enzymes, makes these unique antibodies attractive candidates for the production of useful pharmaceutics (Lauwereys et al., 1998).
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et al., 1993). That group reported finding Ig+ B cells in splenic white pulp at embryonic day 45 to 47 (gestation is 150 days in sheep). By embryonic day 77, B cells occupy 20% of the splenic pulp. B cells then decrease in number, T cells enter the spleen by embryonic day 90, and the architecture of the spleen begins to appear like a secondary lymphoid organ. By embryonic day 100, B cells have colonized the ileal Peyer’s patch (IPP), a lymphoid organ that many suggest is an equivalent of the bursa in chicken. IPP is comprised of discreet follicles, and each follicle is oligoclonal with regard to B cells. Press et al. (1993) suggested that B cells leave the spleen during fetal development and migrate to and populate the IPP. These early observations are supported by the observation that RAG1 and RAG2 expression is detected in bovine and ovine fetal spleen but not in bone marrow. Recently, Davidyuk (2001) studied the temporal development of B cells from the spleen and IPP of ovine fetuses between embryonic day 40 and birth. She found a clear pattern of B-cell development in the spleen where much, if not all, gene rearrangement occurs, followed by migration to the IPP, where diversification occurs. Diversification occurs at a low level during the third trimester of fetal life, and the rate of diversification increases dramatically around the time of birth. Therefore, it appears as though immature B cells with rearranged VDJ/VJ genes exit the fetal spleen and home to the ileal Peyer’s patch, where further diversification occurs either through templated somatic mutation (gene conversion) or untemplated somatic hypermutation. B cells in cattle appear to follow similar developmental patterns (Lucier et al., 1998; Jackson, 2002). Many questions remain about the temporal order of Ig gene rearrangements, the site(s) where rearrangement occurs, and the site and precise mechanisms by which further diversification occurs. These studies have been hampered by the tractability of using large domestic animals and the availability of timed fetal material. Additionally, the relative lack of resources available for genome sequencing in domestic animals has delayed the acquisition of sequence data necessary to determine precisely whether gene conversion, somatic hypermutation, or both drive antibody diversity in these animals.
CH Genes Sites of B Lymphopoiesis and Ig Gene Diversification Artiodactyls, like lagomorphs and avians, differ not only in the manner in which diversity is generated but also in the site(s) at which this diversification occurs. As discussed above, B cells with new combinations of V(D)J gene rearrangements are generated in the bone marrow of human and mouse throughout life. In cattle and sheep, B cells are not generated in the bone marrow. Several years ago, Landsverk and colleagues suggested that sheep fetal spleen is a site of early B-cell development and proliferation (Press
Artiodactyls, like other vertebrates, have IgM, IgG, IgA, and IgE immunoglobulin–constant region genes. However, it has long been thought that artiodactyls lacked a gene encoding IgD (Butler et al., 1996a). As described above, it was suggested that IgD evolved recently and was present only in rodents and primates, but data demonstrating an IgD gene in teleosts proved this incorrect. The recent cloning and characterization of the Cd gene in cattle, sheep, and pigs (Zhao et al., 2002) suggest to us that the status of IgD needs to be reexamined in vertebrates. Not only is an IgD gene
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present in cattle, sheep, and pigs, but at least in cattle, a short Sd region is also present, which, in theory, could mediate switch recombination. The occurrence of switch recombination involving Sm and Sd was demonstrated, suggesting that, at least in cattle, it may be possible for B cells to express IgD exclusively. Another interesting feature of the cattle and sheep IgD genes is the striking degree of similarity shared by the IgM and IgD CH1 domain. Zhao et al. (2002) suggested that bovine and ovine IgD arose through a gene duplication in which the dCH1 exon was duplicated from the mCH1 exon. The porcine dCH1 domain does not share the same degree of homology with the porcine mCH1 domain, suggesting that this duplication may be limited to sheep and cattle. The IgD data are quite recent, and more clarification is needed before a coherent view of IgD expression in artiodactyls can emerge. Although these data suggest that IgD is expressed in artiodactyls, synthesis of protein has yet to be demonstrated. It is critical to determine if IgD can be expressed on B cells and if, as suggested from mRNA data, B cells expressing only IgD exist in sheep and cattle. Several interesting and important questions remain to be addressed in this area.
OTHER MAMMALS Dog Canine Ig genes have not been well studied. Only two canine Ig gene constant region loci, Ca and Ce, have been cloned and characterized (Patel et al., 1995). The canine Ce locus encompasses 1.7 kb of DNA and contains four exons encoding a predicted protein product that is 57% and 49% identical to those of the human and murine Ce genes, respectively. The canine Ca locus consists of three exons spanning 1.5 kb and encodes a predicted protein product with 70% and 65% identity to those of the human and murine Ca genes, respectively. Interestingly, the dog has two types of Peyer’s patches, duodenal/jejunal and ileal (HogenEsch et al., 1987; HogenEsch and Felsburg, 1990, 1992). Twenty-five to twenty-eight duodenal and jejunal Peyer’s patches share a morphology characteristic of secondary lymphoid organs. The single ileal Peyer’s patch is 26 to 30 cm long in 4- to 6month-old dogs and forms a complete ring around the distal ileum. It comprises about 80% of the surface area of all the Peyer’s patches combined (HogenEsch et al., 1987). The ileal Peyer’s patch contains large follicles with small domes, which are characteristic of primary lymphoid tissues. B cells from the ileal Peyer’s patch secrete primarily IgM after in vitro stimulation (HogenEsch and Felsburg, 1992). Unlike the duodenal and jejunal Peyer’s patches, the ileal Peyer’s patch undergoes involution after the dog attains sexual maturity. These features suggest that the ileal Peyer’s patch
might serve as a site of primary Ig gene diversification and B cell expansion.
Cat The feline Ig genes are largely uncharacterized. Cho et al. (1998) analyzed partial cDNA clones of the Cm and Cg genes obtained by RT-PCR amplification of PBL RNA. A 691-bp region of the Cm gene, containing the CH1, CH2, and part of the CH3 domains, encodes a predicted protein sequence that is 59% and 55% identical to the corresponding regions encoded by the human and mouse Cm genes, respectively. A 384-bp region of the Cg gene, containing the CH2 domain and part of the CH3 domain, encodes a predicted protein sequence that is 66% identical to the corresponding regions encoded by the human and mouse Cg genes.
Horse The Ig classes of the horse are complex and are encoded by six Cg genes, as well as Cm, Ca, and Ce genes. The heavy chain constant region genes are aligned, as determined by heterohybridoma deletion analysis, in the order Cm, Cg1–6, Ce, Ca (Wagner et al., 1997, 1998; Overesch et al., 1998). The Cg1, Cg3, and Cg4 genes encode the IgGa, IgG(T), and IgGb isotypes, respectively, which were previously identified serologically. To date, only the Ce gene has been characterized at the nucleotide level (Navarro et al., 1995; Marti et al., 1995; Wagner et al. 2001). The Ce locus contains four exons spanning 1.6 kb and encoding a deduced protein sequence of 424 amino acids. Three Ce alleles have been reported, ranging from 95.9 to 99.8% identity at the deduced amino acid level and differing primarily in the Ce1 and Ce2 domains (Navarro et al., 1995; Wagner et al., 2001). Although the horse has both l and k light chain loci, the antibody repertoire utilizes almost exclusively l light chains (Gibson, 1974). The l locus is estimated by Southern analysis to contain approximately twenty to thirty Vl gene segments and four Cl genes (Home et al., 1992). Three closely related Vl gene segments appear to be preferentially utilized and can rearrange to any one of three Cl genes; the fourth is probably a pseudogene (Home et al., 1992). Horse l light chains exhibit extensive V-J junctional variation, including amino acid substitutions, insertions, and deletions. The horse k locus contains at least twenty Vk gene segments, as estimated by Southern analysis (Ford et al., 1994). A single Ck gene lies about 2.9 kb downstream of five Jk gene segments, three of which appear to be functional. One Jk gene segment is nonfunctional because it lacks RSS, and a second might be nonfunctional because of a short, 20-bp, instead of 23-bp, RSS spacer. The Jk-Ck intron contains a highly conserved enhancer (Ford et al., 1994). The reason for the disproportionate use of l light chains in the antibody repertoire is not known.
28. Immunoglobulin Genes and Generation of Antibody Repertoires in Higher Vertebrates
CONCLUSION We have focused on the Ig genes and development of the antibody repertoire in species that generate their primary antibody repertoires through the extensive somatic diversification of V(D) J heavy and light chain genes. Sitnikova and Su (1998) performed a phylogenetic analysis of VH and VL genes from several species of amniotes and found that the germline VH and VL genes of species that utilize somatic diversification to expand their primary antibody repertoire generally belong to a single group of VH and VL genes. In contrast, the VH and VL genes of mouse and human, which use combinatorial joining of multiple V, (D), and J gene segments to generate the primary repertoire, comprise multiple VH and Vk groups. It is possible that species in which the germline genes belong to only a single group and therefore have a less diverse repertoire of V gene segments have evolved to utilize somatic hypermutation, gene conversion, or both for developing a highly diversified primary antibody repertoire. It is also possible that adopting gene conversion as a mechanism for diversifying Ig genes biases the VH gene segments toward a single family because gene conversion requires a high degree of homology between donor and recipient sequences. Further, the species with less diverse Vgene segments all utilize GALT as the site for B-cell expansion and generation of the primary antibody repertoire. The sites of B-cell development and the generation of the primary antibody repertoire are shown in Table 28.1 for four of the best-studied GALT-species (chicken, rabbit, sheep, and cattle). We predict that several other species, including swine, dogs, horses, and possibly camelids will be added to this list. Notably, chicken, rabbits, sheep, and cattle utilize different GALT sites to generate the primary antibody repertoire, including the bursa, ileal Peyer’s patches, and appendix. Furthermore, in these species the Ig genes can diversify by gene conversion and/or somatic hypermutation. Although exogenous antigens are not required for these processes in chicken, intestinal flora is required in rabbit, not only for GALT development but also for somatic diversification of the Ig genes. In the other species, some somatic diversification may occur before birth; however, considerably more diversification occurs after interaction with the intestinal flora. It is likely that the somatic diversification in GALT that leads to the primary antibody repertoire is not antigen driven, but instead occurs as a polyclonal response either to an exogenous agent, as occurs in rabbit, or possibly to an endogenous agent, such as a retroviral superantigen in species such as chicken, in which exogenous molecules are not required. Further studies are needed to elucidate the mechanism by which both GALT development and the somatic diversification of the Ig genes are initiated.
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29 The Zebrafish Immune System LISA A. STEINER1, CATHERINE E. WILLETT2, AND NADIA DANILOVA1 1
Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts, USA 2 Phylonix Pharmaceuticals, Cambridge, Massachusetts, USA
In the last decade or so, research on zebrafish has mushroomed. A milestone was the publication, in a special “zebrafish issue” of Development, of the results of two largescale genetic screens that described thousands of mutations in hundreds of genes affecting development, and showed that the mutational approach in zebrafish is feasible (Driever et al., 1996; Hafter et al., 1996; and other articles in the same volume). Included were mutations in embryogenesis and in the development of organs such as the brain and the hematopoietic, cardiovascular, and gastrointestinal systems. Conspicuously missing from the list of organs examined in these studies were those of the immune system. There are several reasons for this omission. The cells and organs of the zebrafish immune system are not visible in living fish, and the effects of mutations affecting immunity are unlikely to lead to a detectable phenotype during the embryonic and larval periods when the mutants were scored. At the time these screens were carried out, the organs of the immune system had not been described and no probes were available for identification. Since then, much has been learned about the immune system in zebrafish, and it has been shown that organs such as the thymus can be visualized by special techniques, (e.g., in situ hybridization) or the utilization of transgenes containing reporter constructs. The status of genetic screens related to the immune system is considered in a later section. In addition to the possibility of identifying genes involved in organogenesis, the transparency of the zebrafish larvae facilitates the tracking of cell populations. This has been beautifully illustrated in the work of Herbomel and colleagues, who have followed the migrations of a population of macrophages in the early embryo by microscopic techniques (see Phagocytic (Myeloid) Cells, P. 461). The transparent zebrafish embryo also lends itself to fate
The first task of authors of an article on zebrafish immunity embedded in a text devoted mainly to B cells and Ig genes might be to respond to the inevitable question “why zebrafish?” The immune systems of mice and humans, as well as numerous other vertebrates, including a variety of fish, have already been examined in some detaill. Why turn to a new species? The zebrafish, Danio rerio, is particularly well suited among vertebrates for a genetic analysis of development, as was first recognized by George Streisinger in the late 1960s (Streisinger et al., 1981; Grunwald and Eisen, 2002). The embryos and larvae are transparent, allowing the development of organs to be followed visually, and the free-living embryos develop quickly, independent of maternal influence. Genetic analysis is practical since brood sizes are large (females lay hundreds of eggs at a time) and many fish can be conveniently housed in the laboratory. It is the combination of good embryology and good genetics that makes the zebrafish a particularly useful model for developmental analysis (Grunwald and Eisen, 2002; Nüsslein-Volhard and Dahm, 2002). In contrast to techniques in which the activity of a known gene is disabled, a random mutational approach offers the possibility of identifying previously unrecognized genes. Despite genome sequencing and gene finding methodologies, important genes remain to be discovered. The haploid zebrafish genome contains about 1.7 ¥ 109 base pairs (approximately half the size of the human genome) divided among 25 chromosomes. There is substantial synteny between the zebrafish and human genomes. However, partial genome duplication may have occurred early in the teleost lineage, so that some genes that are unique in mammals have two copies in zebrafish (Postlethwait et al., 1999; Postlethwait et al., 2000). This can complicate genetic analysis.
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mapping. In one application, a protected or “caged” fluorescent marker is injected into early cleavage embryos and then “uncaged” in a particular location at a later time by directed light; the fate of cells labeled at the uncaging step can then be evaluated. It might be possible to follow suitably labeled cells of the B or T lineage as they migrate from sites where they are generated to the organs where they differentiate. Studies of the zebrafish also benefit from the considerable resources that are becoming available, such as genomic and cDNA libraries and EST databases. Maps of the genome are available and constantly being refined; the genome sequence is expected to be mostly complete within the next 2 years. Nevertheless, there are also obstacles to studies of the immune system in this species. Missing is the wealth of information about the cellular and molecular aspects of immunity that has become available, after decades of intense research, in the mouse and human. We may, however, extrapolate from the limited available data that new insights into the adaptive and innate immune systems will be forthcoming from studies of the zebrafish. In this chapter, we endeavor to summarize the present state of knowledge of the zebrafish immune system and its development. We consider the major elements of adaptive immunity (e.g., B and T lymphocytes and their antigen-specific receptors), the MHC, as well as some of the cells and molecules of innate immunity. For perspective on the early development of lymphoid cells and organs, we include an initial section presenting an overview of vertebrate and zebrafish hematopoiesis. We also include discussion of research directed at identifying mutants in developmental pathways, which has been a focus in zebrafish research. In a final section, we consider mutant phenotypes in zebrafish that resemble human disease syndromes. A number of references are indispensable for anyone working with zebrafish or interested in becoming more familiar with research in this organism. The Zebrafish Book by Monte Westerfield (Westerfield, 2000) has long been the
major guide for researchers in the field. Two volumes in Methods in Cell Biology contain a summary of zebrafish cell biology and genetics (Detrich et al., 1999). Another excellent collection of articles surveying basic methodology useful for research with zebrafish, as well as a list of mutants identified though April 2002, has recently been published (Nüsslein-Volhard and Dahm, 2002). Much information of interest to the “zebrafish community” is assembled in the web site ZFIN maintained by the Zebrafish International Resource Center at the University of Oregon: http://zfin.org/ (Sprague et al., 2001; 2003).
HEMATOPOIESIS Hematopoiesis during Vertebrate Development Hematopoiesis is the process of generating the various blood cell lineages, erythroid, myeloid and lymphoid, from a common stem cell. The nature and location of hematopoiesis changes during embryogenesis. In vertebrates, hematopoietic lineages are derived from the embryonic mesodermal layer. During the initial wave of hematopoiesis, called primitive hematopoiesis, only embryonic red blood cells (RBCs) and some myeloid cells are generated. In mammals, chicken, and Xenopus laevis, primitive hematopoiesis occurs in extra-embryonic tissue derived from the ventral mesoderm surrounding the yolk sac (reviewed by Orkin, 1995; Robb, 1997) (Figure 29.1). As development progresses, primitive hematopoiesis is replaced by definitive hematopoiesis, in which fetal and adult RBCs, as well as all myeloid and lymphoid lineages, are produced (Cumano et al., 1996; Medvinsky and Dzierzak, 1996). The initial site of definitive hematopoiesis is in tissue within the embryo, also derived from mesoderm: the para-aortic splanchopleura and the related aorta-gonad mesonephros (AGM) in mammals (Godin et al., 1995; Med-
FIGURE 29.1 Comparison of sites of primitive and definitive hematopoiesis during embryogenesis. Sites of primitive hematopoiesis are in red and definitive sites in green. The diagram for mouse is based on Dzierzak and Medvinsky, 1995; Godin et al., 1995; for chicken on Dieterlen-Lievre et al., 1993; and for frog on Turpen et al., 1997.
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vinsky and Dzierzak, 1996) and chicken (Dieterlen-Lievre et al., 1993), and the dorso-lateral plate in Xenopus (Chen and Turpen, 1995) (Figure 29.1). In each case, the hematopoietic cells are located within or near the ventral wall of the dorsal aorta. It is not clear whether these cells originate from the vessel endothelium or from surrounding mesenchyme (reviewed by Godin and Cumano, 2002). In the mouse, definitive hematopoiesis shifts to the fetal liver and eventually to the bone marrow, where it continues through adulthood (Zanjani et al., 1993; Clapp et al., 1995). Different progenitors are responsible for primitive and definitive hematopoiesis. The relationship between these progenitors has been much debated (reviewed by Cumano and Godin, 2001; Yoder, 2001; Godin and Cumano, 2002; Orkin and Zon, 2002). In the past decade, evidence has accumulated that yolk sac progenitors responsible for primitive hematopoiesis are not capable of generating definitive lineages or of long-term repopulation of irradiated hosts, whereas intra-embryonic definitive progenitors possess both properties and are considered true hematopoietic stem cells. The intra-embryonic stem cells appear before the onset of circulation, suggesting that they arise independently of yolk sac progenitors (Cumano et al., 1996; Nishikawa et al., 1998). Studies in Xenopus show that yolk sac progenitors and intra-embryonic hematopoietic stem cells are derived from different blastomeres (Ciau-Uitz et al., 2000). The
overall evidence suggests that the two progenitor populations arise independently.
Hematopoiesis During Zebrafish Development Hematopoiesis in zebrafish is fundamentally similar to that in other vertebrates. Many homologs of genes known to be required for hematopoiesis in other organisms have been cloned in zebrafish (Figure 29.2) (Detrich et al., 1995; Gering et al., 1998; Liao et al., 1998; Thompson et al., 1998). Common progenitors of both blood cells and vascular endothelium are initially located in bilateral stripes in two regions of the embryo (trunk and cephalic) derived from the lateral mesoderm, beginning approximately 11 hours post-fertilization (hpf), as first shown by the expression of the scl and GATA-2 genes (Detrich et al., 1995; Amatruda and Zon, 1999; Gering et al., 1998). (Formation of somites in zebrafish begins at 10 hpf.) As development proceeds, and before the beginning of blood circulation at about 25 hpf, the stripes in the trunk converge medially into the intermediate cell mass (ICM) where the initial (primitive) differentiation of embryonic erythrocytes occurs (Al-Adhami and Kunz, 1977; Detrich et al., 1995; Gering et al., 1998; Willett et al., 1999) (Figure 29.3). Although the ICM lies within the embryo, it is thought to be the functional equivalent of the extraembryonic blood island of higher vertebrates (Ama-
FIGURE 29.2 Zebrafish homologs of genes involved in the specification of hematopoietic lineages. The scheme shown is based on studies in the mouse. Some aspects are still being debated, for example whether there is a lymphoid progenitor lacking myeloid potential, as depicted here (see discussion in Spangrude, 2002). The genes shown have been cloned in zebrafish and their expression patterns examined. The assignment of zebrafish genes to cells is based both on expression patterns in zebrafish and data from mouse; it is meant only to organize the limited information about the zebrafish genes into the available scheme and is not to be taken literally. Omitted from the diagram are NK cells and dendritic cells, neither of which has been identified in zebrafish. References for zebrafish genes: scl (Gering et al., 1998; Liao et al., 1998); runx1 (Kalev-Zylinska et al., 2002); cbfb (Blake et al., 2000); fli1, flk1, lmo2, c-myb (Thompson et al., 1998); GATA1, GATA2 (Detrich et al., 1995); tie-1, tie-2 (Lyons et al., 1998); pu.1 (Lieschke et al., 2002); Ikaros (Willett et al., 2001); mpo (myeloperoxidase) (Bennett et al., 2001; Lieschke et al., 2001); globins (Detrich et al., 1995; Chan et al., 1997); c-fms (Parichy et al., 2000; Herbomel et al., 2001); L-plastin (Herbomel et al., 1999); rags (rag1 and rag2) (Willett et al., 1997a; Willett et al., 1997b); Igm (Danilova et al., 2000b; Danilova and Steiner, 2002); lck, GATA-3 (Trede et al., 2001); and TCRa (Danilova et al., 2000b; Lam et al., 2002).
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FIGURE 29.3 Sites of hematopoiesis in zebrafish embryo. Genes associated with hemangiogenic progenitors are expressed in the lateral mesoderm in two stripes. In the lateral mesoderm of the trunk (TLM), the stripes converge medially to form the intermediate cell mass (ICM). Primitive erythrocytes differentiate from the cells of the anterior ICM, whereas genes associated with multipotent hematopoietic progenitors are expressed in the posterior ICM. Embryonic macrophages are derived from the caudal lateral mesoderm (CLM) before 24 hpf. Definitive hematopoiesis is thought to originate in cells within or near the dorsal aorta (DA) from 24 to 72 hpf. Cells in the ventral tail (VT) express genes associated with hemangiogenic progenitors until day 4. Lymphoblasts are seen in the thymus (Th) from 65 hpf and express Ikaros from 72 hpf. See color insert.
truda and Zon, 1999). Early macrophages originate in the cephalic mesoderm anterior to the heart field (Figure 29.3), as discussed in a later section. Erythroid differentiation progresses in the anterior part of the ICM, as indicated by the expression of GATA-1 (Detrich et al., 1995; Thompson et al., 1998) and embryonic red blood cells (RBC) are seen here at 20 hpf (Detrich et al., 1995; Willett et al., 1999). Cells in the posterior ICM continue to express genes associated with hematopoietic progenitors for several more hours (GATA-2, c-myb, scl, cbfb) (Detrich et al., 1995; Gering et al., 1998; Liao et al., 1998; Thompson et al., 1998; Blake et al., 2000). These genes are also expressed in a region caudal to the posterior ICM, designated the ventral tail, which has been suggested as a site of definitive hematopoiesis or, more likely, angiogenesis (Gering et al., 1998; Liao et al., 1998; Thompson et al., 1998; Liao et al., 2002). A probable site for definitive hematopoiesis is a region within or ventral to the dorsal aorta, which differentiates from the ICM, and is in a similar location to the AGM of other vertebrates (Figure 29.3). A similar set of genes, c-myb, Ikaros, and runx1, is expressed in the dorsal aorta region of zebrafish (Thompson et al., 1998; Willett et al., 2001; Kalev-Zylinska et al., 2002) and in the mammalian AGM (Mucenski et al., 1991; Wang et al., 1996; North et al., 1999). Lineage tracing or transplantation experiments might indicate whether or not cells in the dorsal aorta are, in fact, the source of definitive hematopoietic lineages.
ADAPTIVE IMMUNITY IN ZEBRAFISH: ORGANS AND MOLECULES Immunity in Teleosts: A Brief Overview Although the investigation of lymphoid organs in zebrafish has begun only recently, studies of these organs in
other teleosts have been carried out for many years. In the last dozen or so years, nucleic acid and antibody probes have become available for the analysis of lymphoid organs that were previously described only anatomically. Genes encoding the antigen-specific receptors of T and B lymphocytes, as well as other genes important for teleost immunity, can be identified by their homology with genes in other organisms. Techniques that have been used include PCR with degenerate primers and cross-hybridization with probes derived from known genes. These approaches, although straightforward in principle, may be difficult in practice because of the sequence divergence that may have accumulated in the more than 400 million years that separate the lineages of ray-finned fish and other vertebrates (Nelson, 1994). Even when application of these methods leads to the identification of a homologous gene, the orthology with known mammalian genes may not be obvious. Nonetheless, these approaches have been productive and have led to the identification of a number of teleost immune system genes, as well as to information about their tissuespecific expression. In this section, we describe the lymphoid organs and antigen-specific receptors in teleosts other than zebrafish. This information forms the background for studies on zebrafish immunity. See also Chapter 27 in this volume.
Teleost Lymphoid Organs In both anatomic and molecular aspects, the immune system in teleosts displays many similarities to, but also differences from, that in the mouse or human. In jawed vertebrates, B and T lymphocytes, with their diverse receptors for antigen, are the fundamental cell types of adaptive immunity, and the thymus is the primary organ for differentiation of the T lymphocytic lineage. However, the organs in which B lymphocytes differentiate are not the same as those in
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the mouse. This is hardly surprising in view of the substantial differences that are found in sites for B lymphopoiesis, even among mammals (see Chapter 28 in this volume). The secondary lymphoid organs in teleosts also have some differences from the familiar spleen–lymph node paradigm. The thymus in teleosts is basically similar in structure and function to that in mammals. Both the teleost and mammalian thymus arise bilaterally in the pharyngeal endoderm. The teleost thymus remains closely associated with each gill chamber, pharyngeal epithelium forming its lower border. In contrast, the bilateral lobes of the mammalian thymus migrate to the upper mediastinum where they fuse and become surrounded by connective tissue. Histologically, the distinction between cortex and medulla is often not as clear in teleosts as in mammals (Zapata et al., 1996), but the two regions can be discerned in some fish (Fishelson, 1995). A detailed analysis of thymic architecture, including growth and involution, was carried out in the carp (Romano et al., 1999). Initially oval in shape, the growing thymus stretches out along the gill chamber, acquiring a conical shape, with a large base just beneath the pharyngeal epithelium. A higher density of thymocytes in the cortex distinguishes it from the medulla. A monoclonal antibody specific for a carp thymocyte subpopulation reacts only with lymphocytes in the cortex, providing additional evidence for its distinction from the medulla (Romano et al., 1999). Teleosts lack bone marrow, and the kidney is generally considered to assume the function of mammalian bone marrow in hematopoiesis and in the differentiation of B lymphocytes (Zapata, 1979; Zapata and Amemiya, 2000). The teleost kidney consists of the pronephros or head kidney and the mesonephros or trunk kidney. Both parts, but especially the pronephros, contain hematopoietic tissue. The lymphopoetic function of the pronephros and mesonephros, initially based on histological observations, received support when it was found that the recombination activating genes, rag1 and rag2, encoding the recombinases responsible for V(D)J rearrangement of antigen-receptor genes, are expressed in these organs in trout (Hansen and Kaattari, 1996; Hansen, 1997). Spleen and gut-associated lymphatic tissues are secondary lymphoid organs in teleosts. Antibody formation may also occur in the pronephros (Fange and Nilsson, 1985), which is therefore a secondary as well as a primary lymphoid organ. Zapata et al. (1996) have noted that teleosts, which generally contain abundant lymphohematopoietic tissue in the kidney, tend to have relatively poorly developed lymphoid tissue in the spleen. There is, however, considerable variation in the content of splenic erythroid and lymphoid tissue among teleost species (Tischendorf, 1985; Rijkers et al., 1980; Fournier-Betz et al., 2000; Pettersen et al., 2000; Stenvik et al., 2001). After antigenic stimula-
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tion, there may be an increase in lymphoid tissue in the spleen (Zapata et al., 1996). In the carp, a cyprinid fish like the zebrafish, a compact spleen contains mainly red pulp (Rijkers, 1981). Lymphoid tissue is fragmented along the intestine and intermixed with liver and pancreas, as well as being present in the kidney (Rijkers et al., 1980). It has been pointed out that, compared to other vertebrates, teleosts have less blood and more lymph (Wardle, 1971). The lymphatic circulation in the mesentery may contribute to a hospitable environment for lymphocytes. Lymphoid cells, in addition to accumulating along the gut, are also found in the skin and gills (Zapata et al., 1996). Neither lymph nodes nor Peyer’s patches are found in teleosts. Immune System Genes in Teleosts It has been known for over 30 years that an IgM-like molecule is the major Ig in teleost serum (early studies were reviewed by Marchalonis, 1977). Despite relatively little similarity in sequence, IgM in teleosts resembles that in mammals in overall features. A notable exception is that the H2L2 units do not form the pentameric aggregates that are typical of mammalian IgM, but associate into a variety of disulfide-bonded polymers up to a tetramer, a structure that was visualized in several cases by electron microscopy (Shelton and Smith, 1970; Acton et al., 1971; Lobb and Clem, 1981; Lobb and Clem, 1983; Warr, 1983; Kaattari et al., 1998). More detailed information about teleost Ig emerged from the analysis of cDNA and genomic clones. For a description of early work on Ig genes in teleosts, as well as in other lower vertebrates, see the review by Litman et al. in the first edition of this book (1989). More recent work on Ig and TCR genes in teleosts has been reviewed ( Wilson and Warr, 1992; Charlemagne et al., 1998; Litman et al., 1999; Bengten et al., 2000; Litman et al., this volume). A prominent difference between mammalian and teleost IgM is that transcripts encoding the membrane form of the teleost m chain are formed by alternative splicing of membrane exons to CH3, rather than to CH4 (Wilson et al., 1990; Bengten et al., 1992; Hordvik et al., 1992; Andersson and Matsunaga, 1993; Lee et al., 1993). Transcripts encoding another heavy chain, which bears considerable resemblance to the mammalian d chain, have also been identified in teleosts (Wilson et al., 1997; Hordvik et al., 1999; Stenvik and Jorgensen, 2000). Curiously, the C regions of these delta chains are chimeric, containing the Cm1 exon followed by Cd–like exons. Gene segments thought to encode a d chain have been identified in the Ig locus of Takifugu rubripes (Aparicio et al., 2002). More than one C region cluster may be present in teleost heavy chain loci (Ghaffari and Lobb, 1999; Bengten et al., 2002; Hordvik, 2002).
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Three light chain isotyptes have been described in teleosts (reviewed by Pilstrom 2002), but relating these to k and l chains of mammals is problematic. An unexpected feature of the Ig gene organization in teleosts is that the light chain genes are arranged in multiple clusters of the type VJ-C (Daggfeldt et al., 1993; Ghaffari and Lobb, 1993), resembling the cluster organization of both light and heavy chain genes in cartilaginous fish. However, the teleost heavy chain genes are disposed similarly to those in mammals, the so-called “translocon” arrangement in which clusters of V gene segments, D gene segments, and J gene segments are grouped separately from each other and from the C gene segments. Genes encoding TCRa and b have been cloned in a number of teleosts (Partula et al., 1994, 1996; Wilson et al., 1998; Wang et al., 2001; Wermenstam and Pilstrom, 2001). Genes encoding TCRd have been identified in the pufferfish, Tetraodon nigroviridis (Fischer et al., 2002) and, very recently, cDNA and genomic DNA encoding TCRa, b, g and d have been identified in the Japanese flounder (Nam et al., 2003); interestingly, three TCR Cd gene segments were identified, one associated with the TCRa locus and two with the TCRg locus. TCRa, b, g and d genes have also been cloned in cartilaginous fish (Rast et al., 1997). The rag genes are expressed in the primary lymphoid organs where B and T lymphocytes differentiate (Schatz et al., 1992). Also expressed in these organs is the gene encoding terminal deoxynucleotidyl transferase (TdT), a templateindependent DNA polymerase responsible for the random addition of nucleotides at coding junctions during V(D)J rearrangement, thereby contributing to junctional diversity (Alt and Baltimore, 1982; Desiderio et al., 1984). Rag and TdT genes were cloned in trout and found to be most highly expressed in thymus and pronephros (Hansen and Kaattari, 1995; Hansen and Kaattari, 1996; Hansen, 1997). Interestingly, TdT is expressed in trout embryos at 20 days (i.e., before hatching). In mice, TdT expression is not seen in the neonatal thymus, appearing only 3 to 5 days after birth (Bogue et al., 1992). In Xenopus laevis, expression of TdT appears to be delayed until metamorphosis (Lee and Hsu, 1994). The Ikaros gene encodes a transcription factor that is expressed in hematopoietic stem cells and in all cells of the lymphoid lineage (Georgopoulos, 1997). A variety of isoforms, important in regulating lymphocyte differentiation and proliferation, have been identified. Ikaros has been cloned in trout and a number of transcripts, some similar to those in mammals and some different, have been identified (Hansen et al., 1997). Trout Ikaros was found to be expressed in lymphoid tissue and, early in development, in yolk sac and embryo.
Identification and Cloning of Genes Characteristic of Zebrafish Lymphoid Cells and Organs Genes Encoding Rag and Ikaros Cloning of rag1 and rag2 provided a logical entry into zebrafish immunity for several reasons: 1) Expression of these genes should identify the primary lymphoid organs, which would then be expected to lead to the identification of the antigen-specific receptors of B and T lymphocytes; 2) the rag genes, especially rag1, are highly conserved among species so that probes or primers derived from other species have a relatively high chance of success in identifying orthologous genes; and 3) The close linkage of rag1 and rag2 in other organisms, and the paucity of introns, means that it might be possible to clone the genes from genomic DNA without requiring a rich tissue source of corresponding mRNA; further, cloning of one of the pair should easily lead to the other. The zebrafish rag locus was cloned and partially sequenced (Greenhalgh and Steiner, 1995; Willett et al., 1997a). As in other species, the two genes are closely linked and convergently transcribed; indeed in zebrafish, pufferfish (Fugu robripes), and trout, the coding regions are more closely spaced than in other vertebrates, being separated by only about 2 to 3 kb; the UTRs may overlap (Hansen and Kaattari, 1996; Willett et al., 1997a; Bertrand et al., 1998; Peixoto et al., 2000). Unlike the lack of introns in the coding regions of rag1 genes in other species, there are two introns in the zebrafish rag1 coding region (Willett et al., 1997a). Introns are found in the same location in rag1 of other Teleostei, but not in other ray-finned fish (Venkatesh et al., 1999, 2001). It seems likely that these introns were acquired before the teleost radiation. cDNA encoding Ikaros has been cloned in zebrafish (Willett et al., 2001). Construction of phylogenetic trees suggested that the sequence is an ortholog of mammalian Ikaros, as it is more similar to Ikaros than to related family members such as Helios, Aiolos, or Eos (Haire et al., 2000a). Genes Encoding IgM Heavy Chain The expression of the IgM heavy chain (Igm) is the most appropriate marker for the identification of B lymphocytes in zebrafish since it is expected to be expressed in all cells of the B lineage, from pre-B cell stage on. A germline VH gene and cDNA encoding zebrafish Igm were cloned and characterized (Steiner et al., 1999; Danilova et al., 2000a). Although the overall similarity in sequence between the zebrafish and mouse or human Cm domains is relatively low, major structural features, which presumably support the immunoglobulin fold, are conserved; for example, the two half-cystine (Cys) residues within each domain that could
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form the intradomain disulfide bridge characteristic of Ig domains and, except for CH3, the Trp between these. In CH3, the replacement of Trp by Ile is unusual since a Trp at this position is a characteristic feature of Ig domains (Kabat et al., 1991). As in other teleosts, the membrane form of the zebrafish m chain appears to be generated by splicing membrane exons to the CH3 exon, rather than to the CH4 exon, as in mammals. Data obtained by Southern blotting are consistent with the presence of a single Cm gene in the zebrafish genome (N.D. and L.A.S., unpublished). Four families of VH genes were identified (Danilova et al., 2000a). On phylogenetic analysis, three of the families cluster mainly with other teleost VH sequences. The tendency of teleost VH regions to form distinct groups was noted previously (Ota and Nei, 1994; Andersson and Matsunaga, 1996). The fourth family, although most closely related to teleost VH regions, falls into a broad group that also includes VH regions of heavy chains from mammals as well as from other lower vertebrates. Genes Encoding Light Chains cDNAs encoding three isotypes of light chain genes have been cloned in zebrafish, each with diversified V segments (Haire et al., 2000b). Phylogenetic analysis indicated that these resembled isotypes previously identified in other teleosts. All three isotypes clustered separately from light chains in non-teleost fish. Similar to transcripts in other teleost species, many of the zebrafish transcripts were found to be aberrant (e.g., germline V) and may not encode functional proteins. Southern blotting suggested that more than one C gene corresponds to each of these light chain isotypes (Haire et al., 2000b). For one of the isotypes, there were multiple hybridizing restriction fragments, consistent with a cluster type of genomic organization (see Immune System Genes in Teleosts, P. 453). As expected, V regions of heavy and light chains show greatest diversity of sequence in the CDRs. Although consistent with diversification by a process of somatic mutation, to date only a single VH germline gene segment and a few VL segments are available for comparison (Danilova et al., 2000a; Haire et al., 2000). The anticipated availability of the complete zebrafish genome sequence should clarify the basis for the V-region diversification, although polymorphisms may obscure the issue.
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residues between the second Cys in the Ig domain and the Cys in the connecting peptide is about 20 for human and mouse, but only five for zebrafish and slightly more for other teleosts. Whether this truncation in the teleost Ca has functional consequences is not known. A number of cDNAs encoding TCRa V-regions were also identified and the sequences fell into ten families (Haire et al., 2000b; Danilova et al., 2004). Southern blotting with genomic DNA from a partially inbred (Tübingen) strain of zebrafish revealed a single restriction fragment hybridizing to a Ca probe (N.D. and L.A.S, unpublished). With outbred fish, there were two such fragments (Haire et al., 2000b), possibly polymorphic variants of a single genomic Ca gene segment. Genes encoding TCRb, g, and d have not yet been identified in zebrafish. As part of the ongoing analysis of the zebrafish genome, the TCRa locus has been examined in some detail. A remarkable feature of the locus is the large number of V gene segments. To date, 148 potentially functional nonallelic V gene segments have been identified and classified into 87 families (T. Ota and C.T. Amemiya, personal communication). One contig of ~170 kb contains at least 44 V segments and another contig of ~320 kb contains over 100 V segments, at least 23 J gene segments and a single C gene segment. As in the pufferfish, Tetradon nigroviridis (Fischer et al., 2002), the V-gene segments are in opposite transcriptional orientation to the J and C segments (T. Ota and C.T. Amemiya, personal communication). In Tetradon nigroviridis a Cd gene segment, as well as Dd and Jd segments, are located upstream of the Ja segments and there appears to be some sharing of V gene segments by Ca and Cd (Fischer et al., 2002). In zebrafish, however, no TCRd gene segments have yet been identified; anchored PCR has so far revealed only Ca regions associated with Va regions in 10 Va families that are expressed in both thymus and intestine (Danilova et al., 2004).
Expression of Immune System Genes during Zebrafish Development Gene expression at various developmental stages has been examined mainly by whole-mount in situ hybridization and by RT/PCR. Expression of Ikaros and Rag
Genes Encoding TCRa Chains To date, the only TCR genes that have been cloned in zebrafish are those encoding TCRa (Steiner and Hohman 1999; Haire et al., 2000b). As in other teleosts, the TCRa C domain (108 amino acid residues) is truncated relative to that in mammals (about 140 residues). The number of
Ikaros was found to be expressed in the ICM of the zebrafish embryo, a site where primitive hematopoietic precursors are generated (Willett et al., 2001; see Hematopoiesis during Zebrafish Development, P. 451). At 48 hpf, Ikaros expression was seen between the dorsal aorta and tail vein, a presumptive site for definitive hematopoiesis
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in zebrafish. Diffuse expression was also seen in the pharyngeal region at this time. At 72 hpf, Ikaros is expressed in the thymus (Willett et al., 2001). In adults, the major sites of expression are the thymus and pronephros, with variable amounts in the spleen, in general agreement with Ikaros expression in trout (Hansen et al., 1997). Details of rag expression in zebrafish lymphoid tissue at various developmental stages are presented in the following two sections. It has been reported that rag1 and rag2 are both expressed in zebrafish olfactory sensory neurons, raising the possibility that they play a role in the expression of olfactory receptor genes (Jessen et al., 1999, 2001). The expression of these genes in olfactory tissue was demonstrated by several methods. However, whether or not rag expression has any role in olfactory receptor gene choice remains an open question. Analysis of large regions of genomic DNA has failed to yield clear evidence for the association of recombination signal sequences with olfactory receptor genes in human, mouse, and zebrafish genomes (Glusman et al., 2000; Dugas and Ngai, 2001; Lane et al., 2001; Kratz et al., 2002; Lane et al., 2002). Moreover, there have been no reported olfactory deficits in rag1 or rag2 mutant mice (Peter Mombaerts, personal communication). Further work, perhaps with homogeneous populations of sensory neurons expressing the same olfactory receptor gene, is needed to elucidate whether gene rearrangements occur in olfactory neurons. Thymus and T Cell Development The zebrafish thymus undergoes dramatic changes in size and shape as it develops. The overall morphology and the distinctions between regions within the thymus, which appear to resemble those observed in carp (Romano et al., 1999), have been studied by microscopy and by in situ hybridization with rag1 and TCRa probes (Willett et al., 1997b; Willett et al., 1999; Danilova et al., 2000b; Lam et al., 2002). Although known only in rough outline, the thymus in zebrafish appears to follow a developmental program similar to that in the mouse. The thymic primordium was identified in zebrafish at 54 hpf as a small pocket of cells ventral to the otic vesicle (Willett et al., 1997b). At 60 hpf, by electron microscopy, the region is seen to consist of two layers of epithelial cells, one layer lining the pharyngeal cavity and the second comprised of thymic epithelial cells; 5 hours later the first lymphoblasts are seen (Willett et al., 1999). At 3 to 31/2 days, Ikaros is expressed in the thymus, as are GATA-3 and the tyrosine protein kinase, lck (Trede et al., 2001; Willett et al., 2001). Also at about this time Foxn1 (whnb), the zebrafish ortholog of the nude gene in mice, is expressed in thymic epithelial cells (Schorpp et al., 2002). Lymphoblasts in the thymus begin proliferating and expressing rag1 between 86 and 92 hpf (Willett et al., 1997b; Willett et al., 1999; Trede et al.,
2001). Expression of TCRa is observed from about 4 days (Danilova et al., 2000b; Trede et al., 2001). Rag1 expression in the thymus at 7 days, detected by whole-mount in situ hybridization and by fluorescence of a rag1-gfp construct, is shown in Figure 29.4A and 4B. In situ hybridization with TCR and rag1 probes can be used to delineate cortical and medullary regions in the thymus. In zebrafish, a distinction between rag1-positive and TCRa-positive regions was discerned at about 1 week (Lam et al., 2002; Danilova et al., 2004). At this time the thymus, which is adjacent and just dorsal to the pharynx, is oval in shape. Most of the area stained by TCRa probes is also stained by rag1 probes; a small region not stained by rag1 presumably represents the forming medulla. At 3 weeks, the thymus is positioned on top of the third pharyngeal cartilage. A dorsal extension is formed, and the thymus begins a period of rapid growth, mainly in the dorsal direction. The epithelial cells in the thymus also differentiate for several weeks (Willett et al., 1999), suggesting that the environment for T-cell maturation may be changing during this time. At 6 weeks, the thymus has a columnlike extension on a base that remains dorsal to the pharynx. At this time, there is a clear demarcation between a rag1-positive region at the periphery and rag1-negative region elsewhere (Danilova et al., 2000b; Lam et al., 2002). With age, the thymus shrinks and is again confined to the pharynx (Lam et al., 2002, Danilova et al., 2004). TCRa expression was detected in peripheral tissues at 9 days, at about the time the cortical–medullary distinction becomes evident, consistent with emigration of mature T cells from the thymus. In adult zebrafish, T cells can be seen in virtually all organs. In young adults, they are especially numerous in the pronephros and mesonephros, where they form aggregates, consistent with the proliferation following activation by antigen. Thus, the zebrafish kidney may carry out functions performed by lymph nodes in mammals. In the
FIGURE 29.4 Rag1 expression in thymus at 1 week. (a) Whole-mount in situ hybridization with rag1antisense RNA probe labeled with digoxigenin; detection is with Fab of anti-digoxigenin conjugated to alkaline phosphatase and chromogenic substrate (Willett et al., 1997b). (b) Fluorescence of a reporter construct consisting of gfp inserted into the rag1 promoter region (Jessen et al., 1999). Photograph courtesy of J. Jessen.
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intestine, T cells are scattered between epithelial cells and in the lamina propria of the villi. This is in contrast to B cells, which are often found in aggregates in the lamina propria near crypts (Danilova and Steiner, 2002). This pattern of T- and B-cell distribution in the intestine of zebrafish resembles that described in mammals (e.g., German et al., 1999; Waly et al., 2001, reviewed by MacDonald and Spencer, 1994). T cells are also found in the spleen and in the gut mesentery. Scattered T cells are also seen in skin and a few in muscle. B-Cell Development Initially, the thymus was the only organ in which rag1 expression was noted when larval fish were examined by whole-mount in situ hybridization. In particular, such expression was not seen in the pronephros, the presumed site for B-cell differentiation in fish (Willett et al., 1997b), nor were lymphoid cells detected here until 3 weeks (Willett et al., 1999). These findings suggested that B lymphopoiesis might be delayed in zebrafish, relative to T lymphopoiesis. Determining the sequence of cDNA encoding V regions of zebrafish Igm permitted an investigation of the timing of gene rearrangement and expression. VDJ rearrangements were detected in DNA extracted from whole zebrafish as early as day 4, and expression of membrane Igm was detected in RNA derived from whole fish by day 7, suggesting that cells of the B lineage are already present. These findings posed the challenge of identifying sites where B cells are located in the period between 4 days and 3 weeks, as no organs for B lymphopoiesis other than the pronephros had previously been described in teleosts. Re-examination of fish for early sites of rag1 expression by whole-mount in situ hybridization revealed, in addition to the thymus, a small stained spot in the right dorsal region of the abdomen, consistent in location with the pancreas (Danilova and Steiner, 2002). This spot was barely visible at 4 days and gradually became more prominent. The identity of the stained organ as the pancreas was confirmed by staining with an insulin probe. In situ hybridization on sections of 10-day-old fish revealed rag1, as well as Igm, staining in a region surrounding an islet of Langerhans. Both rag1 and Igm were seen to be expressed in the pronephros beginning at 19 days (Danilova and Steiner, 2002), in agreement with the appearance of lymphoid cells in this organ at about this time (Willett et al., 1999). There was no evidence for expression of these genes in the liver, the site for B-cell development in fetal mouse and human. In adult zebrafish, Igm expression is prominent in the pronephros and mesonephros (Danilova and Steiner, 2002), consistent with previous observations of the role of the teleost kidney in B-cell development and in antibody production (see Teleort Lymphoid Orgens, P. 452). In the pronephros, Igm-positive cells form clumps, reminiscent of
clusters of B cells in mammalian bone marrow (Jacobsen and Osmond, 1990). That B cells differentiate in the adult kidney is supported by finding that rag1 is also expressed in the pronephros and mesonephros (Willett et al., 1997a; Danilova and Steiner, 2002). In the intestine, Igm-postive cells were found in the lamina propria, often in aggregates that assume a follicular shape (Danilova and Steiner, 2002). Igm-positive and rag1-positive cells were seen in the mesentery along the intestine, near vessels, apparently commingled with pancreatic tissue. The zebrafish spleen contains mostly red pulp, but some Igm expression is also seen, especially in older fish (N.D. amd L.A.S., unpublished). Substantial numbers of Igm-expressing cells were found in the thymus, mostly in the cortex (Danilova and Steiner, 2002).
GENETIC APPROACHES Genetic Screens The purpose of screens for mutant phenotypes is to identify genes that were previously unknown (“forward genetics”, i.e., phenotype to genotype). The main methods that have been used for generating mutations in large-scale genetic screens of zebrafish are: 1) chemical mutagenesis with ethylnitrosourea and 2) the insertion of exogenous DNA in the form of a virus. Diploidy is a major obstacle to the genetic analyses of zebrafish or any higher organism. Streisinger introduced techniques for producing homozygous diploid fish, which carry an induced mutation on both chromosomes, thereby facilitating the detection of recessive mutations (Streisinger et al., 1981). However, homozygosity has associated difficulties (see below), and mutations can also be detected by inbreeding, as has been done in many recent studies. The initial large-scale screens of chemically mutagenized fish identified mutations in genes affecting many aspects of embryonic development (Development, 1996, vol. 123). These screens were designed to examine organs that form within the first few days of development, mainly by visual assessment of the embryo. About fifty genes had been cloned by early 2002 (Mullins, 2002), most by a candidate gene approach, which is less arduous than positional cloning. The anticipated completion of the zebrafish genomic sequence should greatly facilitate the cloning of other mutant genes. The laboratory of Nancy Hopkins has recently completed a large-scale screen of mutants generated by the insertion of a genetically engineered retrovirus into genomic DNA (Amsterdam et al., 1999; Golling et al., 2002). Although insertional mutagenisis is inefficient relative to chemical mutagenesis, and yielded fewer mutants, the genes are much more easily cloned as they are tagged by the inserted DNA. In a report in June 2002, the genes for 75 mutants were
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described; about 1/5 were in genes whose biochemical functions are not known (Golling et al., 2002). It is estimated that by the time the analysis of the results of the insertional screen are complete, about 450 mutated genes will have been identified. Mutations Related to the Immune System In the last few years, several genes related to the zebrafish immune system have been characterized and provide a framework for screens seeking to identify immune system defects. To identify mutations that may pertain to the immune system, one can either examine known mutants having phenotypes related to immunity or carry out a new screen with the primary purpose of identifying such phenotypes. Characterization of Known Mutants Mutations in hematopoiesis may affect lymphopoiesis. Since RBCs are visible in the circulation beginning at about 24 hpf, deficiencies in their number or morphology can be observed easily. Two spontaneous mutations, spadetail (spt) (Kimmel et al., 1989) and cloche (clo) affect both primitive and definitive hematopoiesis (Fouquet et al., 1997; Liao et al., 1997; Thompson et al., 1998; Kalev-Zylinska et al., 2002; Lieschke et al., 2002). The cloche mutation, which functions upstream of spadetail, also affects the development of vascular endothelial cells (Stainier et al., 1995; Thompson et al., 1998). Another spontaneous mutation, bloodless (bls), affects primitive erythropoiesis, but not early macrophage development; initiation of lymphopoiesis and rag1 expression in the thymus is delayed in bls mutant embryos (Liao et al., 2002). Although all of these mutations affect hematopoiesis, none of them is specific to lymphoid lineages. A number of mutations in hematopoiesis were identified in the screens of chemically mutagenized fish (Ransom et al., 1996; Weinstein et al., 1996). Eight mutants having either no or few RBCs in the circulation were evaluated for the presence of lymphocytes in the thymus by whole-mount in situ hybridization with a rag1 probe (C. E.W., unpublished results). All displayed wildtype staining, indicating that lymphopoiesis was not affected. Since the stroma of the thymus is derived from the pharyngeal endoderm, mutations affecting development of the jaw and pharynx (Piotrowski et al., 1996; Schilling et al., 1996) might be expected to disrupt formation of the thymus. The jaw is a prominent structure, and underdevelopment or malformation is easily visualized at day 4 by staining cartilage with Alcian blue. Several jaw mutants were found not to express rag1 in the pharyngeal region, presumably because the thymus was missing (compare Figure 29.5A and 29.5B) (C.E.W. and T. Piotrowski, unpublished results). A detailed analysis of the mutant phenotypes suggests that
rag1 expression in the thymus correlates with complete development of the third arch. Those mutants with no pharyngeal rag1 expression also have defects in many structures of the jaw and are embryonic lethal. In a screen designed to identify genes that regulate anterior–posterior patterning in the heart field, a mutant, casanova, was found to lack endoderm (Alexander et al., 1999). This mutant does not express Foxn1 (whnb), a gene that is expressed in the epithelial component of the thymus in wildtype larvae (Schorpp et al., 2002). However, the mutant is deficient in all endodermal derivatives, and the extensive defects are not confined to the pharyngeal endoderm. Primary Screens for Mutations Affecting Lymphocyte or Thymus Development The mutants described in the preceding section had initially been identified as having defects in hematopoiesis or pharyngeal development. Since these phenotypes might also include deficiencies in lymphopoiesis or thymus formation, some of the mutants were also screened for rag1 expression in the thymus. A more directed approach would be to carry out a primary screen for mutants in rag1 expression. To evaluate feasibility, ninety insertionally mutagenized fish were screened (C.E.W. and J. J. Cherry, unpublished results). Two mutants having no rag1 expression were identified. These were grossly deficient in jaw morphology and were identified independently in the large-scale screen of insertionally mutagenized fish because of this phenotype (Allende et al., 1996; Gaiano et al., 1996). In each case, the gene mutated by the insertion was cloned; loss of nar results in a complete absence of pharyngeal arches and loss of pes results in a very small head and reduced arches. Other primary screens for lack of rag1 expression in the thymus are in progress utilizing chemically mutagenized fish (Trede and Zon, 1998; Schorpp et al., 2000). These screens have taken an approach different from that used in the large chemical screens, in which mutants were brought
FIGURE 29.5 Lack of rag1 expression in pharyngeal region in jaw mutant. Several chemically induced jaw mutants were screened for expression of rag1 by whole-mount in situ hybridization on day 6. (a) Wildtype fish shows rag1 expression in thymus; (b) Mutant tn20c shows no rag1 expression in region of thymus. Arrowheads indicate position of thymus. (Most of the dark material is pigment; oval objects are eyes).
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to homozygosity by inbreeding to the F3 generation. Instead, homozygous diploid fish are screened. Briefly, eggs from mutagenized females are mixed with UV-inactivated wildtype sperm to initiate development; the fertilized eggs are then briefly subjected to high pressure to inhibit the second meiotic division, thus generating diploid embryos (Streisinger et al., 1981). The benefit of this approach is that recessive mutations can be detected already in the F2 generation, with a consequent substantial savings in time and space for housing fish. There are, however, several drawbacks: 1) loci distant from the centromere may be heterozygous; 2) expression of defective alleles unrelated to the induced mutation, or epigenetic effects, may obscure the identification of a mutant phenotype, and 3) homozygous fish may develop poorly and have limited viability. A number of mutants deficient in rag1 expression in the thymus have been identified, and cloning of the mutated genes is in progress (Trede et al., 2001). Target-Selected Mutations To identify the function of a known gene in mice it is possible to introduce targeted mutations into the germline by utilizing embryonic stem cells (“reverse genetics,” genotype to phenotype). Such cell lines are not available in zebrafish and gene replacement has not yet been effected in this species, although efforts to accomplish this are under way (Ma et al., 2001; Lee et al., 2002). However, an alternative method for inactivating any gene whose sequence is known, target-selected mutagenesis, has recently been put into practice (Wienholds et al., 2002). In this procedure, mutagenized male fish are crossed with wildtype females. Genomic DNA is isolated from male progeny and screened for mutations in a specific gene by nested PCR-amplification of the target gene and subsequent DNA sequence analysis. Sperm from the same male fish are cryopreserved to allow recovery of the mutation. In vitro fertilization followed by inbreeding is performed to recover a line carrying the homozygous mutation. The rag1 gene was used to test the method; fifteen rag1 mutants out of 2,679 mutagenized F1 fish were detected by resequencing. Fish homozygous for one of these mutations, which introduced a stop codon, were found to be deficient in V(D)J recombination (Wienholds et al., 2002). Surprisingly, the rag1-deficient fish reach adulthood in a normal fish facility and are fertile, although perhaps more fragile than wildtype fish. They may be a useful resource for studies of infection and innate immunity in zebrafish. Using this technique, it is possible to isolate mutations in any gene of interest.
Transgenesis Transgenic fish lines can be created to express a marker, such as green fluorescent protein (GFP) or b-galactosidase,
under the regulation of specific promoters. The procedure for generating transgenic zebrafish is well established (Westerfield et al., 1992; Lin et al., 1994). Briefly, plasmid DNA is injected into the cytoplasm of embryos at the one- or twocell stage, and the progeny are screened for inheritance of the transgene. The insertion of a reporter such as gfp into a gene of interest simplifies the screening for progeny carrying the transgene since GFP is visible in living fish. Transgenic lines using organ-specific promoters expressing GFP have been generated for hematopoietic cells (GATA-1, GATA-2) (Long et al., 1997; Meng et al., 1997), lymphocytes (rag1) (Figure 29.4B) (Jessen et al., 1999), pancreas (insulin) (Huang et al., 2001), central nervous system (a1 tubulin) (Goldman et al., 2001), blood vessels (Tie-2, Fli1) (Motoike et al., 2000; Lawson and Weinstein, 2002), and platelets (R. Handin, Brigham and Women’s Hospital, Boston, personal communication), among others. There are several applications for such transgenic lines, for example, to study gene regulation, normal organ development, and the effect of drugs on development. They can also serve as a convenient alternative to in situ hybridization in genetic screens. Expression of a transgene bearing a fluorescent marker can be used to isolate specific cell types by sorting. For example, a gene involved in regulating erythropoiesis was identified by screening a cDNA library made from GATA-1/GFP expressing cells purified by fluorescence-activated cell sorting (Long et al., 2000a). Transgenic lines can also serve as disease models, particularly for diseases caused by dominant, single gene mutations. A transgenic zebrafish overexpressing a transcript encoding a human translocation associated with myeloid leukemia (RUNX1-CBF2T1) displays a phenotype similar to the human disease (Kalev-Zylinska et al., 2002). Such zebrafish lines will be valuable in understanding the developmental biology and genetics of some human diseases.
Morphants Although gene replacements and targeted deletions are not yet feasible in zebrafish, it is possible to prevent the formation of a specific gene product by degrading mRNA or interfering with translation. One method utilizes stable synthetic antisense oligonucleotides, called morpholinos (a morpholine moiety replaces ribose), as sequence-specific inhibitors of translation (Nasevicius and Ekker, 2000). An entire issue of Genesis (Volume 30, Issue 3, 2001) was devoted to this approach. A morpholino knockdown was used to demonstrate the function in hematopoiesis of the zebrafish homolog of runx1/aml1/cbfa2. The zebrafish gene, in contrast to the mouse gene, appears to be required for primitive as well as definitive hematopoiesis (KalevZylinska et al., 2002). Similarly, a morpholino knockdown was used to show that Biklf, a Kruppel-like transcription
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factor, is required for primitive erythropoiesis (Kawahara and Dawid, 2001).
Characterizing Gene Expression Patterns Large-scale screens of cDNA expression can be used to identify genes having temporal or spatially regulated expression patterns. The laboratory of C. and B. Thisse is developing a gene expression database with the aim of describing patterns of gene expression during embryogenesis. To date, over 12,000 cDNAs have been analyzed and 3,200 patterns of expression identified, including genes expressed within the vasculature, the ICM, and germ cells (Thisse and Zon, 2002, B and C. Thisse, personal communication). Another application is to identify genes preferentially expressed in an organ by techniques such as subtractive hybridization. Using this approach, an endogenous retrovirus, strongly expressed in the zebrafish thymus, has been identified (Shen and Steiner, 2004).
MAJOR HISTOCOMPATABILITY COMPLEX (MHC) Molecules encoded by the MHC are essential for the function of the adaptive immune system and are found in all jawed vertebrates. Studies of the MHC of zebrafish and other teleosts, largely by the group of Jan Klein, have been informative about the evolution of the genes in this region. In tetrapods, MHC class I and class II regions are closely linked (Trowsdale, 1995), but in teleosts the class I and II loci are in different linkage groups (Bingulac-Popovic et al., 1997; Sato et al., 2000). Therefore, the term “complex” to encompass these loci is a misnomer in these species. Indeed, in zebrafish, class II genes are situated on at least three different chromosomes (although only one appears to contain functionally active genes) (Graser et al., 1998; Kuroda et al., 2002). In two different genera of shark, representatives of cartilaginous fish, the class I and II clusters are linked, as in tetrapods, thus suggesting that the lack of linkage in teleosts is not an ancestral, but a derived characteristic (Ohta et al., 2000). Evidence has recently been presented in support of the hypothesis that the separation of the class I and II loci occurred by translocation of the class II locus to another chromosome in a teleost ancestor (Kuroda et al., 2002). Linkage relationships in the zebrafish genome have helped to clarify the puzzle posed by the location, in tetrapods, of the MHC class III genes (e.g., encoding C2, C4, tumor necrosis factor (TNF) a , steroid 21-hydroxylase), which are situated between the class I and class II linkage groups, but are unrelated in structure to the genes in these regions (see discussion by Klein and Sato, 1998). The puzzle was whether the location of these genes indicates a related function (as implied by the term “class III”) or is acciden-
tal. In zebrafish, genes encoding C4 are not linked either to class I or class II genes, strongly supporting the view that their presence in the MHC region of tetrapods is accidental and without functional significance (Samonte et al., 2002). A number of genes involved in the generation, transport, and loading of peptides onto class I molecules (e.g., LMP or PSMB, TAP), are located in the class II region in mammals, an arrangement that seems paradoxical because these genes are needed for the function of MHC class I, not class II genes. In zebrafish, these genes are instead closely linked to class I genes (Takami et al., 1997; Michalova et al., 2000). Presumably, an inversion in the evolution of tetrapods has placed these genes in the class II region (Kuroda et al., 2002).
INNATE IMMUNITY The innate immune system is a form of host defense found in all multicellular organisms. For jawed vertebrates, innate immunity is critical in providing resistance to pathogens in the early period after infection, before the adaptive system comes into play, and it provides the effector mechanisms required for full implementation of an adaptive response. Exposure to antigen in an adaptive response leads to the large-scale amplification of cells expressing receptors that bind specifically to that antigen. The receptors are generated by the rearrangement of genes encoded in the germline and are clonally distributed. In contrast, innate immunity depends on the immediate function of a smaller repertoire of constitutive “pattern recognition receptors” encoded by genes that do not rearrange and that recognize conserved moieties present on microbial surfaces but not in higher eukaryotes (Janeway and Medzhitov, 2002). When innate receptors such as Toll-like receptors (TLR), mannose-binding lectin (MBL), and scavenger receptors encounter conserved pathogen-associated molecular patterns such as lipopolysaccharide, peptidoglycans, and mannans, the host responds by upregulating genes encoding inflammatory cytokines and antimicrobial peptides. Among the cells that bear innate immune receptors are monocytes/macrophages, granulocytes (neutrophils, eosinophils, and basophils), dendritic cells, and natural killer (NK) cells. Macrophages and some granulocytes are found in zebrafish, but dendritic cells or NK cells have not yet been identified. The innate system is necessary not only to provide protection before the adaptive response is launched, but also for interplay with the adaptive system. For example, an antibody’s V region (adaptive) can bind determinants on the surface of a pathogen, while its C region binds to Fc receptors (innate) on phagocytic cells, thus facilitating the ingestion and subsequent killing of the pathogen. Alternatively, antibody binding to a pathogen can trigger the complement cascade, leading to the lysis of the pathogen, as well as
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the release of molecules that promote inflammation and phagocytosis.
Cellular Effectors of Innate Immunity Phagocytic (Myeloid) Cells The developmental biology of zebrafish myeloid cells has been reviewed recently (Crowhurst et al., 2002). The transparency of the zebrafish embryo has been a particular advantage in studies of macrophage development utilizing video-enhanced differential interference contrast microscopy and lineage tracing of cells bearing visible markers (Herbomel et al., 1999). Using these tools, it was possible to show that early macrophages arise in the anterior part of the embryo, at a site distinct from the posterior site, from which erythroid cells arise. Macrophage progenitors were recognized already at the 13-somite stage (15 hpf) when they emerge from the ventro-lateral mesoderm anterior to the cardiac field; by 24 hpf, they have migrated to the yolk sac, where they differentiate. Some of the macrophages invade the mesenchyme of the head, while others enter the blood circulation. Dissemination of these macrophages occurs before any other leukocytes appear in the embryo. These early macrophages migrate from the mesenchyme to epidermis, retina, and brain (Herbomel et al., 2001). To test the functional capacity of the early macrophages, bacteria were injected into embryos at about 24 hpf, well before functional lymphocytes appear (Herbomel et al., 1999). The bacteria adhered to the macrophages and were rapidly cleared from the circulation. After phagocytosis of bacteria, the macrophages showed signs of activation to the point of engulfing healthy erythroblasts. Evidently, before adaptive immunity comes into play, phagocytic cells may provide protection against bacterial infection; they also appear to eliminate apoptotic cells by phagocytosis (Herbomel et al., 2001). Evidence for apoptotic activity of macrophages had been observed in the pronephros at 2 weeks (Willett et al., 1999). Early macrophages in the mesenchyme of the head, which differentiate before the blood circulation is established, have also been identified in mammals, birds, and Xenopus (Ohinata et al., 1990; Cuadros et al., 1992; Lichanska and Hume, 2000). As in zebrafish, these early macrophages do not pass through a monocytic stage in their development, and they retain proliferative potential, characteristics that differentiate them from macrophages appearing at later developmental stages, (reviewed by Shepard and Zon, 2000). The initial wave of macrophages appears to follow a developmental program that is separate from erythropoiesis, and is also different from later (definitive) myelopoiesis in all vertebrates examined (Figure 29.3) (Lichanska and Hume, 2000; Herbomel et al., 2001).
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Myeloid commitment was evaluated by examining the expression of genes in this lineage. The early macrophages express the zebrafish homolog of the gene encoding the leukocyte-specific actin-binding protein, L-plastin (Herbomel et al., 1999; Bennett et al., 2001; Herbomel et al., 2001). The zebrafish ortholog of c-fms, encoding the receptor for macrophage colony-stimulating factor, was also expressed in cells of the early macrophage lineage (Parichy et al., 2000; Herbomel et al., 2001), as it is in mice (Lichanska et al., 1999). Macrophages from fish mutant in this gene (Parichy et al., 2000) behave normally initially, but fail to emigrate from the yolk sac to colonize embryonic tissues (Herbomel et al., 2001). At about 60 hpf, coincident with a decrease in the expression of L-plastin, macrophages in the brain and retina undergo a phenotypic transformation into cells designated “early microglia,” which express high levels of apolipoprotein E (Herbomel et al., 2001). Additional evidence for the early anterior site of myeloid differentiation in zebrafish was obtained from the expression of pu.1, encoding a member of the Ets family of transcription factors (Lieschke et al., 2002), which, in mice is known to be required for the differentiation of myeloid and lymphoid, but not erythroid, cells (Scott et al., 1994; McKercher et al., 1996). The pu.1 gene was expressed in the anterior site at 12 hpf, before any expression was detected in the posterior part of the zebrafish embryo (Lieschke et al., 2002). The distinction between the anterior “myeloid site” and the posterior “erythroid site” was supported by analysis of a mutant, spadetail, in which pu.1 was expressed only in the anterior site. Moreover, myeloid cells but no erythroid cells were found in this mutant (Lieschke et al., 2002). Fate mapping with a fluorescent marker also indicated that cells labeled in the anterior pu.1-expressing site developed characteristics of macrophages, whereas cells labeled in the posterior hematopoietic site often ended up as circulating erythrocytes (Lieschke et al., 2002). Two types of granulocytes have been described in zebrafish, a neutrophil (heterophil) and another type, variously described as an eosinophil (Jagadeeswaran et al., 1999; Lieschke et al., 2001) or a basophil/eosinophil (Bennett et al., 2001). Cells with the morphology of granulocytes are seen in the kidney and blood (Al-Adhami and Kunz, 1977; Jagadeeswaran et al., 1999; Willett et al., 1999). Neutrophilic granules in zebrafish, like those in mammals, contain a homolog of myeloperoxidase, which was first seen to be expressed in the posterior ICM beginning at 18 hpf (Bennett et al., 2001; Lieschke et al., 2001). In 6-day larvae, granulocytes accumulate at a wound site, suggesting functional capability (Lieschke et al., 2001). Cytotoxic Cells Cytotoxic cells, both cytotoxic T lymphocytes (CTLs) and natural killer (NK) cells, play important roles in immune
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defense. These two types of cells share many properties, such as certain membrane markers and effector mechanisms, and it is thought that they are derived from common lymphoid progenitors. However, CTLs express rearranged antigen-specific receptors (i.e., TCR) whereas NK cells have nonrearranging receptors. CTL are effectors of the adaptive immune system. NK cells are essential elements of the innate immune system, although they do also interact with the adaptive system, and some NK cells are dependent on antibodies for their activity. No cytotoxic cells have as yet been described in zebrafish, but a variety of studies on cytotoxicty have been carried out with other teleosts, including the cloning of cytotoxic cell lines (reviewed by Shen et al., 2002). A receptor whose ligation triggers cytotoxic activity has been cloned in catfish, and its probable ortholog has been identified in zebrafish (Jaso-Friedmann et al., 1997, 2002). The activity of NK cells is regulated by a balance of signals from activating and inhibitory receptors. During infections by viruses and other intracellular pathogens, NK cells are activated by cytokines, such as interferons a/b or interleukin-12, and respond by producing g-interferon and those other cytokines needed to control the infection (reviewed by Lanier, 1998). A key to understanding NK function was the observation that killing is directed only against cells that have altered or absent expression of major histocompatability (MHC) genes, for example due to transformation by oncogenes or viruses. A number of types of receptors have been identified on both human and mouse NK cells that, upon recognition of class I MHC molecules, transmit signals that prevent target-cell killing. The killer cell inhibitory receptors (KIR) (reviewed by Long, 1999; Ravetch and Lanier 2000; Barten et al., 2001), are glycoproteins having two or three extracellular Ig-like C2 domains with features of both V and C domains (Williams, 1987) and cytoplasmic tails with two immunoreceptor tyrosine-based inhibition motifs (ITIMs). Many of these receptors are not highly conserved in evolution, perhaps because of variability in their ligands, and this may lead to difficulty in identifying them in lower vertebrates. Until recently identified only in primates, KIR have now been described in bovines and rodents (McQueen et al., 2002; Hoelsbrekken et al., 2003). A family of receptors, designated novel immune-type receptors (NITR), has recently been identified in the pufferfish, Spheroides nephelus, and in zebrafish. These bear some resemblance both to antigen-specific receptors and to the KIR. The NITR typically have two extracellular Ig-like domains, one V and one designated V/C2, and an ITIMcontaining cytoplasmic tail (Rast et al., 1995; Strong et al., 1999; Litman et al., 2001; Yoder et al., 2001). Evidence for possible function of these genes was obtained by demonstrating that cross-linking of an epitope-tagged NITR transfected into a human NK cell line inhibited signaling in
the presence of a tumor target (Yoder et al., 2001). Ligands binding to the NITR in vivo have not been identified as yet. The NITR are an interesting addition to the large numbers of cell surface molecules, many on lymphoid and myeloid lineages, having Ig-like domains, and in some cases, also ITIM-like (or related activating) motifs (Barclay et al., 1997; Ravetch and Lanier, 2000).
Complement The complement system plays an important role in innate as well as adaptive immunity. The complement cascade can be activated in three ways: 1) the classical pathway, initiated by antigen–antibody complexes, and by two antibody independent pathways; 2) the alternative pathway; and 3) the lectin pathway. The main functions of the complement system are to promote phagocytosis, initiate inflammation, and lyse pathogens. The three complement activation pathways appear to be present in teleosts (reviewed by Holland and Lambris, 2002). The lectin pathway resembles the classical pathway except that antibody is not required for initiation. Instead, the pathway is triggered, as part of an innate response, by the multivalent interaction of a collectin, MBL, or a ficolin (see next section) with sugar arrays on microbial surfaces (reviewed by Matsushita and Fujita, 2001; Fujita, 2002; Wallis, 2002). MBL and ficolins associate in the serum with MBL-associated serine proteases (MASPs). One of these, MASP-2, resembles C1s both structurally and functionally, and is responsible for the cleavage of C4 and C2. cDNAs encoding homologs of mammalian MBL have been isolated from three members of the cyprinid family: zebrafish, carp, and goldfish (Vitved et al., 2000). Interestingly, amino acid residues associated with binding to mannose or glucose are replaced in these teleosts by residues that are associated in other lectins with binding to galactose (Drickamer, 1992). Perhaps selective pressures on teleosts from pathogens bearing galactose-rich surface molecules account for this structural feature (Vitved et al., 2000). Genes encoding molecules in the MASP/C1r/C1s family have been identified in carp, consistent with the presence of the lectin pathway in teleosts (Endo et al., 1998; Nakao et al., 2001). In mammals, components C2 of the classical pathway and Bf of the alternative pathway are similar in structure, gene organization, and function, playing similar roles in the classical and alternative pathways, respectively. The genes encoding these proteins are closely linked in the MHC class III region, and it has been suggested that they arose by duplication from a common ancestor. Genes encoding homologs of Bf and C2 in mouse and humans have been identified in several teleosts, including zebrafish (Kuroda et al., 1996; Seeger et al., 1996; Gongora et al., 1998; Sunyer et al., 1998; Nakao et al., 2002). From sequence considera-
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tions alone, it has not been possible to assign these unambiguously to either C2 or Bf. Interestingly, one of the two identified Bf/C2 molecules in trout has been shown to function in both the alternative and classical pathways (Sunyer et al., 1998). This observation raises the possibility that the gene duplication leading to distinct C2 and Bf proteins may have occurred after the emergence of teleosts in vertebrate evolution. C3, which can react with numerous cell-surface and serum proteins, is a key component shared by the three complement pathways. Although a single gene encodes C3 in mammals, multiple forms of C3 are present in teleosts, apparently due to gene duplication as well as polymorphism. These isoforms may vary in binding efficiency for diverse complement-activating surfaces, the C3 repertoire contributing to immune defense against a range of pathogens (Sunyer et al., 1996; Sunyer and Lambris, 1998). Three genes encoding C3 have been identified in zebrafish; phylogenetic analysis suggests that the duplication events that produced the three zebrafish C3 loci took place after the separation of the lineages leading to the trout and zebrafish (Gongora et al., 1998). Genes encoding C4 have also been identified in zebrafish and, unlike C4 genes in other species, are linked to genes encoding a2-macroglobulin (A2M), a component of the innate immune system that clears proteinases from tissue fluids (Samonte et al., 2002). The linkage of these genes in zebrafish supports the possibility that they originated by tandem duplication. The components C3, C4, and C5, as well as A2M, show significant similarity in sequence and are believed to be encoded by genes that diverged from a common ancestor (Sottrup-Jensen, 1987). C5 has been identified in trout but not yet in zebrafish (Nonaka et al., 1981; Franchini et al., 2001). The components C6 to C9 also remain to be identified in zebrafish.
Pattern Recognition Receptors The function of the innate immune system depends on receptors that recognize conserved molecular patterns found on microbial surfaces, but not in higher eukaryotes. Two groups of proteins capable of recognizing such patterns are the collectins (e.g., MBL) and ficolins. Present in plasma and on mucosal surfaces, these lectins bind carbohydrate structures on pathogens, leading to complement activation and/or phagocytosis (Lu et al., 2002; Holmskov et al., 2003). Collectins and ficolins both have collagen-like N-terminal domains, but they differ in the type of C-terminal lectin domain. In tertiary structure, they resemble the complement component C1q. To date, the only member of these families described in teleosts, including zebrafish, is MBL (Vitved et al., 2000). Pattern recognition receptors that have been the subject of much recent interest are the TLR, which are involved in
innate defense in most multicellular organisms (Medzhitov, 2001). In mammals, there are at least ten of these receptors, and each appears to have a distinct function. The ligands for these receptors are quite diverse, but most are conserved microbial products such as lipopolysaccharide, doublestranded RNA, flagellin, and peptidoglycan. The TLR are transmembrane proteins with an extracellular leucine-rich repeat domain and a cytoplasmic domain, also present in the cytoplasmic tail of the type I receptor for interleukin-1 (IL1), which induces signaling through the NF-kB pathway. Ligand binding leads to the production of molecules that function in host defense, such as inflammatory cytokines, costimulatory molecules, and antimicrobial peptides. It has been only a few years since the initial description of the mammalian TLR (Medzhitov et al., 1997), and work on these receptors in other vertebrates is just getting underway. Several entries in the GenBank database have been identified as zebrafish TLR. There are also entries for salmon and flounder TLR. In addition, there is an entry for a zebrafish cDNA similar to Toll interacting protein.
Cytokines A great variety of cytokines (proteins made by cells that affect the behavior of other cells) have important roles in innate as well as adaptive immunity. Considerable progress has been made recently in the cloning of homologs of mammalian cytokine genes in teleosts; the data up to 2001 have been reviewed (Magor and Magor, 2001). A number of the teleost cytokine genes are present in two copies, probably related to gene duplication that has occurred in this lineage (Postlethwait et al., 1999, 2000). Interleukins (IL) are cytokines synthesized by leukocytes, especially blood monocytes and tissue macrophages. Members of the IL-1 family and their receptors are important components of mammalian inflammatory responses. Genes encoding homologs of mammalian IL-1b have been identified in a number of teleosts, including zebrafish (reviewed by Bird, 2002). cDNA encoding a full-length type I IL-1 receptor, including the characteristic TLR signaling domain, has been cloned in salmon (Subramaniam et al., 2002). Several database entries suggest that other interleukins and receptors are present in zebrafish. Tumor necrosis factor alpha (TNF) and lymphotoxin (LT) are cytokines, initially identified as products of macrophages and lymphocytes, respectively, that cause lysis of certain types of cells, especially tumor cells. They and their receptors are now known to be members of a large family with roles in inflammation, host defense, apoptosis, autoimmunity, and organogenesis (Locksley et al., 2001). Recently, homologs of TNF were cloned in flounder and trout (Hirono et al., 2000; Zou et al., 2002). Some TNF receptors have “death domains,” whose interactions with different cytoplasmic signaling proteins can lead to apoptosis or to induc-
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tion of transcription factors c-Jun and Nf-kB. A zebrafish homolog of a TNF receptor containing a death domain has been cloned (Bobe and Goetz, 2001). Interferons are cytokines that were initially identified by their ability to interfere with viral replication. Type I interferons include interferon a, originally known as leukocyte interferon (actually a family of proteins) and interferon b, originally known as fibroblast interferon. They are induced upon viral infection or exposure to double-stranded RNA and, by reacting with a shared receptor, initiate synthesis of a variety of anti-viral proteins. The only well characterized interferon cloned in any lower vertebrate to date is a type I interferon in zebrafish, which was shown to have anti-viral activity (Altmann et al., 2003). Type II interferon (interferon g) is produced by NK cells and some T cells, has numerous activities, especially stimulation of macrophages, but has not been cloned in lower vertebrates. Chemokines are small cytokines that regulate chemotaxis. They are synthesized by a variety of cells types, often macrophages or T cells, and are divided into sets according to structural features, particularly the spacing of Cys residues that form (usually) two disulfide bridges. Some chemokines may have roles in developmental processes, as described for zebrafish brain (Long et al., 2000b). Chemokine receptors are members of the rhodopsin superfamily, having seven transmembrane segments and signaling through coupled G proteins. A considerable number of homologs of mammalian chemokines and their receptors have been cloned in teleosts (reviewed by Magor and Magor, 2001). Recently, homologs of IL-8, the first known chemokine, have been cloned in flounder and trout (Lee et al., 2001; Laing et al., 2002).
INFECTION Viral, bacterial, and fungal infections of zebrafish have been described, but few studies have been carried out on the role of the immune system in infection. In general, fish that are stressed by poor environmental conditions are susceptible to infection, and many infections are due to opportunistic organisms. The infections that are known to occur in zebrafish, as well as their prevention and treatment, have been reviewed (Astrofsky et al., 2002a). Important routes for pathogen entry in fish are the gills and gastrointestinal tract, but generally not the skin (Evelyn, 1996). Infection with Mycobacteria is the most common chronic disease affecting tropical fish in aquaria and has been a major problem in zebrafish facilities. Three species of Mycobacteria were isolated in one study of laboratory infection (Astrofsky et al., 2000). Infection can involve multiple organs and, once established, is very difficult to eradicate. Chronically infected fish grow poorly and do not reproduce well. Traditionally, diagnosis is by culture of the
organism, which is a lengthy procedure because of their slow growth, and by histology. Diagnosis has also been accomplished by PCR with primers based on the sequence of bacterial ribosomal 16S RNA (Talaat et al., 1997, Astrofsky et al., 2000). As noted earlier, phagocytosis of injected bacteria can be observed in zebrafish embryos at about 24 hpf, before any lymphocytes are present. It was subsequently shown that injection of embryos with Mycobacterium marinum causes a systemic infection that includes the aggregation of macrophages into granuloma-like structures. These resemble granulomas formed in adults, both in histological features and in ability to activate certain macrophage genes (Davis et al., 2002). The migration of macrophages in response to infection is unaffected in fish mutant in c-fms, although their migration during development is severely affected (see Phagocytic (Myeloid) Cells, P. 461). Interestingly, in the presence of infection, macrophages that have been transformed into early microglia appear to leave the brain and are redirected to the site of infection (Davis et al., 2002). Other pathogens causing disease in zebrafish are protozoa such as Tetrahymena, the related Ichthyophthirius multifiliis, and a variety of trichodinids (Astrofsky et al., 2002a, b). Fungi can cause opportunistic disease in zebrafish, and nematode and dinoflagellate infections are common (Dykstra et al., 2001; Astrofsky et al., 2002a). Experimental infections with streptococci and Listeria have been studied (Menudier et al., 1996, Neely et al., 2002). Infectious hematopoietic necrosis virus and infectious pancreatic necrosis virus cause diseases of major importance in salmon and trout (Wolf, 1988; LaPatra et al., 2000). In a study of experimental infection with these viruses in zebrafish, they were found to cause transient toxicity in hematopoietic precursors and terminally differentiated red cells; however, the fish recovered quickly (LaPatra et al., 2000). To date, there have been few if any studies specifically directed to the role of adaptive immunity in combating infections in zebrafish. The availability of fish mutant in rag1, and therefore lacking mature B and T lymphocytes (see Target-Selected Mutations, P. 459), may be useful in such studies.
CONCLUDING REMARKS: ZEBRAFISH AND HUMAN DISEASE In the years since a genetic analysis of zebrafish development was first proposed, many studies have confirmed the feasibility of this approach and its broad applicability to understanding the principles of vertebrate biology. It has become increasingly appreciated that similar genes and pathways form a foundation for all organisms, invertebrate
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as well as vertebrate. The TLR pathway, initially discovered for its role in Drosophila development, is now known to function in innate immunity of both insects and mammals (Medzhitov, 2001). The role of the Hox genes in specifying shared features of a body plan is another prominent example (Akam, 1989). The work of Christiane Nüsslein-Volhard and Eric Wieschaus (1980) demonstrated that a mutational analysis of Drosophila development uncovered basic principles that also apply to vertebrates. The similarity of zebrafish, Xenopus, chick, and mouse in many aspects of embryogenesis was pointed out by Charles Kimmel, a colleague of Streisinger in the early work on zebrafish (Kimmel, 1989). It is therefore no surprise that many zebrafish mutations, especially those in organ development, resemble the gene defects that cause human disease. Zebrafish mutations that appear to be models for human diseases have been reviewed (Dooley and Zon, 2000; Shin and Fishman, 2002; Thisse and Zon, 2002). The ease of identifying defects in RBC formation has facilitated the identification of a large number of mutants in erythropoiesis. For example, the sauternes mutant has a microcytic hypochromic anemia (Ransom et al., 1996); positional cloning identified the affected gene as encoding an enzyme required in the first step of heme biosynthesis (Brownlie et al., 1998), which is defective in congenital sideroblastic anemia. Another mutant defective in heme biosynthesis resembles a human porphyria (Wang et al., 1998). The riesling mutant (Ransom et al., 1996), is characterized by spherocytosis and severe anemia; the affected gene, sptb, was identified by a candidate gene approach as the ortholog of a human gene encoding b-spectrin (Liao et al., 2000). Mutant phenotypes resembling cardiomyopathies and arrhythmias have been described (Shin and Fishman, 2002). Mutations affecting the kidney, gastrointestinal tract, brain, eye, and ear may also resemble human disorders. Recent screens have been designed to identify mutants in genes controlling cell cycle, cell proliferation, and apoptosis, functions that may be related to carcinogenesis (Amatruda et al., 2002). Transgenic zebrafish lines may be useful models for human disease, as discussed earlier. To date, only one zebrafish mutation is known to cause a specific defect in adaptive immunity, the rag1 mutation discussed in an earlier section. The mutant, a potential model for severe combined immunodeficiency, was not identified by a traditional phenotypic screen, but by screening mutated genomes for sequence changes in the rag1 gene. As mentioned previously, the cloning of genes whose mutation disrupts thymus development is ongoing in at least two laboratories (Trede and Zon, 1998; Schorpp et al., 2000). The zebrafish is also finding increasing use in drug discovery (Shin and Fishman, 2002). Many thousands of chemicals can be screened for their effects on living embryos. Small molecules that permeate the embryo may disrupt specific developmental pathways and, in some cases, lead to
phenotypes that resemble mutations (Peterson et al., 2000, 2001). Since the addition of chemicals is easily controlled, the effects on the timing of specific developmental events can be assessed. Chemical screens may be useful if more than one gene is involved in a particular function. The covalent combination of ligand and target may offer a route toward isolation of the target. The effects of chemicals on gene expression of mutated as well as wildtype embryos can be evaluated. In general, chemical screens can complement genetic screens in the dissection of developmental pathways and in providing models of human disease.
Acknowledgments We thank the NIH for support via grant R01 AI-08054. We also thank Drs. C. O’Farrelly and L.S. Lerman for reading the entire manuscript and providing many helpful comments.
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30 The Origin of V(D)J Diversification SUSANNA M. LEWIS
GILLIAN E. WU
ELLEN HSU
Program in Genetics and Genomic Biology, Hospital for Sick Children Research Institute, Toronto, Canada
Biology Department, York University, Toronto, Canada
Department of Physiology and Pharmacology, The State University of New York Health Science Center at Brooklyn, New York, USA
The vertebrate immune system can respond to a broad, ever-changing spectrum of pathogens, and V(D)J recombination plays a central role in achieving this stunning versatility. The evolutionary origin of V(D)J recombination and the selective forces that later pushed the process along are questions that have intrigued investigators ever since the discovery that Ig and TCR genes rearranged. The first part of this chapter focuses on the currently favored hypothesis that the V(D)J recombination machinery was brought into being through the “exaptation” (Gould and Vrba, 1982) of a DNA transposon. The second part discusses germline V(D)J recombination events taking place after exaptation and the possibility that such activity was instrumental in creating the different Ig and TCR loci. The third part of the chapter goes further back in time, to consider the nature of the ur-V gene, and more global questions about selective pressures for diversity that existed before V(D)J recombination was possible.
posase became the RAG1/2 recombinase. To begin the discussion of this version of the origin of V(D)J recombination it is well to define three concepts: the V(D)J transposon, lateral transfer, and exaptation. A DNA transposon is defined by its ability to hop from one DNA site to another (reviewed by Haren et al., 1999). The minimal competent (or “autonomous”) DNA transposon contains a transposase gene bounded by transposase recognition sites that are usually a part of larger inverted terminal repeats (ITRs; Figure 30.1). Although a transposon may contain other genes (Figure 30.2A), their main function requires just a transposase gene flanked by ITRs. In transposition, the transposase protein cuts at the outside of each ITR to liberate the element (and its own gene) from its former position in the host genome. The ends of the transposon, while still gripped by the transposase, are then inserted into a new piece of DNA that has been captured and brought into the transposase:DNA complex. The breakthrough observation that transformed the notion of a V(D)J transposon from speculation to testable hypothesis was the discovery that the V(D)J recombinase proteins RAG1 and RAG2 can carry out DNA transposition in vitro (Agrawal et al., 1998; Hiom et al., 1998). To be specific, these proteins acting on RSS motifs can excise a linear DNA segment, capture a new target DNA, and physically connect the RSS 3¢ ends into the new DNA site. “Horizontal” or “lateral” transfer endows one organism with the genetic material of another, according to processes (transduction, conjugation, transformation) that bypass the usual vertical transmission from parent to offspring (Bushman, 2002). In extreme cases, trans-kingdom jumps can occur, as exemplified by the genetic traffic occurring between Agrobacterium and plants that results in the for-
THE ALIEN SEED Somatic site-specific DNA rearrangement is such an incongruent developmental strategy in higher eukaryotes that many of the first papers describing the process speculated that the recombination apparatus may have had a transposon origin (Sakano et al., 1979; Siu et al., 1984). V(D)J recombination is thought to have arisen when a mobile element (we can call this the “V(D)J transposon”) integrated into a V-like gene in an early vertebrate, perhaps as long as 450 million years ago (Du Pasquier and Flajnik, 2003). Thereafter, the mobile element was dismembered: Its termini became the RSSs and the element-encoded trans-
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nition sites (the RSSs). The task RAG1/2 now performs in B- and T-cell differentiation is a radical departure from its original design. Tracing the twists and turns of the exaptation of the V(D)J transposon provides a case study that gets to the heart of key questions in molecular evolution. To anticipate the next big moves in this area, four main classes of evidence regarding the origin of V(D)J recombination are summarized below. These are: clues based on recognition site sequence, clues from “transposase” gene structure, clues involving transposase protein structure, and clues derived from overall mechanistic properties (Figure 30.2A,B).
Recombination Site Similarities
FIGURE 30.1 Top. An intact DNA transposon contains a gene (or genes) encoding a transposase bracketed by inverted terminal repeats (ITRs). The ITRs contain transposase binding sites and a conserved transposon-specific terminal sequence defining the cleavage position. For some transposons the ITRs are very small and the three features (ITR, binding site, and terminal sequence) are essentially superimposed. The “transferred strand” is indicated: Upon transposition, usually only the 3¢ ends of the cut out element are attached to the new DNA site by transposase (see also Figure 30.5) Bottom. The exapted elements after their conversion into the V(D)J recombination apparatus. RSSs may correspond to the ITRs or just the terminal portion thereof. The RAG1/2 genes may correspond to a two-component transposase.
mation of crown galls (Bushman, 2002). Horizontal transfer is suspected where family trees based upon one particular trait don’t make sense when compared to evolutionary relationships established in other ways (for a recent discussion, see Syvanen, 2002). In the case of V(D)J recombination, horizontal transfer is not invoked for this reason: There is no confusing reticulation in family trees. Instead the notion of horizontal transfer comes up as an explanation for the “sudden” appearance of the V(D)J recombination apparatus in vertebrates (Schluter et al., 1999). There are no identifiable precedents for the RAG1, RAG2 genes in any animal representing any radiation more ancient than cartilaginous fish (Du Pasquier and Flajnik, 2003; but see note added in proof). The lack of precedent forms, rather than any positive evidence for horizontal transfer, is the basis for invoking the possibility that the V(D)J transposon came into the vertebrate lineage from elsewhere, perhaps even from bacteria. “Exaptation” describes the process of using pre-formed components to create a new trait (Gould and Vrba, 1982; Brosius and Gould, 1992). This is in contrast to “adaptation,” where incremental improvements are selected to build a structure over time. Exaptation is an integral part of the story of the V(D)J recombination origins (Figure 30.1). It is quite clear that a “V(D)J transposon,” if it ever existed as such, is no longer intact—the genes encoding the elements transposase (RAG1/2) are no longer flanked by their recog-
The DNA motif that is manipulated by a site-directed recombinase provides a signature of the cognate enzyme. Family relationships between enzymes are reflected in the general layout and, to some extent, in the DNA sequence similarities of their recognition sites. For V(D)J recombination, the small nonamer-spacer-heptamer motif of an RSS (5¢ GGTTTTTGT-12/23-spacer CACTGTG) is sufficient to specify recombination with cleavage occurring just 3¢ of a heptamer-terminal G (Figure 30.3). Over the years, the recombination sites for many sorts of recombinases have been scrutinized for evidence of RSS-like features. To put the recombination site comparisons into context, it helps to first mention an example that doesn’t work out particularly well. Early on, there was a suggested parallel between the RSS sequence and the recognition site for HIN, an enzyme that conducts site-specific inversion of a DNA segment in Salmonella (Simon et al., 1980). One of the two interacting HIX sites recognized by the HIN recombinase, HIXL, can be aligned in a suggestive way with an RSS sequence (Figure 30.3). However, even though it’s possible to make a sequence match, the heptamer-like identities in HIXL lie outside the DNA sequences actually targeted by HIN. This implies that similarity between the HIX site and FIGURE 30.2 Selected anatomical and mechanistic features of DNA transposons. Characteristics that match those of RAG1/2-mediated transposition or V(D)J recombination are underlined (central column). Example transposons are chosen for being well studied structurally, biochemically, or by molecular genetics. Mu, Tn916, Tn5, Tn7, and Tn10 are bacterial elements; the rest are found in higher eukaryotes. Most of the transposons listed are “cut-and-paste transposons”; Mu is a bacteriophage that moves by co-integrate formation and Tn916 is a conjugative transposon. Information is from Beall and Rio, 1997; Bushman, 2002; Dawson and Finnegan, 2003; Handler, 2002; Haren et al., 1999; O’Keeffe et al., 1999; Plasterk et al., 1999; Rubin et al., 2001; Tsai and Schatz, 2003; Turlan and Chandler, 2000, or as cited specifically in the text. For some properties (such as in situ inversion) the reported behavior is atypical for the listed transposon. “Spacer type termini” means that left and right ends differ in the size of a spacer located between terminal and subterminal repeats, analogous to the different spacers between the heptamer and nonamer repeats in a 12- or 23-RSS.
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FIGURE 30.3 Comparison between the leftmost copy of the pair of sites targeted by the bacterial HIN invertase (HixL) and a 23-RSS. The recognized sequences for HIN and RAG1/2 are shaded (i.e., the “acctgtg” sequence in italics is not part of the HIN recognition site). Portions of HixL that bear matches to the nonamer and heptamer of an RSS are indicated by vertical striping. The difference in overall architecture is seen by comparing the cleavage sites (black triangles).
the RSS heptamer motif is fortuitous. Furthermore (see triangles in Figure 30.3), the overall organization of the HIXL recognition site and an RSS is different: HIN cleaves in the middle of the HIXL target sequence whereas the RAG proteins cut at the outside edge. By contrast, the similarity between RSSs and DNA sequences found at the termini of a nematode element called Tc6 is more provocative (Dreyfus and Emmons, 1991; Dreyfus and Gelfand, 1999). Tc6 belongs to a wellstudied eukaryotic DNA transposon superfamily called “Tc1/mariner.” Members of the Tc1 family are found in many different eukaryotes including fish, insects, worms, and mammals (Plasterk et al., 1999). The architecture of the transposon end is like that of an RSS in that transposaseassociated breaks occur at the edge, rather than in the middle of their targets. With respect to actual sequence similarities, the original publications pointed to a suggestive (though incomplete) five-of-seven identity to an RSS heptamer. The presence of a nearby (nonamer-like) A/T-rich stretch in Tc1 family members was also noted (Dreyfus and Emmons, 1991; Dreyfus and Gelfand, 1999) (Figure 30.4). The Tc6-to-RSS matches might seem tenuous because any RSS sequence terminating in “CTG,” as in the recombination site of Tc-6 and other Tc1 family members, is virtually nonfunctional for V(D)J recombination (Hesse et al., 1989). Also the spacing between heptamer-like and any nonamer-like sequences in the Tc1-type element ends is not very RSS-like. However mutability and variation of the recognition site is thought to be a requisite for the maintenance and spread of DNA transposons (Lampe et al., 2001). Therefore any extant descendant of a transposon thought to date back 450 million years would be expected to have retained little of the progenitor’s actual recognition sequence. The fact that some Tc1/mariner superfamily members (Figure 30.2) can have very small ITRs within the size range of an RSS, that there are at least some potential similarities in DNA sequence, and that the position of the cleavage site relative to the recognition motif for RSSs and
FIGURE 30.4 Comparison between transposon ends and an RSS. Shown are the terminal sequences at the right ends of the autonomous elements Tc1 (K01135) and Tc3 (from G.I.R.I at http://www.girinst.org/index.html) and the nonautonomous elements, Tc6 (X55356) and MsqTc3 (AF274037). All but MsqTc3 (Shao et al., 2001) are nematode elements. MsqTc3 has an almost perfect match to the RSS heptamer. Double underlining indicates a cut-site proximal GTG sequence important for RSS function. Open triangles show the position of the recombination site at the transposon ends. Strand breaks occur a few nucleotides within the recognition sequence on the 5¢ (nontransferred) strand, and exactly at the edge of the recognition sequence on the 3¢ (transferred) strand in Tc1 (Plasterk et al., 1999). The extended ITRs and binding sites for the Tc1-family transposases are not shown. Vertical strips indicate the regions positionally equivalent to nonamer and heptamer of an RSS. Identities are in boldface capital letters. The wavy underline shows the A-tract in Tc1 family members positionally equivalent to the T-tract nonamer element of a 23-RSS. A similar A-forT–substituted sequence, also indicated by wavy underlining, can act as a 23-RSS as in D-D joining at the chicken heavy chain locus. Neither right end nor left end comparisons reveal nonamer-like matches at the appropriate spacing for a 12-RSS (dashed box; left end not shown).
ITRs correspond, can be taken as encouraging evidence of a possible evolutionary connection. A fairly recently described member of the Tc1/mariner family from mosquitoes demonstrates that, in principal, conversion of a Tc1-type end to an RSS-like sequence is possible, or in other words, “you can get there from here” (Figure 30.4). The mosquito transposon MsqTc3 (belonging to the Tc3 subgroup of the Tc1/mariner superfamily) is different from other Tc1-type transposons in that it has an extra TG appended to its recognition site. This modification happens to greatly improve the proposed RSS similarity, creating a six-of-seven heptamer match (CACAGTG in MsqTc3; CACTGTG in an RSS). The new heptamer identity includes the critical GTG sequence, and RSSs with a MsqTc3 heptamer sequence are near-perfect in terms of function (Hesse et al., 1989). Moreover, in MsqTc3 an A-tract is located at a 23-bp spacing from the “heptamer.” This A-tract would constitute a noncanonical A for T substitution in the nonamer; however, such a noncanonical nonamer still has significant function in V(D)J recombination (Agard and Lewis, 2000). In a real immune system, in vivo recombination of an RSS with an A for T substituted nonamer appears
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to support D-to-D joining at the chicken IgH locus (Figure 30.4) (Ratcliffe and Jacobsen, 1994). In summary, although the Tc1-type element ends are not precisely like an RSS, there are examples that aren’t such a bad match: The indications of a Tc1/mariner connection are, if anything, more persuasive than might have been hoped for (Dreyfus and Emmons, 1991; Dreyfus and Gelfand, 1999).
Clues Based on the RAG1 and RAG2 Genes: Linkage and Structure The RAG1 and RAG2 genes are linked in animals representing a variety of taxa: chickens, rabbits, humans, mice, frogs, pufferfish, zebrafish, and sharks (Oettinger, 1992; Schluter and Marchalonis, 2003). For eukaryotes, the tight and conserved linkage of unrelated genes is highly unusual. In contrast, for bacteria, this type of arrangement is common, where many times the genes for proteins belonging to the same metabolic pathway can be found together in co-regulated operons. The unusual linkage between RAG1 and RAG2 is therefore suggestive of horizontal transfer from a prokaryotic source (Oettinger, 1992). More generally though, linkage may reflect the advantage gained by grouping any set of functionally related genes when these genes are disseminated by lateral transfer [the “selfish-operon” theory (Lawrence and Roth, 1996)]. Against lateral transfer possibilities (from either eukaryote or prokaryote) is the argument that the linked, tail-to-tail arrangement of the RAG1 and RAG2 genes instead evolved in eukaryotes to satisfy some regulatory necessity. To date, no evidence to support the latter possibility has been provided. Focusing in particular upon the RAG1 to RAG2 intergenic region, potential regulatory motifs have been identified by inspection, but none is confirmed experimentally (Bertrand et al., 1998; Peixoto et al., 2000). Along these lines, an interesting suggestion was that convergent transcription of the RAG genes in trout might allow an overlap of the 3¢ untranslated regions, and potentially, coordinate expression via an RNA antisense mechanism (Hansen and Kaattari, 1996). However, a similarly tightly packed RAG locus in zebrafish does not in fact template overlapping transcripts (Lily Changchien and Ellen Hsu, unpublished observations), so that even a ray-finned fish–specific role for RNA interference is questionable. Although promoter elements 5¢ to RAG2 may govern aspects of RAG1 gene expression in mice (Yu et al., 1999) (and by extension, possibly in other animals as well), this too seems to provide an inadequate explanation for conserved locus structure. If co-regulation were exceptionally important, one might expect that experimentally unlinked RAG genes would be ineffective in V(D)J recombination. Functional studies of proteins templated by fully unlinked RAG1 and RAG2 genes in pre-B cells indicate that this is not the case (Silver et al., 1993).
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Thus, for the time being, the best explanation for the apparently universal linkage of the RAG1 and RAG2 genes is that these genes are seasoned travelers, and were introduced into an early vertebrate by horizontal transfer. This interpretation is of course fully consistent with a transposon scenario, from either prokaryote or eukaryote, supposing only that the ensuing 450 million years has not been long enough to erase evidence of co-transfer. A specific prediction can be made, which is that if through molecular paleontology it is possible to identify a RAG-containing transposon, we might expect to find RAG1 and RAG2-like progenitor genes adjacent to one another and in a tail-to-tail configuration. The RAG1 and RAG2 coding sequences are each fully contained within a single exon in all animals investigated to date excepting ray-finned fish (Venkatesh et al., 1999). In trout, pufferfish, and zebrafish, among others, the RAG1 coding sequence is interrupted by two introns (Venkatesh et al., 1999; Willett et al., 1997). These introns are considered late changes that must have been added after the ray-finned fish lineage had branched off from those leading to sharks and tetrapods (Venkatesh et al., 1999). Based on current knowledge then, we can infer that at their first appearance, the RAG1 and RAG2 genes lacked introns within their coding regions. The small number of introns in RAG1 and RAG2 is to be interpreted with caution. The observation might support the argument that the RAG genes were transferred directly from a prokaryotic source because introns are rare in bacteria. However there certainly are examples of eukaryotic DNA transposases that have few introns (Figure 30.2), and it is also possible too that introns in the 5¢ noncoding region of the RAG1 and RAG2 genes are in fact a universal feature (Hansen and McBlane, 2000).
Clues Based on the RAG1 and RAG2 Proteins: Domains and Motifs No extensive homologies between the RAG proteins and any other protein currently in the public database come to light by simple BLAST alignments. Comparisons that look more closely at a combination of predicted structure and the presence of small functional motifs, however, can find suggestive similarities. At this level of scrutiny, the evidence is likely to reflect deep evolutionary origins in addition to possibly highlighting the characteristic features of the invading V(D)J transposon. A GGRPR amino acid cluster in RAG1 is important for binding the RSS nonamer sequence in vivo and in vitro as well as for V(D)J recombination (Difilippantonio et al., 1996; Spanopoulou et al., 1996). The same amino acid sequence is found in the HIN recombinase, for which there is structural information indicating contacts with the minor groove of the (“nonamer”-like) T tract in the HIX site (Feng et al., 1994) (Figure 30.3). (R)GRP(R) motifs termed “AT
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hooks” appear in a variety of DNA binding proteins, where they are proposed to work as a minor groove tether, acting along with either a homeodomain-like helix-turn-helix or other DNA-binding fold (Aravind and Landsman, 1998). The motifs within both RAG-1 and HIN can be considered divergent AT hooks (L. Aravind personal communication). Notably, members of the Tc1/mariner family transposases have a “GRPR” sequence positioned near a predicted helixturn-helix motif (Plasterk et al., 1999). Parallels between RAG1 and HIN, with respect to the AT hook feature, may yet have some significance with respect to the evolution of V(D)J recombination. However, it is also possible that the AT hook similarities here are more relevant to abovementioned deep evolutionary relationships. AT hook-like motifs are combined with DNA or RNA-binding folds in quite ancient lineages of proteins (Iyer et al., 2003). A special class of experimentally generated RAG1 mutants retain the ability to bind DNA without being able to carry out cleavage (Fugmann et al., 2000; Kim et al., 1999; Landree et al., 1999). Three amino acids—two aspartic and one glutamic—within RAG1 are critical for making cuts in RSS DNA and have been suggested to comprise a “DDE” motif. The DDE pattern of amino acids, occurring with a characteristic order and spacing is seen in enzymes that carry out metal-dependent phosphoryl transferase reactions, including proteins like retroviral and bacterial integrases and DNA transposases (reviewed in Haren et al., 1999). As more fully discussed in the original papers (Fugmann et al., 2000; Kim et al., 1999; Landree et al., 1999), the order and spacing of the D, D, and E amino acids in RAG1 doesn’t unambiguously establish a relationship to the DDE motif in DNA transposons. In fact, attempts to predict the identity of the catalytically important D, D, and E amino acids within RAG1 and RAG2 by extrapolating from Tc1/mariner type transposases (Dreyfus et al., 1999; Plasterk et al., 1999) failed, as was later demonstrated by mutational analyses. Protein structure studies are needed to resolve whether the catalytic amino acids in RAG1 constitute a diverged DDE motif, nevertheless, work to date establishes that some or all of the catalytic site lies within RAG1. RAG2 was also subjected to detailed mutational analyses without revealing specific defects in cleavage for any variant (Landree et al., 1999). A large N terminal portion of RAG1 can be removed without much impact upon the protein’s ability to carry out V(D)J recombination on plasmid substrates. Within the nonessential region, there is a RING finger motif that is evolutionarily conserved in RAG1 proteins (Bellon et al., 1997). Some proteins with RING fingers are E3 ubiquitin ligases (Weissman, 2001), and a recent study demonstrated in vitro ubiquitin ligase activity for RAG1 (Yurchenko et al., 2003). Ubiquitination of a target protein will mark it for degradation or, in other cases (as for some transcription factors) the modification is activating (Fang et al., 2003). Obviously a
determination of the in vitro substrate for the RAG1 ubiquitin ligase activity, possibly be a histone or the RAG1/2 proteins themselves, will have a number of implications regarding evolutionary origins. That is, the ubiquitin ligase activity for RAG1 might have been acquired secondarily during exaptation, or conversely, the ubiquitin ligase function may have been a feature of the original V(D)J transposon-encoded transposase. In the latter case RAG1/2 was evidently well used to functioning in a eukaryotic context at the time it came into the vertebrate genome. This is because RING fingers are a uniquely eukaryotic feature, and as such their presence in a protein argues against a prokaryotic origin. The former life of RAG2 and how it came into its current role as a crucial component of the V(D)J recombination machinery is an important aspect of the evolution of V(D)J recombination. Some transposases interact with additional element-encoded proteins in mobilizing DNA. For example, conjugative transposases work with transposon-encoded “xis” proteins in order to exit from one integration site prior to landing in another (Scott and Churchward, 1995). The transposase encoded by the fruit fly P element competes with an element-encoded inhibitor that helps maintain transposition at a sublethal levels for the host, indirectly ensuring that the P element itself will endure (Lee et al., 1998). However, the situation that seems to most closely parallel that of RAG1/2 is exemplified by the bacterial transposon, Tn7. This transposon requires two proteins to cleave its termini, each performing different duties, and each indispensable for transposition (Biery et al., 2000). If RAG2 is similarly part of a two-unit transposition machinery, there is a notable paucity of candidates among known eukaryotic DNA transposons (Figure 30.2). Analyses that combined basic local alignment with the occurrence of clusters of hydrophobic amino acids revealed series of “kelch” repeats in RAG2 (Callebaut and Mornon, 1998). Kelch repeats fold into a propellerlike structure that supports intermolecular protein–protein and protein–DNA interactions. The three-dimensional propeller structure, if assumed for RAG2, reveals a suggestive spatial clustering of naturally occurring RAG2 mutations in humans (Corneo et al., 2000; Gomez et al., 2000). A “PHD” (plant homeodomain) motif occurs near the C terminus of RAG2 as well (Callebaut and Mornon, 1998). The latter is located in a region of the protein that is not essential for V(D)J recombination, but if removed from RAG2, the efficiency of in vivo V-to-D rearrangement is altered (Akamatsu et al., 2003; Liang et al., 2002). Some PHD domains are suspected E3 ubiquitin ligases (Fang et al., 2003) but this has not been demonstrated experimentally for RAG2. The presence of kelch repeats and/or a PHD domain may be helpful in identifying RAG 2-like proteins in candidate V(D)J transposons. The fact that no extensive homologies exist between either of the RAG proteins and any other proteins currently
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in public databases is significant. Even though, as discussed above, there is some evidence to suggest that a Tc1/marinertype transposon donated the RSS sequences, any evidence that a Tc1/mariner family element also donated the RAG1/2 genes is lacking. It may be then that the “V(D)J-transposon” was really two separate entities, one an RSS transposon and the other a RAG1/2 transposon. (see note added in proof)
Clues Based on Modus Operandi Although some DNA recombinases carry out site-specific recombination and others carry out transposition, there are only a few descriptions of a machinery equally capable of both operations. Site-specific recombination and transposition are each initiated by the creation of two DNA breaks, but diverge thereafter. Site-specific recombinases make a pair of breaks in DNA and reshuffle the four broken ends. Transposases make a pair of breaks in DNA, randomly grab an unbroken segment of DNA, and insert the chopped-out piece within the captured segment. This difference divides the two processes: If a site specific recombinase were to act as a transposase, it would have to be able to capture an additional segment of DNA. If a transposase is to act as a sitespecific recombinase, it must be able to connect the ends at the exit site to the ends of the liberated transposon. Transposons are usually barred from re-entry into their exit sites due to the chemical mechanism by which the element’s 3¢ ends become contiguous with the DNA at the new site. This is through an iso-energetic phosphoryl transfer to the element’s 3¢ ends (Mizuuchi, 1992), meaning simply that a transposase needs an intact phosphodiester bond to complete the transposition cycle: a pre-cleaved end won’t work. RAG1/2 can perform as a transposase or a site-specific recombinase, with an ease not observed in other enzymes. The latter capability (i.e., V(D)J recombination) is carried out with significant assistance from ubiquitous DNA repair functions in T and B cells. However, it is useful to consider the properties of a hypothetical “two-way” transposon (Figure 30.5), based upon the observed behavior of purified RAG1/2 in vitro (Melek et al., 1998). A two-way transposon could manage (as simplistically conceived on paper) by a sharing two key features of the RAG1/2 cleavage cycle: 1) the first nick made in the act of transposition occurs on the “nontransferred” strand (corresponding to the first break at the RSS), and 2) the second strand is broken via a hairpin formation (Figure 30.5) (McBlane et al., 1995; van Gent et al., 1996). The liberated transposon fragment is blunt-ended, with an exposed 3¢OH. Because of these two special properties, subsequent to formation of double-strand breaks at the transposon ends, an (unbroken) phosphodiester bond is available at the exit site that could support site-specific recombination as an alternative outcome (Figure 30.5), to transposition. Transposons lacking either of these two key
FIGURE 30.5 Hypothetical “two-way” transposons. A dual activity for the RAG1/2 progenitor is suggested by the in vitro results of Melek et al. (1998). The relationship between transposition, involving transposon exit followed by capture of a new target site DNA, and site-specific recombination, involving transposon exit two breaks followed by rejoining of ends is illustrated. Dark triangles show the positions of the strand breaks that define and distinguish the two possible (post-exit) outcomes.
features (i.e., that make the first nick on the transferred strand like Tn10, or that don’t form a hairpin intermediate, see Figure 30.2) are not properly endowed for “two-way” capability. For the hypothetical two-way transposons, the latent ability to perform site-specific recombination would be difficult to fully discard during evolution. A recent observation involving a member of the mariner family Mos1 is suggestive. Like RAG1/2, the Mos1 transposase makes the first break on the nontransferred strand (reviewed in Turlan and Chandler, 2000), but, unlike RAG1/2, the second break is not accompanied by hairpin formation (Dawson and Finnegan, 2003). Under certain circumstances, however, it has been shown that Mos1 transposase can catalyze the formation of an alternative, hairpinmediated DNA break just a few bp outside the element end (Dawson and Finnegan, 2003). Given that the order of strand breaks and the formation of hairpin structures is favorable, conceivably the mariner-like element Mos1, if pushed hard enough, could be capable of site-specific recombination. Such dual function may already in fact have been observed in vivo for another Tc1/mariner element. The nematode transposon, Tc1, like RAG1/2 and Mos1, initiates transposition with a break in the nontransferred strand. When artificially introduced into mammalian (COS) cells, Tc1 is reported to carry out in situ, site-specific inversion (Li et al.,
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1998). Although transposable elements of many kinds can cause aberrant DNA inversions of flanking sequences when they exit, a tidy, simple flip of the Tc1 element, as was described in the report, is exceptional and warrants further investigation. Any two-way transposon has an Achille’s heel. Transposition can be foiled if the element can be forced to become reconnected to the hairpin breaks it just made at its exit site (by realizing a site-specific recombination outcome). A DNA transposon that can’t hop is barred from the possibility of spread (and persistence) through matings between polymorphic individuals. As a key step in exaptation, limiting the RAG1/2 transposase to its site-specific recombination capacity could have been accomplished by pre-established host functions. As demonstrated by biochemical studies, RAG1/2 is ready and able to transfer the RSS end back into the hairpin structures it just made (Melek et al., 1998). Perhaps with very little tampering, the progenitor transposon could be made to do this as a matter of course. The war between host genomes and invading elements has been fought many times throughout evolution, and perhaps for the type of engagement envisioned here, it may have been won sometimes by the host as in the exaptation of the V(D)J transposon and sometimes by the element as in the still-thriving hAT family (Rubin et al., 2001). Essential but nonlymphoid components of the V(D)J recombination apparatus, such as DNA-PKcs and Artemis, may have been part of an anti-transposon defense mechanisms dating back to the original integration of the V(D)J progenitor transposon and may still carefully guide the outcome to the present day. Experimental evidence for such devices and counterdevices may come from further study of the behavior of transposons introduced into foreign contexts (such as the worm-to-mammalian transfer for Tc1), or from the examination of the capabilities of RAG1/2 in mutant genetic backgrounds or in nonvertebrate eukaryotes (Clatworthy et al., 2003).
Current Questions: Exaptation Although lateral transfer and exaptation of RAG1/2 is now a reputable notion (Bushman, 2002; Smit, 1999), as can be seen from the above discussion, much remains to be fleshed out in terms of how exaptation was achieved and what the exapted entity looked like in the first place. The rough “before” and “after” sketch in Figure 30.1 shows that the RSS motifs are not part of a larger ITR and are dissociated from any transposase genes. A fundamental question is whether both the RSSs and the RAG1/2 genes were ever in fact contained in one and the same mobile element. Mobile elements in general are highly accident-prone and any genome contains abundant examples of such elements in various states of decay (Bushman, 2002). Errors in transposition can scramble or delete the transposase, leaving dis-
membered ITRs. When this happens, different portions of the dissected transposon have different likelihoods of enduring. ITRs-minus-transposase can still move if transposase encoded by an intact element co-existing elsewhere in the genome is available. Transposase-minus-ITRs however, is doomed. Even if the transposase is still competent, unless it does something of positive value for the genome it inhabits (and causing transposition of other defective elements does not fit the bill), the transposase cannot move (or expand in copy number) and faces extinction. These considerations suggest that, for a RAG1/2containing transposon, the RAG1/2 transposase must have been put to work performing gene assembly as soon as the transposon was dismembered. As discussed later, the exemplary consequences with respect to gene diversification provide a ready explanation for the survival of the RAG 1/2 locus to the present day, despite the catastrophic loss of its ITRs. Another significantly different possibility regarding decay and exaptation is that the RSSs and the RAGs could have entered the genome at different times. The RSS elements might be remnant ITRs that were fortuitously mobilized by the transposase of an unrelated transposon. In this case, “older” RSS motifs could have been borne in on a transposon that has long disappeared, with a subsequent history of uneventful vertical transmission before the “newer” RAG1/2-bearing transposase appeared on the scene (via horizontal transfer). An even more radical idea is that the RSS sequences could have been generated de novo [possible mechanisms are discussed in (Feschotte et al., 2003; Hughes and Coffin, 2002)]. Examples that illustrate these possibilities are found in the mobilization of miniature inverted repeat elements (MITEs) and the Mariner-like elements that appear to drive their movements in plants (Feschotte et al., 2003; Jiang et al., 2003). Conceivably then, the process of exaptation might even have begun to work on the RAG-bearing transposon before the element became completely dismembered. A last possibility is that the RAG gene were already legless remnant when first inserted into the vertebrate a genome. In general, the issue of how DNA transposons invade a different (i.e., reproductively separate) species is a central, but unsolved question (Bushman, 2002; Yoshiyama et al., 2001). Most cut and paste DNA transposons completely lack any equipment that could possibly support interspecies transfer. Nevertheless they get transmitted, injected or transduced into foreign genomes somehow. An infectious agent, pseudotyping the transposon, can provide the missing function. For vertebrates, a retroviral-like entity is an attractive choice (Bushman, 2002). The RAG transposon may have inserted into a retro element in the species of origin, or a partial transcript might have been snatched by reverse transcription, thus entering a new host, while cloaked in “retro” livery.
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THE EVOLUTION OF BCR AND TCR LOCI Once the conversion of V(D)J transposon to V(D)J recombination apparatus was accomplished, new possibilities for genome rearrangement were opened up and likely were instrumental in forming the different Ig and TCR loci. Figure 30.6 sketches out the RAG-mediated manipulations that are either known or postulated to have modified the antigen receptor loci during vertebrate evolution. As shown, a preliminary step (Figure 30.6A) must have been duplication of the region containing a RSS-interrupted gene (Figure 30.6B). This must be assumed in order to create a substrate for creating different loci and “trying out” different configurations. Segmental duplications, unrelated to V(D)J recombination, are both ancient and ongoing throughout evolution (Bailey et al., 2002). Duplications are undefined with respect to gene segment, locus boundaries, or mechanism of formation. Duplication is likely to either include just a subregion of a locus (Matsuda et al., 1990) or can involve an entire functional V(D)J (C) unit (as is postulated to have given rise to the multiple Ig clusters in cartilaginous fish) (Ventura-Holman and Lobb, 2002). Thereafter, various modifications of the duplicated copies could be accomplished by germline RAG activity as outlined below.
Germline Coding Joint Formation: V(D)J Gene Segment Fusions Compelling evidence for germline V(D)J recombination comes from examining the Ig locus structures in sharks. Shark IgH genes occur as repeats of a simple V-D-J (CH) recombination modules: likewise IgL genes are multiply reiterated V-J (CL) clusters (Du Pasquier and Flajnik, 2003). By examining the coding sequences, it can be possible to estimate when two related modules might have arisen through an ancestral duplication event. If one of the clusters is rearranged, it is possible to know roughly when recombination occurred. In the nurse shark, it was shown that modular V-JC copies had become germline-joined within the last 7 million years (Lee et al., 2000). Thus, in some species, germline V(D)J recombination that creates fused VJ and VDJ genes (Figure 30.6C) is not simply a theoretical possibility but still, perhaps, ongoing. Germline joining may have occurred in other species that don’t have highly redundant recombination modules. In such circumstances the event would likely be counterselected. Germline V(D)J assembly is expected to elimiate gene segments significantly reducing the potential for generating diverse T- and B-cell repertoires. Evidence of germline joining has nevertheless been observed in a nonredundant locus in the chicken (Reynaud et al., 1989). Pseudo V genes in the IgH locus are quite obviously germline-joined, being contiguous with D-like sequences
FIGURE 30.6 Germline rearrangement gives rise to different locus configurations. (a) A transposon splits an ur-V gene, placing RSSs adjacent to the interrupted coding sequences. (b) Duplications of the interrupted unit take place through non–V(D)J-related mechanisms. At least one duplication event preceded a germline recombination event, but generally both germline recombination and expansion of unrecombined (and recombined) gene segments can be concurrent. (c-i) A standard V(D)J recombination event forms a coding joint (“CJ”), giving rise to germline-joined copies (see text). (c-ii) A standard V(D)J recombination event involving inverted gene segments generates a signal joint (“SJ”). If signal joint formation is accompanied by N region insertion, this would create a D segment. (c-iii) A nonstandard V(D)J recombination outcome generates hybrid joints (“HJ”). This changes the RSS affiliations of gene segments, and as discussed in the text, could drive the divergence of various Ig and TCR loci in evolution.
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and some having recognizable J segments at their 3¢ ends. In chickens, the germline-joined copies were likely preserved because of the augmented diversity they provided as donors for somatic gene-conversion at the IgH locus (Reynaud et al., 1989). Another example of germline joining is found in the channel catfish, where a duplicated and joined copy of the IgH locus is found (Ghaffari and Lobb, 1999; Ventura-Holman and Lobb, 2002).
Germline Signal Joint Formation: The Creation of D Segments V(D)J recombination creates a reciprocal product to coding joints, termed signal joints. These are comprised of RSS-to-RSS junctions, and are retained in the chromosomes of B and T cells after V(D)J recombination if, as is possible at some loci, rearrangement involved inverted gene segments. Germline V(D)J recombination would be expected to produce a chromosomally retained signal joint from similar inversional joining events. As shown, if this signal joint includes an N-region insertion, it becomes structurally identical to a D segment (Figure 30.6C-ii) (Lewis et al., 1988). Thereafter, positive selection for diversity (as discussed in the next section) would fix such changes in a population. There is no reason to suppose that D segments were invented only once, or only rarely, by germline V(D)J recombination as suggested in some discussions (e.g., Glusman et al., 2001). Chromosomal signal joints will form from any inversional recombination event, and N-region insertion into signal joints is not uncommon (Lieber et al., 1988). The idea that the three-segment loci are derived from simpler kappa or lambda-like loci is a viable notion given current views of the ages of the various isotypes (Flajnik and Rumfelt, 2000).
Germline Hybrid Joint Formation: RSS Swaps and Locus “Speciation” Confusing and conflicting family relationships between Ig and TCR genes (Du Pasquier and Flajnik, 2003) get even worse when the RSS affiliations of the gene segments are taken into consideration. For example, the V genes of kappa and lambda are more similar to each other (and to IgH) than they are to TCR V genes, yet VH, Vl, and Vb all have 23spacer RSSs, whereas Vk has a 12-spacer RSS. Likewise JH and Jk have 23 RSSs, whereas 12 RSSs occur at both Jl and Jb. Some of this complexity can be deconvoluted by schemes that generate one locus structure from another by germline V(D)J recombination. Figure 30.6C-iii illustrates how any RSS configurations can have been derived from another by germline hybrid joint formation (Kokubu et al., 1988; Lewis et al., 1988). RSS swapping is seen in vivo in lymphoid cells. This occurs
when hybrid joint formation takes place instead of the usual (and productive) coding joint–signal joint outcome of V(D)J recombination (Lewis et al., 1988; Morzycka-Wroblewska et al., 1988; Sollbach and Wu, 1995; VanDyk and Meek, 1992). Germline hybrid joint formation, similarly resulting in an RSS swap, would set aside the affected gene segments for separate evolution (a sort of molecular “speciation” event). This is because the swapped units can now profitably rearrange in somatic cells only when they undergo intralocus recombination (with a swapped partner). Interlocus recombination, conducted according to the 12/23 rule would instead result in nonproductive V-V and J-J joining. In conclusion, the above schemes have a compelling simplicity as an explanation for much of the variety seen today in locus configuration. It is a reasonable presumption that the journey from V(D)J transposon to the full combinatorial diversity apparatus was promoted in part by germline V(D)J joining. It is interesting to note here that all evidence of germline activity seems to have come about as the result of RAG1/2 behaving as a site-specific recombinase, not a transposase. This fits with the idea that mechanisms in the first invaded host prevented transposition and that such mechanisms have been durable—an impressive feat considering the transpositional capability of the RAG1/2 proteins observed even to this day (Agrawal et al., 1998; Hiom et al., 1998).
Next Steps For the inquisitive, the drive to learn more about the alien seed and its impact upon the evolution of V(D)J recombination will spur the development of imaginative new ways for experimentally exploring different molecular evolution. An exciting challenge is the reconstruction of stages in exaptation. This is not complete science fiction: Computational tools that screen whole genomes for mobile elements have been shown to work and are constantly being improved (Bao and Eddy, 2002; Rubin et al., 2001; Smit and Riggs, 1996). New families of mobile elements, and new members of known families are constantly being identified and placed on a time line that extends back 60, 100, and even 500 million years (Rubin et al., 2001). Trolling operations could one day include a RAG1/2 transposon in the catch. At this point, what are the best guesses as to the properties of the alien seed? Perhaps the most telling feature we might hope to see in the progenitor transposon will be that it encodes a two-protein transposition apparatus, the genes for which will be closely linked in a tail-to-tail arrangement. Further guidance is likely to come from both structural and biochemical studies of the RAG proteins. The way the proteins enwrap their DNA target, what the catalytic site looks like, how the RAG1 and RAG2 proteins contact one another, whether (and what type) of an AT hook is present etc., will all be illuminating. Even if an identified candidate transpo-
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son contains a “dead” transposase gene, similarities that remain to RAG1/2 deduced from a conceptual translation can be properly weighted and evaluated. Further studies on the effects of nucleotide co-factors (Tsai and Schatz, 2003), and the identification of substrates for RAG1’s proposed E3 ligase function (Yurchenko et al., 2003) will help to fill in the picture of what RAG1 is, what function RAG2 serves, and how transposition of the precursor element might have been regulated or targeted. The big question is whether one mobile element contributed both RAG1/2 genes and a pair of RSSs, or whether the structures were independently acquired from two different mobile elements, (or entirely nontransposon sources). Once that is sorted out, a whole new realm of experimental investigation will be laid wide open. The process of exaptation, and the role that “host” functions (DNA-PKcs, Artemis, and TdT) might have played in redirecting and/or foiling the transposition reaction can then be approached experimentally, in a test tube.
CONSIDERATIONS ON THE UR-V GENE Rearrangement Generates Sequence Length Diversity The previous sections have described the possible transposon origin for V(D)J recombination and the subsequent sculpting of the Ig and TCR gene organization by germline rearrangements during evolution. If we postulate that the first rearrangeable antigen receptor gene indeed arose from one incident of splitting through transposon insertion, several questions come to mind—how and why was this one particular insertion selected for, and what might have been the nature of the archaic “ur-V gene”? The ur-V gene would have encoded a domain—an autonomous folding unit—of the Ig superfamily, whose members include cell surface molecules (Ig, TCR, MHC, b2-microglobulin, Thy-1, CD4, CD8) as well as cell adhesion and muscle components (for a review, see Harpaz and Chothia, 1994; Williams and Barclay, 1988). Four sets of domain structures have been distinguished: V (variable), I (intermediary), C1, and C2 (constant). These structures are differentiated from each other by sequence patterns and lengths but they all share the immunoglobulin fold, which is formed by two b-pleated sheets that are packed face-toface and consist of anti-parallel b-strands. The disulfide bridge characteristic of Ig-related molecules is not necessary to the formation of this sandwich structure. Figure 30.7 shows ribbon diagrams of the V, I, and C structures. V domains are distinguished by the presence of two strands, C¢ and C≤. The antigen combining portions are mainly situated in loops, for example, those connecting B to C strand (CDR1), F to G strand (CDR3). In the prototype Ig gene,
FIGURE 30.7 Immunoglobulin superfamily domain structures. Ribbon diagrams of “I” set based on telokin structure (Harpaz and Chothia, 1994); “V” set, based on Fab NEW VH; “C2” set, based on human CD4 domain 2; and “C1” set, based on b-2 microglobulin (Barclay et al., 1993). The region between C and E strands varies among the sets. The I set is apparently the oldest structure. V set differs in the addition of C≤ strand and C2 set by the deletion of D strand. C1 set structures appear only late in evolution, in gnathostomes, and differ from I set by the absence of C¢.
the hypothetical transposition event would have disrupted the region encoding the loop between F and G strands (Figure 30.7). The relationships between the four Ig superfamily structural sets are not entirely clear as yet, but the I set is probably the earliest and the C1 the most recent in evolution. Du Pasquier (2000) analyzed a receptor tyrosine kinase in the sponge Geodia cydonium, concluding that it possessed the V frame that is common to V and I set structures (Harpaz and Chothia, 1994). With completion of the Caenorhabditis elegans genome, an analysis was made to identify the repertoire of the Ig superfamily domains in the nematode. Out of 64 proteins, all 488 domains were of the I set, establishing that this was the ancestral domain structure (Teichmann and Chothia, 2000). Among molecules reported in later invertebrates, such as molluscs and arthropods, domains with V or C2 features, together or alone, could be distinguished—for example, fibrinogen-related proteins (FREP) from the snail or amalgam in Drosophila (Du Pasquier, 2000; Seeger et al., 1988).
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However, TCR and Ig domains consist of V and C1 type domains, and C1 domains of any kind have only so far been found in jawed vertebrates, where there already exist the rearranging antigen receptor genes (Du Pasquier and Chretien, 1996). C1 domains in fact are not only part of antigen receptors but also other immune response– associated molecules like class I and class II MHC, b2microglobulin, and tapasin, all of which also apparently have no homolog earlier than in cartilaginous fishes (for a review see Flajnik and Kasahara, 2001). Du Pasquier (2000) concluded that a transition to C1 may have occurred in the primitive vertebrate, and that searches in jawless fishes for C1-set molecules may provide clues to the identity of the archaic target of the transposition, which was connected with a C1 domain or acquired a C1 domain by exon shuffling as its function evolved. We go on to reason that the ur-V gene probably was one member of a multigene family. Since the immediate effect of the transposition would have been disruption of the target gene, redundancy of the gene function would have prevented immediate elimination of the to-be founder gene. Among multiple duplicated members that compensated for its loss of function, the split gene could have hitchhiked along, eventually to be eliminated by recombination or possibly altered by gene conversion, as has been observed in the “birth-and-death” evolution of multigene families (Ota and Nei, 1994). Fixation of this novel but null (because silenced) mutant could have been by chance, but recruitment for function must have arisen soon after the integration event. Had there not been selection for the split gene, it would have acquired unfavorable changes in the coding region or the RSSs would have mutated out of recognition for the RAG recombinase. It has been observed that TCR V pseudogenes with frameshift mutations tend to accumulate additional changes, possibly because reversion to a functional gene is less apt to occur than for V pseudogenes with point mutations that generate premature termination: The transposition in this case is effectively a frameshift mutation. The selecting feature would have included RAG activity, and this activity would have to be expressed in somatic cells. In the somatic cell, the RAG proteins would excise the RSSs, along with the intervening DNA, that had been incorporated into the ur-V gene in the germline. With this removal, the cleaved DNA ends at the exit site would be rejoined, most likely by RAG-independent endogenous repair mechanisms. Even discounting enzymatic and genetic elaborations, like N-region addition by TdT and the D gene segments, heterogeneity can be generated at the new junction. The opening of hairpin DNA as formed by RAG recombinase action (McBlane et al., 1995), followed by limited exonuclease trimming and ligation, all could be carried out by functions that are part of the endogenous nonhomologous end-joining pathway of repair (reviewed in Gellert, 2002).
The ubiquity and nonlymphoid cell–specific nature of the proteins involved in reconnecting the ends is illustrated by coding joint formation in RAG-transfected NIH 3T3 fibroblasts (Kallenbach et al., 1992). The net effect of the removal of the intervening DNA from the split gene is that the reformed gene sequences, when ligated in frame, encode proteins with both sequence and length heterogeneity. Although somatic mechanisms change DNA sequence—mutation and gene conversion among them—none generate sequence length diversity at high frequency. Gene conversion will incorporate insertions or deletions, but will do so according to the set of templates available in the germline. Somatic hypermutation can create deletions and duplications (in/del) in V regions, but does so with low efficiency in functioning genes (Goossens et al., 1998; Wilson et al., 1998). Whereas in/dels occur at different locations throughout nonproductive V(D)J exons, in functional rearrangements they reside mainly in the CDR loops (de Wildt et al., 1999). Sequence length variability not only alters the chemical nature but, more drastically, the physical shape of the region involved. The loops best support this type of variability, without interfering with the main-chain folding that creates the Ig fold, they and illustrate it by being the least conserved feature in sequence alignments of evolutionarily related proteins (Williams and Barclay, 1988). The RAG recombinase activity is site-specific, and this ensured localization of sequence length variability at one tolerated place and on a regular basis. The subsequent creation of D gene segments [possibly via germline joining (Figure 30.6Cii)] to further increase heterogeneity at the CDR3 loop supports the idea that junctional diversity is a very strong selecting feature for rearranging genes (Lee et al., 2002). Thus, generation of the unusual molecular heterogeneity provided by RAG action may have been the selecting factor for the split gene.
Hypermutation in Evolution If the novel advantage of CDR3 heterogeneity was greater sequence diversification, the original gene function of the unsplit genes would have also required this feature. We speculate that there may have already existed a diversifying mechanism—the archaic nonrearranging V genes possessed the ability to mutate in somatic cells. Rearrangement and somatic mutation both exist in cartilaginous fishes (sharks, skates, rays), representatives of the earliest jawed vertebrate. So far no evidence shows that one mechanism preceded the other in the evolution of the adaptive immune system. The recognition motifs and enzymatic machinery involved in the two processes point to mechanistically unrelated pathways that arose independently (Flajnik, 2002). Changes that appear to be the result of gene conversion–like events have been observed in shark NAR
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and Ig L chain mutants (M. Flajnik, personal communication; Lee et al., 2002), and for this discussion gene conversion and somatic hypermutation will be bracketed together. It has been proposed that the two are mechanistically related (Maizels, 1995), and an experimental demonstration of the emergence of Ig hypermutation in the chicken cell line that had favored gene conversion can be likewise interpreted (Sale et al., 2001). There are few studies on somatic hypermutation in coldblooded vertebrates. Some somatic mutation seems to occur in the red-eared turtle VH, but this was deduced by a comparison of clonally related cDNA sequences (Turchin and Hsu, 1996). Studies in the amphibian Xenopus reported GCbiased point mutations at low frequency (Wilson et al., 1992). No studies have been made in any bony fish. Initial reports of mutation in skate L chain and horned shark H chain showed that somatic mutation was present in early vertebrates but gave the impression that, as in Xenopus, its presence was not extensive (Anderson et al., 1995; HindsFrey et al., 1993). More recent studies on nurse shark identified small families of germline genes and correlated them with rearranged cDNA sequences, demonstrating that somatic hypermutation was an important mechanism for antigen receptor diversification in sharks (Diaz et al., 1999; Greenberg et al., 1995; Lee et al., 2002). The new antigen receptors (NAR) in nurse shark, a third class of rearranging genes showed extensive somatic hypermutation, averaging 6% substitutions (Greenberg et al., 1995). A comparison of the transmembrane with the secretory NAR forms suggested that mutations, plentiful in the latter, were acquired with antigen stimulation (Diaz et al., 1998). A study of one of the three nurse shark Ig L chain types that showed 0.03% substitution frequency in perinatal animals but >8% in adults (Lee et al., 2002), also suggested antigen-dependent changes. Preliminary studies on mutants from two nurse shark H chain clusters (Chen and Hsu, unpublished results) showed that the mutation patterns for H chain was the same as in L chain sequence: point mutations and tandem mutations, the latter consisting of 2 to 5 bp contiguous changes. A second nurse shark L chain type was also investigated; all three functional gene clusters hypermutated, and with the same pattern for point mutations (55% transitions) and tandem mutations (37% transitions) (Fleurant and Hsu, unpublished results). If cis-acting elements influencing hypermutation are present on different clusters of nurse shark H chains, two L chain types, and NAR genes, then these factors existed before their divergence. If they are composed of multiple parts, as they are at the mammalian kappa L chain locus (Betz et al., 1994; Sharpe et al., 1991), one presumes it is easier to lose the cis-acting factors than to acquire them. The original antigen receptor genes may have possessed these elements, and in the course of their derivation Ig genes retained them, but TCR genes lost them.
The notion that hypermutation is an ancient mechanism whose regulatory elements are genetically inherited with the structural genes is supported by an examination of other cell surface receptors present in the shark and skate. It is more probable that NAR, with clusters each consisting of one V, three D, and one J gene segments and a set of five C exons, was derived from Ig rather than TCR. The V region resembles neither Ig nor TCR by phylogenetic analysis (Greenberg et al., 1995), since it must have undergone extensive changes to adapt the NAR homodimer structure, but four of the five NAR C exons are homologous to the heavy chain of another Ig-like molecule, IgW (Flajnik and Rumfelt, 2000). IgW clusters carry carry six C exons but also bona fide VH gene segments indistinguishable from those of IgM clusters. Their products combine with L chain (Bernstein et al., 1996; Greenberg et al., 1996). Like NAR and IgM, IgW is also secreted and shows signs of hypermutation (M. Flajnik, personal communication). Thus, all the genes that have been shown to mutate are Ig (IgM H chain, clusters from two L chain types) or derived from an Ig-like ancestor (NAR and IgW). Because IgM and IgW clusters are believed have multiple chromosomal locations (Anderson et al., 1994), they must individually harbor cis-acting elements for mutation. If the ur-V genes could hypermutate, why have mammalian-type adaptive immune responses not been observed in nonvertebrates? Actually, even in shark, despite the extensive substitution that has been demonstrated in NAR, L chain, and H chain genes, studies in the literature report that shark antibody responses do not seem to increase in affinity and that memory in the form of a secondary response is problematic to demonstrate consistently (for a review, see Hsu, 1998). Thus, the existence of Ig somatic mutants in a vertebrate cannot always demonstrably be connected with an improved antibody response of the sort defined in rabbits, humans, or mice. Perhaps we should be looking for another parameter affected by mutation. However, the site for the proliferation of activated and hypermutating B cells exists only in birds and mammals. Concentrating the daughter clones in a germinal center results in competition for antigen and selection for those with the mutated, higher affinity receptors. The absence of a locale for efficient selection in cold-blooded vertebrates has been suggested as the reason for the apparently poor affinity maturation of their antibody responses (Wilson et al., 1992).
An Innate Defense V Gene It has been proposed that a transposition event involving a RAG transposon and an archaic V gene initiated the creation of rearranging antigen receptors and therefore the basis of adaptive immunity. We suggest that the prototype V genes were part of a multigene family that diversified by hyper-
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mutation, and that the introduction of a recombining mechanism that generated loop length variability greatly augmented diversification in a way more revolutionary than base substitutions. This was the selecting factor for the split gene created by the RAG transposon. Attempts to identify the prototype antigen receptor genes in jawless fish have so far been unsuccessful in experiments ranging from isolation of serum proteins obtained after immunization to efforts in cross-hybridization and PCR. Perhaps the difficulty in identifying the prototype V gene is that, with the acquisition of junctional diversification, the rest of the sequence may have evolved to accomodate or enhance the useful new structures generated by rearrangement. Hence, the modern vertebrate V gene is sufficiently diverged so that its actual sequence no longer serves as much more than a general guideline to searches in jawless fish, protochordates, or invertebrates. Large sets of nonrearranging V-like genes have recently been isolated from amphioxus (Cannon et al., 2002), but so far no immune-type function has been ascribed to them. We have speculated that the ancient V gene mutated and may have had some sort of defense function. Although in itself no proof of any immune function, somatic mutation should be a fairly restricted phenomenon. One candidate for a cousin of the Ig prototype may be the fibrinogen-related proteins (FREP) isolated from the hemolymph of freshwater snails, Biomphalaria glabrata. FREPs are a family of molecules consisting of one to two Ig superfamily V set– like domains connected with a carbohydrate-binding lectin domain (Du Pasquier, 2000; Zhang et al., 2001). The number of different cloned FREP sequences from one subfamily greatly exceed the five to eight bands apparent by genomic Southern blotting (see discussion in Du Pasquier and Smiith, 2003), and it may be possible that some kind of somatic diversification is producing the sequence heterogeneity (S.-M. Zhang, C.M. Adema, T.B. Kepler, E.S. Loker, personnal communication). The FREPs do not rearrange in the manner of Ig/TCR. They are a multigene family consisting of sets of tandemly arranged V domain–like exons with fibrinogen exons. The V genes are diversified in the germline, the fibrinogen domains can be construed as serving a C region-type function, and some genes appear to be upregulated in response to infection by flatworm parasites such as Schistosoma mansoni and Echinostoma paraensi (Adema et al., 1997). These characteristics suggest that some similar kind of ancestral innate defense gene in a primitive vertebrate could have been the target of the postulated RAG transposition.
Acknowledgments The authors would like to thank L. Du Pasquier for his comments and advice throughout. We are also grateful to J. Brosius, M. Flajnik, and S. Loker for sharing their insights. Additionally the following individuals
answered many and varied questions and/or to provide preprints and manuscripts prior to publication: L. Aravind, T. Baker, S. Desiderio, M. Gellert, A. Hassanin, J. Hansen, M. Haynes, J. Hughes, V. V. Kapitonov, M. Lieber, J. J. Marchalonis, M. Oettinger, R. Plasterk, D. Schatz, L. Steiner, B. Venkatesh, and C. Willet. Work in the authors’ laboratories was supported by the National Cancer Institute of Canada (S.M.L), the Canadian Institutes for Health Research (S.M.L. and G.E.W) and the National Science Foundation (E.H.). Note added in proof: An exciting new development (Dr. Jonathan Rast, personal communication) is the discovery of several RAG1-like sequences in the genome of Strongylocentrotus purpuratus (purple sea urchin). Conceptual translations of sequences obtained from the Sea Urchin Genome Project (Human Genome Sequence Center of the Baylor College of Medicine http://hgsc.bcm.tmc.edu/projects/seaurchin/) can be aligned to a large region of the human RAG1 sequence from approximately aa 470 to 1000 with 30% identity.
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31 Antibody Structure and Recognition of Antigen ERIC J. SUNDBERG AND ROY A. MARIUZZA Center for Advanced Research in Biotechnology, W. M. Keck Laboratory for Structural Biology, University of Maryland Biotechnology Institute, Rockville, Maryland, USA
The VH and VL domains each contain three segments, or loops, which connect the b-strands and are highly variable in length and sequence among different antibodies (Wu and Kabat, 1970). These so-called complementarity-determining regions (CDRs) lie in close spatial proximity on the surface of the V domains and determine the conformation of the combining site. In this way, the CDRs confer specific binding activity to the antibody molecule. The central paradigm of antibody–antigen recognition is that the threedimensional structure formed by the six CDRs recognizes and binds a complementary surface (epitope) on the antigen. Although CDR loops are hypervariable and confer binding specificity to the antibody, it is not necessary that all six CDR loops interact with a given antigen. Antibodies to smaller antigens, such as haptens and peptides, commonly do not utilize all six CDRs (Chitarra et al., 1993; Wilson and Stanfield, 1993), whereas anti-protein antibodies nearly always do. Camelid antibodies that have no light chains (Hamers-Casterman et al., 1993) but can, nonetheless, bind protein antigens with nanomolar affinities using as few as two CDR loops (Decanniere et al., 1999) are a clear exception to this generality. Framework regions are commonly invoked in antigen recognition to varying degrees, and can comprise up to 15% of the buried surface area of an antibody–antigen complex (Wilson and Stanfield, 1994). The VHCDRs, and VHCDR3 in particular, generally make more extensive contacts than VLCDRs, and the geometrical center of the interface tends to lie near VHCDR3. There exists a strong correlation between residues that do not form contacts with antigen and those residues that are important in defining the canonical backbone structures of the CDR loops (Chothia et al., 1989). These residues tend to pack internally and are therefore less exposed on the antibody combining site surface.
Antibodies may be regarded as products of a protein engineering system for the generation of a virtually unlimited repertoire of complementary molecular surfaces. This extreme structural heterogeneity is required for recognition of the infinite array of antigenic determinants presented in nature. Here we discuss broadly the structure of antibodies and their specific recognition of antigens, the binding energetics of antibody–antigen interactions, the structural basis of the antibody maturation process, and limitations to antibody affinity and specificity for antigens.
A STRUCTURAL FRAMEWORK FOR MOLECULAR RECOGNITION Antibody molecules (Figure 31.1) are composed of two identical polypeptide chains of approximately 500 amino acids (the heavy or H chains) covalently linked through disulfide bridges to two identical polypeptide chains of roughly 250 residues (the light or L chains). Based on amino acid sequence comparisons, the H and L chains may be divided into N-terminal variable (V) and C-terminal constant (C) portions. Each H chain contains four or five domains (VH, CH1, CH2, CH3 ± CH4 depending on the antibody isotype) of two anti-parallel b-sheets, whereas each L chain consists of two such domains (VL, CL). The VL and CL domains are disulfide-linked with the VH and CH1 domains, respectively, to form the Fab region of the antibody, which is linked through a hinge region to the Fc domain, formed by noncovalent association of the CH2–4 domains from both chains. All of the b-sheet domains are structurally very similar and belong to the “immunoglobulin fold” superfamily (Amzel and Poljak, 1979), a structure that is not unique to antibodies but is utilized by numerous immunoreceptors.
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FIGURE 31.1 Structural overview of the murine IgG2a monoclonal antibody, Mab231. (a) Structure of the intact antibody, including two light chains each composed of a variable (VL) and a constant (CL) immunoglobulin (Ig) domain (red) and two heavy chains each composed of a variable (VH) and three constant (CH1, CH2, and CH3) domains (blue). The two hinge regions are highlighted within the dotted oval, revealing the source of structural asymmetry within the intact antibody. (b) Ribbon diagram of a single Ig domain, VL, of Mab231 highlighting its anti-parallel b-sheet secondary structure. The amino- and carboxy-termini are marked, as well as the complementarity determining region loops, CDR1 (yellow), CDR2 (blue), and CDR3 (green). (c) Molecular surface of the antibody combining site of Mab231 formed by the intersection of the apical regions of VL and VH. The CDR loops provide a nearly contiguous surface for antigen recognition. VLCDR1 (yellow); VLCDR2 (blue); VLCDR3 (green); VHCDR1 (magenta); VHCDR2 (cyan); VHCDR3 (red). See color insert.
Antibody–antigen complexes exhibit a high degree of both shape and chemical complementarity at their interacting surfaces (Conte et al., 1999). The application of an algorithm to quantitate shape complementarity in protein–protein interfaces (Lawrence and Colman, 1993) to oligomeric proteins or protease–protease inhibitor complexes gives shape correlation (Sc) values ranging from 0.70 to 0.76 on a scale of 0 (topologically uncorrelated) to 1 (perfect geometrical fit). For antigen–antibody interfaces, Sc values of 0.64 to 0.74 are obtained, indicating poorer average shape correlation—albeit a better topological correlation than for other classes of nonobligatory heterocomplexes (Jones and Thornton, 1996). These differences in shape complementarity very likely reflect the particular biological context in which each type of interface is selected. Thus, protease–protease inhibitor and oligomeric interfaces have co-evolved to optimize the fit (and presumably the affinity) between the interacting components, whereas antibodies must bind antigens not previously encountered during the evolutionary history of the immune system. Indeed, in a process termed affinity maturation, somatic hypermutation of antibody genes is believed to increase affinity by improving complementarity between antigen and antibody. The structural basis for antibody affinity maturation is discussed later in this chapter. The combined solvent-accessible surfaces buried in antiprotein antibody–antigen complexes range from approximately 1400 Å2 to 2300 Å2, with roughly equal contributions from antigen and antibody, whereas smaller antigens, such as haptens and peptides, generally bury less overall surface area when bound to antibody. The surface topography of the antigen-contacting surface, as well as other general structural features, of antibodies can vary significantly according to antigen size (MacCallum et al., 1996). Although the percentage of the antigen surface buried in the interface with antibody is always high and their surfaces complementary, the antibody contact surface becomes more concave as the antigen becomes smaller. Thus, although the combining sites of antibodies that recognize large protein antigens are generally planar, and are often more planar than a number of other types of protein–protein interfaces (Jones and Thornton, 1996), antibodies that recognize medium-sized antigens, such as peptides, DNA, and carbohydrates, often have a grooved antigen-contacting surface, while even smaller antigens (haptens) are recognized by antibodies with distinct cavities (Webster et al., 1994). A common feature of anti-peptide antibody–antigen interactions is a b-turn motif of the peptide buried deeply into the combining site. A number of examples exist for this general recognition scheme, including type I b-turns (Rini et al., 1992), type II b-turns (Stanfield et al., 1990), and multiple tight turns (Garcia et al., 1992b). Over the entirety of the antibody combining site the distinctions in planarity become less clear.
31. Antibody Structure and Recognition of Antigen
This is also the case for unbound antibodies, which alludes to the degree of conformational change that is induced increasingly in the antibody as antigen size is reduced (see below). The amount of surface area on the antibody molecule buried by antigen decreases with antigen size, as less of the antibody surface is utilized to envelop the smaller antigens. Large antigens often contact antibody residues at the edge of the combining site and interact with the more apical portions of the CDR loops, while the interactions of smaller antigens are more restricted to the central portion of the antibody-combining site (MacCallum et al., 1996). A number of studies have sought to elucidate patterns in the amino acid composition in the antibody combining site (Davies and Cohen, 1996; Janin and Chothia, 1990; Kabat et al., 1977; Lea and Stuart, 1995; Padlan, 1990) and have revealed the antibody–antigen interfaces to be significantly richer in aromatic residues, particularly tyrosine and tryptophan, than the average protein surface (Conte et al., 1999). They are depleted in the charged residues aspartate, glutamate, and lysine, but are enriched in arginine. The relative abundance of arginine and aromatic residues in the antibody paratope should not be completely unexpected, however, as thermodynamic hotspots in binding are generally weighted towards these amino acid types, perhaps because they are capable of making multiple favorable interactions (Bogan and Thorn, 1998; DeLano, 2002). Structures of antibody–antigen complexes illustrate the importance of bound water molecules in mediating antigen–antibody interactions. Indeed, with only one exception (Muller et al., 1998), water molecules have been localized in the interfaces of each of the antigen–antibody complexes whose crystal structures have been determined at sufficiently high resolution (<2.5 Å) to allow the identification of ordered waters with a reasonable degree of accuracy (Bhat et al., 1994; Faelber et al., 2001; Fields et al., 1995; Kondo et al., 1999; Li et al., 2000; Mylvaganam et al., 1998). It therefore appears that water molecules are required to correct imperfections in antigen–antibody interfaces by improving the fit between the proteins and by neutralizing unpaired hydrogen-bonding groups. Bound waters, acting as molecular adaptors, may compensate for the lack of evolutionary optimization of antigen–antibody interfaces, compared to other protein–protein interfaces in which the interacting surfaces may have co-evolved to maximize complementarity (e.g., oligomeric proteins). In detailed structural analyses of the FvD1.3–HEL complex (Bhat et al., 1994; Braden et al., 1995; Braden et al., 1998), it has been shown that many water molecules from the free antibody and antigen structures are positionally conserved in the complex, that there is a recruitment of water molecules from the bulk solvent to the complex interface as demonstrated by a net gain of water molecules in the complex structures relative to the individual component structures, and that
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water molecules in the interface, regardless of their positional origin, increase the shape and chemical complementarity of the interacting surfaces.
THE ROLES OF CONFORMATIONAL FLEXIBILITY IN ANTIGEN RECOGNITION There exist a number of flexible regions within antibody molecules whose conformational plasticity is a contributing factor to the recognition of antigen. These regions include the hinge region between the Fab and Fc domains, the juxtaposition of the VL and VH domains, and the CDR loops. Conformational flexibility in the antigen epitope may also factor into the molecular recognition process. Recent structural studies have helped to elucidate some of the ways in which flexibility affects antigen recognition and are discussed below. The hinge region between the Fab and Fc domains is composed of three parts, including an upper hinge that permits Fab rotation and movement, a core hinge that allows for the possibility of interchain disulfide bonding, and a lower hinge that provides for Fc movement. Crystallographic studies of intact antibodies and Fc-Fc receptor complexes (Garman et al., 2000; Harris et al., 1992; Harris et al., 1997; Harris et al., 1998; Saphire et al., 2002; Sondermann et al., 2000) have elucidated representative structures for all these hinge region parts, revealing significant structural diversity and flexibility throughout the hinge region. This variability in the hinge angle, defined by the angles between the Fc major axis and the Fab major axes, results in a wide range of available conformations among intact antibodies as well as asymmetry within antibody molecules (Figure 31.1A). In the expanding database of unliganded and antigenbound structures of the same antibody, a number of conformational changes have been observed in mature antibodies upon binding antigen. This has allowed the identification of some trends in the induced-fit mechanism utilized for antigen recognition. In general, it appears that for smaller antigens, notably peptides and DNA, antibody plasticity is more pronounced than for protein antigens, although associations involving the latter commonly involve a nominal degree of molecular flexibility and cannot necessarily be classified as typical “lock-and-key” interactions. Atomic movements induced by antigen binding fall into a number of distinct categories, from gross domain movements to specific fluctuations in side chain rotamer positions. Flexibility in the junction between the V and C superdomains, as defined by the Fab elbow angle, has been observed with essentially equal frequency in both liganded and unliganded antibody crystal structures. Furthermore, two otherwise identical Fabs residing in the same asymmetric
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unit of a crystal have exhibited different elbow angles, revealing that changes in elbow angles are as likely to be the result of crystallographic artifacts as they are from specific antigen binding (Rini et al., 1993; Tormo et al., 1994). Thus, while elbow angle changes induced upon antigen binding were once argued to be a key component of B-cell receptor signaling, the lack of dependence of elbow angle changes on antigen binding has essentially invalidated this theory. It is also likely that, as a general rule, elbow angle diversity is irrelevant to antigen recognition. For most structural comparisons of antibodies in their antigen-bound and free forms, some degree of rearrangement of the two V domains from the heavy and light chains has been observed. One of the most striking examples of this phenomenon is for the anti-HIV-1 gp120 peptide Fab50.1 (Stanfield et al., 1993), in which a relative reorientation of the VL and VH domains of 16.3° was observed upon binding of the peptide antigen. Other anti-peptide antibodies have revealed more modest VL/VH domain rearrangements, including the antiviral capsid protein VP2 peptide Fab8F5, for which there was a 3.5° rotation (Tormo et al., 1994), and the anti-influenza virus hemagglutinin peptide Fab17/9, in which VL/VH relative domain rotations upon complexation were not significantly larger than for the C domains (Rini et al., 1992). An antibody specific for a single-stranded DNA fragment, FabBV04-01, has displayed a relatively large V domain rearrangement of 7.5° upon binding its antigen (Herron et al., 1991). Another large, 8.5°, relative VL/VH domain rotation has been observed for Fab13B5 (BerthetColominas et al., 1999; Monaco-Malbet et al., 2000). This also forms a complex with a helix-turn-helix motif in the HIV-1 capsid protein p24. Arguably, this epitope may be considered more as a continuous peptide than a protein epitope, because the helix-turn-helix extends from the antigen surface, and the interaction is characterized by a buried surface area similar to other peptide–antibody complexes. A smaller magnitude V-domain rearrangement has been observed also for FvD1.3 upon binding its protein antigen, HEL (Bhat et al., 1994). In other antibodies that bind protein antigens, relative rotations of the VL and VH domains in the antigen-bound complex were not significantly different than in their unbound forms (Faelber et al., 2001; Li et al., 2000). The relative V-domain rotation upon antigen binding also seems to bear some correlation to the buried surface area between these domains (Stanfield et al., 1993), a feature that is independent of antigen specificity. Two types of backbone movements within the antibody combining site have commonly been observed upon antibody–antigen complex formation—concerted movements of multiple residue segments of CDR loops and more heterogeneous rearrangements of CDR residues. Upon binding antigen, heavy chain CDR loops in the anti-peptide Fab8F5 undergo essentially rigid-body movements in which the unli-
ganded loop conformations are conserved, while changes in the main chain conformation of the light chain are not significant (Tormo et al., 1994) (Figure 31.2A). The largest backbone displacement, greater than 7 Å, occurs for the VHCDR3 residue Tyr102. The culmination of concerted heavy chain CDR movements towards the light chain reduces the volume of the antigen binding site by some 3% relative to the unbound Fab8F5. Other examples of segments of CDR loops moving en masse towards antigen have been observed (Stanfield et al., 1990). In Fab17/9, a significant rearrangement of the VHCDR3 loop is induced by the binding of its peptide antigen (Figure 31.2B), for which the largest backbone changes are 5 Å (Rini et al., 1992). The restructuring of CDR loop regions from both the heavy and light chains of the anti-DNA antibody FabBV04-01 have also been observed (Herron et al., 1991). Induced CDR loop movements upon antigen binding seem to be less extreme for anti-protein antibodies. Generally, these are small, concerted displacements of less than 3 Å (Bhat et al., 1994; Braden et al., 1994; Faelber et al., 2001; Li et al., 2000; Mylvaganam et al., 1998; Prasad et al., 1993) (Figure 31.2C, D). Molecular flexibility is not limited to the antibody side of the reaction, as a number of structural studies have shown varying degrees of protein plasticity for antigens upon binding. HEL can be crystallized in several space groups (Harata, 1994; Kurinov and Harrison, 1995; Ramanadham et al., 1990). A comparison of the structures reveals significant flexibility in several loops at the molecular surface, including a number of Ca atom displacements greater than 3 Å between HEL molecules from different space groups. Between crystal structures of HEL bound to different antibodies, some main chain movements become more pronounced. Relative to the D1.3–HEL complex (Bhat et al., 1994), Gly102 and Asn103 of HEL are displaced some 8 Å in complexes with the anti-HEL antibodies HyHEL-10 (Padlan et al., 1989) and D11.15 (Chitarra et al., 1993). In the HyHEL-63/HEL complex (Li et al., 2000), residues 99 to 104 of this same loop region of HEL have a root mean square deviation of 6.8 Å relative to their positions in HEL complexed with D1.3 (Bhat et al., 1994). Molecular movement in this HyHEL-63 complex is highlighted by a peptide flip at residue Asp101 that allows the formation of five hydrogen bonds between this residue and the antibody. A number of other, less extreme, examples of antigen plasticity have been documented for HEL upon antibody binding (Davies and Cohen, 1996), but the excessive flexibilty of this particular loop region may contribute to those characteristics of this surface that make it an especially favorable epitope, as evidenced by the large number of anti-HEL antibodies that share this region as part of their recognition surfaces. Indeed, structural plasticity has been observed frequently in the hotspot regions of protein–protein interfaces (DeLano, 2002). Smaller conformational changes are seen in the HIV-1 capsid protein p24 upon binding Fab13B5
31. Antibody Structure and Recognition of Antigen
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FIGURE 31.2 Conformational changes induced by antigen binding. (a) Concerted movement of the Fab8F5 CDR H3 loop induced upon binding its peptide antigen. The unbound Fab structure is in green, the bound Fab structure in blue, and the peptide antigen in yellow. VHSer101 in the bound form makes two hydrogen bonds to Lys157 of the peptide, and VHTyr102 is displaced by more than 7 Å between the unbound and bound Fab8F5 molecules. (b) Atomic rearrangement of the CDR H3 loop of the anti-peptide Fab17/9. The color scheme is the same as in (a). Side chains of residues in contact between the antibody and antigen in the bound complex are shown. Contrary to many anti-peptide antibodies, anti-protein antibodies generally exhibit relatively small conformational changes upon binding antigen as shown in panels (c) and (d) for the anti-HEL antibody FabHyHEL63. (c) Superposition of the CDR H3 loops of FabHyHEL63 bound to HEL (blue) and three different unbound forms: solved in the C2 spacegroup (green); one molecule from the asymmetric unit of the free antibody solved in the P1 spacegroup (red); and the second molecule of the asymmetric unit of the free antibody solved in the P1 spacegroup (yellow). (d) Superposition of the CDR H2 loops of FabHyHEL63 bound to HEL (blue) and three different unbound forms. The color scheme is the same as in (c). See color insert.
(Berthet-Colominas et al., 1999; Monaco-Malbet et al., 2000). Localized to the turn portion of the helix-turn-helix motif, the flexibility of the antigen is highlighted by a 4 Å displacement of the carbonyl oxygen of Pro207 that points in a direction opposite that of its unbound form to adapt to the molecular environment of the antibody. Due to the high degree of flexibility of uncomplexed peptides (Dyson and Wright, 1995), the Fab13B5-p24 complex, with its continuous peptide-like epitope, may present the best current measure for the role of peptide antigen plasticity in antibody recognition. Increased antigen flexibility, however, is not always beneficial to epitope recognition by antibodies. To produce mimics of the N-terminal sequence of transforming growth factor alpha epitope recognized by the monoclonal antibody tAb2, peptides required cyclicization to constrain their conformations to those that are suitable for binding (Hahn et al., 2001).
BINDING ENERGETICS OF ANTIBODY–ANTIGEN INTERACTIONS Most mature antibodies have affinities for their specific antigens in the range of 107 to 108 M-1, although many antibodies that recognize carbohydrates and bacterial polysaccharides fail to reach affinity levels of 106 M-1. It has been proposed (Foote and Eisen, 1995) that, due to diffusion rates and the residence time required for antibody internalization controlling on- and off-rates, there exists an affinity ceiling for antibody–antigen interactions of approximately 1010 M-1. Presumably antibodies with antigen affinities above this threshold would not be further advantaged over their lower affinity counterparts in the antibody selection process in vivo. The existence of this affinity ceiling has been demonstrated for antigen-specific B-cell transfectants. More important, an affinity window for effective B-cell
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response has been revealed for which a minimum affinity of 106 M-1 and half-life of 1s were required for detectable Bcell triggering that reached a plateau for affinities beyond 1010 M-1 (Batista and Neuberger, 1998). Not surprisingly, when primary response antibodies exhibit affinities for their specific antigens approaching this affinity ceiling, they neither require nor undergo further affinity maturation (Roost et al., 1995). Throughout the effective affinity window, the efficiency of antibody-mediated presentation of antigens to T cells is controlled by the off-rate of the antibody–antigen interaction with slower off-rates correlated to increased signaling (Guermonprez et al., 1998). Beyond this quantitative correlation between affinity and response, there exists qualitative variability in the B cell response in which some signaling responses, such as Ca++ mobilization and phosphorylation of Syk and Iga, are significantly affinity dependent whereas others, including phosphorylation of Lyn, are less correlated to antibody–antigen affinity (Kouskoff et al., 1998). All the above studies were performed with soluble antigens. The effective affinity window, however, appears to shift to a range of lower affinities, with an affinity ceiling of ~106 M-1, when the antigen is in particulate form, presumably due to avidity effects. Conversely, the range of the affinity window for extraction of antigen from a noninternalizable surface remains quite broad, with an affinity ceiling similar to that of soluble antigens (Batista and Neuberger, 2000). Antigens in these nonsoluble forms are thought to more closely mimic the properties of antigens in vivo. As the overall affinity of antibody–antigen interactions can vary by several orders of magnitude, so too can the kinetics of these interactions. In a number of kinetic analyses of anti-protein antibodies (England et al., 1997; Gerstner et al., 2002; Rajpal and Kirsch, 2000; Xavier et al., 1999), both association and dissociation rates vary by greater than two log-fold. The kinetics of antibody–antigen interactions are also commonly temperature dependent. In some cases this may be indicative of the structural plasticity involved in antigen binding. Indeed, the binding kinetics of several anti-HEL antibodies have been shown to conform to a two-state model describing induced fit, with distinct association steps for molecular encounter and docking (Li et al., 2001; Lipschultz et al., 2000). The thermodynamics of antibody–antigen interactions have also been investigated for a number of systems. With such an apparently important role for water molecules in the molecular interface, it comes as no surprise that the formation of many of these complexes reflects an enthalpically driven process with some compensating negative entropy component, such as determined by isothermal titration calorimetry for the FvD1.3–HEL association (Bhat et al., 1994). In fact, there is a strong correlation between decreases in water activity and association constants, as determined through binding analyses performed in the pres-
ence of co-solutes with polarities lower than that of water (Goldbaum et al., 1996). Although other antibody–protein antigen (Kelley et al., 1992) and antibody–carbohydrate antigen (Sigurskjold et al., 1991) interactions also appear enthalpically driven, this may not be the general rule for antibody–antigen associations, in particular due to the limited number of antibody–antigen systems whose thermodynamics have been rigorously determined. In accordance with the significance of water activity on antigen recognition, both antibodies binding to protein as well as antibodies binding to hapten antigens have shown a thermodynamic dependence on the solvent pH and ionic strength (De Genst et al., 2002; Gibas et al., 1997; Omelyanenko et al., 1993; Xavier et al., 1999).
MAPPING THE ANTIBODY–ANTIGEN ENERGETIC LANDSCAPE THROUGH MUTAGENESIS Although assessing the binding energetics of numerous antibody–antigen associations can reveal some of the generalities of the binding phenomena for this particular class of interactions, a more complete description of the antibody–antigen energetic landscape is revealed through the perturbation of these interactions and quantification of the resulting changes in the factors that regulate binding. Evaluation of the effects of various mutagenesis techniques, including alanine-scanning (Dall’Acqua et al., 1996; Dall’Acqua et al., 1998; Goldman et al., 1997; Lang et al., 2000; Pons et al., 1999), double mutant cycles (Dall’Acqua et al., 1998; Goldman et al., 1997; Pons et al., 1999), random mutagenesis (Lang et al., 2000), shotgun scanning (Vajdos et al., 2002), alanine shaving/molecular grafting (Jin and Wells, 1994), and sequence plasticity analysis (Gerstner et al., 2002), to perturb associations on a number of antibody–antigen systems has greatly enhanced our understanding of these interactions. For the purposes of highlighting the unique and diverse energetics of antibody–antigen interactions, we will limit our mutagenesis discussion to alanine-scanning mutagenesis and double mutant cycles analysis of the D1.3 antibody interactions with its cognate antigen, HEL, and the anti-idiotypic antibody, E5.2 (Dall’Acqua et al., 1996; Dall’Acqua et al., 1998; Goldman et al., 1997) (Figure 31.3). As with other mutagenesis techniques, these analyses can only provide information on the effects of changes to amino acid side chains and in no way reveal any information on the structural nature or energetic magnitude of main chain interactions. Alanine-scanning mutagenesis has been utilized to determine the energetic contributions to the complex formation of individual residues from both sides of the interface in the D1.3–HEL and D1.3–E5.2 complexes (Dall’Acqua et al., 1996; Dall’Acqua et al., 1998; Goldman et al., 1997).
31. Antibody Structure and Recognition of Antigen
Alanine substitutions into the antibody-combining site of D1.3 have revealed that only three residues in VLCDR1 and VHCDR3 (VLW92, VHD100, and VHY101) make significant energetic contributions to the binding of HEL (Dall’Acqua et al., 1996). When HEL residues in the D1.3–HEL interface were mutated to alanine and their affinities for wildtype D1.3 measured, significant changes in binding were observed for substitutions at only four contact positions (Gln121, Ile124, Arg125, and Asp119) (Dall’Acqua et al., 1998). Therefore, for both the D1.3 and HEL sides of this interface, only small subsets of the total contacting residues appear to account for a large portion of the binding energy. In the D1.3–E5.2 complex, residues located in each of the D1.3 CDR loops contribute significantly to binding, with the energetically most important residues coming from VHCDR2 (W52A and D54A) and VHCDR3 (E98A, D100A, and Y101A) (Dall’Acqua et al., 1996). Of the residues common to the D1.3 interfaces with HEL and E5.2, only five appear energetically important in the D1.3–HEL complex, whereas twelve make significant energetic contributions to D1.3 binding to E5.2. On the E5.2 side of the interface, the most destabilizing alanine substitutions are located in VHCDR3 (VHY98 and VHR100b), whereas seven other E5.2 mutations also resulted in significant effects on binding
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(Goldman et al., 1997). Thus, in contrast to the D1.3–HEL complex, interaction of D1.3 with E5.2 is regulated energetically by nearly all contact residues, even though a number of relative hotspots are clearly present. The functional surfaces of D1.3 involved in binding HEL and E5.2 can be mapped onto the three-dimensional structures of the complexes (Bhat et al., 1994; Braden et al., 1996a; Fields et al., 1995) (Figure 31.4). With the exception of VLW92, which lies at the periphery, the residues of D1.3 most important for binding HEL (VHY101, VHD100, VLY32, and VHE98) are located in a contiguous patch at the center of the combining site. Residues at the periphery make only minor contributions to the binding energy. A similar pattern is observed for the D1.3–E5.2 complex, with the most important residues (VLY32, VHW52, VHD54, VHE98, VHD100, and VHY101) forming a central band of key contacts. Only two D1.3 substitutions, VHD100A and VHY101A, significantly affect the binding to both HEL and E5.2. Thus, a single set of antibody contact residues on D1.3 can bind two antigens (HEL and E5.2) in energetically distinct ways. The residues of HEL most important for binding D1.3 (Asp119, Gln121, Ile124, and Arg125) form a contiguous patch located at the periphery of the surface contacted by
FIGURE 31.3 Structure of antibody–antigen complexes. (a) Ribbon diagram of the FvD1.3–HEL complex. HEL (yellow), D1.3 VL domain (green), and D1.3 VH domain (blue). Residues of HEL and D1.3 involved in interactions in the antigen–antibody interface are cyan and red, respectively. Heavy (H) and light (L) chain CDRs 1–3 are numbered. (b) Diagram of the FvD1.3–FvE5.2 complex. D1.3 VL domain (green), D1.3 VH domain (blue), E5.2 VL domain (yellow), and E5.2 VH domain (gray). Residues of D1.3 and E5.2 in contact in the structure are red and cyan, respectively. D1.3 and E5.2 heavy (H) and light (L) chain CDRs are labeled 1–3, with an asterisk (*) denoting CDRs from E5.2. See color insert.
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FIGURE 31.4 Energetic maps of antigen–antibody interfaces. (a) Space-filling model of the surface of D1.3 (left) in contact with HEL and of the surface of HEL (right) in contact with D1.3. The two proteins are oriented so that they may be docked by folding the page along a vertical axis between the components. Residues are color-coded according to the loss of binding free energy upon alanine substitution: red, >4 kcal/mol; yellow, 2–4 kcal/mol; green, 1–2 kcal/mol; blue, <1 kcal/mol. VL and VH residues are labeled in white, and VL residues are denoted by an asterisk (*). (b) Model of the surface of D1.3 (left) in contact with E5.2 and of the surface of E5.2 (right) in contact with D1.3. Residues are colored and labeled as in (a). See color insert.
the antibody (Dall’Acqua et al., 1998). Hotspot residues on the D1.3 side of the interface generally correspond to hotspot positions on the HEL side. Similarly, functionally less important D1.3 and HEL residues tend to be juxtaposed in the antigen–antibody interface. Residues of D1.3 and E5.2 important in complex stabilization are also found located opposite one another in molecular interface. This complementarity of functional epitopes has been observed in other protein–protein interactions (Clackson and Wells,
1995). Analysis of the D1.3–E5.2 and D1.3–HEL systems shows that both polar (e.g., D1.3 residues VHD54, VHE98, and VHD100) and nonpolar residues (e.g., D1.3 residues VLW92 and VHW52) play a prominent role in complex stabilization and that there is no clear segregation of polar residues at the periphery of the interface and of nonpolar amino acids at the core. On the basis of these studies, two broad categories of antibody–antigen protein–protein interfaces may be defined: 1) those in which ligand binding is
31. Antibody Structure and Recognition of Antigen
mediated by a small subset of contact residues and 2) those in which the free energy of binding arises from many productive interactions distributed over the entire protein–protein interface. In addition, each of these categories may be further subdivided into: 1) those that resemble cross-sections through folded proteins in which hydrophobic residues are in the interior, and hydrophilic ones at the periphery and in which productive binding is mediated largely by the former; and 2) those in which polar and nonpolar residues are evenly distributed throughout the interface and in which both residue types make comparable contributions to complex stabilization. Due to unpredictable disruptions of molecular interactions outside of the interaction of interest, the strength of an interaction between two amino acid residues in a protein or protein–protein complex cannot necessarily be measured by simply mutating one of them (Ackers and Smith, 1985; Fersht, 1988). Thus, comparing the binding of a wildtype protein with that of a mutant in which a side chain has been truncated gives an apparent binding energy that is generally greater than the incremental binding energy attributable to that side chain. A more sophisticated approach to dissecting the energetics of pairwise interactions makes use of double mutant cycles (Ackers and Smith, 1985; Serrano et al., 1990). Double mutant cycles have been constructed for amino acid pairs in the D1.3–HEL (Dall’Acqua et al., 1998) and D1.3–E5.2 (Goldman et al., 1997) interfaces to measure interaction energies (DDGint) for interacting, proximal, and distant side chains, as judged from the crystal structures of the complexes (Bhat et al., 1994; Braden et al., 1996a; Fields et al., 1995). In the D1.3–HEL complex, only three of the ten residue pairs in direct contact in the crystal structure exhibited significant coupling energies. Indeed, with the exception of one proximal residue pair, none of the remaining residue pairs revealed coupling energies exceeding experimental error and, in this way, they are energetically indistinguishable from the residue pairs that do not form direct contacts. The broad distribution of energetically important residues in the D1.3–E5.2 interface, as revealed by alanine-scanning mutagenesis (Dall’Acqua et al., 1996; Goldman et al., 1997), is mirrored in the results of double mutant cycle analysis of this interface (Goldman et al., 1997). All residue pairs tested exhibited significant coupling energies, including both electrostatic and hydrophobic atomic interactions. Even proximal and distant residues pairs in the D1.3–E5.2 complex exhibited coupling energies significantly greater than experimental error. Small-magnitude energetic coupling between amino acid residues separated by large distances has likewise been observed in protein folding (Green and Shortle, 1993; LiCata and Ackers, 1995) and protein–protein interaction (Schreiber and Fersht, 1995) systems. One possible explanation for this phenomenon is that the mutations may introduce solvent
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rearrangements in the D1.3–E5.2 interface, such as described for complexes between mutants of D1.3 and HEL (Braden et al., 1996a; Fields et al., 1995; Sundberg et al., 2000); these localized molecular changes may result in global perturbations in electrostatic fields or vibrational modes within the interface. It is clear from double mutant cycle analysis of the D1.3–HEL and D1.3–E5.2 complexes that neither direct atomic contacts within an antibody–antigen interface necessarily imply energetically productive interactions nor do energetically nonproductive interactions always arise from residues separated by some distance within such an interface. The findings from these energy analyses reveal that the D1.3–HEL and D1.3–E5.2 interactions differ markedly. In the D1.3–HEL complex, most residue pairs in direct contact in the crystal structure exhibit no significant energetic coupling. With one exception, none of the hydrogen bonds in this interface makes significant net contributions to complex stabilization. These include hydrogen bonds that, on the basis of donor–acceptor distance and relative orientation of the interacting groups, would be predicted to be strong, as well as those expected to be weak. On the contrary, in the D1.3–E5.2 interface, nearly all residues within 4 Å of each other show significant coupling (>1.0 kcal/mol). With the exception of weak interactions between certain noncontacting residues, the relative strengths of coupling energies in the FvD1.3–FvE5.2 interface are broadly consistent with expectations based on the three-dimensional structure. Thus, the highest coupling energy (4.3 kcal/mol) was measured for a charged–neutral pair forming a buried hydrogen bond. For residues interacting through solvent-exposed hydrogen bonds, coupling energies were approximately 1.7 kcal/mol, regardless of whether a neutral–neutral or charged–neutral pair was involved. Interactions formed by solvent-mediated hydrogen bonds were energetically neutral, whereas coupling energies of about 1.5 kcal/mol were measured for residues engaged only in van der Waals contacts. These results demonstrate that considerable caution should be exercised when attempting to estimate the strengths of specific interactions in an antigen–antibody or any other protein–protein interface on the basis of three-dimensional structures alone.
ACCOMMODATION OF MUTATIONS IN THE ANTIBODY–ANTIGEN INTERFACE Alanine-scanning mutagenesis and double mutants cycles of the D1.3–HEL interface have demonstrated that it is remarkably tolerant to mutations that, on the basis of the three-dimensional structure of the wildtype complex, might be expected to have pronounced effects on affinity. For example, the truncation of HEL residue Asp18 to alanine
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should result in the loss of a direct hydrogen bond to the side chain of D1.3 VLTyr50 (Asp18HEL Od2-Oh D1.3 VLTyr50), and the loss of seven van der Waals contacts to this residue. Nevertheless, the affinity of HEL D18A for D1.3 (4.5 ¥ 107 M-1) is nearly identical to that of the wildtype (8.0 ¥ 107 M-1), corresponding to a DDGint of only 0.3 kcal/mol. The crystal structure of the FvD1.3–HEL D18A complex at 1.5 Å resolution (Dall’Acqua et al., 1998) reveals that the mutation does not cause significant conformational changes at the site of the mutation or in the overall structure. Instead, the loss of complementarity in the D1.3–HEL interface resulting from replacement of Asp18HEL by alanine is compensated for by the stable inclusion of additional water molecules and by local rearrangements in solvent structure (Figure 31.5). Similar mechanisms may explain the tolerance of the D1.3–HEL interface to mutations at other solvent-accessible sites on the antigen, such as Ser24HEL and Lys116HEL, which also make multiple contacts with D1.3.
Solvent rearrangements, including the incorporation of additional interface waters, have also been observed in structural studies of other site-directed mutants of FvD1.3 in complex with HEL, including VLY50S, VHY32A, and VLW92D (Fields et al., 1996; Ysern et al., 1994). In these cases, however, the mutations are only partially compensated by the solvent rearrangements, since the VLY50S, VHY32A, and VLW92D mutant fragments bind HEL with 10-, 4-, and 1,000-fold lower affinities, respectively, than the original antibody. Thus, there appears to be a wide range in the extent to which solvent rearrangements in an antibody–antigen interface can accommodate mutations, which probably depends on the nature of the local environment. In this respect, it is interesting to note that VLTyr50 and Asp18HEL are juxtaposed in the complex structure. This suggests that these two residues define a site in the interface that, perhaps because of its peripheral location, is particularly suited for the stable incorporation of new waters to occupy cavities or channels created by side chain truncations. In other cases, seemingly conservative mutations have been found to greatly affect antigen–antibody binding. For example, the substitution of lysine for arginine at HEL position 68 in the HyHEL-5/HEL interface produced a 1,000fold decrease in affinity. Comparison of the crystal structures of the mutant and wildtype complexes revealed only small rearrangements in the vicinity of the mutation, with little change in buried surface area. A bound water molecule replaces the two nitrogens of the guanidinium group of the arginine and partially compensates for the loss of two salt bridges between Arg68HEL and HyHEL-5 VHGlu50 (Chacko et al., 1995). The net consequence of the mutation is a loss of hydrogen bonding, since the side-chain amino group of the lysine cannot form the same bonding arrangement as the guanidinium group of the arginine.
CROSS-REACTIVITY AND SPECIFICITY OF ANTIGEN RECOGNITION
FIGURE 31.5 Solvent rearrangement in an antigen–antibody interface induced by a mutation in the antigen. (a) Schematic representation of the FvD1.3–HEL Asp18->Ala complex in the vicinity of the mutation. (b) Schematic showing the same region in the wildtype FvD1.3–HEL complex. Water molecules present in both structures are labeled WAT1–4. WATa, WATb, and WATc are additional waters in the FvD1.3–HEL Asp18->Ala interface.
Although antibodies are commonly highly specific for a single antigen, it is not at all uncommon for antibodies to cross-react with many, structurally similar, yet distinct, antigenic molecules. In some cases, antibodies can bind better to antigens not used in challenging the immune system than to the original immunogen, a phenomenon known as heteroclitic binding. The monoclonal antibody (mAb) D11.15, which was raised against HEL, interacts with higher affinity with several other avian lysozymes. The structural basis for cross-reactivity in the D11.15–lysozyme system has been probed by energetic and structural analysis (Chitarra et al., 1993). FvD11.15 binding to eight different avian lysozymes was tested, and all exhibited high affinity for the antibody; two of these, pheasant egg-white lysozyme (PHL) and guinea fowl egg-white lysozyme (GEL), exceeded the affin-
31. Antibody Structure and Recognition of Antigen
ity of the interaction with HEL. The crystal structures of PHL, GEL, Japanese quail egg-white lysozyme (JEL), and the FvD11.15–PHL complex were also determined. The affinity of JEL for FvD11.15 is slightly lower than that of HEL (1.5 ¥ 109 M-1 versus 4.0 ¥ 109 M-1), which is likely derived from two amino acid differences in the loop region from residue 100 to 104 that forms part of the epitope in the FvD11.15–PHL complex. Whereas residue 102 is a Gly in HEL, it is a Val in JEL, whereas residue 103 is an Asn in HEL and a His in JEL. The result of these changes is a displacement of the 100–104 loop region by 7.5 Å into a conformation that would likely clash sterically with the VHCDR3 loop of FvD11.15 (Figure 31.6A). Conversely, two amino acid differences between PHL and HEL, at residues 113 (Asn in HEL, Lys in PHL) and 121 (Gln in HEL, Asn in PHL), confer higher affinity to the FvD11.15–PHL complex relative to the complex with HEL by two orders of magnitude. The crystal structure of FvD11.15–PHL reveals that the major structural difference in these two complexes is that Lys113 in PHL makes several nonpolar contacts with VHTyr57 (Figure 31.6B). Another anti-HEL antibody, D1.3, binds only its immunogen and one other avian lysozyme, bobwhite quail egg-white lysozyme (BEL), with high affinity. Much of the sequence variability between the eight lysozymes tested occurs at HEL residue Gln121. For the highly cross-reactive D11.15, lysozyme residue 121 is located at the periphery of the antigenic epitope. Conversely, for the highly specific D1.3, this residue is located centrally to the binding interface and acts as a hotspot in binding for the D1.3–HEL complex (Dall’Acqua et al., 1996). One of the avian lysozymes that binds poorly to D1.3, turkey egg-white lysozyme (TEL), has been investigated structurally (Braden et al., 1996b). HEL and TEL differ only at residue 121, which is a His in TEL, with a concomitant decrease in affinity by two orders of magnitude, due primarily to a reduction in the on-rate of the interaction. Whereas Gln121 of HEL makes two hydrogen bonds to VL domain main-chain atoms, TEL His121 makes only one hydrogen bond to the antibody light chain and induces a peptide flip between residues VLW92 and VLS93, a conformational change that is likely responsible for the slower on-rate of the interaction (Figure 31.6C). Anti-idiotopic antibodies (Pan et al., 1995; Poljak, 1994) recognize an antigenic determinant that is unique to an antibody or group of antibodies, or idiotope. An idiotope is defined functionally by the interaction of an anti-idiotopic antibody (Ab2) with an antibody (Ab1) bearing the idiotope. Conventional Ab2 antibodies recognize idiotopes outside of the antibody-combining site paratope, whereas internal image Ab2 antibodies are able to mimic the molecular surface encountered by Ab1, thereby mimicking stereochemically the antigen specific for Ab1. Numerous efforts have been made to use these molecular mimics as thera-
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peutics, similar to vaccines. Several structural studies have been performed detailing a diverse range of idiotope–antiidiotope interactions, including analysis of the anti-idiotypic antibody system that recognizes angiotensin II (AII), an octapeptide hormone. A mAb (Ab1) raised against AII was used to obtain anti-Ab1 polyclonal antibodies (Ab2s) that were, in turn, used to obtain an anti-anti-Ab1 mAb (Ab3) (Budisavljevic et al., 1988). In this system, Ab1 binds specifically to AII, Ab2s to Ab1, Ab3 to Ab2s and, presumably due to molecular mimicry, Ab3 to the original antigen, AII. The crystal structure of the Ab3–AII complex (Garcia et al., 1992b) and sequence comparison of the binding determinants of Ab1 and Ab3 (Garcia et al., 1992a) have shown that the original antibody and its anti-anti-idiotypic antibody are superimposable. Contact residues of Ab3 are highly conserved in the Ab1 sequence, even though the two antibodies are derived from distinct germline genes. Thus, although the structures of Ab2s could not be determined due to their molecular diversity, these polyclonal antibodies would appear to satisfy the stereochemical requirements for molecular mimicry at the atomic level, thereby producing an internal image of the antigen. As discussed earlier, the D1.3 antibody binds to two structurally distinct ligands—its cognate antigen, HEL, and the anti-idiotypic antibody E5.2—and these interactions exhibit molecular mimicry (Figure 31.3). The crystal structures of the complexes formed by FvD1.3 with both HEL (Bhat et al., 1994) and FvE5.2 (Braden et al., 1996a; Fields et al., 1995) have been determined to high resolution. FvD1.3 contacts HEL and FvE5.2 through essentially the same set of combining site residues and most of the same atoms. Of the eighteen FvD1.3 residues that contact FvE5.2 and the seventeen that contact HEL, fourteen are in contact with both FvE5.2 and HEL. These fourteen FvD1.3 residues make up 75% of the total contact area with FvE5.2 and 87% of that with HEL. Furthermore, the positions of the atoms of FvE5.2 that contact FvD1.3 are close to those of HEL that contact FvD1.3. Six of the twelve hydrogen bonds in the FvD1.3–FvE5.2 interface are structurally equivalent to hydrogen bonds in the FvD1.3–HEL interface.
MODEL SYSTEMS FOR PROBING FUNDAMENTAL RULES OF MOLECULAR INTERACTIONS Although our understanding of antibody–antigen interactions is not complete, it has attained a level that justifies their use as a model system for probing some of the fundamental rules governing molecular recognition. Studies that involve extensive mutagenesis combined with rigorous structural and energetic analysis have the potential to yield great strides in our efforts to understand the numerous phenomena, such as van der Waals interactions, hydrogen bonding,
FIGURE 31.6 Cross-reactivity of antibodies. (a) Interaction of FvD11.15 (VH domain in blue, VL domain in green) with PHL (yellow) and JEL (red). The left panel is a close-up view of the encircled region in the right panel, highlighting the relative displacement of the 100–104 loop region between PHL and JEL, resulting in a steric clash between JEL residues Val102 and His103 with the FvD11.15 VH domain. (b) Interaction between FvD11.15 [same color scheme as in (a)] with PHL (yellow) and HEL (red). The left panel is a close-up view of the encircled region in the right panel and highlights the productive interactions that are made between FvD11.15 VHTyr57 and PHL Lys113 (four hydrogen bonds, indicated by dotted lines). Conversely, productive interactions between FvD11.15 VHTyr57 and HEL Asn113 are largely absent (one hydrogen bond, not shown for clarity) and is likely the reason for the binding affinity discrepancy between the two antigens. (c) Hydrogen bonding between FvD1.3 residues VLTyr32, Phe91, Trp92, and Ser93 with HEL Gln121 (left panel) and TEL His121 (right panel), the only amino acid difference between these two antigens. HEL Gln121 makes three hydrogen bonds (indicated by dotted lines) to the main chain nitrogen atom of Ser93, the main chain oxygen atom of Phe91, and the phenyl ring of Tyr32. All three hydrogen bonds are lost in the FvD1.3–TEL complex; however, a peptide flip between FvD1.3 residues Trp92 and Ser93 results in a new hydrogen bond between the TEL His121 side chain and the main chain oxygen atom of Trp92. See color insert.
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31. Antibody Structure and Recognition of Antigen
hydrophobic packing, shape and charge complementarity, cooperativity and plasticity, that are known to regulate molecular interactions. One such study has taken advantage of the relative accommodation of mutations of the hotspot VLTrp92 residue in D1.3 to estimate the magnitude of the hydrophobic effect in an antigen–antibody interface (Sundberg et al., 2000). By replacing VLTrp92 with residues bearing increasingly smaller side chains, and determining the crystal structures and thermodynamic parameters of binding for each of the resulting mutant FvD1.3–HEL complexes, a correlation between the binding free energy and the apolar surface area corresponding to 21 cal mol-1 Å-2 has been demonstrated. This value is in excellent agreement with transfer free energy values for small hydrophobic solutes (Chothia, 1976; Eisenberg and McLachlan, 1986; Ooi et al., 1987) and is lower than the hydrophobic stabilization energy for folding (Eriksson et al., 1992; Kellis et al., 1989; Matsumura et al., 1988; Shortle et al., 1990; Xu et al., 1998; Yutani et al., 1987) and theoretical estimates of the interfacial free energy of protein–protein interactions (Nicholls et al., 1991; Sharp et al., 1991). Furthermore, changes in binding free energy are derived almost entirely from changes in the solvent entropy, demonstrating that the exclusion of solvent from the molecular interface at position VL92 is the predominant energetic factor in the formation of this protein complex. Notably, residues at position VL92 are partially solvent-exposed at the periphery of the interface. This may contribute to the agreement between the magnitude of the effective hydrophobicity measured for this protein–protein interaction and transfer free energy values of hydrophobic solutes. A similar mutational analysis of a hydrophobic hotspot residue in the center of a protein–protein binding interface may yield hydrophobicity values closer to those for protein folding stabilization due to cavity formation at the interface, or values closer to those measured for the partially solvent-exposed residue VL92, due to accommodation by rearrangement and addition of interfacial water molecules.
TOWARD A STRUCTURAL BASIS OF ANTIBODY AFFINITY MATURATION The function of the immune system is dependent on the recognition of essentially any antigenic material, yet the structural diversity of antigens greatly outweighs the genetic diversity encoded by immune system genes. Thus, molecular recognition of diverse antigens is accomplished by producing in two ways antibodies with high affinity and specificity for almost any antigen. Recombination and imprecise joining of antibody gene segments at the pro- or pre-B cell stage focuses molecular diversity at the center of the antibody combining site and results in germline antibodies of relatively low affinity and specificity (Tonegawa,
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1983). This junctional diversity in the primary repertoire can produce CDR loops of different lengths and varying structures (Chothia et al., 1992; Tomlinson et al., 1995). It has been shown that the presence or absence of certain VHCDR3 junctional amino acids can determine the affinity maturation pathway of an antibody by biasing subsequent amino acid replacements by somatic hypermutation (Furukawa et al., 1999). These effects are correlated to the structure and flexibility of the VHCDR3 loop in the germline antibodies (Furukawa et al., 2001). The somatic hypermutation of antibody V-regions spreads the structural diversity generated by gene segment recombination to regions at the periphery of the binding site (Tomlinson et al., 1996). Selective expansion of antibody clones on the basis of antigen affinity produce mature antibodies that are high in both affinity and specificity (Rajewsky, 1996). Somatic hypermutation is primarily a point mutation process in gene regions that are highly conserved in the primary repertoire, and it can result, at times, in codon insertions or deletions (Tomlinson et al., 1996). Structural and energetic studies comparing germline and mature antibodies bound to the same antigen have advanced our understanding of the effects of somatic hypermutation on antibody affinity maturation. The mature Fab48G7 and its germline counterpart, Fab48G7g, both bind a nitrophenyl phosphonate transition-state analog, but with a 30,000-fold difference in affinity, primarily due to a decrease in the dissociation rate (Wedemayer et al., 1997a). The sequence differences between the Fabs are limited to nine somatic hypermutations, six in VH and three in VL, located up to 15 Å from the bound hapten. The crystal structures of the unliganded germline Fab48G7g and its complex with hapten (Wedemayer et al., 1997a) reveal large conformational changes induced upon antigen binding. Conversely, crystal structures of the mature Fab48G7 (Patten et al., 1996; Wedemayer et al., 1997b) in its free and hapten-bound forms exhibit very few conformational changes upon complex formation. Relative rotations of the VL and VH domains of only 0.44° were found for the mature Fab structures, whereas the germline Fab structures had a relative VL/VH domain rotation of 4.6° between free and bound species. Importantly, the conformational changes induced upon binding antigen by Fab48G7g are later observed in the mature Fab structure even in the absence of antigen (Figure 31.7). It appears, at least in the case of the Fab48G7 system, that the affinity maturation process is driven in large part by a mechanism of pre-organizing the antibody combining site into a conformation that is favorable for binding its hapten antigen. Through the introduction of forward and back mutations in the germline and mature Fabs by site-directed mutagenesis, and measurements of binding affinities, the effects of the nine somatic hypermutations on the affinity maturation pathway of Fab48G7 have been deconstructed (Yang and
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FIGURE 31.7 Affinity maturation via preorganization of the antibodycombining site in Fab48G7. Superposition of the CDR loops of the free germline Fab48G7 (green), the antigen bound germline Fab48G7 (red), the unliganded mature Fab48G7 (blue), and the liganded mature Fab48G7 (yellow). The conformational changes invoked upon antigen binding by the germline antibody are commonly replicated in the mature antibody, especially for VLCDR1, VLCDR2, VLCDR3, and VHCDR3. See color insert.
Schultz, 1999). In this system, the effect on binding of the individual mutations was either positive or neutral, yet their additive changes in affinity were not equal to the overall change in affinity between the germline and mature Fabs. Double mutations revealed a high degree of cooperativity between mutations, not only between individually neutral mutations but also between even the two most positive individual mutations. Cooperativity between somatic hypermutations, however, does not appear to be a required mechanism for affinity maturation. For Fab39-A11, which catalyzes a Diels-Alder reaction, only two somatic mutations exist between the germline and mature counterparts, of which only one contributes the majority of binding affinity to mature Fab (Romesberg et al., 1998). Another catalytic antibody, AZ-28, which catalyzes an oxy-Cope rearrangement, has six somatic mutations, five of which contribute to differences in affinity between germline and mature antibodies in a strictly additive way (Ulrich et al.,
1997). In the affinity maturation of an anti-protein antibody, FvD1.3, the five somatic hypermutations have also been shown to be energetically additive (England et al., 1999). In this system, changes in antigen affinity are dominated by the only mutated amino acid in direct contact with the antigen, HEL. The number and cooperativity of somatic hypermutations appears to be dependent on the affinity differences between the germline and mature antibodies. Although the affinity discrepancy between Fab48G7 and Fab48G7g is 30,000fold (Wedemayer et al., 1997a), FabAZ-28, with only five significant somatic mutations, has an antigen affinity only 40-fold greater than its germline counterpart (Ulrich et al., 1997). Furthermore, Fab39-A11 and Fab39-A11g, with only one significant amino acid difference, both bind nine haptens, for most of which the difference in affinity is within an order of magnitude (Romesberg et al., 1998). Germline and mature FvD1.3 also differ by only five amino acids and by 60-fold in affinity (England et al., 1999). If one considers that mature antibodies must break a minimum affinity threshold for antigen binding through a limited number of somatic mutations to be functional in vivo, then it follows that the number of somatic mutations will increase as the difference in affinities between germline and mature antibodies gets larger. Cooperativity between the somatic mutations will be utilized in cases where the affinity maturation process must overcome extreme germline–mature affinity discrepancies. It is important to note that nearly all studies of affinity maturation to date have been confined to antibodies specific for haptens, rather than proteins, which constitute the major class of biological antigens. Whereas haptens characteristically bind in a cleft between the CDR3s of the H and L chains, protein antigens generally occupy the entire antibody-combining site, contacting all six CDRs. Moreover, the physicochemical properties of haptens are obviously different from those of the complex arrays of amino acids that form epitopes on protein surfaces. Recently, the crystal structures of four closely related anti-HEL antibodies (HyHEL8, HyHEL10, HyHEL26, and HyHEL63), representing different stages of affinity maturation, were determined bound to the same site on HEL (Li et al., 2003). These X-ray snapshots of the affinity maturation process reveal that enhanced binding is achieved not through the formation of additional hydrogen bonds, salt bridges, or van der Waals contacts, nor by a net increase in total buried surface area, but by the burial of increasing amounts of apolar surface (at the expense of polar surface), accompanied by improved shape complementarity. The increase in hydrophobic interactions, which can fully account for the 30-fold affinity improvement in these anti-HEL antibodies, is the consequence of subtle, yet highly correlated, structural rearrangements in antibody residues at the periphery of the interface with antigen, adjacent to
31. Antibody Structure and Recognition of Antigen
the central energetic hotspot, whose structure remains unaltered. In particular, VHCDR1 and VHCDR2 undergo rigid body displacements of up to 3 Å over the course of affinity maturation; this increases the amount of apolar surface buried at the VH–HEL interface from 460 Å2 in the lowestaffinity antibody (40% of the total buried surface at this interface) to 602 Å2 in the antibody with highest affinity (50% of the total). Concomitant with these changes, the Sc index of the VH–HEL interface increases from 0.69 to 0.78, in parallel with affinity, whereas the VL–HEL interface is unchanged. Thus, increasing hydrophobic interactions and improving the fit at peripheral sites that have not been optimized for binding, and whose plasticity and ability to accommodate mutations render them permissive to such optimization, constitute effective strategies for maturing anti-protein antibodies. Some of the energetic factors involved in the preorganization of mature antibodies through somatic hypermutation of germline antibodies have been elucidated recently using surface plasmon resonance techniques. Using this method, different binding characteristics of the same complex at various temperatures provide information about the relative enthalpic and entropic contributions to the interaction. The affinities of panels of early primary and secondary response mAbs for a model synthetic 40-mer peptide were determined at two temperatures (Manivel et al., 2000). Although the effects of temperature on the dissociation step of the interaction were similar for mAbs in both panels, opposite temperature effects on association were observed for each panel of mAbs. For primary mAbs, complex association was enthalpically highly favorable but entropically unfavorable, whereas dissociation was enthalpically unfavorable and entropically favorable. The equilibrium binding for primary mAbs was enthalpically driven, with a large entropic cost of complex formation, resulting in relatively low affinity. Conversely, in secondary mAbs, association was enthalpically unfavorable but the entropic costs had been reduced dramatically. Because the dissociation step of the reaction was similar to that for primary mAbs, equilibrium binding in the secondary mAbs was essentially independent of enthalpy effects, and instead, was driven by entropic changes. Thus, the relative high affinity of the secondary mAbs is derived exclusively from the nearly complete abolishment of any entropic costs of complex association, in comparison to the primary mAbs. Although these experiments seem to confirm the idea of antibody affinity maturation through paratope preorganization, at least for an anti-peptide antibody, it is intriguing to note that the increased affinities in the antihapten Fab48G7 and the anti-protein FvD1.3 systems derive nearly entirely from decreases in the dissociation phases of the reactions (England et al., 1999; Wedemayer et al., 1997a). Although similar experiments examining enthalpy and entropy effects on antigen binding to germline and mature Fab48G7 and FvD1.3 have not been performed, it is
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likely that these types of experiments would reveal that these complexes are stabilized due to large entropic barriers to dissociation in the mature versus germline antibodies. Although a significant amount of experimental findings have been generated in investigating the structural basis of affinity maturation by somatic hypermutation, much of the data appear to be contradictory. It seems clear that no one model explains the affinity maturation process, as a great deal of variability exists in the location of energetically important somatic mutations, the cooperativity amongst these mutations, and the entropic control of antigen association and dissociation. It may be that the rules governing antibody affinity maturation of antibodies will be distinct according to antigen size and type (whether hapten, peptide, or protein), to affinity differences between germline and mature antibodies, or to some other as yet undetermined factor.
BREAKING THE AFFINITY CEILING THROUGH IN VITRO EVOLUTION Although there appears to be an affinity ceiling for antibodies in vivo (Batista and Neuberger, 1998; Roost et al., 1995), production by in vitro evolution of Fv fragments with antigen affinities well above this affinity threshold has implicated further conformational refinement as the basis for extending improvements in antigen affinity. A variant of Fab4-4-20 has been produced in vitro by yeast display that has femtomolar affinity for the hapten fluorescein (Boder et al., 2000), well beyond the in vivo affinity ceiling. This antibody contains ten mutations, nine of which are in the VH chain, six of which are in the CDR H3 loop, and only one of which is in contact with the antigen as modeled using the Fab4-4-20-fluorescein structure (Herron et al., 1994). Phage display affinity maturation of another anti-hapten antibody, Fab17E8 (Arkin and Wells, 1998), has also demonstrated the importance of mutations in residues beyond the zone of antigen-contacting residues in improving the affinity of mature antibodies. Several anti-protein antibodies have exhibited susceptibility to affinity maturation through point mutations in either their VL or VH CDR3 loops, including 16-, 14-, and 8-fold affinity improvements in anti-erbB2 (Schier et al., 1996), -VEGF (Chen et al., 1999), and -gp120 (Barbas et al., 1994) antibodies. It is likely that these affinity-evolved antibodies that bind antigen above the in vivo affinity ceiling accomplish this feat by a similar structural preorganization of the antibody-combining site as by somatic hypermutation, although experiments showing this have yet to be performed. Thus, it seems that antibodies might indeed have the intrinsic in vivo ability to break the affinity ceiling through further rounds of somatic hypermutation, but do not do so only because it would be superfluous to the functioning of the immune system.
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CONCLUSION Extensive structural and energetic analyses of antibody–antigen interactions have revealed and described many of the fundamental aspects of antigen recognition by antibodies. The architecture of an antibody molecule is clearly conducive to the recognition of the vast diversity of molecules that may be encountered during the immune response. The close juxtaposition of hypervariable CDR loops from two Ig domains allows for high affinity antigen binding through shape and chemical complementarity of the antibody-combining site and the antigenic epitope. Antibody–antigen interfaces have proved to be structurally diverse, displaying a range of structural attributes from those that resemble folded protein cores to those that have highly distributed densities of the polar and apolar residues. Although significant flexibility in antibody molecules likely does not contribute to antigen recognition per se, structural plasticity in regions of antibody molecules within and proximal to the antibody combining site can be a key determinant in antigen recognition, where VL-VH domain rotations and CDR loop rearrangements roughly correlate in magnitude to the size of the antigen. The binding affinities of mature antibodies for their specific antigens fall into a defined range due to diffusion rates and the residence time required for antibody internalization. Although a structural basis of antibody affinity maturation is slowly emerging, the universality of preorganization of the antibody combining site and the relative impact on antigen recognition of alterations to antibody–antigen complex association and dissociation rates remain to be determined.
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Ulrich, H. D., Mundorff, E., Santarsiero, B. D., Driggers, E. M., Stevens, R. C., and Schultz, P. G. (1997). The interplay between binding energy and catalysis in the evolution of a catalytic antibody. Nature 389, 271–275. Vajdos, F. F., Adams, C. W., Breece, T. N., Presta, L. G., de Vos, A. M., and Sidhu, S. S. (2002). Comprehensive functional maps of the antigenbinding site of an anti-ErbB2 antibody obtained with shotgun scanning mutagenesis. J Mol Biol 320, 415–428. Webster, D. M., Henry, A. H., and Rees, A. R. (1994). Antibody-antigen interactions. Curr Opin Struct Biol 4, 123–129. Wedemayer, G. J., Patten, P. A., Wang, L. H., Schultz, P. G., and Stevens, R. C. (1997a). Structural insights into the evolution of an antibody combining site. Science 276, 1665–1669. Wedemayer, G. J., Wang, L. H., Patten, P. A., Schultz, P. G., and Stevens, R. C. (1997b). Crystal structures of the free and liganded form of an esterolytic catalytic antibody. J Mol Biol 268, 390–400. Wilson, I. A., and Stanfield, R. L. (1993). Antibody-antigen interactions. Curr Opin Struct Biol 3, 113–118. Wilson, I. A., and Stanfield, R. L. (1994). Antibody-antigen interactions: new structures and new conformational changes. Curr Opin Struct Biol 4, 857–867. Wu, T. T., and Kabat, E. A. (1970). An analysis of the sequences of the variable regions of Bence Jones proteins and myeloma light chains and their implications for antibody complementarity. J Exp Med 132, 211–250. Xavier, K. A., McDonald, S. M., McCammon, J. A., and Willson, R. C. (1999). Association and dissociation kinetics of bobwhite quail lysozyme with monoclonal antibody HyHEL-5. Protein Eng 12, 79– 83. Xu, J., Baase, W. A., Baldwin, E., and Matthews, B. W. (1998). The response of T4 lysozyme to large-to-small substitutions within the core and its relation to the hydrophobic effect. Protein Sci 7, 158– 177. Yang, P. L., and Schultz, P. G. (1999). Mutational analysis of the affinity maturation of antibody 48G7. J Mol Biol 294, 1191–1201. Ysern, X., Fields, B. A., Bhat, T. N., Goldbaum, F. A., Dall’Acqua, W., Schwarz, F. P., Poljak, R. J., and Mariuzza, R. A. (1994). Solvent rearrangement in an antigen-antibody interface introduced by sitedirected mutagenesis of the antibody combining site. J Mol Biol 238, 496–500. Yutani, K., Ogasahara, K., Tsujita, T., and Sugino, Y. (1987). Dependence of conformational stability on hydrophobicity of the amino acid residue in a series of variant proteins substituted at a unique position of tryptophan synthase alpha subunit. Proc Natl Acad Sci U S A 84, 4441–4444.
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32 Monoclonal Antibodies from Display Libraries JAMES D. MARKS Departments of Anesthesia and Pharmaceutical Chemistry, San Francisco General Hospital, San Francisco, California, USA
Until approximately 10 years ago, monoclonal antibodies were produced using hybridoma technology (Figure 32.1A). Despite their utility, however, monoclonal antibodies from hybridomas share a number of characteristics that can limit their use as reagents, diagnostics, and therapeutics. Monoclonal antibodies are time-consuming to make—initial injection and periodic boosts can require weeks to months, and hybridoma construction and screening require additional time. The fusion process is relatively inefficient, typically resulting in the generation of a limited number of unique antibodies. In addition, the immune response can be biased towards certain “immunodominant” epitopes, making it difficult or impossible to produce monoclonals with the precise specificity desired for a particular aim. Furthermore, production of antibodies against proteins conserved between species may be difficult or impossible. The maximum binding affinities of monoclonal antibodies— limited by B-cell biology and seldom more than nanomolar [1–3]—may also be inadequate for many diagnostic and therapeutic applications. For therapeutic applications, murine antibodies are immunogenic when administered to humans, resulting in the (human anti-mouse antibodies) (HAMA) response [4, 5]. This leads to an increase in clearance of the foreign antibody, a decrease in efficacy with repeat administration, and the potential for allergic reactions or serum sickness. Unfortunately, the adaptation of hybridoma technology to produce human monoclonals has been largely unsuccessful [6]. To reduce immunogenicity, murine monoclonals can be “chimerized” (by grafting the murine V-domains onto human constant domains) [7–9] or “humanized” (by grafting the murine complementarity determining regions onto human framework regions) [10]. This is laborious and not necessarily straightforward. For chimerization or humanization, the DNA encoding the antibody heavy (VH)
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and light (VL) chain variable region genes must be cloned from the hybridoma cell for expression as an Fv, Fab, or single chain Fv (scFv) antibody fragment (Figure 32.2). Such validation is necessary to ensure that the correct VH and VL genes have been isolated. Mutations introduced by the somatic hypermutation machinery into the regions where the primers anneal may make PCR amplification difficult or impossible, thus necessitating another amplification approach such as RACE or oligoligation PCR [11–13]. Cloning the correct VH and VL can also be complicated by the presence of several immunoglobulin transcripts, some of them arising from the fusion partner [14]. PCR may also introduce mutations coding for stop codons or destabilizing amino acids, thus necessitating the sequencing of multiple clones. Furthermore, the V genes may be cut internally by restriction enzymes, especially when cloned sequentially using hexanucleotide recognition sites [15, 16]. Once the V genes have been successfully cloned, the antibody fragment should be expressed and characterized biophysically and biochemically prior to chimerization or humanization. The most rapid method for expression is in Escherichia coli (Figure 32.3). However, expression levels in bacteria vary considerably [17] due to antibody fragment toxicity and poor folding kinetics [18]. These differences are sequence dependent, differ dramatically between antibodies, and in many instances result in a failure to produce adequate quantities of antibody fragment for further in vitro characterization [18]. Thus, despite the large number of well characterized hybridoma cell lines, very few of these antibodies can be successfully reconstructed as antibody fragments that express at high levels in E. coli. Thus, secretion in a eukaryotic cell line is necessary to prove the antibody fragment is functional and for biochemical and biophysical characterization [19, 20].
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Copyright 2004, Elsevier Science (USA). All rights reserved.
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FIGURE 32.1 Generation of monoclonal antibodies using hybridoma technology and phage display. Hybridoma technology (Panel A, left): The naïve mouse generates a primary repertoire of more than 106 rearranged VH and VL genes (colored bars) in B cells, coding for antibodies that are displayed as membrane-bound molecules. Immunization causes antigen-driven proliferation and somatic hypermutation (“stars” within V genes). To make hybridomas, B cells are harvested from the spleen or marrow and fused with immortal myeloma cells (wrinkled edges) to generate immortalized, antibody-secreting hybridomas. Hybridomas are screened by ELISA for antigen binding and the monoclonal antibodies produced in tissue culture. Phage display (Panel B, right): For phage display, B cells are isolated from immunized mice (as in panel A) or naïve or immunized humans. Heavy and light chain V genes (shaded bars) are amplified by PCR and assembled as single-chain Fv antibody genes (scFv). Alternatively, rearranged V genes can be generated entirely in vitro from cloned V segments and synthetic oligonucleotides. The repertoire of scFv genes are cloned into a phage display vector, where the encoded scFv proteins (colored ovals) are displayed as fusion proteins to one of the phage coat proteins. The phage contain the appropriate scFv gene within. Multiple rounds of selection with immobilized antigen allows isolation of even rare antigen-binding phage antibodies, which are identified by ELISA. Native scFv can be expressed from E. coli and purified for characterization and use in assays. See color insert.
Many of the above limitations can be overcome by taking advantage of recent advances in biotechnology to produce monoclonal antibodies directly using a variety of different display technologies (see ref. [21] for review). These technologies include phage display, yeast display, ribosome display, and a number of other less frequently used tech-
nologies. Display technologies have three characteristics in common: 1) A method for generating antibody gene diversity; 2) a method for linking the antibody genotype with the expressed antibody phenotype; and 3) a method for isolating rare antigen binding antibodies (and their genes) from a majority of nonbinding antibodies. In 2002, the first mono-
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FIGURE 32.2 Antibodies and antibody fragments. IgG antibody consists of a pair of heavy and light chains. Each chain consists of the antigen binding variable domains (VH and VL) and one or more constant domains (CH1, CH2, CH3, and CL). Each V or C domain contains an intramolecular disulfide bond (S-S). A single glycosylation site (CHO) exists in the CH2 domain of the heavy chain. The VH and VL domains contact antigen via the amino acids in the complementarity determining regions (CDRs). A number of antibody fragments can be expressed efficiently in bacteria. These include the Fab, Fv, scFv, and diabody. The Fv is the minimal antigen binding unit, consisting of the VH and VL domains. Since the VH and VL domains of a Fv may dissociate at physiologic concentrations, the two domains can be stabilized via a linker, yielding a single-chain Fv (scFv). Shortening the linker to less than approximately seven amino acids prevents pairing of VH and VL domains on the same molecule. This results in the pairing of complementary V domains from two scFv molecules, yielding the dimeric diabody, which has two antigen binding sites, The Fab consists of the VH and VL domains, with the CH1 and CL domains. In the IgG, the two Fabs are connected to the Fc via the flexible hinge.
clonal antibody produced using phage display technology was approved by the United States Food and Drug Administration for human use (Humira, an anti-TNF-alpha antibody for rheumatoid arthritis). More than 10 other monoclonal antibodies produced using display technologies are in various stages of clinical trials [22]. Such clinical successes illustrate the importance of these approaches for generating therapeutic antibodies. The earliest display technology, and the one most frequently employed, is phage display (reviewed in ref. [21, 23–26]). Antigen-specific antibody fragments are directly selected from antibody fragment gene repertoires expressed on the surface of bacteriophages, viruses which infect bacteria (phage display) [27, 28]. The antibody V genes are already cloned and almost invariably express at high level in bacteria [29, 30]. Higher affinity antibody fragments can be selected from phage antibody libraries created by mutating the antibody fragment genes of initial isolates [31, 32]. Moreover, the approach can be used to produce human antibodies, which are difficult to produce using conventional
hybridoma technology, and to produce antibodies without immunization, thus allowing the production of antibodies to conserved proteins [29, 33]. Since the technology utilizes selection, rather than just screening (as for hybridomas), it is possible to directly select for desired biologic functions, such as the triggering of receptor-mediated endocytosis or gene delivery resulting in transgene expression [34–36]. Since phage display is the most mature of the display technologies, we cover it first and in the most detail. We then describe yeast and ribosome display and discuss their relative strengths and weaknesses compared to phage display.
OVERVIEW OF ANTIBODY PHAGE DISPLAY Phage display mimics the strategy used by the natural humoral immune system to produce antibodies in vivo (reviewed in references [21, 23–26]). This strategy has three key components: 1) the generation of millions of different
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FIGURE 32.3 Constructs for E. coli secretion or display of antibody fragments. (a) Constructs for secretion of scFv, Fv, and Fab antibody fragments: rbs = ribosome binding site; leader = bacterial secretion signal directing the expressed protein to the bacterial periplasm; VH = heavy chain variable domain; VL = light chain variable domain; linker = flexible scFv linker between VH and VL domains; CH1 = heavy chain constant domain 1; CL = light chain constant domain. (b) Constructs for phage display of scFv and Fab antibody fragments: For scFv display, the scFv gene is genetically fused to the phage minor coat protein gene III. For Fab display, one chain (here the heavy chain VH-CH1) is secreted into the periplasm, whereas the other chain (here the VL) is fused to gene III.
antibody molecule genes by recombination and combinatorial pairing; 2) expression of the antibody gene repertoire on the surface of B-lymphocytes, where the antibody functions as an antigen receptor; and 3) antigen-driven selection of antigen binding B lymphocytes for proliferation and differentiation (Figure 32.1A). The B lymphocyte serves an essential role by providing a physical linkage between surface antibody (phenotype) and the gene encoding the antibody (genotype). Three technical achievements made it possible to reproduce the process of immune selection in vitro using phage display (see Figure 32.lB for an overview of phage display technology). First, it proved possible to express the antigen binding VH and VL domains of antibodies in E. coli, either as Fab, Fv, or scFv antibody fragments (see Figures 32.2 and 32.3 for Fab, Fv, and scFv) [37–40]. Second, large and diverse repertoires of Fab or scFv genes could be generated using the polymerase chain reaction. The antibody fragment repertoires can be built from VH and VL genes obtained from B lymphocytes, either before or after immunization, or from cloned V gene segments rearranged in vitro [41]. Third, the antibody fragments can be expressed on the surface of viruses (phage) that infect E. coli [27, 42]. This is accomplished by cloning the antibody gene into a phage vector so that it is genetically fused with a gene that encodes a protein expressed on the phage surface. The resulting phage has the antibody on its surface, anchored to the phage via the coat
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protein, and contains the gene encoding the antibody inside the phage. Thus, the phage mimics the function of the B lymphocyte, providing a physical linkage between phenotype on the surface and genotype within. Antibody fragment gene repertoires can be cloned into phage vectors, resulting in the creation of phage antibody libraries [43]. Because of the high transformation efficiency of E. coli, it is possible to create libraries of millions to billions of different antibodies. Phage antibodies binding a specific antigen can be separated from nonbinding phage antibodies by selection on antigen. Phage are incubated with immobilized antigen, nonbinding phage are removed by washing, and bound phage are eluted. A single round of selection will result in a 20- to 1000-fold enrichment for binding phage [27]. Eluted phage are used to infect E. coli, which produce more phage for the next round of selection. Repetition of the selection process makes it possible to isolate the binding phage present at frequencies of less than one in a billion. Antibody phage display has resulted from concurrent progress in prokaryotic expression of antibody fragments, PCR cloning of antibody gene repertoires, and display of peptides and proteins on filamentous bacteriophages. These three areas are reviewed in detail in the following sections. Subsequent sections describe specific applications of antibody phage display, including the use of phage display to: 1) bypass hybridoma technology; 2) bypass immunization; and 3) produce affinity mature antibodies.
PROKARYOTIC EXPRESSION OF ANTIBODY FRAGMENTS Efforts to express full-length antibodies in the cytoplasm of E. coli by co-expression of heavy and light chains were initially met with limited success. When a mouse heavy (m) and light (l) chain on two compatible plasmids were introduced into E. coli and expressed off the trp promoter, the proteins were found as insoluble inclusion bodies in the cytoplasm. Antigen binding activity was only recovered after solubilization and refolding of the aggregated material [44]. Similarly, expression of the heavy and light chains of a murine anti-CEA antibody in the cytoplasm of E. coli yielded only insoluble aggregates that required solubilization and refolding to detect antigen binding [45]. Presumably, these results were due to the difference between the reducing environment of the prokaryotic cytoplasm versus the oxidizing environment of the eukaryotic secretion pathway, where disulfide bonds can form. A breakthrough was achieved when researchers expressed antibody fragments fused to bacterial signal sequences that directed their secretion into the periplasmic space of E. coli. Mark Better and colleagues constructed a
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biscistronic operon that expressed genes encoding the two chains of a human–mouse chimeric Fab; each polypeptide chain was directed to the periplasm by a bacterial signal sequence that was subsequently cleaved by the bacterial signal peptidase (Figure 32.3). In the periplasm, the two chains folded correctly and formed a fully functional Fab [37] (Figures 32.2 and 32.3). Similarly, Skerra and Plukthun secreted the VH and VL genes of a mouse monoclonal antibody into the periplasm where they correctly folded to form a fully functional Fv fragment [38] (Figures 32.2 and 32.3). Although Fvs are the smallest antibody subunit that retain antigen binding, their utility is limited by instability. At typically utilized concentrations, Fv dissociate into VH and VL domain, with the KD for dissociation ranging between 10-4 to 10-8 M [46–50]. Differences in the VH-VL Kd result from differences in residues composing the b sheets that make up the VH-VL interface [51]. Although many of these interface residues are conserved, 25% of the interface result from residues in the hypervariable complementarity determining regions (CDRs), which comprise the antigen binding loops [51] (Figure 32.2). Several strategies have been utilized to stabilize the Fv fragment [49] including engineering a disulfide bond into the VH-VL interface [49, 52]. The most commonly employed strategy, however, has been to physically link the VH and VL into a continuous polypeptide chain, to form a “single-chain Fv” (scFv) [39, 40] (Figures 32.2 and 32.3). Both VH-linkerVL and VL-linker-VH orientations have been shown to yield functional scFv and, as long as the linker is of sufficient length, many different linker sequences are tolerated [17, 40, 53, 54]. Shortening of the linker to less than approximately 8 amino acids prevents intramolecular pairing of the VH and VL domains and forces intermolecular pairing. This results in a stable homodimer called a diabody (Figure 32.2) [55, 56]. These homodimers have two antigen binding sites and can exhibit increased functional affinity due to avidity, like IgG, when the antigen is multivalent [57, 58]. Expression of two different scFv genes with shortened linkers within the same E. coli can yield bispecific diabodies capable of binding two different antigens simultaneously [55, 59]. scFvs generally retain the specificity and affinity of the antibody from which they were derived [60]. Furthermore, the crystal structure of a Fab and its related scFv have been solved, and only minor difference in the antigen binding sites were observed [61].
GENERATION OF ANTIBODY GENE REPERTOIRES USING THE POLYMERASE CHAIN REACTION Before the advent of the polymerase chain reaction (PCR), cloning antibody genes was a laborious process,
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requiring the creation and screening of genomic or cDNA libraries. PCR has streamlined the process considerably, permitting the construction of large antibody fragment gene repertoires (see Figure 32.4 for overview). For PCR amplification, first strand cDNA is obtained by reverse transcription of mRNA with constant region primers [primers that anneal in the CH gene (Cg1, Cg2, Cg3, Cg4, Cm, Ce, Ca or Cd) or the CL gene (Ck or Cl)]. Since the sequences of the constant domain exons are known [62], primer design is straightforward. For creation of Fabs, the VH-CH1 (fd) and light chain (VLCL) are amplified. Design of primers that anneal to the 3¢ end of these genes is straightforward, since the constant regions have been sequenced [62]. For Fv or scFv, only the rearranged VH and VL are amplified. Design of PCR primers for the 3¢ end of rearranged murine [43] or human [29] VH and VL gene is also straightforward, since primers can be based on the J gene segments, which have been sequenced. These primers can also be used for first-strand cDNA synthesis instead of constant region primers. Design of primers for the 5¢ end of the V gene was thought to be less straightforward due to the sequence variability of different V-genes. In the earliest attempt to use PCR to amplify V-genes, N terminal protein sequencing was done on purified antibody from a hybridoma and the sequence used to assign the VH and VL (Vl or Vk) gene families [62]. The VH and VL gene assignments were used to design degenerate primers based in FR1 [63]. A generally applicable approach was taken by Orlandi et al [64]. The nucleotide sequences of murine VH and VL genes were extracted from the Kabat database [62], aligned, and the frequency of the commonest nucleotide plotted for each position. Conserved regions were identified at the 5¢ region of the VH and VL genes and the sequences used to design degenerate oligonucleotide primers containing restriction sites for directional cloning. Additional groups have designed sets of universal V-gene primers containing internal or appended restriction sites suitable for amplification of murine [65–70], human [29, 71–74], chicken [75], and rabbit [76] V genes. Fv, Fab, and single chain Fv genes are typically constructed by sequential cloning of VH (VH-CH1) and VL (VLCL) genes [39, 40]. Alternatively, PCR splicing by overlap extension (PCR assembly) of VH and VL genes has been used to construct scFv genes [29, 43] (Figure 32.4). Using PCR assembly, only two restriction sites are required to clone the scFv. These can be appended to the 5¢ and 3¢ end of the scFv gene cassette, and expression systems have been described where octanucleotide cutters can be utilized [29]. This markedly decreases the likelihood that a restriction enzyme will cut internally in the V-gene. Sequential cloning requires four restriction sites and increases the chances of restriction enzymes cutting internally [16, 77].
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FIGURE 32.4 PCR cloning of antibody V-gene repertoires as single-chain Fvs (scFv), (a) mRNA is isolated from peripheral blood, spleen, or bone marrow, and antibody genes are reverse transcribed (by using reverse transcriptase) using IgG, IgM, k, and l constant-region specific primers, creating 1st strand cDNA. (b) The VH and VL variable region genes are amplified in a series of PCR reactions, using primers specific for each of the known heavy and light chain Vgene families. (c) scFv assembly by PCR. PCR amplified VH and VL genes are pooled, together with a short “linker DNA,” which overlaps the 3¢ and 5¢ ends of the VH and VL regions respectively, and PCR amplified to yield one continuous DNA fragment. A final PCR reaction (not shown) adds flanking restriction sites to the assembled repertoire for cloning into the phage display vector.
ANTIBODY PHAGE DISPLAY Phage display technology exploits several features of the filamentous E. coli bacteriophages M13, fd, and f1. These are single-stranded DNA viruses with genomes of approximately seven thousand base pairs; the viral particle is a filament that contains one copy of the viral genome encased in a protein coat. This coat is composed primarily of 2,700 copies of the gene 8 protein pVIII; in addition there are several minor coat proteins including the gene 3 product pIII, which is found in three to five copies at one end of the viral particle, pVII, and pIX (Figure 32.5). pIII has a threedomain structure; the domains are separated by glycine rich regions (presumed to be flexible linkers). The replication of the phage begins with binding of the pIII protein to the Fpilus of a male (F+) E. coli bacterium, which leads to entry of the viral genome into the bacterial cytoplasm. The Nterminal domain of pIII binds to the pilus, and the second
domain mediates penetration of the phage genome into the bacteria. Once inside the bacterium, the viral genome is copied into a double-stranded replicative form (RF), which serves as a template to make more copies of the singlestranded viral genome for packaging. The intragenic region of the phage genome contains a signal for the newly created genomes to be packaged and extruded from the host cell. During extrusion, the phage DNA becomes coated with pVIII and pIII, forming a new phage particle that can begin a new round of infection. In 1985, George Smith showed that when DNA encoding a peptide was cloned as a gene fusion with gene III, the resulting phage “displayed” the peptide on its surface as a fusion with pIII [42]. Phage displaying the peptide could be enriched from populations of wildtype phage by affinity chromatography using a monoclonal antibody specific for that peptide. By infecting bacteria with the enriched eluted phage, more phage could be grown and subjected to another
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FIGURE 32.5 Different formats for antibody phage display. Wildtype phage (far left) has four coat proteins, pIII, pVIII, pVII, and pIX. pIII display (middle two phage). ScFv (red and yellow) are fused to the minor coat protein pIII. In true phage vectors (left), scFv are displayed on all three to five copies of pIII. In phagemid display, scFv display is typically single copy per phage particle, due to competition from wildtype pIII from the helper phage. pVIII display (far right phage) theoretically yields multiple scFv/phage particle. Wildtype pVIII must be supplied to make phage. See color insert.
round of selection. Repetition of the selection process makes it possible to isolate binding phage present at frequencies of less than one in a billion. Moreover, the same technique could be used to enrich peptide phage with high affinity for ligand from medium- or low-affinity phage. In 1990, McCafferty et al. demonstrated that it was possible to display a functional antibody fragment on the surface of filamentous phage using a scFv fragment of the anti-lysozyme Mab D1.3 [27] (see Figure 32.5). A scFv-pIII fusion protein was expressed in the phage fd vector fdCAT1, and it was demonstrated that the displayed scFv retained antigen binding and specificity. Phage displaying the scFv could be enriched from a mixture with wildtype phage by affinity chromatography on a lysozyme column. Enrichment factors of 103 for one round of selection and 106 for two rounds of selection were achieved.
Vectors for the Display of Antibody Fragments on Filamentous Phage A large number of vector systems have subsequently been described for the display of antibody fragments, and some of the more important ones are summarized in Figures 32.5 and 32.6. The vectors differ primarily with respect to the type of antibody fragment displayed: Fab [28, 69, 78–81] or scFv [27, 43, 82], the fusion partner pIII [27, 28, 43, 69, 79, 81, 82] or pVIII [78, 80], and whether the vector is a phage [27, 43, 80] or phagemid [28, 69, 78, 79, 81, 82]. Vectors for display of scFv have a single leader sequence and two or four cloning sites for insertion of the scFv genes (Figures 32.3 and 32.6). Vectors for display of Fab have two leader sequences and two pairs of restriction sites for the sequential cloning of VH and VL genes (Figure 32.3). Fab fragments
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are displayed by fusing one of the chains to a phage coat protein and secreting the other chain into the bacterial periplasm, where the heterodimer forms [28]. The most widely used vectors result in fusion with the minor coat protein pIII [27, 28, 79, 83–85] (Figure 32.5). Alternatively, antibody fragments can be displayed as pVIII fusions [78, 80, 86] (Figure 32.5). This theoretically results in many more copies of antibody fragment per phage (up to twenty-four copies per phage [78]. In fact, the toxicity of the antibody fragments and competition with wildtype pVIII can result in lower levels of display than for pIII fusions in phagemid vectors. More recently, the display of antibody fragments on pVII and pIX has been reported [87, 88]. Display of antibody fragments on pIII in phage vectors also leads to multicopy display, with each of the three to five pIII proteins having an antibody fragment fusion (Figures 32.5 and 32.6). Since there is no competition with wildtype pIII, as with phagemid systems (see below), pIII display in a phage vector leads to a greater amount of pIII fusion on the phage, compared to phagemid systems [89]. Multivalent display leads to an increase in the apparent affinity (avidity) when the phage antibody is selected on antigen immobilized on a solid support. This leads to more efficient selection than for monovalent phagemid display [89]. For some antigens, the efficiency of selection is so great that binding antibodies can be identified after a single round of selection [89]. This facilitates the automation of the selection process, since no amplification in liquid culture is required. Avidity can also permit the selection of very low affinity antibody fragments that would not be selected with monovalent display. With respect to selections directly on cells, the avidity resulting from pIII phage display may also lead to more successful selections [90]. In addition, it has been shown that cell surface receptor cross-linking, which can occur with multicopy antibody fragment display, can lead to more efficient phage internalization than monovalent display [34, 35]. Thus selections for antibodies that trigger receptormediated endocytosis may be more efficient with pIII phage antibody libraries. However, multivalent display makes it more difficult to discriminate between phage with only minor differences in affinity [79] and may result in the isolation of lower affinity antibodies [89]. Use of a phagemid vector results in “monovalent” antibody fragment display (Figures 32.5 and 32.6). Phagemids are plasmids that contain the phage f1 intragenic region (and therefore can be packaged in a phage-like particle) but lack other essential phage genes and therefore cannot independently produce phage. Essential phage genes are provided in trans by infection of phagemid-containing E. coli with a helper phage. The helper phage has a weakened packaging signal, and thus the phagemid genome is preferentially packaged into the phage particle. In phagemid vectors, antibody fragment-pIII fusion expressed from the phagemid DNA competes with wildtype pIII expressed from the helper
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FIGURE 32.6 Features of phage display vectors. A representative phage and phagemid vector are shown. (a) Phage vector fd-tet-Sfi/Not (right panel) and (b) phagemid vector pHEN1 (left panel). Both vectors display scFv (VH-linkerVL) as fusions to the amino terminus of the pIII protein. Both vectors have leader sequences (pelB or gene III) to direct the expressed fusion protein to the bacterial periplasm. In phage fd-tet-Sfi/Not, all copies of pIII are scFv fusions, leading to three to five copies displayed per phage particle. With phagemid pHEN1, expression in supressor strains of E. coil allows the amber codon following the scFv-tag to be read as a glutamine, thus causing the scFv to be fused to the pIII protein. In phagemids, both wildtype pIII (from the helper phage) and fusion pIII (from the phagemid) compete for inclusion in the viral particle. In nonsuppressor E. coil strains, the scFv is expressed as a soluble protein with a myc epitope tag for detection of binding in ELISA.
phage genome. The resulting phage have on average less than one copy of antibody-pIII fusion–phage (see Figure 32.5). Monovalent phage display leads to more efficient selection of phage on the basis of binding affinity. For example, in a single round of selection using a mixture of two Fab that differed 100-fold in affinity (Kd~10-7 M vs 10-9 M), the enrichment ratio was 253-fold for monovalent display on pIII versus only 5.5-fold for multivalent pVIII display [79]. Phagemid vectors also yield higher transformation efficiencies compared to phage vectors, making construction of large phage antibody libraries easier. As a result of these two features, the most widely used systems for anti-
body phage display use phagemid vectors with display on pIII. Analysis for phage binding to antigen can be performed in an ELISA format using anti-phage antibodies [43]. Analysis for binding is simplified by including an amber codon between the antibody fragment gene and gene III [28] (Figure 32.6). This makes it possible to easily switch between displayed and soluble antibody fragment simply by changing the host bacterial strain. When phage is grown in a supE suppressor strain of E. coli, the amber stop codon between the antibody gene and gene III is read as glutamine and the antibody fragment is displayed on the surface of the
32. Monoclonal Antibodies from Display Libraries
phage. When eluted phage is used to infect a nonsupressor strain, the amber codon is read as a stop codon and soluble antibody is secreted from the bacteria into the periplasm and culture media [28]. Binding of soluble scFv to antigen can be detected by ELISA using epitope tags, such as c-myc [29] or E-tag [91] incorporated into the vectors 5¢ to the amber codon. The inclusion of a hexahistidine tag makes it possible to easily purify native scFv without the need for subcloning [30].
USE OF PHAGE DISPLAY TO BYPASS HYBRIDOMA TECHNOLOGY Phage display can be used to produce monoclonal Fab or scFv antibody fragments from immunized mice and humans. The primer systems described above have been validated to produce diverse murine or human VH (VH-CH1) and VL (VLCL) gene repertoires from spleen, bone marrow, or peripheral blood lymphocytes [29, 43, 70, 71]. The VH and VL genes can be randomly spliced together using PCR [29, 43] (Figure 32.4), or cloned sequentially to create scFv or Fab gene repertoires displayed on the surface of phage. In the first example, an scFv phage antibody library was created from the V genes of a mouse immunized with the hapten phenyloxazolone [43]. After cloning, a 2 ¥ l05member scFv phage antibody library was created in a phage (fd) vector. Binding phage antibodies were selected on a phOx-BSA column. After two rounds of selection, the majority of clones analyzed produced phOx binding scFv. More than 20 unique scFvs were isolated. The Kd of the highest affinity phage antibodies (1.0 ¥ l0-8 M) were comparable to the affinities of IgG from hybridomas constructed from mice immunized with the same hapten. Similar panels of scFv have been obtained using phage display and mice immunized with EGF receptor [92] and botulinum neurotoxin type A [70]. This approach has also been used to produce monoclonal chicken [75] and rabbit [76] antibody fragments using species-specific primers. A greater number of examples exist of phage libraries constructed from the V genes of immunized humans. In most of these examples, Fab libraries were constructed. Variable-region genes were obtained either from peripheral blood lymphocytes, bone marrow, lymph nodes, and rarely, from spleen. In the area of infectious disease, human monoclonal scFv or Fab antibody fragments have been isolated from immunized volunteers or infected patients against tetanus toxin [79], botulinum neurotoxin [93, 94], HIV-1 gp120 [95], HIV-1 gp41 [96], hepatitis B surface antigen [97], hepatitis C [98], respiratory syncytial virus [99, 100], and hemophilus influenza [101]. Autoimmune antibodies have been isolated from patients with systemic lupus erythematogus (SLE) (anti-DNA) [102], Hashimoto’s disease (anti-thyroid) [103], myasthenia gravis (anti-acetylcholine
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receptor) [104], as well as many other autoimmune diseases. Recently, there has been interest, and success, in generating antibodies to self antigens, such as tumor antigens, from libraries constructed from patients with disease [105] or after vaccination with tumor antigen [106, 107]. Immune libraries allow the exploitation of the affinity maturation process that occurs by somatic hypermutation. In addition, immunization amplifies the number of lymphocytes producing binding antibodies, as well as their RNA content. This is particularly true for IgG-producing plasma cells. Thus, libraries are enriched for V genes from antigen binding antibodies. However, it is important to keep in mind that this may be offset by the fact that the original VH and VL pairings are lost during library construction, when the lymphocytes are lysed for RNA isolation. With achievable library sizes, the probability of recovering the original VH and VL pairings is small. Techniques for “in-cell” PCR have been devised to retain the original pairings [108], but this has not been shown possible for large library constructions. The fact that these immune libraries yield high-affinity binders is probably due to the fact that in many cases, a binding light chain can be replaced by a homologous light chain with recovery of antigen binding. Such “chain promiscuity” has been observed in immune libraries and during chain shuffling, when the binding light chain is replaced by a library of light chains [31, 43]. After light chain shuffling, the frequency of binding antibody fragments can be as high as 10% [31]. Antibodies from immune phage antibody libraries share a number of characteristics. Typically, a panel of antibodies are obtained. Some of these are clonally related, being derived from the same VDJ rearrangement, with differences due to somatic hypermutation. Others are clearly derived from different VDJ rearrangements. The antibodies are highly specific for the antigen used for selection, and they bind antigen with affinities typical of IgG produced from hybridomas derived from immunized mice or humans. Moreover, the ability to select rather than screen for binding properties (see below) permits the isolation of antibodies to rare epitopes [109–111], cell surface markers [106], or unstable antigens [112], which have proven difficult to produce using hybridoma technology. The antibody fragments can be used for the same purposes as IgG derived from hybridomas, including western blotting [113], epitope mapping [96], cell agglutination assays [114, 115], cell staining, FACS scaning [116], and immunohistochemistry [107]. The use of immune phage antibody libraries allows the isolation of a large number of scFv or Fab of high specificity and affinity. This approach, however, is subject to the limitations associated with immunization. First, with many antigens, the immune response is directed to only a few immunodominant epitopes [96, 117]. This may result in a failure to isolate the precise specificity desired for a partic-
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ular aim. Furthermore, production of antibodies against proteins conserved between species may prove difficult or impossible. Production of nonimmunogenic human antibodies is also limited by the inability to immunize with many antigens, especially those of human origin. Finally, for each desired specificity, a new library must be constructed— a relatively time consuming process.
USE OF PHAGE DISPLAY TO BYPASS IMMUNIZATION As an alternative to immune phage antibody libraries, monoclonal antibody fragments can be produced without prior immunization by displaying very large and diverse scFv or Fab gene repertoires on phage [29]. This theoretically results in the ability to isolate antibodies to any desired specificity from a single phage antibody library. In the first example, rearranged human VH, Vk, and Vl gene repertoires were amplified from the mRNA of peripheral blood lymphocytes obtained from two nonvaccinated volunteers [29]. Diverse scFv gene repertoires were obtained by randomly joining the VH and VL gene repertoires with DNA encoding the 15–amino acid linker sequence (G4S)3. The human scFv gene repertoire was cloned into the phagemid pHEN-1 to create a nonimmune phage antibody library of 3.0 ¥ 107 members. This is similar to the size of a primary human or mouse B-cell repertoire. From this single nonimmune library, scFvs were isolated against more than twenty different antigens, including a hapten, three different polysaccharides, and sixteen different proteins [29, 33, 114, 118, 119]. The scFvs were highly specific for the antigen used for selection and had affinities typical of the primary immune response, with dissociation equilibrium constants ranging from 1 uM to 15 nM [29, 33, 118]. Using this approach, it was possible to isolate scFv that bound self proteins; for example, human scFv that bound human thyroglobulin, immunoglobulin, tumor necrosis factor, CEA, ErbB-2, and blood group antigens [29, 33, 114, 118, 119]. The ability to isolate anti-self specificities may result from the generation of specificities that had been deleted from the B-cell repertoire. These new specificities would arise from the random reshuffling of VH and VL genes that occurs during library construction. Alternatively, antiself specificities could arise from the 10 to 30% of human B cells that may be making low-affinity anti-self antibodies at any time [120]. Larger or more diverse phage antibody libraries should theoretically provide higher affinity antibodies against a greater number of epitopes on all antigens used for selection [121, 122]. One approach to increase diversity is to rearrange V genes in vitro using cloned V-gene segments and random oligonucleotides encoding part of the antigencombining site [24, 113, 116, 123–125]. Using this
approach, it is possible to ensure that the potential diversity is large enough so that a library will only contain a single member of each sequence. In contrast, there may be considerable duplication of sequences using V-gene segments rearranged in vivo. The design of semisynthetic libraries is based on structural, genetic, and mutagenesis data demonstrating that the determinants of antibody specificity reside in the six CDRs (reviewed in ref. [126]). Five of the six CDRs are encoded in a small number of germline gene segments [127–129] and have limited sequence variability and main chain conformations (canonical structures) [128, 130, 131]. The exception is VH CDR3, which is generated by recombination of three gene segments (V, D, and J). The addition of random nucleotides (N-segment addition) at the V–D and D–J junctions leads to tremendous sequence and structural diversity. Not surprisingly, this CDR contributes a disproportionate number of antigen-contacting amino acid side chains and binding energy [132–134]. Thus, synthetic antibody libraries have focused the diversity in VH CDR3. For the first semisynthetic phage antibody library, a VH gene repertoire was constructed from fifty-one cloned germline human VH gene segments [127] and oligonucleotides designed to yield a VH CDR3 of five or eight residues, in which five residues were of random sequence. The repertoire was cloned into a vector containing a single rearranged Vl light chain gene to create an scFv phage library of 2.2 ¥ 107 members [24]. This library yielded large panels of monoclonal scFv to hapten antigens, but not to protein antigens [24]. Increasing the length of the synthetic VH CDR3 to between four and twelve residues resulted in a library from which antibodies could be isolated to protein antigens [113]. The affinities of the antibodies, however, were not reported. Subsequent examples of semisynthetic antibody libraries have used a larger number of germline VH and VL genes [116, 125, 135] or have been constructed from single VH and VL germline segments where multiple CDRs are partially diversified [136]. The greatest improvements in the utility of nonimmune libraries has resulted from increasing library size several orders of magnitude. This yields libraries capable of yielding panels of high-affinity antibody fragments against any antigen. At least five published examples of such libraries exist, with sizes ranging between 109 and 1011 members [30, 74, 125, 137, 138]. These include both scFv [30, 74, 137, 138] and Fab libraries [125, 138], constructed using naturally occurring V genes or semisynthetic V genes. For two of these libraries, recombination either between or within vectors was used to overcome the limitations imposed on library size by transformation efficiency [74, 125]. From these five libraries, panels of antigen-specific antibody fragments have been isolated against more than fifty different antigens, including haptens and proteins. Affinities of the antibody fragments are typical of the secondary immune
32. Monoclonal Antibodies from Display Libraries
response (Kd ranging between 4.1 ¥ 10-8 M and 3.0 ¥ 10-10 M). For example, we have constructed a 6.7 ¥ 109 member scFv phage antibody library from the VH and VL genes of healthy humans [137]. An average of 9.2 scFv (with Kd as high as 3.7 ¥ 10-10 M) were isolated to ten different protein antigens. Antibodies from nonimmune libraries have been used successfully for western blotting [113], epitope mapping [117], cell agglutination assays [114], cell staining, and FACS [90, 116]. The nonimmune libraries described above represent the current state of the art—the ability to generate high-affinity human monoclonal scFv or Fab antibody fragments to any antigen within weeks and without immunization.
A COMPARISON OF DIFFERENT PHAGE ANTIBODY LIBRARY TYPES AND APPLICATIONS
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It is presently uncertain whether nonimmune libraries are best constructed from V genes rearranged in vivo (natural or naïve libraries) or in vitro (semisynthetic). A theoretical advantage to semisynthetic libraries is that only a single copy of each antibody exists in a library. Furthermore, Vgene segments can be selected that express well in E. coil, thus providing a higher number of functional antibodies and reducing selection biases based on expression efficiency [136]. The use of synthetic CDRs, however, assumes that the VH CDR is a random peptide, which is not the case [139]. It is thus likely that a significant number of synthetic VH CDR3s (and other synthetic CDRs) do not fold properly, or do not make useful binding pockets, compared to those constructed in vivo from V-, D-, and J-gene segments. Semisynthetic libraries are also more difficult to construct, requiring the cloned V-gene segments. At present, it appears that library size is a more important parameter than the source of the rearranged V genes.
Immune vs. Nonimmune Libraries
Fab vs. scFv Libraries
The choice of whether to generate immune or nonimmune phage antibody libraries will depend on the uses intended and the skills of the investigator. Compared to nonimmune libraries, immune libraries yield a larger number of antibody fragments having higher average affinity. Immune libraries are also easier to construct, since significantly smaller libraries (as few as one million members) can be utilized. In addition, immune libraries are the only possibility for probing the in vivo immune response to a particular pathogen or disease. Immune libraries, however, suffer from the drawback that a new library has to be constructed for each new antigen desired. In addition, for many antigens, the immune response is directed to only a few immunodominant epitopes [96, 117]. This may result in a failure to isolate the precise specificity desired for a particular aim. Furthermore, the production of antibodies against proteins conserved between species may prove difficult or impossible. Production of nonimmunogenic human antibodies is also limited by the inability to immunize humans with many antigens, especially those of human origin. In contrast, nonimmune libraries need to be made only once, can provide antibodies to any antigen (including those that are conserved or “self”), and can serve as a source of therapeutic human antibodies. Compared to immune libraries, however, fewer antibodies will be obtained, and the affinities are typically not as high. Moreover, useful libraries are of a size (more than one billion members) that has proved difficult to produce except in a limited number of laboratories. The use of recombination to unite VH and VL repertoires may simplify the construction of large nonimmune libraries [74]. Once the hurdles of constructing very large libraries are overcome, nonimmune libraries are likely to assume increasing importance for antibody generation.
The choice of whether to produce scFv or Fab libraries depends partly on the intended use. For construction of fusion proteins, such as immunoadhesions or immunotoxins, the single gene format of the scFv is an advantage. This is especially true for targeted gene therapy approaches, in which the scFv gene is fused to a viral envelope protein gene [140, 141] or the gene for a DNA binding protein [142]. scFv also appear to be the preferred antibody fragment for intracellular immunization, a technique in which the antibody gene is delivered intracellularly to achieve phenotypic knockout [143, 144]. In theory, Fab libraries may be preferred where the final product will be a complete antibody, since in some instances removal of the scFv linker might alter antigen binding properties. In practice, too few examples exist in which either Fab or scFv have been retroengineered into complete antibodies to draw conclusions, although a number of scFv have been converted to IgG without loss of affinity or specificity [117]. Technical issues also influence the choice of antibody fragment type for library construction. In general, scFv libraries are easier to construct. The genes are smaller, amplify more easily using PCR, and clone with higher efficiency. scFvs are also generally less toxic to E. coli and fold more efficiently. This leads to a lower deletion rate of insert from phage libraries and higher expression levels of native antibody fragment. This makes subsequent characterization easier. A disadvantage of scFvs include the tendency of some scFv to dimerize and aggregate. Dimerization and aggregation occur when the VH domain of one scFv molecule pairs with the VL domain of a second scFv molecule [55, 145]. The tendency of scFv to dimerize is sequence dependent, with some scFvs existing as stable monomer [33, 115, 118] and others as mixtures of monomeric and oligomeric scFv
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[113, 115, 135, 145]. Such multimerization does not occur with Fab, where the constant domains and intermolecular disulfide bond help stabilize the molecule. This increased stability of Fabs may be important in applications such as the construction of antibody arrays.
STRATEGIES FOR SELECTION OF PHAGE ANTIBODIES One of the advantages of display technologies for antibody generation is that binding antibodies are “selected” rather than first identified by screening. This allows the tailoring of selection methods to optimize the isolation of antibodies that have the properties sought for a particular application. In its original implementation, selection involved: 1) the binding of phage expressing a relevant antibody to antigen immobilized on a solid surface, 2) removal of irrelevant nonbinding phage by washing, and 3) elution of specifically bound phage (See Figure 32.1). The first phage antibody libraries were selected on columns using antigen covalently linked to a matrix such as cyanogen bromide activated Sepharose [27, 43]. The simplest technique, however, has been to adsorb the antigen of interest onto a polystyrene surface, such as an ELISA plate well [73] or immunotube [29]. One way to increase enrichment during selection is to use a “specific eluant”, such as a high concentration of soluble antigen [32, 43], competitive elution with a known ligand [146], or cleavage of a susceptible linker in the antigen-support linkage [147]. Adsorption of protein antigens to plastic may lead to partial or complete antigen denaturation. This can lead to the disappearance of certain epitopes and the exposure of other cryptic epitopes. Thus, antibodies selected on surfaceabsorbed antigens may not react with the native antigen; for example, in solution or on the surface of cells [118, 148]. The problem of antigen denaturation can be overcome by selecting on soluble antigen in solution. The antigen can be chemically tagged, for example by biotinylation, and captured along with bound phage using avidin or streptavidin magnetic beads [32, 149], Alternatively, the antigen can be genetically tagged with a hexahistidine tag with capture on Ni-NTA agarose [70], or expressed as GST or maltosebinding protein fusions, with capture on either glutathione or maltose columns. Not only does this approach increase the likelihood of obtaining antibodies that recognize native antigen, it is also useful for limiting the antigen concentration to the selection of antibodies on the basis of affinity. Selecting in solution also reduces the avidity effect of dimeric scFv, thus biasing for the selection of stable monomeric scFv [149]. A number of approaches have been used to generate antibodies to one particular desired epitope. In general, specific elution strategies have been used for this purpose. For
example, if antibodies binding only one specific epitope on an antigen are desired, phage that bind that epitope can be specifically eluted with a monoclonal antibody [146] or ligand known to bind to that epitope. Alternatively, undesired epitopes can be “masked” using monoclonal antibodies to direct the selection towards yet undiscovered epitopes [110, 150, 151]. A related approach is to use solution-phase competition to direct the selection to epitopes that differ between two related molecules. For example, antibodies binding only activated complement C5a were isolated by performing selections on immobilized activated C5a in the presence of unactivated C5 in solution [152]. This approach can thus be employed to select for “neoepitopes,” which are formed upon protein activation. Selections can be performed on more complex mixtures of antigen, including intact cells, as long as measures are taken to prevent the enrichment of phage antibodies that bind to nonrelevant cell-surface antigens. Cell-surface antigen-specific antibodies have been isolated by selecting phage antibody libraries on adult erythrocytes [114], fetal erythrocytes [90], lymphocytes [116], melanoma cells [106, 107], and breast tumor cells [36]. For highly expressed antigens, specific antibodies may be isolated by direct selection on cells [114]. For antigens expressed at lower density, it is frequently necessary to deplete the library of binders to other antigens using a closely related cell type [36, 114]. Multivalent phage antibody libraries may be more useful for cell selections, due to the more efficient selection resulting from avidity [89, 90]. It has also proved possible to directly select phage antibodies that trigger receptor-mediated endocytosis [36], since phage antibodies binding internalizing cell surface receptors can undergo receptor-mediated endocytosis [34]. Such antibodies may prove particularly useful for the intracellular delivery of therapeutic molecules [153, 154]. Bacterial or mammalian cells can also be used as “living columns” for antibody selection, by either fusing the antigen gene to a cell-surface protein gene [155], or by transfecting the cell with the gene for a cell-surface antigen [156].
INCREASING ANTIBODY AFFINITY USING PHAGE DISPLAY Phage display is a powerful tool for increasing antibody affinity to values not achievable using hybridoma technology [1, 3]. To increase affinity, the antibody sequence is diversified, a phage antibody library is constructed, and higher affinity binders are selected on antigen [31, 157– 159]. Two considerations that must be addressed to successfully apply this approach are: 1) where to introduce mutations into the antibody fragment gene; and 2) how to efficiently select rare higher affinity antibodies from more frequent lower affinity antibodies.
32. Monoclonal Antibodies from Display Libraries
Where and How Should Mutations Be Introduced? When designing mutant phage antibody libraries, decisions must be made about how and where to introduce mutations. The considerations below also apply to the generation of diversity for libraries constructed using either yeast or ribosome display (see below). One approach is to randomly introduce mutations, thus apparently mimicking the process of somatic hypermutation in vivo. Random mutations can be introduced using chain shuffling [31, 43], in which the VH or VL gene of a binding antibody is replaced with a V-gene repertoire. For example, chain shuffling was used to increase the affinity of a scFv for a hapten by 300-fold [31]. Light chain shuffling resulted in a 20-fold increase in affinity to 16 nanomolar. A further 15-fold increase in affinity, to 1.1 nM, was achieved by conserving the new light chain and VH CDR3, then shuffling the VH gene segment. As with in vivo affinity maturation, the kinetic basis for the increase in affinity [31] was largely due to a decrease in the dissociation rate constant (koff) [160]. Random mutations can also be introduced using error-prone PCR [32] or mutator strains of E. coli [161]. The random introduction of mutations is simple and easy to perform and has resulted in large increases in affinity (greater than 100-fold) for hapten-binding antibody fragments [31, 161]. Results with protein binding antibody fragments, however, have been more modest (<10-fold) [32, 149]. Moreover, the relatively random distribution of mutations in higher affinity clones provides little useful information about where to direct additional mutations to further increase affinity. As an alternative to random mutagenesis, mutations can be specifically introduced into the CDRs that form the contact interface between antibody and antigen. Targeting mutations to the CDRs has been shown to be a very effective approach for increasing antibody affinity, including the affinity of protein binding antibodies. Yang et al. increased the affinity of an anti-HIV gp120 Fab 420-fold (to a Kd = 1.5 ¥ 10-11 M) by mutating four CDRs in five libraries and combining independently selected mutations [158]. Similarly, Schier et al. increased the affinity of an anti-ErbB-2 scFv more than 1200-fold (to a Kd = 1.3 ¥ 10-11 M) by sequentially mutating VH and VL CDR3 [159]. Complete randomization of an amino acid position typically requires the use of the nucleotide sequence NNS, where N = A, G, C, or T; and S = G or C. Since this generates thirty-two possible nucleotide sequences, randomization of only five amino acid positions will generate (32)5 or 3.4 ¥ 107 possible sequences. This number is at the limit of the size of phage libraries that can be conveniently produced. Thus only four to five amino acids can be mutated at a time if the entire sequence space is to be sampled. This is only a fraction of the number of amino acids in the CDRs,
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which can exceed fifty residues. One approach to reduce the number of CDR residues that need to be sampled is to perform molecular modeling on a homologous Fab or Fv crystal structure [159]. For all the CDRs except VH CDR3, modeling permits the identification of CDR residues that have a structural role—either in maintaining the main chain conformation, or packing at the VH-VL interface [51, 127, 128, 130]. The mutation of residues with predicted structural roles results in a reselection of the wildtype amino acid residue [91, 159]. Thus, these residues should not be randomized. Instead, the amino acids selected for mutagenesis should be those whose side chains are predicted to be solvent accessible. We have also observed complete conservation of glycine and tryptophan residues within CDRs [91, 159]. Glycine residues are typically key residues in turns, and the chemical properties of tryptophan make it a frequent structural or high-energy contact residue [162]. Thus, when randomizing CDRs conservation of these two residues should be considered. With respect to the initial CDRs to select for mutagenesis, mutation of the VH and VL CDR3 has yielded greater increases in affinity than mutation of other CDRs [158, 159, 163]. Affinity maturation can be performed sequentially, with each subsequent library constructed from the sequence of the highest affinity mutant. Alternatively, to save time, a parallel strategy can be used in which different regions of the molecule are randomized independently, higher affinity mutants are isolated, and mutations are combined. With respect to results from phage antibody libraries, some mutations are additive [158, 159], while others are not [149, 159]. Therefore, a sequential strategy may be more prudent. Alternatively, it is possible to scan many residues at a low mutation frequency (parsimonious mutagenesis [164]) to identify those residues that modulate affinity and those structural and functional residues that are conserved [91]. Residues identified as modulating affinity could then be completely randomized in a subsequent library.
Selection of Higher Affinity Phage Antibodies The efficient selection of higher affinity phage antibodies is less than straightforward due to sequence-dependent differences in phage antibody expression and in toxicity to E. coli. These differences can lead to selection for increased expression levels, or decreased toxicity, rather than for higher affinity. In the case of scFv phage antibodies, selection is also complicated by the tendency of some scFvs to dimerize [55, 145]. Dimeric scFv can form on the phage surface by noncovalent association of the V domains of the scFv-pIII fusion with the V domains of native scFv in the periplasm. Native scFv appears in the periplasm both from incomplete suppression of the amber codon between the scFv gene and gene III, as well as by proteolysis. Dimeric
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scFvs exhibit increased apparent affinity due to avidity and are preferentially enriched over monomeric scFv when selections are performed on antigen immobilized on a solid phase [135, 149]. Thus, selections must be carefully designed to ensure enrichment is based on affinity rather than expression level, toxicity to E. coil, or avidity. Selection for monomeric higher affinity antibody fragments is optimal when selections are performed in solution; for example, using biotinylated antigen with subsequent capture on streptavidin-coated magnetic beads [32, 149]. For the initial round of selection, an antigen concentration greater than the Kd of the wildtype antibody is used to capture rare or poorly expressed phage antibodies. In subsequent rounds, the antigen concentration is reduced to significantly less than the desired Kd [149].
ALTERNATIVE ANTIBODY DISPLAY TECHNOLOGIES In recent years, additional display technologies have been developed to provide alternatives to phage display for the in vitro generation and evolution of antibodies. The two most widely used are yeast display and ribosome display (reviewed in refs. [21, 165]). These two display technologies are described in more detail here.
Yeast Display For yeast display, antibody genes have been fused to the C-terminus of the Sachromyces cell surface protein AgaII (Figure 32.7) [166]. AgaII forms a pair of disulfide bonds with AgaI, yielding a cell surface–displayed antibody. To date, only scFv antibody fragments have been displayed on yeast, with a single yeast cell displaying up to 100,000 scFv. Yeast displaying binding scFv are isolated from nonbinding yeast using soluble antigen, to avoid avidity effects. The most elegant form of selection has involved sorting the yeast based on the intensity of antigen staining using flow cytometry (Figure 32.7). By manipulating the antigen concentration, it is possible to finely select antibodies on the basis of affinity [167, 168]. In fact, the major reported use of yeast display has been to increase antibody affinity. Most approaches have used error-prone PCR or DNA shuffling to introduce mutations, followed by selection of the highest affinity scFv by flow cytometry [169, 170]. Using this approach, it proved possible to increase the affinity of an scFv more than 1,000-fold to 48 femptomolar [170]. This is probably the highest affinity antibody reported in the literature. More recently, the successful generation and selection of a nonimmune human scFv library displayed on yeast has been reported [171]. Despite the lower transformation efficiency of yeast compared to bacteria, a library of 109
FIGURE 32.7 Yeast display of scFv antibody fragments. For yeast display (left panels), scFv genes are cloned as Cterminal fusions to the Aga2 gene, with a c-myc epitope tag at the scFv C-terminus for detection of expression. On the yeast cell surface, Aga2 disulfide bonds to Agal leading to scFv display on the cell surface. HA = HA epitope tag; c-myc = myc epitope tag. Binding yeast can be selected by flow cytometry (right panel). Binding of fluorescent antigen is displayed on the X-axis and binding of fluorescent anti-myc antibody (to quantitate expression) is displayed on the Y-axis. Populations of non-scFv expressing yeast, no antigen binding, low affinity antigen binding, and high affinity antigen binding can be clearly seen. See color insert.
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32. Monoclonal Antibodies from Display Libraries
members was created by scaling up the transformation process. Since flow sorting libraries of such size is not practical due to sorter speeds, the first two rounds of selection were performed using antigen coupled to magnetic beads. Subsequent rounds of selection were performed using flow cytometry. Multiple, different scFvs were isolated to nine different antigens used for selection, with affinities as low as 3 nM. There is no reason why yeast display would not prove similarly useful for the generation of antibodies from immune libraries, though no reported examples exist. The potential advantages of yeast display compared to phage display include: 1) the ability to select (sort) with fine affinity discrimination; and 2) the ability to measure antibody fragment affinity with the displayed antibody. This obviates the need for antibody fragment expression and purification. Potential disadvantages include the lower transformation efficiencies of yeast compared to bacteria.
Ribosome Display For ribosome display, scFv antibody gene repertoires encoded by mRNA are translated in vitro in cell free systems (Figure 32.8) (reviewed in ref. [165]). The lack of a stop codon in the antibody gene repertoire results in the mRNA
FIGURE 32.8 Ribosome display of scFv antibody fragments. ScFv DNA is generated by PCR and then transcribed in vitro to generate mRNA. mRNA is translated in vitro, the absence of a stop codon in the scFv mRNA results in continued attachment of the mRNA to the ribosome, along with the translated scFv. Ribosomes displaying antigen-binding scFv are isolated from nonbinders by affinity chromatography. The ribosome is then disrupted, freeing the mRNA, which is converted back to DNA by RT-PCR. This last step can introduce random mutations into the scFv gene, mimicking the random introduction of mutations by the somatic hypermutation machinery. These steps are repeated to isolate and affinity mature antigen binding scFv. See color insert.
remaining attached to the ribosome along with the translated sequence (the scFv). Thus, the ribosome provides a physical linkage between genotype and phenotype, taking the place of the phage. Antigen binding scFv are selected on immobilized antigen, and the mRNA encoding the scFv genes amplified by RT-PCR. Transcription and translation of the selected scFv provide the display library for the next round of selection. The major advantage of ribosome display is that no cloning is required. mRNA can be transcribed from scFv DNA repertoires created by PCR and can yield libraries much greater in size than those created by cloning. Antigenspecific scFv can be isolated from nonimmune repertoires using ribosome display. Since the RT-PCR amplification process introduces random mutations due to polymerase infidelity, iteration of the selection process leads to affinity maturation of the selected scFv [172]. Ribosome display has been successfully applied to generate scFv from immune [172] and nonimmune [173] libraries and for the affinity maturation of existing antibodies [173]. By altering the selection conditions, it is also possible to select for increased scFv stability [174].
CONCLUSION Although hybridoma technology has proved useful for the generation of reagent and diagnostic antibodies, the immunogencity of rodent monoclonal antibodies has limited their therapeutic development. This limitation has been largely overcome by the chimerization and humanization of murine monoclonal antibodies. Display technologies promise new and improved approaches for the isolation and optimization of fully human antibodies, with affinities not achievable by rodent immunization. For pre-existing murine monoclonal antibodies, display technologies can be used to obtain antibodies that are entirely human in sequence, but that bind to the same epitope as the murine monoclonal (175). In fact, the first fully human phage antibody approved by the U.S. Food and Drug Administration was a murine monoclonal antibody made human by such chain shuffling. Once antibody fragments are isolated using display technologies, they can be engineered into a range of different antibody formats (see Figure 32.2) depending on the application. For therapeutic antibody fragments, the display technologies offer a means to increase stability and production levels. Antibodies have become an important new class of therapeutic molecules, with thirteen now approved by the FDA. Their utility as therapeutics relates to their general safety, ability to interrupt protein–protein interactions better than small molecules, and their known means of production. The display technologies described here should continue to fill the pipeline of therapeutic applications for monoclonal antibodies.
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33 Humanization of Monoclonal Antibodies NAOYA TSURUSHITA AND MAXIMILIANO VÁSQUEZ Protein Design Labs, Inc. Fremont, California, USA
1994; Khazaeli et al., 1994; Kuus-Reichel et al., 1994). Therefore, major research efforts have focused on the elimination, or at least reduction, of the immunogenicity of rodent monoclonal antibodies in humans.
Since the introduction of hybridoma technology by Kohler and Milstein (1975), monoclonal antibodies have rapidly become one of the most important tools in bioscience, with their utility extending to the therapy of human diseases. Hybridoma cells, created by the fusion of an antibody-secreting B cell with a myeloma cell, produce monoclonal antibodies that are homogeneous in composition and monospecific in antigen recognition. In addition, hybridoma cells are immortal and thus can produce an unlimited supply of the monoclonal antibody. Due to their high affinity and specificity, monoclonal antibodies can be used to detect minute amounts of antigen in complex mixtures or on cell surfaces. It is possible, in theory, to obtain monoclonal antibodies against almost any compound, and, indeed, monoclonal antibodies have been successfully generated against proteins, peptides, carbohydrates, and small organic and inorganic chemicals. The utility of monoclonal antibodies to both the diagnostic and therapeutic aspects of medicine is potentially far reaching. The high sensitivity and specificity of monoclonal antibodies have allowed the successful development of a variety of diagnostic tools, including the detection of disease-associated antigens in body fluid (e.g., a blood test to detect PSA molecules associated with prostate cancer; approved by the FDA in 1986; Ward et al., 2001) and the localization of tumor cells in the body (e.g., nofetumomab, a radiolabeled Fab fragment that recognizes the pancarcinoma glycoprotein antigen Ep-CAM; approved by the FDA in 1996; Breitz et al., 1997). Conversely, despite the generation of a number of antibodies with potential clinical application, such as neutralizing antiviral antibodies, the use of monoclonal antibodies as therapeutics was initially hampered by the fact that murine (and other rodent) antibodies proved to be highly immunogenic in humans (Fagnani,
Molecular Biology of B Cells
MURINE, CHIMERIC, AND HUMANIZED ANTIBODIES Clinical applications of murine monoclonal antibodies in the 1980s met with both encouraging and discouraging results. The first successful use of a monoclonal antibody to treat human disease was reported in 1982. Using a murine anti-idiotype monoclonal antibody, Miller et al. (1982) obtained complete remission of disease in a B-cell lymphocytic lymphoma patient. Although other murine monoclonal antibodies tested also showed limited efficacy in certain indications, the vast majority induced a potent human antimurine antibody (HAMA) response (Fagnani, 1994; Khazaeli et al., 1994; Kuus-Reichel et al., 1994), which resulted in neutralization and rapid clearance of the administered murine antibodies upon subsequent treatment, and occasionally induced anaphylactic shock. For example, the murine monoclonal antibody Muromonab-CD3 (Orthoclone OKT3) that was approved by the FDA for treatment of allograft rejection is highly immunogenic in humans despite its immunosuppressive nature (Hooks et al., 1991; Jaffers et al., 1986). Therefore, even though certain murine monoclonal antibodies exhibited the potential to treat human disease, their successful therapeutic application in humans required a significant reduction of their inherent immunogenicity. Advances in genetic engineering greatly facilitated the development of less immunogenic antibodies. Murine– human chimeric antibodies were produced using recombi-
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nant immunoglobulin genes, in which murine constant region exons were replaced with the corresponding human exons (Morrison et al., 1984). The resulting antibody had murine variable regions joined to human constant regions for both the heavy and light chains. These chimeric antibodies are not only more humanlike in amino acid sequence, and therefore potentially less immunogenic in humans than the original murine antibodies, they also elicit stronger effector functions, since the human Fc region interacts more efficiently with human Fc receptors than does the murine Fc region (Ravetch and Kinet, 1991). Moreover, as the circulating half-life of an antibody is in part a property of the Fc region, chimeric IgG antibodies exhibit longer serum halflives than do murine antibodies in humans. Clinical studies demonstrated that chimeric antibodies are indeed effective in treating human diseases and were, in general, less immunogenic than murine antibodies (Khazaeli et al., 1994; Kuus-Reichel et al., 1994; Mountain and Adair, 1992). However, constructing chimeric antibodies did not fully solve the immunogenicity problem. A HAMA, or human anti-chimeric antibody (HACA), response against the variable region was observed with most chimeric antibodies tested, and in some cases the response was severe (Khazaeli et al., 1994; Kuus-Reichel et al., 1994; Mountain and Adair, 1992). To systematically assess the immunogenicity of chimeric antibodies, Bruggemann and coworkers (1989) monitored the immune responses in mice to a human–mouse chimeric antibody, which was composed of a mouse light chain and a chimeric heavy chain carrying human variable and mouse constant regions, alongside murine and human antibodies. Although the murine antibody was nonimmunogenic in mice, the human–mouse chimeric antibody was immunogenic, albeit to a lesser degree than the human antibody. These results indicated that additional modification of the variable region in chimeric antibodies was required to further decrease immunogenicity. Each of the heavy and light chain variable regions is encoded by a single exon and forms a domain structure belonging to the Ig superfamily. The antigen binding site consists of six polypeptide chain loops, three from the heavy chain and three from the light chain. These are called hypervariable domains or complementarity-determining regions (CDRs) (Chothia and Lesk, 1987; Kabat et al., 1991), and all cluster at one end of the variable region. Although the shape of the antigen-binding site is different in each monoclonal antibody in order to recognize a variety of antigen molecules, the remainder of the variable region, called the framework, possesses the same basic structure (Padlan, 1994). Jones et al. (1986) reasoned that the transfer of the CDRs of a murine antibody onto the framework of a human antibody variable region would impart the specificity and affinity of the parental murine antibody. This technique, termed CDR-grafting, is only occasionally successful in maintaining antigen-binding affinity (Riechmann et al.,
1988; Tempest et al., 1991), because certain framework amino acids are essential for proper conformation and orientation of the CDRs and thus crucial in maintaining the three-dimensional structure of the antigen-binding site. Therefore, simple transfer of the CDR amino acids onto a human variable region framework is frequently not sufficient to maintain the affinity of the original murine antibody towards its antigen. To overcome this problem, Queen and collaborators (1989) used a three-dimensional model of the variable regions to identify framework amino acids interacting with amino acids in the CDRs. Upon grafting CDR amino acids along with these important framework amino acids onto a human framework, which was chosen based on its high homology to the counterpart of the original murine antibody, the resulting engineered antibodies were found to maintain the antigen specificity and most of the binding affinity. The term “humanized antibody” has been applied differently over time, as it literally means a more humanlike antibody. In some cases, murine–human chimeric antibodies were referred to as humanized antibodies. In this text, the term “humanized antibody” is used for a genetically engineered antibody in which the CDR amino acids (all or part of them), often along with one or more framework amino acids, are grafted from a murine antibody to a human antibody.
COMPUTER-GUIDED DESIGN OF HUMANIZED V REGIONS The typical process for antibody humanization includes the following steps: 1) cloning and sequencing of the light and heavy chain variable region genes encoding the rodent monoclonal antibody, 2) design of humanized variable region amino acid sequences, 3) synthesis of variable region genes, 4) expression of the humanized antibody, and 5) characterization of the antigen affinity and specificity of the humanized antibody. Except for designing humanized variable regions, these experimental steps can be achieved by using standard molecular biology techniques. Thus, in this section, we are primarily concerned with point 2, and in particular we describe the methodology for computerguided antibody humanization used in our laboratories. The technology is based on the procedure described by Queen and coworkers (1989), and we use the humanization of the murine anti-Tac monoclonal antibody (anti-CD25) to illustrate the major points. Queen’s method of antibody humanization implemented several important refinements to the CDR-grafting technique. First, a human framework with high homology to the original rodent antibody is chosen to minimize the deformation of the CDRs upon grafting. Second, rodent framework amino acids that could interact with the CDRs or
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directly with the antigen (contact framework amino acids) are identified and retained in the humanized antibody. Third, and often less appreciated, “atypical” (or “rare”) amino acids in the acceptor human frameworks are identified and replaced with “typical” human amino acids at corresponding positions. To achieve these goals, our standard procedure for designing humanized variable regions begins with the construction of a three-dimensional model of the variable domains of the antibody to be humanized. We have used the algorithms developed by Levitt and coworkers (Levy et al., 1989; Zilber et al., 1990) for this purpose, although many other programs or approaches may be used with similar success (e.g., Allcorn and Martin, 2002; Morea et al., 1997). Separately, each of the light and heavy chain variable region (VL and VH, respectively) sequences of the rodent antibody is compared to existing sequences of human antibodies. A highly homologous human sequence is chosen as the source of the framework region for grafting the rodent CDR and contact framework amino acids identified in the model. In the case of anti-Tac, the human antibody Eu was chosen to provide the frameworks of both VL and VH regions for CDR grafting. The VL of Eu belongs to subgroup I of human variable kappa chains, whereas the VH belongs to subgroup I of human variable heavy chains, according to the
definition of Kabat et al. (1991). The amino acid sequence identity in the framework regions between anti-Tac and Eu is 65% in the VL and 67% in the VH. Although humanization of anti-Tac used VL and VH sequences from the same human antibody, this is not required. Once the human variable region frameworks are chosen for CDR grafting, framework amino acids that may directly contact antigen or indirectly influence the conformation of the antigen-binding site are identified through the careful examination of the three-dimensional model structure. Although framework amino acids with any side chain atom located within 5 Å of the CDRs usually fall into this category, it is important to thoroughly inspect the location and orientation of the side chain of each of the contact framework residue candidates in order to minimize the number of rodent-specific amino acids retained in the humanized antibody. In the case of anti-Tac, the spatial positions of the following framework amino acids were such that they were considered likely to contribute to antigen binding: 48 and 60 in the VL, and 27, 30, 48, 66, 67, 94, and 103 in the VH (according to the Kabat numbering system, Kabat et al., 1991; Figure 33.1). Although the transfer of the CDR and contact framework residues may be sufficient to maintain the antigen-binding
(A)
Eu Humanized Mouse
10 20 30 40 50 60 DIQMTQSPST LSASVGDRVT ITCRASQSIN TWLAWYQQKP GKAPKLLMYK ASSLESGVPS DIQMTQSPST LSASVGDRVT ITCSASSSI- SYMHWYQQKP GKAPKLLIYT TSNLASGVPA QIVLTQSPAI MSASPGEKVT ITCSASSSI- SYMHWFQQKP GTSPKLWIYT TSNLASGVPA CDR1 CDR2
Eu Humanized Mouse
70 80 90 100 107 RFIGSGSGTE FTLTISSLQP DDFATYYCQQ YNSDSKMFGQ GTKVEVK RFSGSGSGTE FTLTISSLQP DDFATYYCHQ RSTYPLTFGQ GTKVEVK RFSGSGSGTS YSLTISRMEA EDAATYYCHQ RSTYPLTFGS GTKLELK CDR3
(B)
Eu Humanized Mouse
Eu Humanized Mouse
52 10 20 30 40 50 a 60 QVQLVQSGAE VKKPGSSVKV SCKASGGTFS RSAIIWVRQA PGQGLEWMGG IVPMFGPPNYA QVQLVQSGAE VKKPGSSVKV SCKASGYTFT SYRMHWVRQA PGQGLEWIGY INPSTGYTEYN QVQLQQSGAE LAKPGASVKM SCKASGYTFT SYRMHWVKQR PGQGLEWIGY INPSTGYTEYN CDR1 CDR2 82 70 80 abc 90 100 110 QKFQGRVTIT ADESTNTAYM ELSSLRSEDTAFY FCAGGYGIYS PEEYNGGLVT QKFKDKATIT ADESTNTAYM ELSSLRSEDTAVY YCARG-GGVF DYWGQGTLVT QKFKDKATLT ADKSSSTAYM QLSSLTFEDSAVY YCARG-GGVF DYWGQGTTLT CDR3
113 VSS VSS VSS
FIGURE 33.1 The amino acid sequences of the light (A) and heavy (B) chain ariable regions of the mouse anti-Tac (bottom), humanized anti-Tac (middle), and human Eu (top) antibodies are shown in single letter code. The numbering of amino acid locations is according to Kabat et al. (1991). The CDRs are underlined in the mouse sequences. The single and double underlined amino acids in the humanized sequences indicate the differences compared to the Eu sequences. The single underlined amino acids are mouse-specific residues. The double underlined amino acids are changes to the human consensus residues.
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affinity of the original rodent antibody in the humanized form, additional effort is made to eliminate possible immunogenicity by examining the occurrence of atypical amino acids in the human framework selected as the “acceptor.” Atypical amino acids are defined as those that appear infrequently (or do not appear at all) at each position in the framework when compared to human antibody sequences of the same subgroup in the database. If the new amino acid type is not the same as the rodent residue, then the substitution is made only if it is not considered critical for maintaining CDR structure. In the case of anti-Tac, amino acids at positions 48 and 63 in VL and 89, 91, 94, 103, 104, 105, and 107 in VH were identified to be atypical by this “rareness test,” and substituted with the most common amino acids in the same human subgroup at the corresponding positions (Figure 33.1). This process is particularly important when a mature, rearranged human variable region sequence is used as an acceptor, because the framework sequence could carry potentially immunogenic sequences due to somatic hypermutation, or possibly even sequencing errors. Indeed, a stretch of three amino acids (Glu-Tyr-Asn) at positions 103 to 105 in the VH of the Eu antibody is very unusual at this location (Figure 33.1). The framework amino acids at positions 48 of the VL and 94 and 103 of the VH had already been identified as contact framework residues. The mouse residues corresponding to these three amino acids turned out to be also human consensus amino acids. Therefore, only amino acids at positions 60 of the VL and 27, 30, 48, 66, and 67 of the VH, or six amino acids in total, may be considered to be mouse-specific residues. The resulting humanized antiTac antibody retained the specificity for CD25 and exhibited an affinity within three-fold of that of the murine anti-Tac antibody (Queen et al., 1989). A second engineered antibody with only the CDR amino acids grafted onto the VL and VH of Eu showed no binding to CD25 (our unpublished results), indicating the importance of contact framework residues to maintain the integrity and functionality of the antigen-binding site. To date, we have accomplished more than forty antibody humanizations successfully in our laboratories. Although the method reported by Queen et al. (1989) is still the basis of our current humanization technology, the expansion of the antibody sequence and structure databases during the last decade has enabled us to introduce a number of incremental improvements that have made the design of humanized antibodies more straightforward. In the late 1980s, when the humanization of anti-Tac was performed, the database of known three-dimensional structures of antibodies had only a limited number of entries (eight full binding site structures derived from Fabs, and two light chain-only structures derived from Bence-Jones proteins). Now, this number has increased to over 200, of which about 100 are nonredundant. Thus, there is a much higher probability of finding an antibody of known structure that is highly homologous to the
rodent antibody of interest. The level of homology (or percent of sequence identity) is the best predictor of the quality and accuracy of any homology-based structure model (Chothia and Lesk, 1986). In addition, the level of overall sequence conservation usually correlates closely with the matching of CDR canonical structures (Chothia and Lesk, 1987). Therefore, it is now possible to generate more accurate three-dimensional models of the variable regions of the antibody to be humanized. The growth of databases of human antibody sequences has also made it easier to find ideally matched human frameworks as acceptors for grafting of the CDR sequences of any given rodent antibody. From the practical point of view of antibody humanization, the current database of human antibody sequences seems to represent most of the typical human antibody framework sequence repertoire. Access to the catalog of human germline sequences (Matsuda et al., 1998; Zachau, 2000) also makes it possible to apply the “rareness test” (Queen et al., 1989) in a more complete fashion. The amino acids present in the germline variable segments, even if they are atypical or rare at each position, are less likely to be immunogenic in humans, unless the chosen germline sequence is infrequently used in the normal repertoire. The comparison of framework amino acids of an expressed human antibody to the closest human germline variable segment provides reliable guidance in deciding which atypical amino acids should be changed to human consensus residues. Overall, consensus experience suggests that properly humanized antibodies retain the antigen-binding affinity of the original rodent antibodies within a two to three-fold range. Humanized antibodies retaining the affinity and specificity of the original rodent antibodies usually exhibit the same level of biological activities. However, the correlation between affinity and biologic activity is not always congruent. There are examples of antibodies that maintained binding affinity upon humanization, but additional engineering was needed to retrieve full functional characteristics. For example, in our humanization of murine anti-IFN-g antibody AF2 (Thakur and Landolfi, 1999), the first humanized version retained the affinity of the original murine antibody, but was twenty-fold less potent in an in vitro neutralization assay of IFN-g activity than was the murine version. It was ultimately established that the amino acid at position 11 in VH, which is located away from the CDRs and thus had been predicted to have no contribution to binding to IFN-g, is important in the neutralization of IFN-g (Landolfi et al., 2001). Another example of disparate affinity and biological activity is the difference between chimeric and humanized Fab versions of the anti-Her2/neu antibody 4D5. Whereas a humanized Fab form exhibited an antigen-binding affinity similar to that of the chimeric Fab, it severely lost its cytostatic potential (Kelley et al., 1992). Likewise, humanization of a murine antibody
33. Humanization of Monoclonal Antibodies
specific for the RSV glycoprotein F by the “resurfacing” method (described below) fully preserved antigen affinity, but the humanized antibody showed a significant loss of antiviral activity both in vitro and in vivo (Delagrave et al., 1999).
OTHER HUMANIZATION METHODS In this section, we review a number of alternative approaches for antibody humanization, focusing first on design strategies and then on experimental library approaches. Since the initial publications in the late 1980s on antibody humanization (Jones et al., 1986; Queen et al., 1989; Riechmann et al., 1988; Verhoeyen et al., 1988), the total number of humanized antibodies described in the literature has now reached over 100. Although the vast majority of these humanizations have implicitly, or explicitly, followed the design procedures described in the early literature, there are several examples where interesting variations have been introduced into the humanization procedure. The procedure described by Presta and coworkers (Carter et al., 1992), as in the work by Winter and coworkers (Riechmann et al., 1988), uses predefined human framework sequences, instead of choosing homologous human sequences for antibody humanization. Presta and coworkers, following earlier work (Queen and Selick, 1990), chose consensus sequences derived from human subgroups, in particular those of Vk subgroup I and VH subgroup III. This choice corresponds to the most commonly used Vk and VH subgroups in the natural human repertoire of antibodies, leading to the expectation that engineered antibodies based on this approach are less likely to be immunogenic in humans. In the next step of the humanization process, Presta and coworkers used computer-generated models of antibody structures to determine important mouse framework amino acids that need to be retained in the humanized form. In addition, Presta and coworkers changed portions of CDRs from mouse to human in a number of cases (Carter et al., 1992; Presta et al., 1993) with the hope of further reducing immunogenicity without compromising affinity. When the best-fit human framework happens to be from subgroup I for Vk and subgroup III for VH, the sequences humanized by Presta’s approach are likely to differ only by a very small number of amino acids, compared to those made by full application of the Queen method. Presta and coworkers, using this variation, have successfully humanized a number of murine antibodies, including anti-HER-2/neu, anti-CD3, anti-VEGF, and anti-IgE (Carter et al., 1992; Presta et al., 1997; Presta et al., 1993; Shalaby et al., 1992). Padlan has introduced a number of variations for antibody humanization, starting with the idea of the “veneering” of antibody variable domains (1991). The “veneering”
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approach uses statistics collected from the examination of experimentally determined three-dimensional antibody structures to predict the level of surface exposure for each framework amino acid. Using this approach, a mouse variable region sequence to be humanized is first compared to a highly homologous human acceptor framework sequence. Then, any mouse-specific framework amino acids predicted to be surface-exposed are considered to be candidates for replacement with human sequences. In this manner, apart from the CDRs, only the “surface veneer” is human while the remainder is mouse, therefore reducing the risk of deforming the CDR structure and thus impacting affinity or specificity. The veneering approach was used successfully to humanize an antibody specific to human CD18 (Singer et al., 1993). Roguska and coworkers (Pedersen et al., 1994; Roguska et al., 1996; Roguska et al., 1994) also reported a “resurfacing” method for antibody humanization that is conceptually very similar to Padlan’s “veneering” procedure. In the work by Roguska and coworkers (Pedersen et al., 1994), further analysis of the structure and sequence databases was carried out, so that the set of amino acids identified as surface-accessible is slightly different, and more reliably identified, than the set outlined by Padlan. Roguska and coworkers applied their “resurfacing” procedure to several antibodies and showed that the resulting molecules were as active as the original mouse versions (Roguska et al., 1996; Roguska et al., 1994). It should be noted, however, that the level of amino acid conservation between mouse and human sequences tends to be higher in interior than in surface locations. For example, in the original application of Padlan’s “veneering” method (Singer et al., 1993), thirty-eight of the forty-one (93%) VH nonsurface amino acids were already identical between the mouse and best-fit human sequences, whereas the same was true for thirty of the thirty-three (91%) VL nonsurface amino acids. Therefore, at least for this case, there is a relatively small actual difference in humanized sequences between the “veneering” method and those approaches that start with explicit CDR-grafting (Carter et al., 1992; Queen et al., 1989; Riechmann et al., 1988). Beginning with an analysis of the three-dimensional structures of twelve mouse Fab structures, Padlan introduced a “template” or “positional consensus” method, which is considered an advanced version of the “veneering” approach (Padlan, 1992). A version of this procedure was used later in humanization of a murine anti-mucin monoclonal antibody (Couto et al., 1994), and further refinements of the methodology have been described (Couto et al., 1995a; Couto et al., 1995b). Studnicka and coworkers, at about the same time, described a methodology, termed “human engineering,” which is very similar to the “positional consensus” method, and applied it in the humanization of a murine anti-CD5 antibody (Studnicka et al., 1994). In these methods, surface exposure, framework-CDR contacts, and VH–VL interaction in the variable region of a
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mouse antibody are analyzed through an examination of antibody crystal structures. Because of the judgment of each investigator and the actual set of crystal structures examined, the sets of replaceable amino acids, while similar, are not identical in each of the methods (compare Figure 1 of Studnicka et al., 1994, with the listings given in the results sections of Couto et al., 1994, and Couto et al., 1995a). As in the “veneering” and “resurfacing” methods, surfaceexposed framework amino acids are substituted with the counterparts in the acceptor human frameworks of suitable chosen sequences. In addition, amino acids in the mouse framework predicted to have no important role for the formation of CDR structure or VH–VL interaction are considered safe for substitution with human sequences, whereas those matching to positions frequently associated with important structural roles in CDR formation or VH–VL interaction are kept as mouse residues. Although these two methods seem very different from the conventional CDRgrafting followed by analysis of key contact framework residues, the designed sequences may be quite similar to each other. However, there is a tendency for methods based on CDR grafting to lead to humanized frameworks that have fewer amino acids of murine origin. The conventional method (Queen et al., 1989) first produces a threedimensional model of the variable regions, derived from the analysis of known crystal structures, and then, CDRframework contacts are examined. In the “positional consensus” and “human engineering” methods, the CDRframework examination is conducted first, using known crystal structures of antibody variable domains, and then the mouse sequence is compared to the result of this analysis. More recently, Padlan and coworkers (De Pascalis et al., 2002; Tamura et al., 2000) have outlined an approach based on specificity-determining regions (SDRs) (Padlan et al., 1995). In general, the SDRs comprise fewer amino acids than CDRs, so that SDR-grafting, in theory, should lead to less immunogenic humanized antibodies. Unlike CDRs, however, SDRs are not always predictable from sequence alone, although some degree of generalization is possible from an examination of the three-dimensional structures of antibody–antigen complexes (Padlan, 1990; Mian et al., 1991; Martin and Thornton, 1996) and from previous antibody engineering studies on the role of CDR amino acids in antigen binding (Glaser et al., 1992; Kelley and O’Connell, 1993; Iwahashi et al., 1999; Tamura et al., 2000). Therefore, full implementation of the SDR concept may still require a significant amount of experimentation of a trial-and-error nature without a guarantee that more “human-like” CDRs can be found with any given mouse antibody. Nonetheless, for a murine anti–TAG-72 antibody, Padlan and coworkers (Tamura et al., 2000) were able to construct a humanized version in which the L1 and L2 CDRs were entirely derived from the human framework, as was one amino acid in L3
and two more in H2. Interestingly, the resulting antibody lost only three-fold of its affinity towards the antigen. It should also be noted that the early work of Presta and coworkers on the humanization of the anti-HER2 antibody, which led to the development of trastuzumab, anticipated part of the SDR concept, as Presta’s group used “human” amino acids within the CDR portion of L1, L2, and H2 that were not derived from the CDR sequences of the original murine 4D5 antibody (Carter et al., 1992). One interesting development potentially relevant to antibody humanization is the construction of a virtually complete catalog of human germline sequences (Cook and Tomlinson, 1995; Matsuda et al., 1998; Meindl et al., 1990; Williams et al., 1996). The use of germline sequences as acceptor frameworks (Hsiao et al., 1994) may be considered a major improvement over the use of human mature—and therefore somatically mutated—sequences, as such mutations are likely to be unique in each antibody and thus potentially immunogenic in the general human population. However, the early humanization methods largely anticipated this issue stemming from somatic hypermutations in the frameworks by either using consensus variable region sequences (Carter et al., 1992), or by replacing “rare” amino acids in the human frameworks with consensus residues (Queen et al., 1989). Thus, the direct use of human germline sequences as acceptor frameworks only marginally alters the final designed sequences that follow faithfully more conventional humanization methods, and we have found this to be true in a number of cases (our unpublished results). The availability of human germline sequences nevertheless is helpful even in the context of comprehensive approaches to antibody humanization, as already indicated in the previous section. Building on systematic biochemical and structural studies of the humanization of an anti-lysozyme antibody (Foote and Winter, 1992; Holmes and Foote, 1997; Holmes et al., 1998; Holmes et al., 2001), Foote and coworkers described an intriguing variation in the use of germline sequences as acceptors (Tan et al., 2002). They focused on sequence comparison in the CDR and not in the framework regions; in particular, these authors tried to match the canonical classes of CDRs using Chothia’s structure-based definition (Chothia and Lesk, 1987; Chothia et al., 1989) between the mouse sequence of interest and the candidate human germline sequence. This “superhumanization” approach may seem to be a radically different way of choosing human acceptor sequences for humanization, as the framework regions usually have been used in comparing mouse and human sequences to dictate the choice of acceptors. However, because particular combinations of CDR canonical structures and sequences tend to correlate with particular framework sequences (Chothia et al., 1992; Tomlinson et al., 1995), the choice of human acceptor sequences based
33. Humanization of Monoclonal Antibodies
on the “superhumanization” approach does not necessarily provide a significant difference compared to the choice based on the alignment of framework sequences. For example, using the CDR-fit criteria on a database of human germline sequences, Foote and coworkers chose B3 for the acceptor human VL framework and DP-45 for the VH in the humanization of a murine anti-CD28 antibody (Tan et al., 2002). B3 is the sole member of human Vk germline family IV (Schable and Zachau, 1993; Tomlinson et al., 1995), whereas DP-45 belongs to VH germline family III (Tomlinson et al., 1992). When the entire VL and VH sequences of this antibody are compared to the collection of human germline sequences, the matches with the most identical amino acids are B3 for the VL, and DP-65 (family IV) for the VH, respectively. Thus, in this example, “superhumanization” would be identical to standard CDR grafting with a best-fit match to germline sequences in the VL, but different in the VH (DP-45 differs from DP-65 at thirty-one framework positions). Whether a humanized antibody with an engineered VH based on DP-65 would perform better or worse, in terms of binding affinity and biologic activity, than the DP-45–based construct actually made by Foote and coworkers must be tested experimentally. Besides the structure-guided approaches described above, a library approach is also possible to obtain properly humanized antibodies. The “guided selection” method, originally described by Jespers et al. (1994), uses phage display vectors to express antibody fragments (Fab, single-chain Fv, VH, or VL). The light (or heavy) chain of a rodent antibody to be humanized is paired with diverse human heavy (or light) chains to construct a V gene–shuffling library. After selecting antigen-binding clones, the remaining mouse chain is replaced with diverse human corresponding chains in a new V gene–shuffling library, and the selection is repeated. Using this method, Jespers et al. (1994) isolated a humanized anti-TNF antibody that as a whole antibody had antigen affinity comparable to that of the original mouse antibody. The “guided selection” method has since been used for the humanization of several rodent antibodies (Beiboer et al., 2000; Figini et al., 1998; Klimka et al., 2000; Wang et al., 2000). The modified “guided selection” method used by Rader et al. (1998) for the humanization of a murine antiaVb3 antibody requires the grafting of the VH (or VL) CDR3 sequence of a mouse antibody onto diverse human VH (or VL) sequences before constructing V gene–shuffling libraries. As the CDR3 regions are considered critical in determining the affinity and specificity of an antibody, this approach allows the identification of human CDR1, CDR2, and framework sequences that match to the grafted mouse CDR3 sequence in properly forming the antigen binding site. This modified approach was further applied for the humanization of a rabbit monoclonal antibody (Rader et al., 2000).
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The humanization method reported by Rosok et al. (1996) used both structure-guided and library approaches. As in the conventional structure-guided humanization methods, the variable region sequences of a mouse antibody were first aligned with human antibody sequences, and highly homologous human frameworks were chosen for CDR-grafting. Among framework amino acids that were different between the mouse and human sequences, surfaceexposed amino acids were kept as human in the acceptor framework. For buried amino acids, their importance in maintaining the proper structure of antigen binding site was directly tested with a library approach. The CDR-grafted antibody was expressed in the Fab form by phage display and a library containing all the combinations between mouse and human sequences at the potential CDR-contacting, buried framework residues was constructed. After selecting phage clones based on antigen binding, Rosok et al. (1996) obtained humanized anti-Lewis Y antibodies, in which only a portion of buried amino acids originated from the murine sequence, with an affinity equivalent to that of the original mouse antibody. In extending this approach to the humanization of a murine anti-CD40 antibody, Wu et al. (1999) introduced mutations into the light and heavy chain CDR3 sequences in a combinatorial Fab-expressing phage library between mouse and human sequences at positions of potential CDR-contacting framework residues. The selected humanized anti-CD40 antibody exhibited an affinity approximately seventeen-fold higher than that of the chimeric form of the original antibody. Baca’s library approach (1997) is similar to Rosak’s, but optimization of the framework in CDR-grafted antibodies was performed by the random mutagenesis of a small set of framework residues critical for antigen binding. The resulting humanized anti-VEGF antibody had approximately six-fold weaker binding to antigen compared to the chimeric form of the original antibody. Since the initial publications in the late 1980s and early 1990s, humanization of mouse and other rodent antibodies has become increasingly straightforward, and indeed reports of successful applications have appeared at a rapid pace in the scientific and patent literature. By examining the PubMed database, we found about 250 publications between 1986 and 1995 that used the term “humanized antibody.” In 1996 alone, there were 64 such publications, with the numbers increasing to 79, 94, 138, 152, and 174 from 1997 through 2001, respectively. Although many variations in the humanization technique were introduced, the main scheme for successful humanization in the vast majority of reported cases still remains the same: identification of framework amino acids critical to the proper formation of the structure of the antigen binding site in an antibody to be humanized, and their transfer together with CDR amino acids (all or part) onto an appropriately chosen human acceptor framework.
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IMMUNOGENICITY OF HUMANIZED ANTIBODIES Clinical studies have indicated that humanized antibodies are far less immunogenic in humans than are murine antibodies, and in many cases appear to be completely nonimmunogenic (Vincenti et al., 1998; Vincenti et al., 1997). As to the difference between humanized and chimeric antibodies, clearly the general trend is that humanized antibodies are less immunogenic (Fagani, 1994; Stephens et al., 1995; Van Assche and Rutgeerts, 2000). In a study comparing the pharmacokinetics of chimeric and humanized versions of a murine anti-EGF receptor monoclonal antibody in African Green monkeys, the humanized antibody was at least twofold less immunogenic than the chimeric antibody (Mateo et al., 1997). Nonetheless, humanized antibodies are not always completely free from immunogenicity. Depending on target antigens and the sensitivity of assays, weak human antihumanized antibody (HAHA) responses can be detected in subpopulations of human subjects treated with humanized antibodies. The immunogenicity of monoclonal antibodies in humans is a result of the combination of many factors, including the number of nonhuman amino acids retained, structure and aggregation state of the administered antibody, nature of the antigen, dosing, frequency of administration, and immunocompetence of the patients. For example, Fab fragments of murine antibodies are less immunogenic than their whole antibody counterpart because Fab fragments are cleared from circulation faster due to their small size and the lack of interaction with FcRn, a receptor involved in regulating catabolism of antibodies (Ghetie and Ward, 2000). Administered antibodies are also less likely to be immunogenic in immune-deficient, immune-compromised, or immune-suppressed patients than they are in healthy individuals. On the other hand, the frequent administration of high doses of antibodies is more likely to induce immunogenicity. Also, cell-binding antibodies tend to be more immunogenic than non–cell-binding antibodies, particularly those binding to leukocytes or antigen-presenting cells (Gilliland et al., 1999; Isaacs and Waldmann, 1994). The major difference between chimeric and humanized antibodies is the extent of amino acids of rodent origin retained in the variable regions. In addition to CDR amino acids, humanized antibodies typically carry three to six, nonhuman consensus residues derived from the original rodent antibody in heavy chain variable regions, and one to three in light chain variable regions. In contrast, chimeric antibodies carry twenty to twenty-five rodent-derived residues in the framework of both heavy and light chain variable regions. This level of difference in amino acid composition seems to provide the basis for the lower immunogenicity of humanized antibodies. Indeed, this may be quantified by calculating the level of sequence identity between rodent, or humanized, V regions and the closest matching human
germline V-gene sequences (Clark, 2000). We have carried out this analysis for humanized antibodies created in our laboratories and their parental rodent antibodies. For VL, the rodent V regions have, on average, a best-matching human germline that is 69% identical. The humanized versions of the corresponding VLs are, on average, 84% identical. A similar result is observed for VHs, wherein rodent versions are 66% identical to their best-matching human germline sequences, whereas humanized versions are 80% identical. Also of interest is the observation that in no case did humanization lead to a decrease in the level of matching to human germline sequences. Thus, to the extent to which “percent identity” to human germline sequences may be used to predict immunogenicity, humanization, at least as practiced in our laboratories, does lead to antibodies expected to have reduced immunogenicity in humans. Therefore, although chimeric antibodies are not always strongly immunogenic in humans, humanized antibodies provide a safer choice with respect to immunogenicity for clinical development. The goal of antibody humanization is to convert a nonhuman monoclonal antibody to be as humanlike as possible with respect to immunogenicity, but without losing the antigen specificity and binding affinity of the original antibody. Although humanized antibodies are far less immunogenic than the original rodent antibodies, it is difficult to ascertain whether a humanized antibody is different, in terms of immunogenicity, from a native human antibody. To this end, Klingbeil, at Protein Design Labs, conducted an extensive pharmacokinetics analysis in Rhesus monkeys by injecting either human (HSV863; Harfeldt et al., 1997) or humanized (HuFd79D; Co et al., 1991) antibody against herpes simplex virus (unpublished observation). The results in Figure 33.2 show no significant difference in the serum levels between HSV863 and HuFd79D, nor was there any sign of immune response to either one of the administered antibodies, thus indicating that the human and humanized antibodies were indistinguishable with respect to immunogenicity in this study.
HUMANIZED ANTIBODIES APPROVED FOR CLINICAL USE In the United States, 15 monoclonal antibodies (3 murine, 4 chimeric, 7 humanized, and 1 human) have been approved by the FDA for therapeutic use (Table 33.1). Daclizumab, the first approved humanized antibody, is the humanized version of a murine anti-Tac (CD25) monoclonal antibody. It was approved in 1997 for the prevention of renal allograft rejection. In the two phase III clinical studies of 535 patients who underwent kidney transplantation, daclizumab significantly reduced the frequency of acute rejection without increasing the number of serious adverse events associated with HAHA responses as compared with
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TABLE 33.1 Monoclonal antibodies approved for therapeutic use in the United States Generic name
Product name
Antigen
Indication
Form
Approval
Company
Muromonab-CD3 Abciximab* Rituximab Daclizumab Infliximab Basiliximab Trastuzumab Palivizumab Gemtuzumab ozogamicin** Alemtuzumab Ibritumomab tiuxetan*** Adalimumab
Orthoclone OKT3 ReoPro Rituxan Zenapax Remicade Stimulec Herceptin Synagis Mylotarg
CD3 GP IIb/IIIa receptor CD20 CD25 TNFa CD25 HER-2/neu RSV glycoprotein F CD33
Allograft rejection Coronary angioplasty NHL Renal allograft rejection RA, CD Renal allograft rejection Breast Cancer RSV infection AML
Murine Chimeric Chimeric Humanized Chimeric Chimeric Humanized Humanized Humanized
1986 1994 1997 1997 1998 1998 1998 1998 2000
Ortho Biotech Centocor IDEC/Genentech Protein Design Labs/Roche Centocor Novartis Genentech MedImmune Celltech/Wyeth
MabCampath Zevalin
CD52 CD20
CLL NHL
Humanized Murine
2001 2002
Millennium/ILEX IDEC
Humira
TNFa
RA
Human
2002
Omalizumab Tositumomab**** Efalizumab
Xolair Bexxar Raptiva
IgE CD20 CD11a
Asthma NHL Psoriasis
Humanized Murine Humanized
2003 2003 2003
Cambridge Antibody Technologies/Abbott Tanox/Genentech/Novartis Corixa/GlaxoSmithKline Xoma/Genentech
Abbreviations: RSV, respiratory syncytial virus; NHL, non-Hodgkin’s lymphoma; RA, rheumatoid arthritis; CD, Crohn’s disease; AML; acute myeloid leukemia; CLL, chronic lymphocytic leukemia. The murine anti-Ep-CAM monoclonal antibody, Panorex, developed by GlaxoSmithKline and Centocor, was approved in 1995 for treatment of colorectal cancer in Germany. * Produced in the Fab form. ** Conjugated to calicheamicin. *** Conjugated to Indium-111 and Yttrium-90. **** Conjugated to Iodine-131.
Pharmacokinetics ofAnti-HSV Antibodies in Monkeys 100 Antibody Serum Level(mg/ml)
IV injections
SC injections Human Humanized
10
1
0.5 mg/kgdose 0.1 0
50
100
150
200
250
Days
FIGURE 33.2 Either human (HSV863) or humanized (HuFd79D) monoclonal antibody against herpes simplex virus was injected into three rhesus monkeys at 0.5 mg/kg intravenously on days 1, 71, 85, and 99, and subcutaneously on days 184, 197, and 212. The serum level of HSV863 or HuFd79D in each monkey was monitored by ELISA at indicated days. The averages with three monkeys are shown for each of the HSV863 and HuFd79D groups in the figure.
placebo (Bumgardner et al., 2001). Besides organ transplantation, daclizumab has been tested in clinical trials for chronic autoimmune diseases including uveitis (Nussenblatt et al., 1999), psoriasis (Krueger et al., 2000), and ulcerative colitis (Van Assche et al., 2002), and for treatment of graftversus-host disease (Przepiorka et al., 2000). To date, over 16,000 patients have been treated with daclizumab; the consensus is that daclizumab is safe and well tolerated without inducing any major adverse events related to HAHA responses. Following daclizumab, six additional humanized monoclonal antibodies have been approved for treatment of human disease. These include palivizumab for RSV infection, trastuzumab for breast cancer, gemtuzumab ozogamicin for acute myeloid leukemia, alemtuzumab for chronic lymphocytic leukemia, omalizumab for asthma, and efalizumab for psoriasis (Table 33.1). Daclizumab, palivizumab, trastuzumab, alemtuzumab, omalizumab, and efalizumab are all of IgG1 isotype, whereas gemtuzumab ozogamicin is an IgG4 conjugated to calicheamicin, a cytotoxic anti-tumor agent. Besides the indications approved by the FDA, some of these humanized antibodies are also being tested for other diseases. For example, trastuzumab, which blocks the function of the Her2/neu (c-erbB-2) oncogene, is currently being evaluated for treatment of various solid tumors. In addition to the five market-approved humanized antibodies, several humanized antibodies are currently undergoing phase III
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trials. The status of the clinical trials with humanized antibodies can be viewed at the NIH web site (http://www.clinicaltrials.gov/ ).
CONCLUSION Shortly after the introduction of hybridoma technology, it became obvious that monoclonal antibodies could be important human therapeutics. The clinical use of monoclonal antibodies, however, had to wait for advances in genetic engineering. Over the past 15 years, antibody humanization has proved to be a powerful technique to minimize, or even eliminate, the immunogenicity of rodent monoclonal antibodies. Since the basic framework structure of the immunoglobulin variable regions is well conserved among species, humanization technology may be applied beyond rodent antibodies. Nonrodent antibodies, particularly nonmammal antibodies, would recognize antigens (or epitopes) highly conserved between rodents and humans, to which rodents cannot effectively generate antibodies. Thus, the use of nonrodent monoclonal antibodies for humanization could expand the repertoire of antibody-based therapeutics. Currently, over fifty humanized antibodies are being evaluated in a variety of clinical trials. The antigens include cytokines and their receptors, chemokines and their receptors, infectious viruses, microorganisms and their toxins, tumor-specific antigens, growth factor receptors, angiogenic factors, and adhesion molecules. With the advances in genomics and proteomics, the number of target molecules for antibody-based therapeutics is rapidly increasing. Once a rodent antibody with clinical potential is generated by the well-established hybridoma technology, humanization will reduce its immunogenicity for therapeutic use. Humanized antibodies can be used in the native form (IgG1~4, IgM, IgA) or as conjugates with a cytotoxic agent (protein or small molecule toxin) or radionuclide. Furthermore, humanized antibodies can be engineered for better antibodydependent cellular cytotoxicity (ADCC) and complementdependent cytotoxicity (CDC) activities for tumor targeting through genetic modifications in the Fc region. Humanized antibodies are, in general, safe and well tolerated in humans (Baselga, 2001; Leyland-Jones et al., 2001; Niemeyer et al., 2002; Sorrentino and Powers, 2000). With these properties, it is expected that the number of humanized antibodies evaluated in clinical trials will increase dramatically. It is now clear that humanized antibodies represent an important new class of therapeutics. As such, we expect that many of them will be approved for the treatment of otherwise incurable human diseases.
Acknowledgments We are grateful to Bill Benjamin, Paul Hinton, Shankar Kumar, Nick Landolfi, Kanokwan Pakabunto, Cary Queen, Bill Schneider, and J Tso for discussion and comments on the manuscript. We also thank Man Sung Co for multiple discussions on humanization technology and Corine Klingbeil for allowing us to use the unpublished pharmacokinetics data.
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Guild, B. C. (1994). Humanization of murine monoclonal antibodies through variable domain resurfacing. Proc Natl Acad Sci U S A 91, 969–973. Rosok, M. J., Yelton, D. E., Harris, L. J., Bajorath, J., Hellstrom, K. E., Hellstrom, I., Cruz, G. A., Kristensson, K., Lin, H., Huse, W. D., and Glaser, S. M. (1996). A combinatorial library strategy for the rapid humanization of anticarcinoma BR96 Fab. J Biol Chem 271, 22611–22618. Schable, K. F., and Zachau, H. G. (1993). The variable genes of the human immunoglobulin kappa locus. Biol Chem Hoppe Seyler 374, 1001–1022. Shalaby, M. R., Shepard, H. M., Presta, L., Rodriques, M. L., Beverley, P. C. L., Feldmann, M., and Carter, P. (1992). Development of humanized bispecific antibodies reactive with cytotoxic lymphocytes and tumor cells overexpressing the HER2 protooncongene. J Exp Med 175, 217–225. Singer, S. S. II, Kawka, D. W., DeMartino, J. A., Daugherty, B. L., Elliston, K. O., Alves, K., Bush, B. L., Cameron, P. M., Cuca, G. C., Davies, P., et al. (1993). Optimal humanization of 1B4, an anti-CD18 murine monoclonal antibody, is achieved by correct choice of human V-region framework sequences. J Immunol 150, 2844–2857. Sorrentino, M., and Powers, T. (2000). Effectiveness of palivizumab: Evaluation of outcomes from the 1998 to 1999 respiratory syncytial virus season. The Palivizumab Outcomes Study Group. Pediatr Infect Dis J 19, 1068–1071. Stephens, S., Emtage, S., Vetterlein, O., Chaplin, L., Bebbington, C., Nesbitt, A., Sopwith, M., Athwal, D., Novak, C., and Bodmer, M. (1995). Comprehensive pharmacokinetics of a humanized antibody and analysis of residual anti-idiotypic responses. Immunology 85, 668– 674. Studnicka, G. M., Soares, S., Better, M., Williams, R. E., Nadell, R., and Horwitz, A. H. (1994). Human-engineered monoclonal antibodies retain full specific binding activity by preserving non-CDR complementarity-modulating residues. Protein Eng 7, 805–814. Tamura, M., Milenic, D. E., Iwahashi, M., Padlan, E., Schlom, J., and Kashmiri, S. V. (2000). Structural correlates of an anticarcinoma antibody: Identification of specificity-determining residues (SDRs) and development of a minimally immunogenic antibody variant by retention of SDRs only. J Immunol 164, 1432–1441. Tan, P., Mitchell, D. A., Buss, T. N., Holmes, M. A., Anasetti, C., and Foote, J. (2002). “Superhumanized” antibodies: Reduction of immunogenic potential by complementarity-determining region grafting with human germline sequences: Application to an anti-CD28. J Immunol 169, 1119–1125. Tempest, P. R., Bremner, P., Lambert, M., Taylor, G., Furze, J. M., Carr, F. J., and Harris, W. J. (1991). Reshaping a human monoclonal antibody to inhibit human respiratory syncytial virus infection in vivo. Biotechnology (NY) 9, 266–271. Thakur, A. B., and Landolfi, N. F. (1999). A potent neutralizing monoclonal antibody can discriminate amongst IFNgamma from various primates with greater specificity than can the human IFNgamma receptor complex. Mol Immunol 36, 1107–1115. Tomlinson, I. M., Cox, J. P. L., Gherardi, E., Lesk, A. M., and Chothia, C. (1995). The structural repertoire of the human V-kappa domain. EMBO J 14, 4628–4638. Tomlinson, I. M., Walter, G., Marks, J. D., Llewelyn, M. B., and Winter, G. (1992). The repertoire of human germline VH sequences reveals about fifty groups of VH segments with different hypervariable loops. J Mol Biol 227, 776–798. Van Assche, G., Dalle, I., Noman, M., Aerden, I., Swijsen, C., Asnong, K., Maes, B., Ceuppens, J., Geboes, K., and Rutgeerts, P. (2002). A pilot study on the use of the humanized anti interleukin-2 antibody, Daclizumab, in active ulcerative colitis. Am J Gastroenterol. In press. Van Assche, G., and Rutgeerts, P. (2000). Anti-TNF agents in Crohn’s disease. Expert Opin Investig Drugs 9, 103–111.
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34 Human Monoclonal Antibodies from Translocus Mice MARIANNE BRÜGGEMANN Laboratory of Developmental Immunology, The Babraham Institute, Babraham, Cambridge, United Kingdom
The properties of antibodies and the general excitement just over a century ago when it was discovered that serum can neutralize harmful toxins has been judiciously summarised by César Milstein in the previous issue of this book. His most famous scientific discovery was the immortalization of antibody producing B cells, which allowed the production of hybridomas secreting mouse monoclonal antibodies (Köhler and Milstein, 1975). This discovery, which led to a now widely used procedure of generating specific antibodies in rodents following immunization with proteins or cells, was essential to establish the use of monoclonal antibodies for immunotherapy (Waldmann and Cobbold, 1993). However, the therapeutic application of monoclonal and polyclonal antibodies could be greatly improved if they were of human origin, since such antibodies should exhibit markedly reduced immunogenicity in patients (Klingbeil and Hsu, 1999). For this reason, and to allow the study of antibody expression, transgenic mice carrying human immunoglobulin (Ig) genes in germline configuration were first created some 15 years ago (Brüggemann et al., 1989). In this review, I summarize the various strategies used to introduce and express human Ig heavy (H) and light (L) chain genes in transgenic animals and show that a diverse human antibody repertoire can be obtained in a mouse background with silenced endogenous IgH and IgL chain loci. The use of large Ig loci permits hypermutation, classswitching, and good expression levels. However, I also draw attention to present shortcomings and speculate what the future may provide.
recently bacterial artificial chromosomes (BACs) and yeast artificial chromosome (YACs). This has allowed the determination of the sequences of probably all variable (V) and certainly all diversity (D), joining (J), and constant (C) region genes. The human IgH locus is ~1.5 Mb in size, with ~50 functional VH genes, twenty-seven D segments, six functional JH segments, and nine functional CH genes (Cook et al., 1994; Hofker et al., 1989; Corbett et al., 1997; Matsuda et al., 1998). Of the two human L chain loci, the Igl locus is ~1.1 Mb and accommodates fifty-two Vl and seven JlCl genes (Frippiat et al., 1995; Kawasaki et al., 1995). The Igk locus is the largest Ig locus comprising ~3 Mb and carrying a total number of approximately onehundred forty Vk genes, of which seventy-five are functional and a further twenty-one potentially functional, five Jk and one Ck gene (Roschenthaler et al., 2000; Zachau, 2000). Members of the different VH and Vk gene families are interspersed, whereas the Vl families form group clusters. Interestingly, IgH chain as well as Igl chain gene segments (as far as known) are assembled in the same transcriptional orientation, which permits conventional DNA rearrangement and deletion, whereas Vk genes are organized in both transcriptional orientations, which allows deletional and inversional joining (Frippiat et al., 1995; Weichhold et al., 1990). In transgenic mice rearrangement and expression of human Vk genes in either transcriptional orientation has been achieved and expression preference was not observed (Xian et al., 1998).
Assembly of Minigene Constructs HUMAN IG TRANSLOCI
An essential question in developing the transgenic mouse approach was whether the introduced human genes would be rearranged and expressed. Of similar importance in the creation of a human antibody repertoire in the mouse was a
In the last few years, the human Ig loci have been entirely cloned, initially using phage and cosmid vectors and more
Molecular Biology of B Cells
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Copyright 2004, Elsevier Science (USA). All rights reserved.
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flawless interaction of the transgenic Ig chains with the cellular signaling machinery to allow, for example, Ig transport and anchoring in the cell membrane to permit BCR assembly as well as antigen recognition and B-cell expansion. For the expression of human IgH and IgL chain genes on minigene constructs, the V-, (D)-, J-, and C-gene segments in germline configuration were placed in artificially close proximity. The sizes of the different constructs (less than 100 kb) limited the number of VH and Vk genes, which was between one and five, and also the number of functional D segments
for the IgH chain, which was between three and fifteen (Brüggemann et al., 1989 and 1991; Taylor et al., 1992 and 1994; Lonberg et al., 1994; Xian et al., 1998). In the initial IgH chain constructs, a ~15 kb region including DQ52, JH, Em, switch m, and Cm was largely or completely retained, whereas for an Igk construct a Vk was added to the Jk – Ck cluster (Table 34.1). An advantage of maintaining the region between J and C was that the functional activity of the intronic enhancer, such as transcriptional activation of a rearranged IgH or IgL chain, could be secured. Adding an
TABLE 34.1 Size, V gene content and serum titre of mice carrying human Ig transloci. The approximate (usually mean) titre of serum antibody containing human Ig is shown for the different transgenic lines carrying silenced endogenous monse H and/or L chain (k and/or l) loci. Titres in a non-KO (normal mouse antibody) background is indicated (*). Expression in chimeric mice is indicated (§) Translocus
Human Ig chains
Size (kb or Mb)
Functional V genes
Titre (mg ml-1)
Reference
Plasmid-based transgenes IgH loci HuIgH
m
25 kb
2 Vs
20*, 300
HuIgHCOS
m
100 kb
2 Vs
50*, 350
HC1 HC2
m,g1 m,g1
61 kb 80 kb
1V 4 Vs
10 100
Brüggemann et al., 1989; Wagner et al., 1994; Wagner et al., 1994a Brüggemann et al., 1991; Wagner et al., 1994; Wagner et al., l994a Taylor et al., 1992 Lonberg et al., 1994; Taylor et al., 1994
k k k
24 kb 43 kb 15 kb
1V 4 Vs 5 Vs
100 15*, 50
Taylor et al., 1992 Lonberg et al., 1994 Xian et al., 1998
IgH loci J1.3 HuIgHP1–2 HuIgH yH1 yH2
m,d m,d m,d m,d m,d,g2(or g1 or g4)
85 kb 210 kb 240 kb 220 kb 1020 kb
2 Vs 5V 5 Vs 5 Vs ~40 Vs
0.2* 5*, 180 200 1*, 350 700 (IgM), 600 (IgG)
Choi et al., 1993 Wagner et al., 1996 Nicholson et al., 1999; .Mundt et al., 2001 Green et al., 1994 Mendez et al., 1997; Davis et al., 1999; Green, 1999
IgL loci yK1 HuIgk YAC HucosIgk YAC KCo5 yK2 Igl
k k k k k l
170 kb 300 kb 1300 kb 450 kb 800 kb 380 kb
2 Vs 2 Vs ~80 Vs ~26 Vs ~25 Vs 15 Vs
16* 50*, 800 100*, 800
Green et al., 1994 Davies et al., 1993; Xian et al., 1998 Xian et al., 1998; Zou et al., 1996 Fishwild et al., 1996 Mendez et al., 1997 Popov et al., 1999
IgL loci KC1 KCo4 HuIgkML YAC-based transgenes
800 400*, 1500
Chromosome-fragment transgenes IgH loci hCF(SC20)
m,d,g,e,a
>20 Mb
whole locus
10*, 350 (IgM); 3*, 300 (IgG)
Tomizuka et al., 2000; Tomizuka et al., 1997
IgL loci hCF(2-W23) hCF[MH(ES)22-1]§
k l
5–50 Mb near intact chromosome 22
whole locus whole locus
60*, 450 20*
Tomizuka et al., 2000; Tomizuka et al., 1997 Tomizuka et al., 1997
34. Human Monoclonal Antibodies from Translocus Mice
additional CH gene to an IgH chain minigene construct proved difficult, as the repetitiveness of the m switch region affected cloning stability in Escherichia coli. Nevertheless, Taylor et al. (1992) obtained such a construct and showed that switching from Cm to Cg1 is possible. For the derivation of transgenic mice, purified DNA was microinjected into the male pro-nucleus of fertilized mouse eggs usually obtained after superovulation (Gordon and Ruddle, 1983). This resulted in transgenic animals with multiple copies of the human Ig construct. The advantage was that multiple copies permitted better expression and also tandem integration of two or more constructs mixed and injected together. This resulted in head to tail integration of two cosmids and created a ~100 kb IgH locus that rearranged and expressed a combination of segments from both cosmids (Brüggemann et al., 1991). A further increase in size was achieved by the use of the P1 cloning system and the microinjection of three overlapping regions of ~80 kb each (Wagner et al., 1996). Following homologous recombination with each other, ~180 kb of the core region (defined as the minimum number of gene segments [V, (D), J, and C] to allow DNA rearrangement and expression) of the human IgH locus, containing five VH genes, all D and J segments, and Cm and Cd, was reconstituted. For the Igk chain, coinjection of two minigene constructs allowed the integration of a contiguous 43-kb translocus following homologous recombination between V genes (Lonberg et al., 1994). These constructs, some with closely assembled exons and control regions, established that human Ig gene segments in germline configuration could be rearranged and expressed in mouse lymphocytes. The addition of mouse sequences, although potentially beneficial to drive transgene expression, appears to be unnecessary if equivalent human sequences, such as enhancer regions, are included. However, the expression of small human Ig constructs is generally poor, and fully human antibody repertoires have not been obtained in mice that still rearrange and express their own endogenous IgH and IgL chain genes.
Modification and Introduction of YACs Defined larger genomic regions ranging from a few hundred kb to around one Mb have either been directly cloned into YAC vectors or have been isolated from established YAC libraries. Cloning in yeast has the advantage that YACs can be easily manipulated by site-specific integration, which allows targeted addition (retrofitting) or the removal of specific sequences. Moreover, YACs have been extended and modified by mating of yeast clones carrying different but overlapping regions. This has allowed locus extension by homologous recombination between YACs (Markie, 1996). The core region of the human IgH locus (and separately the Igk locus) that included V, (D), J, and C genes has been cloned on YACs up to ~300 kb (reviewed in
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Brüggemann and Neuberger, 1996). Impressive V gene additions by stepwise recombination have been made to IgH and Igk YACs (Mendez et al., 1997; Zou et al., 1996). This resulted in a ~1 Mb human IgH YAC with ~66 VH genes and a ~800 kb Igk YAC with ~32 Vk genes with VHs, Ds, JHs, Cm, and Cd and Vks, Jks, and Ck in authentic configuration (Mendez et al., 1997). The efficiency of the yeast host in homologous integration has also been exploited by extending a 300-kb Igk YAC (Davies et al., 1993) to 1.3-Mb by multiple integration of a 50-kb cosmid with five Vk genes (Zou et al., 1996). However, for the human Igl locus, no cloned core region (Vl-JCls) was available and this had to be constructed. To achieve this, three overlapping cosmids with their 5¢ and 3¢ regions ligated to YAC vector-arm sequences were co-transfected into yeast. YACs with correctly reconstituted Igl core region were identified by Southern hybridization (Popov et al., 1996). Further extension was achieved by yeast mating, which allowed homologous recombination of the overlapping region and resulted in an Igl YAC with the authentic 380-kb region accommodating fifteen functional Vl genes (Popov et al., 1999). This stepwise recombination of YACs (methods described in Markie, 1996) has now been routinely used and allows the creation of human Ig loci with large or near authenticregions over 1 Mb in size. The rearrangement and expression of different V genes from a large gene pool is regarded essential to create antibody repertoires comparable to the spectrum expressed in humans. For this reason, much attention focused on using artificial chromosomes to accommodate large regions to which individual V genes of the different families, either on minigene constructs or in authentic configuration, could be added (Lonberg et al., 1994; Fishwild et al., 1996; Mendez et al., 1997; Popov et al., 1996 and 1999). The introduction of multiple VH and VL genes allowed mice to express different combinations of V-region pairs, which produced extensive antibody repertoires. PCR technology and subcloning procedures could then be used to add the desired Cregion gene, which upon re-expression allowed bulk production of monoclonal human antibodies (reviewed in Maynard and Georgiou, 2000). For this reason, the addition of C-region genes to allow switching from IgM to IgG was regarded as less pressing, particularly in mice that rearranged and expressed Cm with different VH genes diversified by hypermutation (Wagner et al., 1996). Figure 34.1 illustrates the C gene make-up of the different IgH transloci, all of which accommodate a Cm gene that can be initially expressed as surface IgM receptor and subsequently in secreted form. On the larger IgH loci, Cd is included, probably as a matter of convenience rather than to put emphasis on the derivation of IgD antibodies, because Cm and Cd are closely linked and were present on IgH YACs from libraries screened with JH and Cm probes (Choi et al., 1993; Green et al., 1994; Wagner et al., 1996; Brüggemann and
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Figure 34.1 Human IgH C genes and transcriptional enhancers on translocus constructs. C genes and enhancer regions have either been retained in the natural configuration (Cm, Cd, Em and Ed-g3) or have been added 3¢ proximal of another C gene by subcloning procedures (g1, g2 and E3¢a).
Neuberger, 1996). Nevertheless, there are several important considerations regarding the addition of C genes. Switching to IgG allows the cell to enter the recirculating B-cell pool and become established as a long-lived memory B cell, with an increased chance of accumulating high levels of somatic mutation. Furthermore, the creation of designer mouse strains, which express particular Cg genes, would provide the desirable effector functions for each antibody produced. For example, IgG1 and IgG3 antibodies have the ability to fix complement and can initiate extensive hemolytic activity. The strong (IgG1 and IgG3) and weak (IgG2 and IgG4) Fc receptor (FcgR I, II, and III) binding is also useful to initiate or avoid macrophage or NK cell interaction. By the addition of one Cg gene immediately 3¢ of Cd, human IgH loci on YACs have been engineered with predefined isotype activity to produce IgG1 for the destruction of target cells, and IgG2 or IgG4 to essentially block or compete binding without initiating cell lysis (Mendez et al., 1997; Davis et al., 1999). Thus, the selective addition of one particular Cg gene and derivation of mice producing either IgG1, IgG2, or IgG4 antibody repertoires provided a choice
of isotype combining specificity with desired effector functions (Green, 1999). However, threading C genes in artificially close proximity disregards the possible importance of intervening sequences, which may provide as yet unknown control functions critical in securing good expression levels. In the human IgH locus, the Cd–Cg3 interval is such a region, rich with transcription factor binding motifs and with lymphocyte-specific enhancer and silencer activity (Mundt et al., 2001). To gain comprehensive information about IgH locus activation and gene usage, it will be interesting to see how C genes assembled in authentic configuration are expressed. Good expression levels may depend on the presence of one or more transcriptional Ig enhancers. All IgH and IgL chain YACs contain at least one but usually two enhancers; 5¢ of the first C gene and 3¢ of the last C gene. For IgL loci, with their relatively compact structure, transcriptional enhancers are proximal to both Ck and Cl, so that no additional enhancers need be added to these YACs. For the (human) IgH locus, the activity of four enhancers has been described: EDQ52, Em, Ed-g3, and E3¢a (with multiple
34. Human Monoclonal Antibodies from Translocus Mice
sites) (reviewed in Magor et al., 1999; Mundt et al., 2001). Human Em is present on all IgH transloci (see Figure 34.1). As a second control region, the rat or mouse enhancer downstream of the last C gene, E3¢a, has been added 3¢ of Cg (Taylor et al., 1994; Mendez et al., 1997). However, the use of E3¢a, later termed HS1,2, may at best only confer reduced enhancer activity, because it is part of an enhancer complex of at least four transcriptional activators on a 40-kb region (Arulampalam et al., 1997 and refs therein). The necessity to include multiple enhancers on incomplete transloci is debatable, because the presence of Em with or without a second enhancer secures similar expression levels (Brüggemann and Neuberger, 1996). Human, mouse, and rat enhancer sequences have been used to drive human IgH and IgL chain transcription, but there is a lack of information about their individual needs and how their functions compare. For YAC introduction into the mouse germline, three different strategies have been used: DNA purification and microinjection into fertilized oocytes (Fishwild et al., 1996); co-lipofection of embryonic stem (ES) cells with a mixture of size-fractionated YAC DNA in agarose and a selectable marker gene (Choi et al., 1993); and fusion of yeast protoplasts with ES cells (Davies et al., 1993 and 1996). Although very efficient methods for YAC purification and use for microinjection have been described (Schedl et al., 1993), the problem of handling and shearing of large DNA molecules makes it difficult to integrate one complete copy of a human Ig locus (Taylor et al., 1992; Fishwild et al., 1993). Improvements to allow complex (human Ig) loci in their intact form to be transferred into the mouse genome came with the use and manipulation of ES cells (Hogan et al., 1994). Lipofection, or lipid-mediated DNA transfer, of ES cells largely avoids the handling of naked DNA. In addition, large linear molecules on YACs or BACs do not necessarily have to be retrofitted with a selectable marker gene if a co-transfection approach is used (Choi et al., 1993). Unfortunately, a frequently encountered problem is that only a portion of the introduced locus is integrated into the host genome, and the transfectants obtained must be carefully analyzed for the presence and integrity of the introduced genes. Protoplast fusion, as an alternative approach for the transfer of YACs into ES cells, has previously been exploited for YAC integration into other, differentiated, mammalian cells (Pachnis et al., 1990; Pavan et al., 1990). The preparation of YACcontaining yeast spheroplasts does not involve DNA handling or gel separation but the YAC to be transferred must be retrofitted with a selectable marker gene. The method used for protoplast fusion with ES cells is similar to that employed in the generation of hybridomas and has been described in detail (Davies et al., 1996). Usually, only a few clones are obtained, but the approach allows a reliable integration of complete single copy YACs into the mouse genome (Mendez et al., 1997; Nicholson et al., 1999).
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Transfer of Chromosome Fragments Although the introduction of minigene constructs and YACs relies on the integration of the translocus into a mouse chromosome, transferred chromosome fragments can be maintained in the cell as distinct minichromosome(s) (Figure 34.2). Individual chromosomes or their fragments, tagged by transfection with a selectable marker gene (neomycin, hygromycin, or puromycin), have been transferred by the microcell-mediated fusion of human fibroblasts with somatic cell lines such as A9 (mouse fibroblasts), CHO (Chinese hamster ovary cell line), or DT40 (ALV induced chicken tumour line) (Fournier and Ruddle, 1977; Koi et al., 1989; Shinohara et al., 2000). This has allowed the establishment of monochromosomal hybrid libraries with different chromosomal regions maintained under selection. Mouse A9 clones with defined chromosomic-regions, including the human IgH, Igk, and Igl loci and their adjacent loci, which had been identified by chromosomal marker analysis, were introduced into ES cells by microcellmediated fusion. Viable chimeric mice were then derived from these ES cells after injection into eight cell embryos (Tomizuka et al., 1997). The size of the transchromosomal regions obtained varies and may perhaps be only a few percent of that of the chromosome they are derived from but, despite the problem with stability, apparently near-intact human chromosomes have also been identified and transferred (Tomizuka et al., 1997). Apart from the derivation of chimeric mice, important considerations for the use of the chromosome technology are that transchromosomes can be transmitted and maintained in the mouse. Germline
Figure 34.2 Two-color FISH analysis of tail fibroblasts prepared from a 4-week old double chromosomic mouse. Probes identified the human H chain locus (red) and k L chain locus (green). Permission for display of the figure (Tomizuka et al., 2000) was kindly provided by Prof. Ishida and Proc. Natl. Acad. Sci. USA. See color insert.
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transmission was first obtained with a human chromosome 2–derived fragment accommodating the Igk locus (Tomizuka et al., 1997). To secure germline transmission of the IgH locus, a fragment of ~one-fifth of human chromosome 14 was derived to establish a mouse line maintaining an IgH and Igk minichromosome that could be bred with a strain in which the endogenous IgH and Igk L chain loci had been silenced by gene targeting (Tomizuka et al., 2000). To overcome the problem that some transferred chromosomal regions were unstable and not maintained in all mouse cells (somatic mosaicism) and that perhaps larger human artificial chromosomes (HACs) introduced prevented germline transmission, efforts have concentrated on sitedirected chromosome truncation (Kuroiwa et al., 1998 and 2000). The derivation of minichromosomes or HACs has been carried out in recombination-proficient DT40 cells by the site-specific integration of telomeric regions and/or a loxP sequence. The human Igk and CD8A locus are both centromere-proximal, but are on opposite sites of the centromere. The gene order has been identified as CD8A (2p11.2)-centromere-Ck-Jks-Vks (2q11.2) (http://www.ensembl.org/Homo_sapiens). A telomeric sequence linked to the puromycin selection marker was inserted into CD8A, and a loxP sequence next to the hygromycin gene was inserted downstream of the Vk gene cluster. Fusion of these cells with cells that carry a minichromosome with a loxP site in the telomere proximal RNR2 locus (14p12) at the opposite end of the IgH locus (14q32.33) and subsequent transfection with a vector encoding Cre recombinase resulted in a human Igk locus on a ~5 Mb minichromosome (or HAC) comprising an authentic centromere and one authentic and one artificial telomeric region (Kuroiwa et al., 2000). For the human Igl locus, to stabilize transfer and maintenance of this locus on chromosome 22, telomeric and loxP sequences were site-specifically introduced into adjacent loci (Kuroiwa et al., 1998 and 2000). This resulted in a HAC with loxP integration in the HCF2 locus (22q11.22), upstream of the unmodified Igl locus (22q11.23), followed downstream by telomere insertion in the LIF locus (22q12.2) and chromosome truncation. Electroporation of targeting constructs using DT40 cells achieved site-specific integration in ~10% of the clones. Cells harboring site-specific alterations 5¢ and 3¢ of the Igl locus allowed successful fusion with cells that carried a HAC accommodating the IgH locus and loxPmodified RNR2 locus, which after transfection with Cre produced an Igl HAC of ~10 Mb. The strategy demonstrated the successful recombination of two nonhomologous chromosome fragments with added telomere proximal sequences and generated individual stable minichromosomes with a human IgH, Igk, or Igl locus. Although the HACs have been introduced individually into ES cells, a combination of IgH and IgL chain loci has been successfully expressed in mice (Kuroiwa et al., 2000). Furthermore, recent results showed
that Cre-loxP–mediated reciprocal translocation can be used for the joint assembly of several Ig loci to allow the production of transchromosomic calves producing Ig from one HAC accommodating both the human IgH and Igl locus (Kuroiwa et al., 2002).
Replacement of Mouse C Genes In an approach to make use of a mouse Ig locus as the driving force in directing human antibody expression and maturation, mouse C genes were targeted and substituted. This allowed the site-specific integration of human Ck and Cg1 replacing mouse Ck and Cg1 or Cg2a (Zou et al., 1993a; Zou et al., 1994; Pluschke et al., 1998). For homologous integration, two strategies were used. In constructs applied for conventional gene targeting procedures human Ck, adjacent to a selectable marker gene, was flanked by mouse homology sequences and, separately, human g1 next to a marker gene was flanked by mouse g2a sequences. This produced animals that rearranged and expressed chimeric Igk and IgG1 antibodies with human C regions (Zou et al., 1993a; Pluschke et al., 1998). In the strategy used by Zou et al. (1993a) mouse Cg1, excluding the transmembrane exons, was replaced by human Cg1. The use of a construct in which the homology region and selectable marker gene were flanked by loxP sequences allowed their removal by Cre-mediated deletion. This “cleanly” inserted a human g1 C gene into the previous location of mouse g1. This resulted in a mouse producing chimeric human IgG1 in serum at levels similar to those of mouse IgG1 in normal animals. Immunization of these animals mounted a normal immune response and produced a diverse repertoire of chimeric IgH and IgL chains (Zou et al., 1994). The success in targeted replacement of mouse for human genes favorably supports more extensive alterations that may allow the expression of fully human antibody repertoires controlled by the endogenous Ig loci.
THE MOUSE STRAINS The initial experiments showed that introduced human Ig genes in germline configuration could be rearranged and expressed in the mouse. However, although human antibodies were clearly detectable in mouse serum, tested by ELISA, their concentration was relatively low compared to the level of endogenous Ig (summarized in Brüggemann and Neuberger, 1996). Similarly, only a few percent of human IgH+ B cells are normally found in the transgenic mice, whereas the majority of lymphocytes express mouse Ig. Nevertheless, human IgM titers of up to 100 mg/ml have been achieved in some transgenic lines carrying microinjected minigene constructs (Brüggemann et al., 1989; Brüggemann and Neuberger, 1991). However, such rela-
34. Human Monoclonal Antibodies from Translocus Mice
tively high H-chain expression levels have not been shown for IgH YACs introduced via the ES cell route. Furthermore, a comparison of human Ig expression in normal translocus mice indicated that IgL chain loci may be better expressed than IgH chain loci, and that perhaps larger or near complete Ig loci can very efficiently compete with the endogenous mouse locus (Popov et al., 1999). The findings that human Ig loci are expressed in the mouse coincided with the development of the gene targeting technology and the derivation of knockout mice (Capecchi, 1989). Silencing of the mouse Ig loci, first achieved by Rajewsky and co-workers for the H-chain locus and later for the k locus (Kitamura et al., 1991; Zou et al., 1993), proved invaluable to secure human antibody expression without mouse H and k L chain interference. Indeed, the currently used mouse strains that express fully human antibody repertoires have been produced by the integration of human H and L chain (k and/or l) YACs and/or HACs and crossing with animals in which the endogenous H and k L chain loci have been silenced by gene targeting (Lonberg et al., 1994; Mendez et al., 1997; Nicholson et al., 1999; Tomizuka et al., 2000). Several websites illustrate the generation and use of these mice: http://www.babraham.ac.uk, http://www.abgenix.com, http://www.medarex.com and http://www.tcmouse. com.
Locus Stability The integration of two or all three human Ig loci on minigene constructs or YACs, and the silencing of the equivalent endogenous mouse loci, has been achieved by breeding individual transgenic and knockout mice with each other. This established heterozygous and homozygous mouse strains carrying a human IgH, human Igk, and/or human Igl translocus integrated into a chromosome and bred to homozygosity with strains in which the endogenous IgH and Igk loci had been silenced by gene targeting (Lonberg et al., 1994; Mendez et al., 1997; Nicholson et al., 1999). As homozygosity for a combination of all five features (human IgH, human Igk, human Igl, mouse IgH KO, mouse Igk KO) has been readily obtained, one can assume that these loci, transferred by DNA injection into oocytes or YAC integration in ES cells, largely integrate in a random fashion and not at preferred sites (Nicholson et al., 1999). In addition, breeding from homozygous stock results in 100% homozygous male and female offspring, thus implying that no undesired integration—for example, into a sex chromosome—took place. However, although no reports show the actual chromosomal integration sites of minigene constructs or YACs, the maintenance of introduced HACs, as separate single units, has been well documented (Tomizuka et al., 2000). Individual HACs can be maintained in ES cells under selection but their germline transmission and maintenance in somatic cells is significantly reduced
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(Shinohara et al., 2000). Breeding analyses established a variable transmission efficiency that reached an impressive 38% for one particular HAC, compared to the ideal 50% transmission rate of conventional genes in heterozygous configuration. In these mice, the HAC is maintained in a majority of the somatic cells but, as transmission and stability of individual HACs is well below the 50% threshold, it seems to be a significant challenge to breed and establish transchromosomal mouse lines carrying at least one IgH and one IgL chain HAC in the majority of their cells. It appears that human HACs frequently rearrange or segregate inaccurately in mouse ES cells. A reason for this instability appears to be the imprecise separation at mitosis due to poor centromere function (Shen et al., 1997). However, this problem has been overcome when a 4-Mb minichromosome acquired mouse centromeric sequences and was stably maintained in culture. A suggestion by Shen and collaborators was that the addition of minor satellite DNA, a candidate for centromeric DNA, could be used to target HACs transferred into recombination-proficient DT40 cells; this may ensure stability and perhaps the maintenance of multiple HACs per cell in near homozygous configuration. Unfortunately, there are as yet no reports on the use of this strategy for securing the stability of Ig HACs. Both the YAC and HAC technology for the integration and expression of large human loci in the mouse have advantages and disadvantages. The advantage of transferring YACs is that the integration into a mouse chromosome secures perfect stability and transmission and the gene content of the YAC is essentially known from sequence analysis. A disadvantage is that current YACs are hardly larger than 1 Mb, which means that they cannot accommodate complete Ig loci. The transfer of tagged human chromosomes or their fragments is possible but the generation of HACs may be more advantageous, because it allows the maintenance of clearly defined Mb size loci and the removal of regions that interfere with their stability. Nevertheless, maintaining HACs as separate minichromosomes has an essential drawback; the somatic mosaicism results in variable transmission rates that restrict the level of available B cells with the desired multiple features for which every resulting mouse still has to be analyzed. Perhaps a compromise can be found in a combination of the technologies, which every would then allow chromosomal integration and transmission stability of ≥Mb size loci.
Antibody Expression Human antibody expression in mice with fully functional endogenous Ig loci is relatively inefficient since only a low level of largely chimeric Ig is produced. At present, there is little interest in generating human antibodies from such “normal” animals. Nevertheless, such translocus mice could become useful for the derivation of human IgH and IgL
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chain libraries after antigen challenge and in vitro combinatorial selection using phage or ribosome display technology (in vivo/in vitro technologies are compared in Neuberger and Brüggemann, 1997). Although human IgH and IgL chains can be expressed from minigene constructs containing one or a few of each of the V, (D), J, and C genes and one C proximal enhancer, it became clear that better expression and repertoire formation is attained from the integration of larger Ig regions. Here I would like to focus on the four laboratories that have generated four- and fivefeature human antibody expressing mouse strains, or xeno mice, with large integrated IgH, Igk, and/or Igl transloci in an endogenous mouse IgH and Igk knockout background (Lonberg et al., 1994; Mendez et al., 1997; Nicholson et al., 1999; Tomizuka et al., 2000). Human antibody titers for plasmid-based, YAC-based, and chromosomal transloci are listed in Table 34.1. Multifeature mice have been obtained by crossing animals with defined phenotypes created by translocus insertion and, separately, by the silencing of endogenous mouse genes. For human antibody expression in a normal mouse background, this identified a generally low but variable concentration of Ig with human H chain (<50 mg/ml) and a much greater but also variable level of Ig with human L chains (15– 400 mg/ml). In these mice, both human H and human L chains are predominantly associated with mouse chains and form almost exclusively chimeric antibodies. In the knockout background, the level of human Ig reaches in many cases a few 100 mg/ml, which suggests that this expression level is by and large determined by the transferred Ig core region. Improvements in Ig expression can be seen in YAC-based mouse strains, in which human antibody levels reached or even exceeded 1,000 mg/ml. This suggested that larger Ig transloci with more gene segments are potentially better expressed. However, Ig expression in the transchromosomal mice carrying complete H and L chain loci (Tomizuka et al., 1997 and 2000) is not further increased and is only about half the levels of what is found in IgH and Igl YAC mice (Mendez et al., 1997; Popov et al., 1999). A reason for this could be the mitotic instability of the transchromosome and the degree of chimerism of the mice. The fact that the transferred HAC has to be maintained as a separate human chromosome in the mouse cells also raises the possibility that recognition by the cellular machinery is suboptimal, which could affect chromatin re-modeling and locus accessibility (Jenuwein and Allis, 2001). In most cases, antibody titers refer to total human Ig (or IgM or IgG) levels in the mouse. This may be misleading because of the presence of mouse Igl associated with human H chain. Depending on the mouse strain, background, and immunization procedure, a large proportion of the “human” antibodies can contain mouse l L chain and thus be in chimeric configuration (Fishwild et al., 1996; Mendez et al., 1997; Nicholson et al., 1999; Magadán et al., 2002). Although the mouse Igl locus
has been recently silenced by gene targeting (Zou et al., 2003), human Ig mice with all three endogenous mouse Ig loci rendered nonfunctional by gene targeting—and thus potentially not expressing any endogenous mouse Ig—have not yet been obtained by cross-breeding. IgM from translocus mice, where human m is associated with human k or human l L chain, is secreted as a pentamer and almost indistinguishable from the IgM produced in humans. [The mouse J chain, important for IgM assembly, is still present in those animals but could be easily replaced by the human J chain since mice with inactivated endogenous J chain have already been produced (Erlandsson et al., 1998).] The high avidity but generally low affinity of IgM can be advantageous for some applications (Okada and Okada, 1999), but early on there was a desire to obtain mice that express human IgG. The initial experiments adding a Cg1 gene to allow switching from IgM to the desired isotype proved successful and established genomic recombination between the transgene m and g1 switch regions (Taylor et al., 1992 and 1994). Nevertheless, in the H and k L chain transgenic mice, human Ig levels were disappointingly low, albeit detectable by ELISA. The human IgM and IgG1 concentration in serum improved significantly when the animals were crossed with endogenous H and k L chain knockout mice. The addition of VH and Vk genes further increased human antibody levels (Lonberg et al., 1994; Taylor et al., 1994). Perhaps the most notable advantages of an introduced human IgH locus on a 1-Mb YAC have been described by Mendez et al. (1997): good expression levels, extensive VH gene diversity, and switching to the desired isotype—here, IgG2. Although switching from Cm to Cg is achieved, the serum levels of human IgG are lower than those of human IgM and do not represent the ~1 : 10 ratio (0.7–1.7 mg/ml IgM: 9.5–12.5 mg/ml IgG with 34–87% IgG1, 5–56% IgG2, 0.5–12% IgG3, 7–12% IgG4) found in human serum (Frazer and Capra, 1999). The presence of additional C genes and switch recombination may also allow a more efficient selection of highaffinity antibodies diversified by hypermutation. Indeed, there appears to be an advantage of transloci that allow isotype switching. Multiple immunizations of IgM translocus mice appear to be much less effective than consecutive immunizations of IgG producers, which may continue to diversify their repertoire much more efficiently. In translocus mice, with or without Cg addition, the hypermutation rates of VH genes linked to Cm can be relatively poor (Wagner et al., 1994; Lonberg and Huszar, 1995; Nicholson et al., 1999). Nevertheless, upon VDJ joining, combinatorial and junctional diversity create extensive modifications of CDR3. The length of the CDR3 regions, seven to nineteen amino acids, is comparable to those identified in humans and considerably longer than those found in the mouse (Mendez et al., 1997; Nicholson et al., 1999). The low hypermutation levels of the m H chain of perhaps 0.1% could be increased to almost 2% upon IgG expression, even when human H
34. Human Monoclonal Antibodies from Translocus Mice
chains were expressed from relatively small loci of less than 100 kb (Lonberg and Huszar, 1995). The creation of translocus mice by transfer of essentially the same H chain locus on a YAC containing five V genes, Ds, Js, Cm, and Cd, produced two separately derived strains that showed similar expression levels of human IgM. Only one strain readily somatically diversified its VH genes, whereas VH sequences from the other strain showed few nucleotide changes (Wagner et al., 1996; Nicholson et al., 1999). A reason for this could be the difference in the integration site, which may also influence the enhancer or insulator activity of a control region downstream of Cd on this YAC (Wagner et al., 1996; Mundt et al., 2001). Expression from a large human IgH locus with about thirty-four functional VH genes showed the utilisation of eleven of those genes, ten different D segments, and four different JH segments linked to either Cm or Cg2 (Mendez et al., 1997; Davis et al., 1999), with the rearranged VH genes originating from all parts of the V region cluster. Nevertheless VH genes from families VH3 and VH4 were preferentially used. This is similar to the V gene choice in adult humans, which is proportional to family size (Mendez et al., 1997). Sequence analysis of five VH4-31 and three VH4-61 genes spliced to g2 H chains identified two to ten nucleotide changes, which indicated that B lymphocytes expressing human IgG2 were able to participate normally in an immune response. Diverse usage and hypermutation was also found for human Igk and Igl L chain genes introduced on large YACs. Rearranged k L chains utilized all J segments and different V genes from all parts, distal and proximal, of the cluster (Fishwild et al., 1996; Mendez et al., 1997; Xian et al., 1998; Nicholson et al., 1999). Extensive usage was also found for Vl genes, located on different parts of the introduced 380-kb locus, which rearranged and expressed using any of the three functional J-C segments (Popov et al., 1999). The use of human Vl segments in productive and nonproductive rearrangements in translocus mice were similar to those found in the human peripheral blood repertoire (Ignatovich et al., 1999). Illustrated in Table 34.1, human Igk and Ig l L chain loci of several hundred kb in size, most of which contain a large number of V genes, produce expression levels close to or around 1 mg/ml (Zou et al., 1996; Mendez et al., 1997; Xian et al., 1998; Nicholson et al., 1999; Popov et al., 1999). However, in one strain carrying a 300-kb human Igk translocus with two functional Vk genes, the expression levels of ~800 mg/ml are similar to those in mice carrying the same L chain locus, only with more V genes added (Xian et al., 1998). Crossing mice carrying highly productive L chain loci with human H chain mice notably improves H chain expression with a significant increase in Ig levels (Nicholson et al., 1999). In summary, this suggests that multiple parameters are important to secure high expression levels, either determined by the transgene make-up, such as locus core region and V gene content, or by the com-
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bination of other genes (or transgenes) provided by the host. For both H and L chain loci, the efficacy of somatic hypermutation appears to depend on locus size and, for the H chain, also on the presence of C genes that allow isotype switching. In general, there seems to be a broad, but not perfect, correlation with size, in that larger loci are better expressed. B-cell development in a translocus mouse is therefore critically dependent on the makeup of the introduced locus, and repertoire formation upon immunization can be seen as a reliable quality control parameter. In a normal mouse, about 2 days after immunization, some B cells differentiate into IgM+ plasma cells, generally of low affinity, whereas after another 2 to 3 days antigen-specific B cells are detectable in the primary follicles of the spleen, where they proliferate to form germinal centres. These events create essentially two proliferative populations of activated B cells: one leading to the primary antibody response and the other to maturation and memory. It seems plausible that human IgH translocus mice are less proficient in this maturation process, which may be particularly inefficient in mice producing only IgM. Nevertheless, apart from junctional diversity, mutation of the k L chain is achieved in the strains even without H chain mutation or switching and thus supports the notion that somatic diversification of H and L chains is separately controlled (Nicholson et al., 1999). The compactness of the k L chain locus with Vs, Js, Ck, and enhancers of less than 100-kb allows relatively small transloci to work very efficiently (Lonberg et al., 1994). However, when the natural spacing of Vks to J-Ck was discarded on an artificially close minigene construct, expression levels and somatic hypermutation were quite low (Xian et al., 1998). This suggests caution and may imply that the close addition of, for example, one member of each VH gene family may not improve the repertoire or diversity of human H chains from expressed minigene constructs. It is possible that introduced large Ig regions in authentic configuration are activated and processed in a similar manner to the endogenous loci. This may be seen in human Igl translocus mice, where high expression and somatic hypermutation is well achieved, even in a normal mouse background in competition with endogenous L chain (Popov et al., 1999). A likely reason for this is that the YAC accommodating a large part of the Igl locus in germline configuration, with eighteen V genes, all seven J-Cs, and enhancers, contains all necessary control elements and also the added value that J-C proximal Vl genes are preferentially rearranged and expressed in humans. The human Ig loci are expressed in a background in which the endogenous mouse H and k L chain loci have been rendered nonfunctional. Mice with a silenced Igk locus express only Igl L chain antibodies, but their B-cell level is largely retained. However, the silencing of the H chain locus, or both H and k L chain KO, leads to a block in B-
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Figure 34.3 Flow cytometry analysis of bone marrow and splenic lymphocytes from 5-feature mice (Nicholson et al., 1999) and normal mice. Staining was carried out with antibodies recognising specific cell-surface markers which define B-cell development: B220 (pan B-cell), c-kit and CD43 (both, pro and pre B-I), CD25 (pre B-II), and IgM, Igk and Igl on immature and mature B-cells.
cell development. This block can be overcome by the introduction of an IgH translocus, which kick-starts B-cell development and leads to (human) antibody production. This allowed the B-cell recovery illustrated in Figure 34.3 by flow cytometric analysis of cells from a five-feature mouse (human IgH, k, and l translocus in a background with silenced endogenous H and k locus) stained with antibodies against B-cell surface markers that identify pre-B, immature, and mature B-cell populations in bone marrow and spleen (Nicholson et al., 1999). In a mouse KO background, transfer and expression of human IgH or human IgH and IgL transloci containing many
gene segments leads to a partial recovery of the B-cell pool, usually well below 50% (Taylor et al., 1993; Green et al., 1994; Lonberg et al., 1994; Wagner et al., 1994a and 1996; Lonberg and Huszar, 1995; Fishwild et al., 1996; Nicholson et al., 1999). Comparing the size, gene content, and efficiency of single copy YAC transloci (plasmid-based miniloci are difficult to judge because of variable transgene copy numbers) indicated that larger loci are better expressed. This has been confirmed by Mendez et al. (1997), who found that in mice carrying a ~1 Mb human IgH locus in near-authentic configuration, B-cell recovery could reach over 60% (see also Figure 34.3B). Indeed, half to near normal B-cell
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34. Human Monoclonal Antibodies from Translocus Mice
numbers facilitate high expression of human H and L chains and may also significantly reduce any remaining mouse Igl+ B cells—which in four-feature mice are 15 to 28% compared to 48 to 78% of human k expressers (Mendez et al., 1997; Xian et al., 1998; Green, 1999; Nicholson et al., 1999). Interestingly, the introduction of a human Igl locus bred into mice expressing already fully human antibodies increased human IgM levels considerably and reduced the levels of mouse Igl+ B cells to below 5% (Nicholson et al., 1999). Nevertheless, in all currently used translocus strains, chimeric human antibodies have been identified to various degrees. These are, on the one hand, human H chains pairing with mouse l L chains and, separately, chimeric human–mouse H chains. Whereas mouse l can contribute to about half of the human antibodies produced in some strains (Magadán et al., 2002), H chains with a rearranged human VDJ region associated with a mouse C gene appear to be relatively rare and are seen as a consequence of transswitch recombination (Taylor et al., 1994; Lonberg et al., 1994; Lonberg and Huszar, 1995). In these mice, the endogenous H chain locus has been silenced by the deletion of the JH segments, which prevents endogenous DNA rearrangement and the use of a mouse VH. A disadvantage of the various H chain silencing approaches is that V, D, and in some cases J and C genes, are still present and allow their use in trans-switching, trans-splicing, and also translocation events (Brüggemann and Taussig, 1997 and refs. therein).
Immune Responses and Affinity of Fully Human Antibodies Mice expressing human antibodies are immunized in much the same way as conventional mice, but a comparison of immune responses showed that human antibody titers are quite reduced compared to normal nonmanipulated animals (Lonberg et al., 1994; Wagner et al., 1994; Jakobovitz et al., 1995; Magadán et al., 2002). Despite the relatively low titers, mice carrying human Ig loci are capable of mounting antibody responses to a wide range of antigens. Similar to normal mice, increased levels of specific antibodies are visible 2 to 3 weeks after primary immunization and can be further increased by secondary immunization. Serum analysis by ELISA revealed that some antigen-specific antibodies were fully human (human H and L chain) whereas others were chimeric (human and mouse H or L chain). The formation of mixed molecules or polypeptides led to disappointment, which was encountered when some immunizations produced a substantial number of chimeric human antibodies with mouse l L chain (Russell et al., 2000; Magadán et al., 2002). Nevertheless, antigen-specific fully human antibodies were produced and hybridoma technology (background and methods provided by King, 1998) allowed the establishment of a large number of human monoclonal
TABLE 34.2 Fully human monoclonal antibodies H chain
L chain
tetanus toxin progesteron
m m
k k
IGF human IgE PLAP S. pneumonia human tumour cells PLAP human IgM,k human tumour cells progesteron IGF human IgE digoxin human CD4
m m m m m
k k k k k
m m m
l l l
m m m g1 g1
l l l k k
human IL8
g2
k
human EGFR
g2
k
human TNFa
g2
k
IL6 L-selectin GROa CD147 CD4 GCSF
g& g& g& g& g& g&
k k k k k k
Antigen
affinity
Reference Green et al., 1994 Nicholson et al., 1999; He et al., 1999 Nicholson et al., 1999 Nicholson et al., 1999 Nicholson et al., 1999 Russell et al., 2000 Magadán et al., 2002
0.5 nM
a
Magadán et al., 2002 Magadán et al., 2002 a a a
2.5–22 nM 11 nM–27 pM
Ball et al., 1999 Lonberg et al., 1994; Fishwild et al., 1996 0.2–0.9 nM Mendez et al., 1997; Green, 1999 0.8 nM–30 pM Mendez et al., 1997; Green, 1999 0.2–0.8 nM Mendez et al., 1997; Green, 1999 Green, 1999 Green, 1999 Green, 1999 Green, 1999 32–77 pM Ishida et al., 2002 0.2–0.3 pM Ishida et al., 2002
a kindly provided by Mike Taussig, The Babraham Institute, Cambridge UK; & isotype not defined.
antibodies from the described translocus strains. Table 34.2 summarizes their features and chain composition. Expression levels were quite reasonable, from at least a few mg/ml in conventional tissue culture plates, up to 400 mg/l in serum-free, fed-spinner cultures (Ball et al., 1999; Nicholson et al., 1999; Davis et al., 1999; Green, 1999). It appears that the monoclonal antibodies exhibit a good usage of the different V, (D), J, and C genes and, as discussed earlier, that N sequences, in addition to junctional diversity and hypermutation, allow the creation of extensive repertoires. For example, antibodies specific for the epidermal growth factor receptor (EGFR), which may be overexpressed on many types of tumors, showed the preferential use of the closely related VH genes 4-31 or 4-61(8/11) and Vk 018 (8/11) in combination with different D and J segments (Davis et al., 1999). The V genes were diversified by somatic mutation, but most strikingly, as a result, all eight VH genes bore an aspartate residue in CDR1 at position 33. Detailed analysis of the five-feature mice described by
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Nicholson et al. (1999) revealed that the proportion of human antibodies with human k or human l L chain appears to be antigen-driven, but that these mice do not produce significant levels of chimeric human antibodies with mouse l L chain (Magadán et al., 2002; M. Taussig personal communication, and see Table 34.2). Immunization of transchromomal mice resulted in antibody responses that included all Ig subclasses (Yoshida et al., 1999; Ishida et al., 2002). Nevertheless, a problem was encountered in obtaining hybridomas. A reason for this was the instability of the Igk locus–bearing chromosome fragment. However, this inefficiency could be overcome by cross-breeding the transchromosomal mice with mice carrying an Igk YAC (Ishida et al., 2002; Fishwild et al., 1996). The translocus mouse strains allowed the creation of diverse human antibody repertoires of high affinity and desired specificity. But despite this success, their immune responses, Ig levels, and antibody diversity are not as refined as those of a normal mouse. Nevertheless, a number of the human monoclonal antibodies generated appear to have an exceptional high affinity in the picomolar range. Extensive mutation in VH and VL have been described, but it seems that the mouse strains with larger and indeed complete Ig loci (Ishida et al., 2002) may produce a more diverse repertoire and, that simple choice and selection allows the identification of superior antigen-binders. However, despite the success in producing high-affinity, fully human antibodies, and although several human monoclonal antibodies—such as anti-IL8 to treat psoriasis and anti-EGFR to inhibit tumor growth—have shown impressive results in clinical trials (Davis et al., 1999; Yang et al., 2001), none has yet been approved by the FDA (Gura, 2002).
CONCLUSION Human antibodies represent an invaluable and growing class of biotherapeutics, and their production and use will be expanded rapidly in the next few years. From recent advances, it seems that several new technologies will be extensively used to produce high-affinity, fully human antibodies. Whole Ig loci will be routinely transferred into different species of animals, with their own antibody loci silenced by gene targeting or suppressed by selected breeding. The animals will be immunized and essentially provide highly specific, yet diverse human antibody repertoires. This may already be possible in cattle with introduced human Ig loci (Kuroiwa et al., 2002) and may soon produce not just monoclonal antibodies but large amounts of polyclonal Ig. Besides producing high-affinity antibodies through recurring somatic hypermutation, extensive diversity will also be created in some translocus animals by gene conversion (evolutionary processes reviewed in Diaz and Flajnik, 1998). Another approach may use targeted integration into a mouse
Ig locus to replace not just endogenous C genes by human equivalents, but perhaps substitute many of the V genes by the insertion of artificial chromosomes (strategies described by Houdebine, 2002). In other experiments, researcher will make use of highly recombinogenic cells, such as the chicken DT40 cell line, and in vitro selection (http://swallow.gsf.de/dt40.html; Harris et al., 2002). This strategy already allows the selection of antigen-specific antibodies with nanomolar affinities generated by iterative affinity maturation in cell lines that hypermutate their Ig genes constitutively in culture (Cumbers et al., 2002). The integration of human Ig regions into appropriate cells will permit the selection of spontaneous VH and VL mutants, as well as the generation of targeted modification by transfection with DNA or degenerate oligonucleotide mixtures. In yeast, polypeptides displayed on the cell surface have been modified by DNA shuffling, and antibody fragments with femtomolar antigen-binding affinity were obtained (Boder et al., 2000). Cells expressing novel, high-affinity antibodies can be easily selected or identified using flow cytometry and cloning procedures. Indeed mutator strains, which can recombine and/or hypermutate introduced reporter genes with high efficiency, may provide a useful alternative to human antibody production in transgenic animals.
Acknowledgments I thank Drs. Jennifer Smith for comments, Xiangang Zou for FACS data, Mike Taussig for unpublished information, and Isao Ishida for giving permission to use his FISH profile. The work in my laboratory is supported by the Babraham Institute.
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Index
Note: Boldface numbers indicate illustrations.
A AA4.1, B cell development and common lymphoid precursors (CLP), 106 abciximab, 541 accessibility hypothesis, V(D)J recombination and, 128 activation-induced deaminase (AID) affinity maturation and, 342 APOBEC-1 and, 313 B cell development and, 143 CH12F3–2 and, 313 class switch recombination (CSR) and, 289, 290, 307, 311, 313–314, 317, 319–320 deficiency of, 313 digestion circularization PCR (DC-PCR) and, 313 enhancement and inhibition of, 314 estrogen receptor (ER) (AID-ER) in, 313 function of, molecular mechanism for, 313–314, 317 high affinity antibody and, 342 hyper-IgM syndrome (HIGM) and, 313, 410–411 immune response of B cells and, 342 isolation, structure, role of, 313, 317 mismatch repair (MMR) and, 314 mutations and, 314 S region cleavage and, 314 somatic hypermutation (SHM) and, 307, 314, 319–320, 328, 330, 331–332 UNG and, 314 acute lymphoblastic leukemia (ALL), 149–150, 349–352, 350, 355–356 acute myeloid leukemia (AML), 293 adalimumab, 541 adaptive extrafollicular antibody response, 187 adaptive immune receptor origin, lower vertebrate immunoglobulin genes and, 427 adaptive immunity, in zebrafish (Danio rerio), 450, 452–457, 464 adaptive memory (See also memory and memory B cells), 254–255
Molecular Biology of B Cells
adhesion molecules gut lamina propria and, 236 mucosal B cells and, 234–235, 237 adult early B-lineage acute lymphoblastic leukemia, 355–356 ”affinity ceiling,” in vitro evolution, 505 affinity maturation activation-induced deaminase (AID), 342 affinity-based selection after GC reaction ends in, 344 ”affinity ceiling” in, in vitro evolution and, 505 in antibody production and response, 187, 492, 503–505, 504 autoimmune disorders and, 381 CD40 and, 342, 344 class switch recombination (CSR) and, 307 competition between cells and, 343–344 complementarity-determining regions (CDRs) in, 342–344 Fc (FcgRIIB) and complement receptors in, 279–280, 279 framework regions (FR) and, 342 germinal center (GC) in, 342–344 high affinity antibody and, 339–348 immune response of B cells and, 341–342 memory and memory B cells in, 248 phage display and, 522–524 ”selective advantage” concept in, 343 translocus mice and human Mab and, 557–558, 557 agammaglobulinemias (See also autoimmune disorders; immunodeficiency diseases), 148, 403–410 heavy chain gene mutations in, 409 B cell linker protein (BLNK) and, 410 CD79 in, 409, 410 g5/IGLL1 mutations in, 409, 409 Iga mutations in, 410 ITAM in, 410 X-linked (Bruton’s) (XLA), 403–409, 404 agglutinins, cold (CA), autoimmune disorders and, 388–390
563
agnathans (jawless vertebrates), lower vertebrate immunoglobulin genes and, 427–428 AID estrogen receptor (ER) (AID-ER), 313 alemtuzumab, 541, 541 alleles, immunoglobin l genes (IGL), human/mouse, 38 alleles of IgH, transcription of immunoglobulin genes and, 92 allelic exclusion, IgH, 133–134 immunoglobulin assembly and secretion, 262 allergic reactions/serum sickness, 511 allotyping, human IgH and, 15 a4 integrin, B cell development and, 103 a4b1, migration of B cells and, 208, 212, 214 a4b7 migration of B cells and, 212, 215 mucosal B cells and, 236, 237 a5b, migration of B cells and, 214 Alt, Frederick W., 61 amphibians and reptiles, class switching recombination (CSR) in, 422–423 AMuLv, V(D)J recombination and, 134 anaplastic plasmacytoma (PCT-A), 372–373 Von Andrian, Ulrich H., 203 anti-acetylcholine receptor (AchR), 392–393, 393 antibody-dependent cellular cytotoxicity (ADCC), 542 antibody-forming cells (AFC), 340, 345 Fc (FcgRIIB) and complement receptors in, 284 antibody production and response, 187–201, 155, 491–509 adaptive extrafollicular, 187 adaptive, stages of, 187–188, 188 ”affinity ceiling” in, in vitro evolution and, 505 affinity maturation and, 187, 492, 503–505, 504 allergic reactions/serum sickness, 511 antibody–secreting cells (ASCs) and, homing of, 213–215
Copyright 2004, Elsevier Science (USA). All rights reserved.
564 antibody production and response (continued) antigen entrapment and, 188 antigen recognition by, 491 anti-idiotopic antibodies in, 501 in artiodactyls (hoofed, grazing animals), 442–443, 442 B 1 cells in, 187 B cell proliferation and differentiation in, 188 B cell receptor (BCR) regulation of, 147–148, 147, 166, 188 backbone movements and binding of antigen in, 494–495, 495 BAFF and APRIL in, 191 Bcl and, 192 in birds/avians and chickens, 435–436, 435 binding energetics of antibody–antigen interactions in, 495–496 BLIMP 1 and, 190 bound and unbound antibodies in, 492–493 C region in, 491, 493–495 CCL3 and, 189 CD4 and, 189, 190, 194, 196 CD8 and, 194 CD11 and, 188, 190, 191, 197 CD19 and, 176 CD21 and, 188, 194, 195 CD23 and, 194 CD28 and, 196, 197 CD40 and, 189, 191, 194, 196, 197 CD54 and, 195 CD57 and, 194 CD86 and, 196 cFLIP and, 196 chimerization in, 511, 533–534, 540 clones of, sustained survival of, 197 commitment of activated B cells to follicular/extrafollicular growth in, 189–190, 190 complementarity-determining regions (CDR) in, 491, 494, 496–499, 504 conformational flexibility in antigen recognition and, 493–495 cross-reactivity in, 500–501, 502 CTLA 4 and, 196, 197 CXCL12 and, 189, 191 CXCL13 and, 189, 192, 195 CXCR4 and, 191 CXCR5 and, 194, 195 D1.3 and, 494–501 E5.2 and, 497–499 egg white lysozyme (xEL) studies in, 500–501 evolutionary studies in, 505 exponential growth of B cells in, 190–192 Fab and Fc domains in, 493–494 FCgIIR and, 195 follicular dendritic cells (FDC) and, 188, 191, 192, 194, 195 FvE5.2 and, 501 germinal centers (GC) and, 187, 189–197, 193 centroblasts and centrocytes in, 194 organization of, 192–194 primary follicles in, 192–193, 230
Index proliferation, hypermutation, selection in, 192–197 secondary follicles and, 193–194 T cells in centrocyte selection and maintenance of, 195–197 T cells in secondary follicles of, 194 growth and differentiation of CD4 T cells in, 190 heavy chain in, 491 HEL and anti-HEL in, 494–505, 497, 500 heteroclitic binding in, 500 HIV-1 and anti-HIV-1 peptide in, 494–495 how and where B cells encounter antigen in, 188–189 human anti-chimeric antibody (HACA) response in, 534 human anti-humanized antibody (HAHA) response and, 540 human anti-mouse antibodies (HAMA) response in, 511, 533, 534 humanization in, 511, 533–545 HyHEl complexes in, 494, 504 immunoglobulin assembly and secretion, 261 interacting surfaces/chemistry of antibody–antigen interface in, 492– 493 in lagomorphs (rabbits and hares), 437–440, 438 LFA 1 and, 195 light chain in, 491 lymphotoxin and, 195 major histocompatibility class (MHC) II in, 195 mapping of antibody–antigen interactions in, mutagenesis and, 496–499, 497, 498 marginal zone (MZ) B cells and, 188–189 model systems for research in, 501, 503 molecular mimicry in, 497, 501 molecular structure of, 491, 492 monoclonal (See monoclonal antibodies) mutagenesis and, 496–499 mutational accomomodation in antibody–antigen interface and, 499–500 mutations and, 503–505 naive or memory B cells in, 188, 203 pathogenic (See autoimmune disorders) Peyer’s patches and, 189, 203 phage display in, 513–514, 516–519, 517, 522 phases of, 187, 188 plasma cells and, 188, 197 plasmablasts and, 187, 189–191, 197, 253–254 primary cognate interaction of B cells with primed T cells in, 189–190 prokaryotic expression of fragments in, 514–515 somatic hypermutation (SHM) and, 503–505, 511 specificity of antigen recognition in, 500–501, 502 strength of antibody–antigen interaction in, 499
structure of, 491–505 T-dependent/-independent antigens and, 187, 188–189, 210 thermodynamics of antibody–antigen interactions in, 496 three routes of, 187 tonsillar B cells and, 189 TRAF and, 189, 196, 197 V region in, 491, 493–495 VCAM-1 and, 195 VLA4 and, 195 water molecules in, binding of, at interface, 493 XBP 1 and, 190 antibody–secreting cells (ASCs), migration of B cells and, homing of, 213–215 anti-DNA antibodies, autoimmune disorders and, 394 antigen entrapment, in antibody production and response, 188 antigen–presenting cells (APCs), 223–224, 226 antigen receptor genes, in V(D)J recombination, 61, 62–64, 63, 64 antigen receptor signaling regulation (CD19/CD22), 171–186 antigen processing with CD19 in, 176 B1 cells in, 155, 172 calcium mobilization in, 171, 177 CD19 in, 171–172 CD22 inhibition of intracellular signaling in, 177–178, 178 CD72 in, 180–181, 181 CD81 and, 172 CD100 and, 180–181 extracellular domain of CD22 and signaling in, 179–180 FcgRs in, 181, 182 Grb2 and, 177 Ig-like transcript (ILT) and, 182 inhibitory co-receptors on B cells and, 177–182, 178 inhibitory receptors CD22 and FcgRII with CD19, 176 ITIM and, 177, 182 leukocyte Ig-like receptor (LIR) and, 182 ligands of CD19 and, 173–174 Lyn and, 163, 174, 177 MAP kinase activation and, 171, 175, 178, 182 marginal zone (MZ) B cells in, 172–176 myeloid Ig-like receptor (MIR) and, 182 paired immunoglobulin-like receptors (PIR A-B) and, 181–182 PI3K and, 177 PLCg2 and, 175, 177, 182 Shc and, 177 SHIP and, 182 SHP-1 and, 177–178, 181 signal transduction by CD19 in, 173–176, 175 SLP-65 and, 177 STAT1 and, 175 Syk and, 163, 177 thymus-dependent antigens and, 173
565
Index thymus-independent antigens and CD19 in, 172–173 antigens affinity maturation and, 503–505, 504 B cell encounters with, 188–189 B cell receptor (BCR) and, 161 binding energetics of antibody–antigen interactions in, 495–496 conformational flexibility in recognition of, 493–495 cross-reactivity in, 500–501, 502 D1.3 and, 494–501 egg white lysozyme (xEL) studies in, 500–501 Fc (FcgRIIB) and complement receptors in, localization to FDC, co-receptor signaling vs., 281–285 FvE5.2 and, 501 heteroclitic binding in, 500 mapping of antibody–antigen interactions in, mutagenesis and, 496–499, 497, 498 memory and memory B cells in, lifespan and persistence of, 251 molecular mimicry in, 497, 501 mucosal B cells and, 238 mutagenesis and, 496–499, 503–505 mutational accomomodation in antibody–antigen interface and, 499–500 recognition of, 491 self- (See autoimmune disorders) somatic hypermutation (SHM) and, 503–505 specificity of recognition by antibody in, 500–501, 502 strength of antibody–antigen interaction in, 499 thermodynamics of antibody–antigen interactions in, 496 anti-idiotopic antibodies, 501 anti-immunoglobin l genes (IGL), human, leukemia and lymphoma, 40 anti-mitochondrial antibodies, autoimmune disorders and, 390–391 anti-phospholipid antibodies, 394–395 anti-phospholipid syndrome (APS), 394–395 anti-platelet autoantibodies, 391–392 anti-Ro autoantibodies, 393–394 anti-Tac (CD25) monoclonal antibodies, 540–541 anti-thyrotropin receptor (TSHR), autoimmune disorders and, 391, 391 aorta-gonad mesonephros (AGM), hematopoiesis and, 102, 450–452 AP1 and class switch recombination (CSR), 290 API2-MLT/MALT1 fusion gene in marginal cell lymphomas, 353 APOBEC-1, activation-induced deaminase (AID) and, 313, 331 apoptosis B cell development and, 117–118 plasma cells, memory and, 254 suppression of, during lymphoid development, translocations associated with, 352–353
APRIL B cell development and, 118, 191 class switch recombination (CSR) and, 290 Artemis immunodeficiency diseases and, 73, 148 nonhomologous end joining (NHEJ) and, 73–76, 148 artificial substrates for class switch recombination (CSR), 311, 311 artiodactyls (hoofed, grazing animals), 440–444 antibody diversity generation in, 442–443, 442 CH genes in, 443–444 heavy chain antibodies (HCAb) in, 440–441, 441 ileal Peyer’s patch (IPP) in, 443 immunoglobulin genes in higher vertebrates and, 440–444 k light chain genes in, 441–442 l light chain genes in, 441–442 lymphopoiesis and Ig gene diversification in, 443 V, D, J gene segments in, 441 ataxia telangiectasia mutated (ATM) protein, 74, 319 ATF6, immunoglobulin assembly and secretion, 270 autoantibodies (See autoimmune disorders) autoimmune disorders, 10, 381–401, 382 affinity maturation and, 381 anti-acetylcholine receptor (AchR) in, 392–393, 393 antibody subsets in, 382–383 anti-DNA antibodies and, 394 anti-mitochondrial antibodies in, 390–391 anti-phospholipid antibodies in, 394–395 anti-phospholipid syndrome (APS), 394– 395 anti-platelet autoantibodies in, 391–392 anti-Ro autoantibodies in, 393–394 anti-thyrotropin receptor (TSHR) and, 391, 391 CD5 and, 383, 384, 387 CD21 deficiency and, 276, 277 CD35 deficiency and, 276, 277 central tolerance and, 381 clonal anergy concept in, 382 cold agglutinins (CA) and, 388–390 complement and complement receptor deficiency and, 276 complementarity determining regions (CDR) and, 387–388, 390–391, 395 CREST syndrome, 383 Crohn’s disease, 382, 404 diabetes, 404 E2 antigen and, 390–391 environmentally induced, 382 evolution and, 381 exquisitely specific autoantibodies in, 383 Fc (FcgRIIB) and complement receptors in, 275–276, 284–285, 286 FR1 and, 388–390, 389 genetics of, 384–388 Graves’ disease, 391 HEL gene and, 382
hemolytic anemia, 384 horror autotoxicus concept in, 381 human/murine counterpart autoantibodies in, 392–395 IDDM, 382 identification of autoantigen in, 383 idiopathic thrombocytopenia purpura (AITP), 391–392 IgM and, 389–390 induction of damage by passive transfer of autoantibodies in, 383–384 isolation of pathogenic autoantibodies in, 384 MAbs in, 389–390 MAD-2 in, 389–390 memory and memory B cells in, 248 migration of B cells and, 213 molecular and immunochemical characteristics of autoantibodies, in humans, 388–392 molecular charactistics of autoantibodies, in mouse, 384–385 molecular charactistics of V gene repertoire and, in human, 386–388, 387 monoclonal antibodies and, 541 multiple sclerosis, 382 mutations and, 381–382 myasthenia gravis (MG), 392–393 natural, polyspecific autoantibodies in, 382–383 pathogenic autoantibodies in, criteria for defining, 383–384, 385 polymorphisms and, in human genes, 386–388 polymyositis, 383 primary biliary cirrhosis (PBC), 390–391, 391 scleroderma, 383 self-antigen and, 381 self-reactive B cells, negative selection of, 284–285 superantigenic drive on B cells and, 388–390 systemic lupus erythematosus (SLE), 393–394 thyroiditis, 383 uveitis, 382 V genes/regions in, 384–388, 385, 386, 387, 394, 395, 395 V region mutations and, 381–382 V regions and, transgenic mouse studies on, 384 X-linked (Bruton’s) agammaglobulinemia (XLA) and, associated with, 404 autoimmune idiopathic thrombocytopenia purpura (AITP), 385, 391–392 avians (See birds)
B B 1 cells, 172 in antibody production and response, 187 B cell antigen receptor complexes (See B cell receptor) B-cell–attracting chemokine (BCA), class switch recombination (CSR) and, 290
566 B cell development (See also B cell receptors), 101–126, 141–154, 142 accessibility hypothesis in, 128 activation-induced deaminase (AID) gene in, 143 allelic exclusion at H chain locus and, 113, 114 antibody production and, 155 antibody regulation by B cell receptors and, 147–148, 147 aorta-gonad-mesonephros (AGM) region and, 102 B cell receptor (BCR) in (See B cell receptors) B lineage leukemia and, 149–150 B lymphoid follicles (FO) and, 155 B1 cell compartment and, 118 B1 cells in, 155, 172 BAFF and APRIL in, 118–119, 157, 191 Bcl and, 192 blastocyst complementation assay studies in, 103 bone marrow and, 102–103, 117, 141, 143, 150, 212–213 CCL3 and, 189 CD3 and, 147 CD4 and, 194, 196 CD5 and, 157 CD8 and, 194 CD10 and, 142, 149 CD11 and, 188, 190, 191, 197, 207 CD18 and, 207 CD19 and, 101, 142, 149, 163, 165–166, 171–186, 231 CD20 in, 101 CD21 and, 147, 171, 176, 188, 194, 195 CD22 and, 165–166, 171–186, 178, 212 CD23 and, 194 CD27 and, 143 CD28 and, 147, 196, 197 CD31 and, 207 CD34 and, 141–142, 149 CD35 and, 176 CD38 and, 142 CD40 and, 147, 158, 163, 189, 191, 194, 196, 197 CD45 and, 142 CD54 and, 195 CD57 and, 194 CD72 and, 165–166, 180–181, 181 CD80 and, 176 CD81 and, 147 CD86 and, 176, 196 CD100 and, 180–181 chemokines and, 103, 156 cKit and, 141 class switch recombination (CSR), 147 clonal expansion in, 142 commitment of, 106–107, 109, 189–190, 190 common acute lymphoblastic leukemia antigen (CALLA) and, 142 common lymphoid precursors (CLP), 106, 141–142 earliest progenitor cells in, 106–107 myeloid progenitor (pM) cells in, 107
Index ordering of lymphoid progenitors by marker expression in, 106 pL cell compartments in, 106 surrogate light chains (SLC) of the prelymphocyte receptors and, 107 compartmentalization in, 157, 209–213, 239 complement receptor requirements, during stages of, 282–285, 283 complementarity-determining regions (CDR3) in, 144 CTLA4 and, 147 CXCL12 and, 189, 191 CXCL13 and, 157, 189, 192, 195 CXCR4 and, 191 CXCR5 and, 194, 195 dendritic cell (DC) development and, 141, 142 differentiation in, 141–142, 156–157, 234, 237–238, 249, 251, 255, 263–264 E2A protein and, 108, 109, 130 early B cell factor (EBF) in, 108, 109 early stages of, 101–126 ”editing” of receptors in, 115–116, 145–146, 146 embryonic development and, hematopoiesis during, 101–103, 102 erythrocyte development and, 102 EU12 and, 146 Fas and, 157–158 FcRH and, 147 FcRL and, 147 FcRX in, 147 FcgRIIBs in, 181, 182, 276 FREB and, 147 G protein coupled receptors (GPCRs) in, 156, 204 galectin 1 in, 111, 145 growth factors in, 103 heavy and light chain assemblage in, 101, 141–144 hematopoietic stem cells (HSC) and, 141 hemoglobin development and, 102 HLA-DR cells and, 142 hormonal activity and, 107 human immunoglobulin genes in, 143–144 ICAM-1 and VCAM-1 in, 157 IgA and, 149 IL7 and, 109–110, 141, 148–149 immature B cells and, 116–117 immune response and, 155 immunodeficiency diseases and, 148–149 immunoreceptive tyrosine-based activiation motif (ITAM) in, 147, 177, 182 immunoreceptor tyrosine-based inhibitory motif (ITIM) in, 147 inhibitory co-receptors (See also CD22), 177–182, 178 integrins and, 103 kl and ll chain rearrangement in, 114–115 in lagomorphs (rabbits and hares), 437–440 l5 and, 109, 111, 127, 142–144, 148 liver-derived B cells and, 102, 141, 143 localization in, 156, 249, 250 lymph nodes and, 155, 187, 188–189, 203, 204
lymphocyte development and, 102 megakaryocyte development and, 102 migration of (See migration of B cells) mH chains and allelic exclusion in, 114 multiple VL-JL rearrangement in, 115–116 mutations and, 103, 117, 148 myeloid cell development and, 102 myeloid progenitor (pM) cells in, 107 natural killer cell development and, 108, 141, 142, 155 negative selection and, anergy and apoptosis in, 117–118 omentum and “milk spot,” 211–212, 212 P1/PDL-1 and, 158 pathogenic autoantibody production by (See autoimmune disorders) Pax5 protein and, lineage and stage-specific Ig enhancers in, 93 Pax5-deficiency pre-B cells and, plasticity of, 109–110 periarteriolar lymphocyte sheath (PALS) and, 117 peritoneal and pleural cavities in, 155, 211–212, 226 pL cell compartments in, 96, 106 platelet development and, 102 pluripotent hematopoietic stem cells (pHSC), 103–106 hemangioblasts as early progenitors of, 104–105 long-term reconstitution potential of, 104 migration and homing in, 104 plasticity vs. stability of, 105–106, 105 pluripotency of, 104 self-renewal ability in, 103–104 stress-induced development of, 105–106 transdifferentiation to and from nonhematopoietic cells, 106 vascular endothelium development and, 104–105 positive selection and B1 cell compartment, 118 pre-B cell formation in, 101 pre-B cell receptors (pre-BCRs) in, 101, 110, 127, 141, 142, 144–145 pre-B-1 cells in, 112 pre-BcR and BcR-independent development in, 118–119 pre-B-II cells in, 112 predetermination of primary vs. secondary response in B cells and, 248 primitive blood cell lineages and, 102 proliferation of, 237–238 PU-1 and, 107–108 pyk-2 and, 156–157 rapid selection of Vk to Jk rearrangement and allelic exclusion in, 116–117 rearrangement of L chain loci and, 114–116 recombination activating genes (RAG1/RAG2), 106, 108, 109, 112, 114, 119, 127, 141, 142, 144–145 recombination signal sequences (RSS) in, 145 relocalization of, 210 retention of, 237–238
Index Sca 1 and, 141 selection of immature B cells for, 117–119, 156–157 SH2 domain containing inositol 5phosphatase (SHIP) in, 147, 182 SHP-1 and, 166, 181 signaling reactions initiated by pre-B cell receptors in, 113 sites of, 143 SL-containing pre-B cell receptor and, 112–113 somatic hypermutation (SHM) and, 330–331 spleen and, 117, 155, 187, 208–209, 208 stem cell development and, 102 steps in, 101 stress-induced, 105–106 subset development (See subset development and function of B cells) surrogate light (SL) chain and, 107, 110–111, 144–145 expression of, 111 ligands for pre-B cell receptors in, 111 structure and assembly of pre-B cell receptor in, 110–111 T cell development and, 108–109, 127, 141, 142, 155 T cell receptor (TCR) genes and, 127, 128, 129, 130, 155 TdT and, 106, 107, 114, 144 Thy 1 and, 141 transcription factors control of lymphoid cell development in, 107–109 PU-1 and, 107–108 T, B, or NK decision by, 108 tumor necrosis factor (TNF) and, 118, 157 12/23 rule in, 128 V(D)J recombination and, 101, 109, 110, 112, 113, 114, 119, 141 Vk gene segment usage in, 115 VkJk rearrangement in, 115, 116 VpreB and, 109–111, 113, 114, 127, 142–144 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 407–408 zebrafish (Danio rerio) and, 457 B cell leukemias and lymphomas (See also mouse B cell lineage lymphomas), 10, 149–150, 354–355, 349–364 apoptosis suppression during lymphoid development, translocations associated with, 352–353 blocks in lymphoid differentiation and, translocations associated with, 349–352, 350 cell cycle regulation in mantle cell lymphoma/myeloma, 355–356 chromosomal translocations in, 349–364 11q23 abnormalities in MLL and, 350–351 immunoglobin l genes (IGL), human and, 40 lymphoid precursor proliferation and, translocations associated with, 354–355 mouse (See mouse B cell lineage lymphomas)
NFKB pathway and translocations associated with, 354 t(11;18)(q21;q21) and API2-MLT/MALT1 fusion gene in marginal cell lymphomas, 353 t(14;18) and activation of BCL2 in follicular lymphoma, 352–353, 370 t(17;19)(q21-q22;13) and E2A-HLF fusion gene in pro-B cell lymphoblastic leukemias of adolescence, 353 t(21;21) (p13;q22), E2A-PBX1 fusion gene and, pre-B cell lymphoblastic leukemias, 351 t(21;21) (p13;q22), TEL-AML1 (ETV6RUNX1) fusion gene and, pediatric early B-lineage acute, 349 t(3;14)(q27;q32) and BCL6 activation in diffuse large cell lymphoma, 351–352 t(4;14)(p16;q34) and activation of FGFR3 and MMSET in multiple myeloma, 356 t(8;14)(q24;q23) and MYC activation in Burkitt lymphoma/B cell leukemia, 354–355 t(9;14)(p13;q32) and PAX5 activation in marginal cell lymphomas, 352 t(9;22)(q34;q11) and BCR-ABL fusion gene in adult early B-lineage acute lymphoblastic leukemia, 355–356 B cell lineage specific activator protein (BSAP), transcription of immunoglobulin genes and, 88 B cell receptor (BCR), 101, 127, 141–145, 156–157, 161–169, 248–249 activation of, 161, 162–163, 163 in antibody production and response, 188 antibody production regulated by, 147–148, 147 antigen defined in, 161 antigen sensititive/specificity and, 166 B cell development and, 101, 142, 143, 145, 156–157 BANK and, 164 BAP29 and BAP31 in, 265, 267 binding of, 161 Btk and, 162, 164 calcium complex initiation in, 164 CD19 and, 163, 165–166 CD22 and, 165–166 CD40 and, 163 CD72 and, 165–166 class switch recombination (CSR) and, 289–290, 290 Epstein-Barr virus (EBV) and, 166, 389, 390 erythropoietin (EPO) molecules/receptors and, 161 evolution of, 481–483, 481 Fc (FcgRIIB) and complement receptors in, 277, 278, 285–286 fine-tuning of, using ITAM, 165–166 glycosyl-phosphatidyl inositol (GPI) anchor and, 166 hematopoietic progenitor kinase (HPK1) in, 165 immune response and, 166, 340 immunodeficiency diseases and, 164
567 immunoglobulin assembly and secretion in, 261, 263, 265 inositol triphosphate (IP3) and, 164 ITAM and, 162–166 linker of activated T cell (LAT) in, 165 Lyn and, 162, 163, 174, 177 memory and memory B cells in, 248–249, 251–252 migration of B cells and, 210 monitoring assembly of, 265 mouse B cell lineage lymphomas and, 365 mucosal B cells and, 230, 231 NADPH-oxidase and, 163, 163 non-T cell activation linker (NTAL) in, 165 phosphatase inhibition in, 162 phosphoinositide 3 kinase (PI3K) and, 164 protein kinase C (PKC) in, 164 protein tyrosine kinases (PTK) and, 162 protein tyrosine phosphatases (PTP) in, 162 redox regulation of signaling in, 162–163 SH2 domain and, 162, 165 SHP-1 and, 162, 166, 177–178 SLAP130 and, 165 SLP-65 (BLNK, BASH), 163–166, 177 structure of, 161–162 Syk coupling with, 162, 163, 177 transient receptor potential (TRP) and, 164 X-linked agammaglobulinemia (XLA), 164 B natural killer cell lymphomas, mouse B cell lineage lymphomas and, 373 B1 cells, 212, 282 B2 cells, 212 B220, 106, 117, 248 B7RP-1, class switch recombination (CSR) and, 290, 287 bacterial artificial chromosome (BAC) lower vertebrate immunoglobulin genes and, 422 Vk genes, human and, 27 Vk genes, mouse and, 31, 33 BAFF, 118–119, 157, 191 BANK, B cell receptor (BCR) and, 164 BAP29, 265, 267 BAP31, 265, 267 base excision repair, somatic hypermutation (SHM) and, 332, 333 base unpairing regions (BURs), transcription of immunoglobulin genes and, 93 BASH, B cell receptor (BCR) and, 163–165 basiliximab, 541 BCA-1 mucosal B cells and, 231 BCAP, lower vertebrate immunoglobulin genes and, 428 BCDX2, somatic hypermutation (SHM) and, 335 Bcl/bcl in antibody production and response, 192 immune response of B cells and, 340 memory and memory B cells in, 251 plasma cells, memory and, 253 BCL, 366, 371 BCL-2, follicular lymphoma and, 352–353, 370 BCL-6, 255, 330, 351–352 Bcl6, immunoglobulin assembly and secretion, 269
568 BCR-ABL fusion gene in adult early B-lineage acute lymphoblastic leukemia, 355–356 Bence-Jones proteins, immunoglobulin assembly and secretion, 266 BENE, Vk genes of human/mouse and, 28 B1 integrin, B cell development and, 103 beyond 12/23 (B12/23) rule, V(D)J recombination and, 64–65 bifunctional genes, V chain (VCBP), 429 BILL cadherin, B cell development and, 111 binding energetics of antibody–antigen interactions, 495–496 BiP, immunoglobulin assembly and secretion, 264–265, 264 birds/avians antibody diversity generation in, 435–436, 435 B cell receptors (BCR) in, 435–436 bursa of Fabricius in, 330, 435–436 CH genes in, 436 chicken Ig genes, organization of, 433–434 class switch recombination (CSR) and, CH gene organization and, 310 complementarity-determining region (CDR) in, 435 gut-associated lymphoid tissues (GALT) in, 433–436 Ig genes in, organization of, 433–434 immunoglobulin genes in higher vertebrates and, 433–436 lymphopoiesis in, limited, 434–435 somatic hypermutation (SHM) and, templated mutation in, pre-immune diversification, 328–329 V(D)J recombination in, 433–435 Birshtein, Barbara K., 289 blastocyst complementation assay studies, B cell development and, 103 BLC, 209, 231 BLIMP-1, 190, 255, 269, 270 BLIN cells, immunodeficiency diseases and, 150 BLNK agammaglobulinemias and, 410 B cell receptor (BCR) and, 163–165 immunodeficiency diseases and, 148 BLys, class switch recombination (CSR) and, 290 body cavity B cells, 211–212 Bona, Constantin A., 381 bone marrow, 102–103, 141, 143 antibody–secreting cells (ASCs) and, homing of, 213–215 B cell development and, 102–103, 117, 150 Fc (FcgRIIB) and complement receptors in, 284 immune response of B cells and, 340 memory and memory B cells in, 249, 250, 251 migration of B cells and, 212–213 plasma cells, memory and, 253 sialic acid and, 212–213 stromal cells and, 150, 150 bone morphogenic protein (BMP), pluripotent hematopoietic stem cells (pHSC) and, 103
Index Bonilla, Francisco A., 403 bony fish (See also fish), 419–422 Brandtzaeg, Per, 223 break and repair pathway, somatic hypermutation (SHM) and, 334 BRIGHT, MAR binding protein, 93 bronchus-associated lymphoid tissue (BALT), mucosal B cells and, 224–225, 236–237 Brüggemann, Marianne, 547 Bruton’s tyrosine kinase (BTK) (See also Xlinked agammaglobulinemia), 403 Btk/btk subset development in B cells, 156 B cell receptor (BCR) and, 162, 164 BTK deficiency, X-linked (Bruton’s) agammaglobulinemia (XLA) and, 148, 368, 405–409, 406, 407 BTrCP/SCF complex, transcription of immunoglobulin genes and, 95–96 Burkitt-like lymphoma (BLL), 366, 371–374 Burkitt lymphoma (BL), 215–216, 354–355, 366, 371, 374, 375 Burrows, Peter D., 141 bursa of Fabricius, 330, 435–436 BXSB, 275
C Cm, class switch recombination (CSR) and, 292–293 C region antibody production and, 491, 493–495 class switch recombination (CSR) and, 307 immunoglobin l genes (IGL), human and, 41, 42–43, 46, 48, 49, 49 immunoglobin l genes (IGL), human/mouse, 38 immunoglobin l genes (IGL), mouse and, 50–51, 52–55, 54–55, 56 immunoglobulin assembly and secretion, 261, 264–265 somatic hypermutation (SHM) and, 327 transcription of immunoglobulin genes and, 83 translocus mice and human Mab and, 547–558 zebrafish (Danio rerio) and, 453–454 C to U deamination, somatic hypermutation (SHM) and, and base excision repair in, 332, 333 C/EBPb class switch recombination (CSR) and, 292, 293 transcription of immunoglobulin genes and, 89 C/G base pairs, somatic hypermutation (SHM) and, 332, 334 C1q and Fc/complement receptors, 275, 276, 281 C2 and Fc/complement receptors, 281 C3 and Fc/complement receptors, 275–276, 280–281 C3d and Fc/complement receptors, 171 C4 and Fc/complement receptors, 275, 276, 281
C5–C9 and Fc/complement receptors, 280 Calame, Kathryn, 83 calcium signaling/transport antigen receptor signaling regulation and, 171 antigen receptor signaling regulation and, mobilization and, 177 B cell receptor (BCR) and, 164 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 Fc (FcgRIIB) and complement receptors in, 278, 278 Ca, class switch recombination (CSR) and, regulatioin, 293 cancer (See also B cell leukemias and lymphomas), Pax5 protein and, lineage and stage-specific Ig enhancers in, 94 CARD, transcription of immunoglobulin genes and, 95 Carroll, Michael C., 275 cartilaginous fish, gene multiplicity in, 417–419 cats, immunoglobulin genes in higher vertebrates and, 444 CBF2T1, 459 CCL19 migration of B cells and, 206–207, 206, 208 mucosal B cells and, 235 plasma cells, memory and, 253 CCL20, migration of B cells and, 211 CCL21 migration of B cells and, 206–207, 206, 208, 213 mucosal B cells and, 235 plasma cells, memory and, 253 CCL22, migration of B cells and, 210 CCL25 migration of B cells and, 215 mucosal B cells and, 236 plasma cells, memory and, 254 CCL28, mucosal B cells and, 236 CCL3 in antibody production and response, 189 migration of B cells and, 210 CCL4, migration of B cells and, 210 CCND1, mouse B cell lineage lymphomas and, 373 CCR10, mucosal B cells and, 236, 237 CCR7 migration of B cells and, 206–207, 206, 209, 210, 214 mucosal B cells and, 232, 235 plasma cells, memory and, 253 CCR9, 215 mucosal B cells and, 236 plasma cells, memory and, 253 CD3 B cell development and, 108, 147 mucosal B cells and, 227 CD3e, lower vertebrate immunoglobulin genes and, 428 CD4, 190 in antibody production and response, 189, 194, 196 B cell development and common lymphoid precursors (CLP), 106
Index mucosal B cells and, 231 CD5 autoimmune disorders and, 383, 384, 387 Fc (FcgRIIB) and complement receptors in, 282 mouse B cell lineage lymphomas and, 366 mucosal B cells and, 227 subset development in B cells, 156, 157 CD8 in antibody production and response, 194 B cell development and, 110 class switch recombination (CSR) and, 311 memory and memory B cells in, 252 CD8A, translocus mice and human Mab and, 552 CD9, lower vertebrate immunoglobulin genes and, 428 CD10 B cell development and, 111, 142, 149 immunodeficiency diseases and, 149 CD11, 207, 290 in antibody production and response, 188, 190, 191, 197 Fc (FcgRIIB) and complement receptors in, 282 mucosal B cells and, 235 CD14, class switch recombination (CSR) and, 290 CD18 migration of B cells and, 207 mucosal B cells and, 235 CD19, 163, 165–166, 171–172 agammaglobulinemias and, 409 antigen processing and, 176 B cell development and, 101, 106, 107, 109, 111, 142, 149 common lymphoid precursors (CLP) and, 106, 107 biochemistry of signal transduction in, 174–176 CD21 complex with, 174 immunodeficiency diseases and, 149 inhibitory receptors CD22 and FcgRII with, 176 ligands of, 173–174 MAP kinases and, 175 memory and memory B cells in, 248, 253 mucosal B cells and, 227, 231 PLCg2 and, 175 role of, in antigen receptor regulation, 171 signal transduction by, 173–176, 175 STAT1 and, 175 subset development in B cells, 156 thymus-dependent antigens and, 173 thymus-independent antigens and, 172–173 transcriptional regulation of expression in, 171–172 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 CD20 B cell development and, 101 plasma cells, memory and, 253 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408
CD21, 171, 176 in antibody production and response, 188, 194, 195 B cell development and, 106, 117, 147 common lymphoid precursors (CLP) and, 106 CD19 complex with, 174 deficiency of, 276, 277 Fc (FcgRIIB) and complement receptors in, 281–286 mucosal B cells and, 231 subset development in B cells, 156 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 CD22, 165–166, 171 calcium mobilization and, 177 CD19 and, 176 extracellular domain of, signaling in, 179–180 Grb2 and, 177 impairment/deficiency of, 178 inhibition of intracellular signaling with, 177–178, 178 ITIM and, 177 ligands and ligand binding in, 179–180, 180 Lyn and, 177 MAP kinases and, 178 migration of B cells and, 212, 214–215 PI3K and, 177 plasma cells, memory and, 253 PLCg2 and, 177 Shc and, 177 SHP 1 and, 177–178 subset development in B cells, 156 CD23 in antibody production and response, 194 B cell development and, 106, 117 common lymphoid precursors (CLP) and, 106 Fc (FcgRIIB) and complement receptors in, 282 CD25 B cell development and, 106, 107, 109, 111 common lymphoid precursors (CLP) and, 106, 107 CD27, 143, 255 B cell development and, 106, 107, 111 common lymphoid precursors (CLP) and, 106, 107 memory and memory B cells in, 248–249, 253 mucosal B cells and, 227, 232 CD28 in antibody production and response, 196, 197 B cell development and, 147 class switch recombination (CSR) and, 290 CD29, mucosal B cells and, 235, 237 CD30, class switch recombination (CSR) and, 290 CD31, migration of B cells and, 207 CD34 B cell development and, 141–142, 149 migration of B cells and, 205
569 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 CD35, 176 deficiency of, 276, 277 Fc (FcgRIIB) and complement receptors in, 281–286 mucosal B cells and, 231 CD38, 142 mucosal B cells and, 232, 236 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 CD40, 163 affinity maturation in, 342, 344 in antibody production and response, 189, 191, 194, 196, 197 B cell development and, 147 class switch recombination (CSR) and, 290, 292, 293, 297, 312 Fc (FcgRIIB) and complement receptors in, 282 high affinity antibody and, 342, 344 hyper-IgM syndrome (HIGM) and, 410 memory and memory B cells in, 255 mucosal B cells and, 227, 231 subset development in B cells, 158 CD43, Fc (FcgRIIB) and complement receptors in, 282 CD44 B cell development and, 109 mucosal B cells and, 236, 237 CD45 B cell development and, 142 lower vertebrate immunoglobulin genes and, 428 plasma cells, memory and, 253 subset development in B cells, 156 CD45R, B cell development and, 117 CD49, mucosal B cells and, 237 CD54 in antibody production and response, 195 mucosal B cells and, 235 CD57 in antibody production and response, 194 mucosal B cells and, 231 CD62L, migration of B cells and, 204–205 CD70 memory and memory B cells in, 249 mucosal B cells and, 232 CD72, 165–166, 180–181, 181 CD79, agammaglobulinemias and, 409, 410 CD80, 176 CD81, 147, 171, 172 CD86, 176, 196 CD95 plasma cells, memory and, 253 somatic hypermutation (SHM) and, 330 CD98, lower vertebrate immunoglobulin genes and, 428 CD99, migration of B cells and, 207 CD100, 180–181 CD120, mucosal B cells and, 235 CD138, plasma cells, memory and, 253 CD154, hyper-IgM syndrome (HIGM) and, 410 CDK6, 255
570 CDR grafting technique, monoclonal antibodies and, 534–536 Celera database, immunoglobin l genes (IGL), mouse and, 51 cell cycle regulation in mantle cell lymphoma/myeloma, 355–356 central tolerance, autoimmune disorders and, 381 centroblastic DLBCL, 370 centroblasts and centrocytes, germinal centers (GC), 194 cFLIP in antibody production and response, 196 Fc (FcgRIIB) and complement receptors in, 282 Cg1, class switch recombination (CSR) and, 292–293, 312 Cg2a, class switch recombination (CSR) and, 290, 293, 312 Cg2b, class switch recombination (CSR) and, 292 Cg3, class switch recombination (CSR) and, 292 CH (constant) region, human IgH, 1 IgH, 12–15, 13 in artiodactyls (hoofed, grazing animals), 443–444 in birds/avians, 436 in lagomorphs (rabbits and hares), 439–440, 439 polymorphisms of, 14–15, 14 structure of genes in, 13–14, 13 switch region in, 14 transcription enhancers in, 13 CH12F3–2, activation-induced deaminase (AID) and, 313 chemokines autoimmune disorders and, 382 B cell development and, 103 gut lamina propria and, 236 immune response of B cells and, 340 innate immunity and, 464 migration of B cells and, 214–215 migration of B cells and, integrin activation triggering by, 205–207 mucosal B cells and, 231, 234–235, 237 subset development in B cells, 156 chickens (See also birds/avians) antibody diversity generation in, 435–436, 435 B cell receptors (BCR) in, 435–436 bursa of Fabricius in, 330, 435–436 CH genes in, 436 class switch recombination (CSR) and, CH gene organization and, 310 complementarity-determining region (CDR) in, 435 Ig genes in, organization of, 433–434 lymphopoiesis in, limited, 434–435 somatic hypermutation (SHM) and, templated mutation in, pre-immune diversification, 328–329 V(D)J recombination in, 433–435 chimerization of monoclonal antibodies, 511, 533–534, 540
Index choriomeningitis virus (CMV), 389, 390 chromatin, transcription of immunoglobulin genes and, structure and, 90, 130–132 chromatin remodeling, class switch recombination (CSR) and, 3¢ IgH regulatory region and, 299–300 chromosomal translocation, (See B cell leukemias and lymphomas) chronic lymphocytic leukemia (CLL), 366, 367, 541 cirrhosis, primary biliary (PBC), 390–391, 391 CIS, mouse B cell lineage lymphomas and, 373 cis elements, somatic hypermutation (SHM) and, 329–330, 485 cKit, B cell development and, 141 class switch recombination (CSR), 289–305, 307–326 3¢ IgH enhancers in, 293–295, 316 activation-induced deaminase (AID) and, 289, 290, 307, 311, 313–314, 317, 319–320 affinity maturation and, 307 in amphibians and reptiles, 422–423 AP1 and, 290 artificial substrates for, 311, 311 ataxia-telangiectasia mutated (ATM) gene in, 319 B-cell–attracting chemokine (BCA) in, 290 B cell development and, 147 B cell receptors (BCR) and, 289–290, 290 B7RP-1 and, 290, 287 BLys and APRIL in, 290 C regions in, 307 C/EBPb and, 292, 293 Ca regulatioin in, 293 CD11 and, 290 CD14 and, 290 CD28 and, 290 CD30, 290 CD40 and, 290, 292, 293, 297, 312 CD8a and, 311 Ce in, 292–293 Cg1 in, 292–293, 312 Cg2a and, 290, 293, 312 Cg2b in, 292 Cg3 in, 292 chicken, frog, bony fish, CH gene organization and, 310 chromatin remodeling and, 3¢ IgH regulatory region and, 299–300 cognate interactions with T cells and, 290 coordinated regulation of transcription, recombination, replication in, 300 CREB and, 293 cytokines in, 312 DNA ligase IV in, 319 DNA processing and end joining after cleavage in, 315–319 DNA-dependent protein kinase (DNA-PK) in, 316 double strand breaks (DSBs) and, 314–315, 316, 319 E2A in, 312 E47 and, 312 Em enhancer and, 291–292, 308–309
evolutionary studies of, 310, 320 exonuclease I and ERCC1 in, 318–319 follicular dendritic cells (FDC) and, 290 G quartet structures in, 315 g-H2AX in, 319 germinal center (GC) and, 289–291, 290 germline transcription (GT) and, 289, 312 histone deacetylase (HDAC) in, 291 human CH gene organization and, 310 human I promoters in, specificity of, 293 hyper-IgM syndrome and, 290 hypersensitive sites (HS) and, 309 I exon transcription in, 291, 312 I promoter interaction with 3¢ IgH enhancers in, 295–297 ICOS and, 290, 287 IFN-g and IFN-a in, 293 IFN-g cells in, 290, 287 IgA in, 293 IgG and, 290, 293 IKKa and, 290, 297 IL4 and IL13 in, 292 immunodeficiency diseases and, 290 inhibition of, 312 insulin receptor substrate (IRS) in, 292 interleukins in, 292, 312 IRF and, 293 isotype specificity in, 312 isotype switching and, 307 Im GT and, 291–292 JAK in, 292, 293 late SV40 factor (LSF) in, 291 LPS and, 312, 320 LR1 and, 291 LTNa/b and, 290 lymphoid-specific factors of Oct, E2A, ets families in, 298 lymphotoxins and, 290 macrophage inflammatory protein (MIP) in, 290 MAPK/JNK pathways and, 290 mechanisms for 3¢ IgH regulatory region mediated regulation of GT in, 295–300, 296 mismatch repair (MMR) and, 316–318, 319 mouse B cell lineage lymphomas and, 365 mouse S region organization and, 308–310, 310 mucosal B cells and, IgA promotion in, 232–234 mutations and, 311 natural killer cells and, 290–291 NEMO and, 290 NF-kB in, 291, 292, 287, 312 NFkB/Rel proteins and, 290 Nijmegen breakage syndrome 1 (Nbs-1) gene in, 319 nonhomologous end joining (NHEJ) and, 297, 310, 311, 315–316, 319 non-Ie promoters and cytokine inducibility in, 292–293 organization of S region and CH genes in, 308–310, 310 palindromic tetramers in, 315 Pax5 as regulator in, 298–299
571
Index polarized effect of 3¢ IgH regulatory region and, 295 promoters of, 289 proteins for DNA repair, processing, joining of S regions in, 316–318 proximal cis regulatory elements for GT in, 291–293 PU-1 and, 292 ”recombinasome” in, 319 RNA-DNA hybrids in, 315 S region cleavage in, 314–315 S regions in, 308–319, 309 secondary structures and, activity of, 315 SHIP and, 291 somatic hypermutation (SHM) and, 289, 307, 319–320, 330, 331 S-S recombination with looped-out CH gene deletion in, 307–308, 308 staggered nick cleavage in, 314–315 STAT in, 292, 293 stem-loop structures in, 315 stromal cell-derived factor (SDF) in, 290 SWAP-70 and, 290, 297 switch sequences as transcriptional stimulatory elements in, 291 T cell-independent (TI) antigens and, 289 TGF-b and, 293 TGF-b inhibitory element (TIE) in, 292 TNFR and, 290 Toll-like receptors and, 290 TRAF and, 290 transcription of S region in, 311 transcription factors controlling 3¢ IgH regulatory elements in, 297–299 transcription of immunoglobulin genes and, 91, 147 unique properties of, 310–311 V regions in, 307 V(D)J recombination and, 289, 292, 307, 312, 315 vasointestinal protein (VIP) and, 293 CLASSIFICATION concept, Immunogenetics Database (IMGT) and, 37–38, 38 clonal anergy concept, autoimmune disorders and, 382 coding flanks, V(D)J recombination and, 65 coding joint (CJ), 61, 72, 72 Cogne, Michel, 289 cold agglutinins (CA), autoimmune disorders and, 388–390 cold hemolytic anemia, 385 Colliers de Perles representations, Immunogenetics Database (IMGT) and, 40, 41, 42, 43 common acute lymphoblastic leukemia antigen (CALLA), 142 common lymphoid precursors (CLP) B cell development and, 106, 141, 142 earliest progenitor cells in, 106–107 myeloid progenitor (pM) cells in, 107 ordering of lymphoid progenitors by marker expression in, 106 pL cell compartments, 106 surrogate light chains (SLC) of the prelymphocyte receptors and, 107
common myeloid precursors (CMP), B cell development and, 106 common variable immunodeficiency (CVI), 149 comparative genomic hybridization (CGH), 366 compartmentalization of mature B cells, 157, 209–213, 239 complement control proteins (CCP), Fc (FcgRIIB) and complement receptors in, 281 complement-dependent cytotoxicity (CDC), 542 complement system (See Fc and complement receptors), 275 complementarity-determining regions (CDR) affinity maturation and, 342–344 antibody production and, 491, 494, 496–499, 504 autoimmune disorders and, 387–388, 390–391, 395 B cell development and, 144 cartilaginous fish, 418 CDR grafting technique in, 534–536 high affinity antibody and, 342–344 immune response of B cells and, 342–344 monoclonal antibody display libraries and, 511, 523, 534–539 phage display libraries and, 523 somatic hypermutation (SHM) and, 327 V(D)J recombination and, 61, 146, 483 Cooper, Max D., 141 cosmid cloning, human IgH and, 2 CpG methylation, V(D)J recombination and, 132 CR1 Fc (FcgRIIB) and complement receptors in, 275, 276 mucosal B cells and, 231 CR2 Fc (FcgRIIB) and complement receptors in, 275, 276 mucosal B cells and, 231 CREB class switch recombination (CSR) and, 293 V(D)J recombination and, 130 CREST syndrome, 383 Crohn’s disease, 228, 382, 404 cross-reactivity, antibody production and, 500–501, 502 CTLA 4 in antibody production and response, 196, 197 B cell development and, 147 CXCL4, plasma cells, memory and, 253 CXCL9 migration of B cells and, 215 plasma cells, memory and, 253, 254 CXCL10 migration of B cells and, 215 plasma cells, memory and, 253, 254 CXCL11 migration of B cells and, 215 plasma cells, memory and, 253, 254 CXCL12 in antibody production and response, 189, 191
migration of B cells and, 206–207, 206, 212–215 mucosal B cells and, 231, 235 plasma cells, memory and, 253, 254 CXCL13 in antibody production and response, 189, 192, 195 migration of B cells and, 206–207, 206, 208–215 mucosal B cells and, 231, 235 plasma cells, memory and, 253 subset development in B cells, 157 CXCR3 migration of B cells and, 215 plasma cells, memory and, 253 CXCR4 in antibody production and response, 191 B cell development and, 103, 106 common lymphoid precursors (CLP) and, 106 migration of B cells and, 206–207, 206, 212, 214 mucosal B cells and, 231, 235 plasma cells, memory and, 253, 254 CXCR5 in antibody production and response, 194, 195 migration of B cells and, 206–207, 206, 208, 210, 212, 214 mucosal B cells and, 232, 235 Cys213, immunoglobulin assembly and secretion, 266 Cys575, immunoglobulin assembly and secretion, 266 Cyster, Jason G., 203 cytokines autoimmune disorders and, 382 class switch recombination (CSR) and, 312 innate immunity and, 463–464 migration of B cells and, 213 pluripotent hematopoietic stem cells (pHSC) and, 104 cytosolic localization, transcription of immunoglobulin genes and, 94 cytotoxic lymphocytes (CTLs), innate immunity and, 462 cytotoxicity antibody-dependent cellular cytotoxicity (ADCC) and, 542 complement-dependent cytotoxicity (CDC) and, 542
D D regions beyond 12/23 (B12/23) rule in, 64–65 creation of, in V(D)J recombination, germline signal joint formation in, 482 transcription of immunoglobulin genes and, 91 translocus mice and human Mab and, 547–558 12/23 rule in, 62, 63, 64–65, 128 V(D)J recombination and, 62 D1.3, antibody production and, 494–501
572 daclizumab, 541, 541 Danilova, Nadia, 449 Davis, Randall S., 141 deafness-dystonia protein gene, X-linked (Bruton’s) agammaglobulinemia (XLA) and, 406 decay activating factor (DAF) regulator, Fc (FcgRIIB) and complement receptors in, 281 defensins, migration of B cells and, 211 delta-1, pluripotent hematopoietic stem cells (pHSC) and, 104 dendritic cell (DC) development, 141, 142, 460 innate immunity and, 460 DESCRIPTION concept, Immunogenetics Database (IMGT) and, 39–40 Dh (diversity) region chromosome 15 and 16 segments of, 5–6, 6 evolution of, 10–11 human IgH and, 1 organization of, 5 Dh-Cu region activation, transcription of immunoglobulin genes and, 91, 92 diabetes, 213, 404 differentation of B cells, 234, 249, 251, 255, 263–264 diffuse large B cell lymphoma (DLBCL), 366, 367, 368, 370–374 diffuse large cell lymphomas (DLCL), 330, 351–352 digestion circularization PCR (DC-PCR), activation-induced deaminase (AID) and, 313 dimerizing proteins, transcription of immunoglobulin genes and, regulation by, 87–88 display libraries (See monoclonal antibodies from display libraries) diversification of B cells (See also V(D)J diversification), 473 DNA binding proteins transcription of immunoglobulin genes and, regulation in, cooperative interactions in, 88–89 DNA breaks, somatic hypermutation (SHM) and, 334–335 DNA-dependent protein kinase (DNA-PK) class switch recombination (CSR) and, 316 nonhomologous end joining (NHEJ) and, 74–75, 76 somatic hypermutation (SHM) and, 335 transcription of immunoglobulin genes and, 95 DNA ligase IV class switch recombination (CSR) and, 319 nonhomologous end joining (NHEJ) and, 74, 76 DNA polymerases, V(D)J recombination and, NHEJ proteins and, 75 DOCK2, migration of B cells and, 216 dogs, immunoglobulin genes in higher vertebrates and, 444 domains, recombination activating genes (RAG1/RAG2) and, 477–479
Index double strand break (DSB) class switch recombination (CSR) and, 314–315, 316, 319 nonhomologous end joining (NHEJ) and, 72–73, 77 recombination activating genes (RAG1/RAG2), 77 somatic hypermutation (SHM) and, 319, 334–335 V(D)J recombination and, 61, 62, 71, 71, 72–73, 77 DQ52 promoter, 129 Dunn, Thelma, 365
E E2 antigen, autoimmune disorders and, 390–391 E2A B cell development and, 108, 109, 130 class switch recombination (CSR) and, 298, 312 transcription of immunoglobulin genes and, 93 V(D)J recombination and, 130 E2A-HLF fusion gene in pro-B cell lymphoblastic leukemias of adolescence, 353–354 E47, class switch recombination (CSR) and, 312 E5.2, antibody production and, 497–499 Ea, V(D)J recombination and, 130 early B cell factor (EBF), 108, 109 “editing” of receptors, in B cell development, 115–116, 145–146, 146 egg white lysozyme (xEL) studies, antibody production and, 500–501 11q23 abnormalities in MLL and, 350–351 ELISA analysis phage display, 518–519 translocus mice and human Mab and, 552–553 embryonic development, hematopoiesis during, 101–103, 102 embryonic stem (ES) cells, translocus mice and human Mab and, 551 encephalomyelitis syndrome, X-linked (Bruton’s) agammaglobulinemia (XLA) and, 403–404, 403 endoplasmic reticulum (ER) (See also plasma cells) ER-associated degradation (ERAD) pathway in, 267–268 ER degradation enhancing mannosidase like molecule (EDEM) in, 268 immunoglobulin assembly and secretion, 263, 264–266, 264 plasma cell generation and differentiatiion in, 268–270, 269 Russell bodies and, 268 stress response and Ig production in, 269–270 unfolded protein response (UPR) in, 269–270, 270 enhancers of Ig genes, lower vertebrate immunoglobulin genes and, 426–427, 426
Epstein-Barr virus (EBV), 166, 211, 389, 390 Epstein-Barr virus-induced chemokine (ELC), mucosal B cells and, 235 ER-associated degradation (ERAD) pathway, 267–268 ERCC1, class switch recombination (CSR) and, 318–319 erythrocyte development, B cell development and, 102 erythropoietin (EPO) molecules/receptors, B cell receptor (BCR) and, 161 ets class switch recombination (CSR) and, 298 transcription of immunoglobulin genes and, regulation by, 88–89, 93 Em intronic enhancer class switch recombination (CSR) and, 291–292, 308–309 lower vertebrate immunoglobulin genes and, 426–427 transcription of immunoglobulin genes and, 83–91, 84, 129 EU12, B cell development and, 146 evolutionary studies adaptive immune receptor origin and, 427 antibody production and, 505 of artiodactyls (hoofed, grazing animals), 440–444 autoimmune disorders and, 381 of avians/birds and GALT, 433–436 B cell receptor (BCR), 481–483, 481 of cats, 444 class switch recombination (CSR) and, 310, 320 of dogs, 444 gut-associated lymphoid tissues (GALT) and, 417, 445 of horse, 444–445 human IgH and, 1 immunoglobulin genes in higher vertebrates and, 433–448 immunoglobulin genes in lower vertebrates (See immunoglobulin genes in lower verterbates) lagomorphs (rabbits and hares), 436–440 somatic hypermutation (SHM) and, 320, 335, 484–485 T cell receptors (TCR), 481–483, 481 ur-V gene in, 483–486 V(D)J recombination and, 473 VH region, human IgH, 10–12 Vk genes of human/mouse and, 33 “exaptation,” V(D)J recombination and, 473, 474, 480 exons, recombination activating genes (RAG1/RAG2), 68 exonuclease I, class switch recombination (CSR) and, 318 extraosseous PCT (PCT-E), 372–373
F Fab domain, antibody production and, 493–494 Fab vs. scFv libraries, 521–522 Fas, subset development in B cells, 157–158
573
Index Fc and complement receptors, 275–287 affinity maturation and, 279–280, 279 antibody-forming cells (AFC) and, 284 antibody production and, 493–494 antigen localization to FDC, co-receptor signaling vs., 281–285 autoimmune disorders and, 284–285, 286 B cell receptors (BCR) and, 277, 278, 285–286 B1 cells and, 282 bone marrow and, 284 C1q and, 275, 276, 281 C2 and, 281 C3 and, 275, 276, 280–281 C4 and, 275, 276, 281 C5–C9 and, 280 CD21 and, 281–286 CD21 deficiency and, 276, 277 CD35 and, 281–286 CD35 deficiency and, 276, 277 complement and complement receptor deficiency and, 276 complement control proteins (CCP) in, 281 complement-dependent cytotoxicity (CDC) and, 542 complement receptors in, 280–281 complement vs. FC receptors in, opposing actions of, 285–286 CR1 and CR2 in, 275, 276 decay activating factor (DAF) regulator in, 281 FC receptors in, 276–280 affinity maturation and, 279–280, 279 calcium-dependent processes and, 278, 278 follicular dendritic cells (FDC) and, 279–280 germinal centers (GC) and, 279–280, 279 Ig enhancement and support in, 280 ITIM pathways and, 276–279 MAP kinases and, 278 memory response and, 280 RIIB expression and signaling in, 276 FcgRIIB deficiency and, 275–276 FcgRIIB in, 275 follicular dendritic cells (FDC) and, 275, 279–281, 285–286 germinal centers (GC) and, 279–280, 283, 285–286 herpes simplex virus 1 (HSV 1) study and, 276, 277 humoral response and complement influence on, 280–281 Ig enhancement and support in, 280 IgG and, 280 immunodeficiency diseases and, 284–286 innate immunity and, 462 ITIM pathways and, 276–279 membrane cofactor (MCP) regulator in, 281 memory B cells and, 284 memory response and, 280 naive B cell activation and, 282–283 plasma cells and, 284 self-reactive B cells, negative selection of, 284–285
SHIP and, 277–278 stages of B cell development and complement receptor requirements, 282–285, 283 FC receptors (See Fc and complement receptors) FCgIIRs, 275 in antibody production and response, 195 antigen receptor signaling regulation and, 181, 182 CD19 and, 176 deficiency of, 275–276 FcRH, B cell development and, 147 FcRL, B cell development and, 147 FcRX, B cell development and, 147 Fearon, Douglas T., 171 Ferrando, Adolfo, 349 FGFR3 and MMSET in multiple myeloma, 356 fibrinogen-related proteins (FREP), 486 fibroblast growth factor (FGF), pluripotent hematopoietic stem cells (pHSC) and, 103 fibronectin and migration of B cells, 214 fish (See also immunoglobulin genes in lower vertebrates) bony fish, Ig heavy chain genes and IgM and IgD, 419–422, 421 class switch recombination (CSR) and, CH gene organization and, 310 complementarity-determining regions (CDRs) in cartilaginous fish, 418 gene multiplicity in, 417–419 IgM in cartilaginous fish, 417–419 IgNAR gene in cartilaginous fish, 417–419 IgW in cartilaginous fish, 417–419 IgX in cartilaginous fish, 417–419 immune response in, 453–454 lymphoid organs in, 452–453 membrane receptor expression and secreted Ig, 419–422 transitional arrangement of recombining elements, 422 V(D)J recombination and, 453–454, 457 flk tyrosine kinases, B cell development and common lymphoid precursors (CLP), 106 flk-1, pluripotent hematopoietic stem cells (pHSC) and, 105 folding, immunoglobulin assembly and secretion, 261–262, 263 follicle-associated epithelium (FAE), mucosal B cells and, 224, 226, 229 follicular B cell lymphoma (FBL), mouse B cell lineage lymphomas and, 370, 374 follicular dendritic cell (FDC) in antibody production and response, 188, 191, 192, 194, 195 antigen localization to, co-receptor signaling vs., 281–285 class switch recombination (CSR) and, 290 Fc (FcgRIIB) and complement receptors in, 275, 279–281, 285–286 immune response of B cells and, 340–341 memory and memory B cells in, 251 migration of B cells and, 209 mucosal B cells and, 229–231, 230 follicular lymphoma, 352–353, 370
FR1, autoimmune disorders and, 388–390, 389 framework regions (FR), 342 FREB, B cell development and, 147 Fredrickson, T.N., 374 frog, class switch recombination (CSR) and, CH gene organization and, 310 FTY720 and stopping of B cell migration, 209–210, 216 fucosyltransferase T-VII, migration of B cells and, 211 FvE5.2, antibody production and, 501
G g-H2AX class switch recombination (CSR) and, 319 somatic hypermutation (SHM) and, 335 G protein coupled receptor (GPCR) migration of B cells and, 204 subset development in B cells, 156, 204 galectin 1, B cell development and, 111, 145 GATA-1 genes, in zebrafish (Danio rerio) and, 459 GATA-2 genes B cell development and, 107 in lower vertebrate immunoglobulin genes and, 428 in zebrafish (Danio rerio) and, 451–452, 459 Geha, Raif S., 403 gemtuzumab ogogamicin, 541, 541 gene conversion, VH region, human IgH, 11, 12 gene expression profiling, mouse B cell lineage lymphomas and, 374–375, 374, 375 GeneCards, immunoglobin l genes (IGL), human/mouse, 38 genetic engineering, monoclonal antibodies and, 533–534 Genome Database (GDB), immunoglobin l genes (IGL), human/mouse, 38 germinal center (GC) (See also high affinity antibody) affinity-based selection after GC reaction ends in, 344 affinity maturation in, 342–344 in antibody production and response, 187, 189–197, 193 antibody-forming cells (AFC), 345 antibody–secreting cells (ASCs) and, homing of, 213–215 Burkitt lymphoma (BL) and, 375 centroblasts and centrocytes in, 194 class switch recombination (CSR) and, 289–291, 290 differentiation and dispersion of B cells in, 231–232 exit of B cells froom, 232 Fc (FcgRIIB) and complement receptors in, 279–280, 283, 285–286 formation of, molecular interactions in, 229–231 high affinity antibody and, 339–344 immune response of B cells and, 340–344 memory and memory B cells in, 248, 249, 345
574 germinal center (GC) (continued) mouse B cell lineage lymphomas and, 365 mucosal B cells, IgA and, 234 organization of, 192–194 positive selection and plasma cell induction in, 231–232 precursor cell seeding in, 345 primary follicles in, 192–193, 230 proliferation, hypermutation, selection in, 192–197 secondary follicles and, 193–194 somatic hypermutation (SHM) and, 330 T cells in centrocyte selection and maintenance of, 195–197 germinal center dendritic cells (GCDC) and mucosal B cells, 231 germline transcription (GT) 3¢ IgH enhancers in, 293–295, 316 C/EBPb and, 292, 293 CD40 and, 292, 293, 297 Ce in, 292–293 class switch recombination (CSR) and, 289, 291, 312 coordinated regulation of transcription, recombination, replication in, 300 CREB and, 293 Ca regulation in, 293 Cg1 in, 292–293, 312 Cg2a in, 293, 312 Cg2b in, 292 Cg3 in, 292 Em and, 291–292 human I promoters in, specificity of, 293 I promoter interaction with 3¢ IgH enhancers in, 295–297 IFN-g and IFN-a in, 293 IgA and IgG in, 293 IL4 and IL13 in, 292 insulin receptor substrate (IRS) in, 292 interleukins in, 292 IRF and, 293 Im GT and, 291–292 JAK in, 292, 293 mechanisms for 3¢ IgH regulatory region mediated regulation of, 295–300, 296 NFkB and, 292, 297 nonhomologous end joining (NHEJ) and, 297 non-Ie promoters and cytokine inducibility in, 292–293 polarized effect of 3¢ IgH regulatory region and, 295 proximal cis regulatory elements for, 291–293 PU-1 and, 292 STAT in, 292, 293 switch sequences as transcriptional stimulatory elements in, 291 TGF-b and, 293 TGF-b inhibitory element (TIE) in, 292 transcription factors controlling 3¢ IgH regulatory elements in, 297–299 V(D)J recombination and, 128, 131, 292 vasointestinal protein (VIP) and, 293 glycam 1 and migration of B cells, 205
Index glycosylation, immunoglobulin assembly and secretion, 262–263 glycosyl-phosphatidyl inositol (GPI) anchor, B cell receptor (BCR) and, 166 golgi organs, immunoglobulin assembly and secretion, transport to, 267 Goodpasture’s syndrome, 275, 385 graft vs. host disease, 541 Graves’ disease, 385, 391 Grb2, antigen receptor signaling regulation and, 177 group and subgroups, immunoglobin l genes (IGL), human/mouse, 38 growth factors B cell development and, 103 pluripotent hematopoietic stem cells (pHSC) and, 103 growth hormone deficiency XLA, 407 gut-associated lymphoid tissues (GALT) (See also immunoglobulin genes in higher vertebrates), 443–448 in avians/birds, 433–436 B cell traffic in, naive and primed, 235–236, 236 evolutionary studies of, 417 gut lamina propria and, 236 immunoglobulin genes in higher vertebrates and, 445 in lagomorphs (rabbits and hares), 438–439 memory and memory B cells in, 249 mucosal B cells and, 224, 225, 226–227, 227, 231, 233, 234–236, 238 mucosal homing molecules beyond the gut, 236–237
H H chain locus, B cell development and, allelic exclusion at, 113, 114 H2AX histone, V(D)J recombination and, 77 hairpin end opening, V(D)J recombination and, 75 HAPPY mapping, human IgH and, 4 Hardie, Deborah L., 187 hares (See lagomorphs) Hashimoto disease, 385 heavy chain antibodies (HCAb), artiodactyls (hoofed, grazing animals) and, 440–441, 441 HEB, V(D)J recombination and, 130 Hedgehogs (Hhs), pluripotent hematopoietic stem cells (pHSC) and, 103 HEL and anti-HEL, 117 antibody production and, 494–505, 497, 500 autoimmune disorders and, 382 helper T cells, migration of B cells and, 203 hemangioblasts, pluripotent hematopoietic stem cells (pHSC) and, as early progenitors of, 104–105 hematopoeitic cell development aorta-gonad mesonephros (AGM) in, 450–451, 452 embryonic stages in, 101–103, 102 intermediate cell mass (ICM) stage in, 451–452, 460
transcription of immunoglobulin genes and, 93 vertebral, 450–452, 450 zebrafish (Danio rerio) and, 450–452, 450, 451 hematopoietic progenitor kinase (HPK1), B cell receptor (BCR) and, 165 hematopoietic stem cells (HSC), B cell development and, 141 hemoglobin development, B cell development and, 102 hemolytic anemia, 384, 385 hen egg lysozyme (See HEL) Hendershot, Linda M., 261 herpes simplex virus (HSV) Fc (FcgRIIB) and complement receptors in, 276, 277 monoclonal antibodies and (HSV863/HuFd79D) and, 540, 541 heteroclitic binding, antibody production and, 500 high affinity antibody (See also germinal center), 339–348 activation-induced deaminase (AID), 342 affinity-based selection after GC reaction ends in, 344 affinity maturation and, 339, 341–342 antibody-forming cells (AFC), 340, 345 B cell immune response and, 339–341 CD40 and, 342, 344 competition between cells and, 343–344 complementarity-determining regions (CDRs) in, 342–344 framework regions (FR) and, 342 germinal center (GC) and, 339–344 hybridoma technology and, 339 memory B cells and, 339, 340–341, 345 mutation and, 339 phage display and, selection of, 523–524 precursor cell seeding in GC and, 345 “selective advantage” concept in, 343 V regions and, 339 high endothelial cell (HEC) deficiency, migration of B cells and, 205 high endothelial venules (HEVs) migration of B cells and, 203–210, 205 mucosal B cells and, 234–235 HIN and HIX sites, in V(D)J recombination, 474, 476, 476 histiocyte-associated (HA) DLBCL, 370–371 histone acetyltransferases (HATs), V(D)J recombination and, 131 histone deacetylase (HDAC) class switch recombination (CSR) and, 291 V(D)J recombination and, 131 HIV-1 and anti-HIV-1 peptide, antibody production and, 494–495 HLA-DR B cell development and, 142 plasma cells, memory and, 253 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 HMG1/HMG2 DNA bending protein, V(D)J recombination and, 65–68, 128, 131
Index homing of pluripotent hematopoietic stem cells (pHSC), 104 Honjo, Tasuku, 307 horizontal or lateral transfer, V(D)J recombination and, 473–474 hormonal activity and B cell development, 107 horror autotoxicus concept, autoimmune disorders and, 381 horse, immunoglobulin genes in higher vertebrates and, 444–445 HOXB4, pluripotent hematopoietic stem cells (pHSC) and, 103 Hsu, Ellen, 473 human anti-chimeric antibody (HACA) response, 534 human anti-humanized antibody (HAHA) response, 540 human anti-mouse antibodies (HAMA) response, 511, 533, 534 human artificial chromosome (HAC) studies, translocus mice and human Mab and, 552, 553 Human Genome Organization (HUGO) Nomenclature Committee, immunoglobin l genes (IGL), human/mouse, 38 human immunoglobin heavy chain (IgH), 1–17, 3 allotyping of, 15 B cell development and, 141–144 CH (constant) region in, 1, 12–15, 13 polymorphisms of, 14–15, 14 structure of genes in, 13–14, 13 switch region in, 14 transcription enhancers in, 13 cosmid cloning in, 2 DH (diversity) region in, 1 chromosome 15 and 16 segments of, 5–6, 6 evolution of, 10–11 organization of, 5 evolutionary studies of, 1 HAPPY mapping in, 4 JH (joining) region in, 1 mapping of Vh locus in, 2 mouse B cell lineage lymphomas and, loci of, 366 pulse field gel electrophoresis (PFG) in, 2 southern hybridization in, 2 structural analysis of, 1–2 V (variable) region in, 1 VDJ recombination in, 1 VH region in, 1, 2–7 5¢ regulatory regions in, 8–9, 8 analysis of, 7–10 B cell and immune response in, 9–10 chromosome 14 and, evolution of, 10 chromosome 15 and 16 segments of, 5–6, 6 classification of segments in, 4, 4 evolution of, 10–12 nonimmunoglobulin genes in, 6–7 open reading frames (ORF) in, 3–4 orphon, evolution of, 10–11 overexpression of, 10 polymorphism of, 4–5
primary repertoire of, 9–10 pseudogenes and gene conversion in, 11, 12 recombination signal sequence in, 9 segment usage and repertoire formation in, 10 subgroups and families of, 7–8, 7 total number of segments, 3–4 yeast artificial chromosome (YAC) studies in, 2 human immunoglobin light chain (IgL) locus, 141–144 mouse B cell lineage lymphomas and, loci of, 366 humanization of monoclonal antibodies, 511, 533–545 humoral immunity/response complement influence on, 280–281 plasma cells, memory and, 252 hybrid joints (HJ), 482 recombination activating genes (RAG1/RAG2), 70 V(D)J recombination and, 61, 70 hybridoma technology for monoclonal antibodies, 511, 512 high affinity antibody and, 339 monoclonal antibody and, 511, 512, 519–520, 533 phage display vs., 519–520 HyHEl complexes, antibody production and, 494, 504 hyper-IgM syndrome (HIGM) (See also immunodeficiency diseases), 313, 410–411 activation-induced deaminase (AID) gene in, 410–411 CD154/CD40 in, 410 class switch recombination (CSR) and, 290 TNFSF5 in, 410 hypermutation (See somatic hypermutation) hypersensitive sites (HS), class switch recombination (CSR) and, 309
I I exon transcription in class switch recombination (CSR), 291, 312 I promoters, interaction with 3¢ IgH enhancers in, 295–297 ibritumomab tiuxetan, 541 ICAM (ICAM-1 and ICAM-2) migration of B cells and, 207, 208, 210, 213 mucosal B cells and, 235 subset development in B cells, 157 Icaros, B cell development and, 108, 109 iccosomes, 231 ICOS, class switch recombination (CSR) and, 290, 287 Id2 natural killer cell development and, 108 T cell development and, 109 IDDM, 382 IDENTIFICATION concept, Immunogenetics Database (IMGT) and, 37 IFN-a, class switch recombination (CSR) and, 293
575 IFN-g, class switch recombination (CSR) and, 290, 293 Ig-like transcript (ILT), 182 Ig promoters, transcription of immunoglobulin genes and, 83–85 Igl, V(D)J recombination and, allelic exclusion and, 136 IgA, 248–249 B cell development and, 149 cell cycle and, mucosal B cells and, 224 class switch recombination (CSR) and, 293 IgA deficiency (IgAD), 149 immunoglobulin assembly and secretion, 267 migration of B cells and, 215 mucosal B cells and, 223, 224, 226–227, 226 class switch recombination and, 232–234 subclass distribution in, 227 stimuli promoting production of, 233–234 Iga B cell development and, 106, 107, 109 common lymphoid precursors (CLP) and, 106 agammaglobulinemias and, 410 Igb B cell development and, 106, 109 common lymphoid precursors (CLP) and, 106 IgD, 238, 248–249 B cell development and common lymphoid precursors (CLP), 106 bony fish and, Ig heavy chain genes in, 419–422, 421 Fc (FcgRIIB) and complement receptors in, 282 IgE, 248–249 IgG, 238 class switch recombination (CSR) and, 287, 290, 293 Fc (FcgRIIB) and complement receptors in, 280 migration of B cells and, 215 IgH (See human immunoglobulin heavy chain) Igk, V(D)J recombination and, 135, 145 IgL (See human immunoglobulin light chain) IGLLl mutations, agammaglobulinemias and, 409, 409 IgM, 248–249 autoimmune disorders and, 389–390 in bony fish, Ig heavy chain genes in, 419–422, 421 in cartilaginous fish, 417–419 developmental control of secretion of, 266, 266 Fc (FcgRIIB) and complement receptors in, 282 immunoglobulin assembly and secretion, 267 mucosal B cells and, 223, 224, 228 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 407–408 in zebrafish (Danio rerio), 453, 454–455 IGMT database, 13 IgNAR gene in cartilaginous fish, 417–419 IgW in cartilaginous fish, 417–419
576 IgX in cartilaginous fish, 417–419 IkB kinase (IKK) complex, 94–96 class switch recombination (CSR) and, 290, 297 subcellular localization and, 94–95 transcription of immunoglobulin genes and, 95–96 Ikaros gene, zebrafish (Danio rerio) and, 454, 456 IL-2, 106, 109 IL-3, 104 IL-4, 292, 312 IL-6, 104, 255 IL-7 B cell development and, 106, 109–110, 119, 141, 148–149 common lymphoid precursors (CLP) and, 106 immunodeficiency diseases and, 148–149 pluripotent hematopoietic stem cells (pHSC) and, 104 V(D)J recombination and, 133 IL-10, 255 IL-13, 292 IL-15, 104 ileal Peyer’s patch (IPP), 443 IMGT database (See ImMunoGeneTics database) IMGT-ONTOLOGY, 37 immune inductive tissue compartments, mucosal B cells and, 223–226 immune response (See also germinal center; high affinity antibody; zebrafish immune system), 155, 339–341 activation-induced deaminase (AID), 342 affinity-based selection after GC reaction ends in, 344 affinity maturation in, 341–342 antibody-forming cells (AFC), 340, 345 B cell receptor (BCR) and, 166, 340 bone marrow and, 340 CD40 and, 342, 344 chemokines and, 340 clonal selection in, 345 complementarity-determining regions (CDRs) in, 342–344 follicular dendritic cells (FDC), 340–341 framework regions (FR) and, 342 germinal center (GC) in, 340–344 high affinity antibody in (See high affinity antibody) integrated view of, future directions in, 344–345 lagomorphs (rabbits and hares) and, secondary, 439 LPS in, 339–340, 339 memory B cells and, 340–341, 345 migration of B cells in, 340 NF-kB/rel proteins and, 93–96 overview of, 339–341 plasmablasts in, 340 precursor cell seeding in GC and, 345 primary vs. secondary response in, 345 “selective advantage” concept in, 343 spleen in, 340
Index translocus mice and human Mab and, 557–558, 557 zebrafish (Danio rerio) and, teleosts, 449, 453–454 immune type receptors (ITRs), V(D)J recombination and, 476 immune vs. nonimmune libraries, 521 immunization, phage display vs., 520–521 immunoblastic DLBCL, 370 immunodeficiency diseases, 148–149, 237, 403–416 l5 and, 148 activation-induced deaminase (AID) and, 313 acute lymphoblastic leukemia (ALL) and, 149–150 agammaglobulinemias, 148, 403–410 atypical XLA in, 405 B cell linker protein (BLNK) and, 410 CD79 in, 409, 410 g5/IGLLl mutations in, 409, 409 growth hormone deficiency XLA, 407 Iga mutations in, 410 ITAM in, 410 m heavy chain gene mutations in, 409 T cell function in XLA, 408 X-linked (Bruton’s) (XLA), 403–409, 404 Artemis and, 73, 74, 76, 148 B cell development and, 148–149 B cell receptor (BCR) and, 164 B lineage leukemia and, 149–150 BLIN cells and, 150 BLNK and, 148 bone marrow stromal cells and, 150, 150 BTK deficiency and, 148, 368 CD10 and, 149 CD19 and, 149 CD21 deficiency and, 276, 277 CD22 and, 178 CD35 deficiency and, 276, 277 class switch recombination (CSR) and, 290 common variable immunodeficiency (CVI), 149 complement and complement receptor deficiency and, 276 DNA-PKcs in, 76 Epstein-Barr virus (EBV) and, 166, 211, 389, 390 Fc (FcgRIIB) and complement receptors in, 284–286 FcgRIIB deficiency and, 275–276 hyper-IgM syndrome (HIGM), 290, 313, 403, 410–411 activation-induced deaminase (AID) gene in, 410–411 CD154/CD40 in, 410 TNFSF5 in, 410 IgA deficiency (IgAD), 149 IL7 and, 148–149 Ku70 and Ku80 in, 76 migration of B cells and, 213 murine models of B cell deficiency in, 411–412, 411 mutations in, 148, 403, 404 nonhomologous end joining (NHEJ) and, 73–77
recombination activating genes (RAG1/RAG2), 69, 148 self-reactive B cells, negative selection of, 284–285 X-linked agammaglobulinemia (XLA), 164 XRCC4 and DNA ligase IV, 76 Immunogenetics Database (IMGT database), 37–59 CLASSIFICATION concept in, 37–38, 38 Colliers de Perles representations in, 40, 41, 42, 43 DESCRIPTION concept in, 39–40 IDENTIFICATION concept in, 37 immunoglobin l genes (IGL), human/mouse, notation in, 37–40 JunctionAnalysis tool in, 40 PhyloGene tree map in, 55, 57 unique numbering in, 39–40 V-QUEST alignment tool in, 40 immunoglobin k (Vk) genes, human/mouse, 27–36 evolution of, 33–34 general features of, 27 sequencing of, 34 Vk genes, human, 27–30 bacterial artificial chromosome (BAC) studies in, 27 elucidation of, 27–28 evolution of, 33–34 general features of, 27 k protein classification in, 29 mapping of, 28, 28 orphons in, 28, 30 polymorphisms in, 30 pseudogenes, relics, repetitive elements in, 28–29 rearranged and expressed genes in, 29– 30 sequencing of, 34 V-J rearrangement in, 29–30 yeast artificial chromosone (YAC) studies in, 27 Vk genes, mouse, 30–33 bacterial artificial chromosome (BAC) studies in, 31, 33 elucidation of, 31 evolution of, 33–34 general features of, 27 genes and pseudogenes in, 31–32, 32 relics and orphons in, 33 sequencing of, 34 yeast artificial chromosome (YAC) studies in, 31 immunoglobin l genes (IGL) alleles in, 38 C regions in, 38 chain type of, 37 CLASSIFICATION concept in, 37–38, 38 configuration of, 37 DESCRIPTION concept in, 39–40 functionality of, 37 gene type in, 37 genes in, 38 group and subgroups in, 38 IDENTIFICATION concept in, 37
Index IMGT-ONTOLOGY system notation and, 37–40, 37 immunoglobin l genes (IGL), human, 37–59 chromosomal localization of IGL locus in, 40–41, 46 IGLC genes in, 41, 42–43, 46, 48, 49, 49 IGLC genes in, 46, 48, 49, 49 IGLJ genes in, 41, 46 IGLJ genes in, 41, 46 IGLV genes in, 41, 46, 48 leukemia, lymphoma, 40 number and repertoire of, 50, 50, 51, 52 organization of, 41–49, 44–45, 47 orphons in, 49, 49 polymorphisms, 49 immunoglobin l genes (IGL), mouse, 50–55 C genes in, 50–51, 52–55, 54–55, 56 chromosomal localization and, 50 J genes in, 50, 52–55, 56 number and repertoire of, 55, 56, 57 phylogenic tree of, 55, 58 pulse field gel electrophoresis (PFGE), 50 RFLP analysis of, 52, 55 strains of mice and, differences in V, 52 V genes in, 50, 51–52, 53, 54 J regions in, 38, 39, 39, 38 locus in, 38 open reading frame (ORF) in, 37 pseudogenes in, 37 rearranged (unproductive) genes in, 37 V regions in, 38, 39, 39, 41 V(D)J recombination and, 62 immunoglobulin assembly and secretion, 261–273 allelic exclusion and, 262 ATF6 and, 270 B cell receptors (BCR) and, 263, 265 BAP29 and BAP31 in, 265, 267 Bence-Jones proteins and, 266 BiP and, 264–265 Blimp1 and, 269, 270 C domain in, 261 CH1 domain in, 264–265 Cys213 and, 266 Cys575 in, 266 degradation of misfolded and unassembled subunits in, 267–268 differential fate of mm and ms during differentiation and, 263–264 endoplasmic reticulum (ER) and, 263–266, 264 ER-associated degradation (ERAD) pathway in, 267–268 ER degradation enhancing mannosidase like molecule (EDEM) in, 268 Fc (FcgRIIB) and complement receptors in, 280 folding and assembly in, 261–262, 263 free light chain (LC) secretion and, 266 glycosylation and, 262–263 golgi organs and, transport to, 267 heavy chain in, 261–273 IgA and, 267 IgM and, 267
IgM secretion and, developmental control of, 266, 266 J chain in polymerization and transcytosis in, 267 lectin and, 267–268 light chain in, 261–273 mechanisms of, 261–264 monitoring assembly of BCRs and, 265 ms2LC2 complexes and, 266 numbers of, in humans, 261 PERK in, 270 plasma cell differentiation and, 268–270, 269 pre-B cells and, 262 pre-B receptor transport in, 265–266 quality control in, 264–267, 264 regulation of, 269 repertoire of, 261 Russell bodies and, 268 stress response in ER and, 269–270 surrogate light chain (SLC) in, 262, 265–266 unfolded protein response (UPR) in, 269–270, 270 V domain in, 261, 262 XPB1 and, 269, 270 immunoglobulin genes in higher vertebrates (See also gut-associated lymphoid tissues; immunoglobulin genes in lower vertebrates), 433–448 in artiodactyls (hoofed, grazing animals), 440–444 antibody diversity generation in, 442–443, 442 CH genes in, 443–444 heavy chain antibodies (HCAb) in, 440–441, 441 ileal Peyer’s patch (IPP) in, 443 k light chain genes in, 441–442 l light chain genes in, 441–442 lymphopoiesis and Ig gene diversification in, 443 V, D, J gene segments in, 441 in avians/birds and GALT, 433–436 antibody diversity generation in, 435–436, 435 B cell receptors (BCR) in, 435–436 bursa of Fabricius in, 330–436 CH genes in, 436 chicken Ig genes, organization of, 433–434 complementarity-determining region (CDR) in, 435 lymphopoiesis in, limited, 434–435 V(D)J recombination in, 433–435 in cats, 444 in dogs, 444 gut-associated lymphoid tissues (GALT) and, 433, 445 in horse, 444–445 in lagomorphs (rabbits and hares), 436–440 antibody diversity in, 437–440, 438 B cell development in, 437–440 CH genes in, 439–440, 439 enhancer regions in, 440 gut-associated lymphoid tissues (GALT) in, 438–439
577 k light chain gene segments in, 437 l light chain gene segments in, 437 polymorphisms of VHa in, 437 secondary immune response in, 439 V(D)J recombination and, 436–438 immunoglobulin genes in lower vertebrates (See also immunoglobulin genes in higher vertebrates), 417–432 adaptive immune receptor origin and, 427 amphibians and reptiles, class switching recombination (CSR) in, 422–423 bony fish and, Ig heavy chain genes and IgM and IgD, 419–422, 421 bony fish, membrane receptor expression and secreted Ig, 419–422 cartilaginous fish and, gene multiplicity in, 417–419 complementarity-determining regions (CDRs) in cartilaginous fish, 418 enhancers of Ig genes in, 426–427, 426 fleshy-finned fish, isotype diversity in, 422 gut-associated lymphoid tissues (GALT) and, 417 IgM in cartilaginous fish, 417–419 IgNAR gene in cartilaginous fish, 417–419 IgW in cartilaginous fish, 417–419 IgX in cartilaginous fish, 417–419 ITIM in, 428 jawless vertebrates and, 427–428 killer-cell immunoglobulin-like receptors (KIRs) in, 428 light chain genes and, diverse structures and organization in, 423–425, 424 lobe-finned fish, transitional arrangement of recombining elements, 422 phylogenic relationships of immunoglobulin classes in vertebrates, 420 promoters of Ig genes in, 425–427 protochordates, V region diversification in, 428–429 T cell receptor (TCR) genes and, 417, 427–428, 428 transcriptional control of, 425–427 V chain bifunctional genes (VCBP) in, 429 immunoreceptive tyrosine-based activiation motif (ITAM) agammaglobulinemias and, 410 B cell development and, 147 B cell receptor (BCR) and, 162–166 immunoreceptor tyrosine-based inhibitory motif (ITIM) antigen receptor signaling regulation and, 182 B cell development and, 147, 177 Fc (FcgRIIB) and complement receptors in, pathways for, 276–279 innate immunity and, 462 lower vertebrate immunoglobulin genes and, 428 inflammatory bowel disease (IBD), 228 inflammatory processes migration of B cells and, 213 NF-kB/rel proteins and, 93–96 infliximab, 541 innate defense V gene theory, 485–486
578 innate immunity, 460–464 cellular effectors of, 461 chemokines in, 464 complement system, 462 cytokines, 463–464 cytotoxic cells, 462 dendritic cells in, 460 innate defense V gene theory and, 485–486 interferons (IFN) in, 464 interleukins in, 463–464 ITIM and, 462 killer cell inhibitory receptors (KIR) in, 462 lymphotoxin in, 464 mannose-binding lectin (MBL) in, 460 MBL-associated serine proteases (MASPs) in, 462 natural killer cells in, 460 novel immune type receptors (NITR) in, 462 pattern recognition receptors in, 463 phagocytic (myeloid) cells in, 461–462 Toll-like receptors (TLR) in, 460, 463 tumor necrosis factor (TNF) in, 464 in zebrafish (Danio rerio) and, 460–464 inositol triphosphate (IP3), B cell receptor (BCR) and, 164 insulin receptor substrate (IRS), class switch recombination (CSR) and, 292 integrins B cell development and, 103 chemokine triggering of, 205–207 migration of B cells and, firm adhesion stage in, 207 interferon g, class switch recombination (CSR) and, 312 interferons (IFN), innate immunity and, 464 interleukins (See also IL-x) class switch recombination (CSR) and, 292, 312 innate immunity and, 463–464 memory and memory B cells in, 251–252 plasma cells, memory and, 254, 255 intermediate cell mass (ICM) stage, hematopoiesis and, 451–452, 460 intronic enhancers, in lagomorphs (rabbits and hares), 440 introns, recombination activating genes (RAG1/RAG2), 68 inverse PCR (IPCR) studies, mouse B cell lineage lymphomas and, 373 inverted terminal repeats (ITRs), V(D)J recombination and, 473, 474 IP10 and migration of B cells and, 215 IRE1, 255 Ire1a/b, immunoglobulin assembly and secretion, 270 IRF, class switch recombination (CSR) and, 293 IRF4, 255 B cell development and, 113 transcription of immunoglobulin genes and, 89 IRF8, B cell development and, 113 isolated lymphoid follicles (ILFs), mucosal B cells and, 224, 227
Index isotype diversity, in fish, 422 isotype switching (See class switch recombination (CSR) isotypic exclusion, 127–140 ITAC and migration of B cells, 215 ITAM (See immunoreceptive tyrosine-based activiation motif) ITIM (See immunoreceptor tyrosine-based inhibitory motif) Im GT, class switch recombination (CSR) and, 291–292
J J chain/J regions beyond 12/23 (B12/23) rule in, 64–65 immunoglobin l genes (IGL) human and, 41, 46 human/mouse, 38, 39, 39, 40 mouse and, 50, 52–55, 56 immunoglobulin assembly and secretion, in polymerization and transcytosis in, 267 mucosal B cells and, 223, 224, 228–229 transcription of immunoglobulin genes and, 83, 91 translocus mice and human Mab and, 547–558 12/23 rule in, 62, 63, 64–65, 128 V(D)J recombination and, 62 JAK, class switch recombination (CSR) and, 292, 293 JAK-STAT activation, pluripotent hematopoietic stem cells (pHSC) and, 103 jawless vertebrates, lower vertebrate immunoglobulin genes and, 427–428 JH (joining) region, human IgH and, 1 junctional adhesion molecules (JAMs), migration of B cells and, 207 JunctionAnalysis tool, Immunogenetics Database (IMGT) and, 40
K k intronic and 3¢ enhancers, transcription of immunoglobulin genes and, 85, 86, 88–89 k light chain gene segments in artiodactyls (hoofed, grazing animals), 441–442 in lagomorphs (rabbits and hares), 437 kL chain rearrangement, B cell development and, 114–115 Kearney, John F., 155 kelch repeats, recombination activating genes (RAG1/RAG2) and, 478–479 killer-cell immunoglobulin-like receptors (KIRs), 428, 462 Kincade, Paul, 101 Kinoshita, Kazuo, 307 Knight, Katherine L., 433 Krangel, Michael S., 127 Ku70, nonhomologous end joining (NHEJ) and, 73–74, 76 Ku80, nonhomologous end joining (NHEJ) and, 73–74, 76
L l5, 109, 111, 127, 142, 143–144, 148 B cell development and, 143–144 common lymphoid precursors (CLP), 106, 107 IGLLl mutations and, agammalobulinemias and, 409, 409 immunodeficiency diseases and, 148 lL chain rearrangement, B cell development and, 114–115 l enhancers, transcription of immunoglobulin genes and, 85, 86, 88–89 l (lambda) light chain genes in artiodactyls (hoofed, grazing animals), 441–442 in lagomorphs (rabbits and hares), 437 L chain genes, B cell development and, 107 L selectins, migration of B cells and, 204–205 lagomorphs (rabbits and hares), 436–440 antibody diversity in, 437–440, 438 B cell development in, 437–440 CH genes in, 439–440, 439 enhancer regions in, 440 gut-associated lymphoid tissues (GALT) in, 438–439 immunoglobulin genes in higher vertebrates and, 436–440 k light chain gene segments in, 437 l light chain gene segments in, 437 polymorphisms of VHa in, 437 secondary immune response in, 439 somatic hypermutation (SHM) in, templated and nontemplated mutation in, antigenactivated, 329 V(D)J recombination in, 436–438 Lamm, Michael E., 223 Lanning, Dennis, 433 late SV40 factor (LSF), class switch recombination (CSR) and, 291 lck, subset development in B cells, 156 LeBien, Tucker, 141 lectin B cell development and, 111 immunoglobulin assembly and secretion, 267–268 LEF-1, V(D)J recombination and, 130 Lefranc, Gerard, 37 Lefranc, Marie-Paule, 37 leukemia (See B cell leukemias and lymphomas) leukocyte function-associated (LFA), mucosal B cells and, 235 in antibody production and response, 195 migration of B cells and, 205, 207, 208, 210, 212, 214 leukocyte Ig-like receptor (LIR), 182 Lewis, Susanna M., 473 LIF, translocus mice and human Mab and, 552 Lig4 (See DNA ligase IV) light chain genes, lower vertebrate immunoglobulin genes and, diverse structures and organization in, 423–425, 424
Index linker of activated T cell (LAT), B cell receptor (BCR) and, 165 LinkOut, immunoglobin l genes (IGL), human/mouse, 38 lipopolysaccharide (LPS), mucosal B cells and, 227 liver-derived B cells, 102, 141, 143 lobe-finned fish, transitional arrangement of recombining elements (See also fish), 422 localization of B cells, 156 LocusLink, immunoglobin l genes (IGL), human/mouse, 38 Look, A. Thomas, 349 looping of DNA, transcription of immunoglobulin genes and, 89–90 loss of imprinting (LOI) phenomenon, mouse B cell lineage lymphomas and, 375 lox, translocus mice and human Mab and, 552 LPS, 339–340 class switch recombination (CSR) and, 312, 320 LR1, class switch recombination (CSR) and, 291 LTa1b2 migration of B cells and, 213, 215 mucosal B cells and, 231 LTNa/b, class switch recombination (CSR) and, 290 lupus (See systemic lupus erythematosus) lymph nodes (See also lymphatic organs) B cell development and, 155, 187, 188–189 memory and memory B cells in, 249, 250 migration of B cells and, 203, 204, 205 mucosal B cells and, 225–226 peripheral lymph node addressin (PNAd) in, 205 lymphoblastic leukemia, 349–352, 350 lymphocyte choriomeningitis virus (LCMV), 253 lymphocyte development, 102 lymphoid follicles, migration of B cells and, 209–210 lymphoid organs antigen–presenting cells (APCs) and, 223–224, 226 apoptosis suppression during development of, translocations associated with, 352–353 bronchus-associated lymphoid tissue (BALT) in, 224–225, 236–237 follicle-associated epithelium (FAE) and, 224, 226, 229 gut-associated lymphoid tissues (GALT) and, 224, 225, 226–227, 227, 231, 233, 234–236, 238, 249, 443–448 isolated lymphoid follicles (ILFs) in, 224, 227 lymph nodes in, 225–226 memory and memory B cells in, 249, 250 migration of B cells to, 203–221, 204 mucosa-associated lymphoid tissue (MALT) and, 223, 225, 227–232, 238 nasopharynx-associated lymphoid tissue (NALT) in, 224–225, 236–237, 249
Peyer’s patches and, 223, 224–230 somatic hypermutation (SHM) and, 330 zebrafish (Danio rerio) and, teleosts, 452–453 lymphoid precursor, proliferation of, translocations associated with, 354–355 lymphoid tumors recombination activating genes (RAG1/RAG2), 71 V(D)J recombination and, 71 lymphomas (See B cell leukemias and lymphomas; mouse B cell lineage lymphomas) lymphopoiesis in artiodactyls (hoofed, grazing animals), Ig gene diversification in, 443 in birds/avians, limited, 434–435 lymphotoxin in antibody production and response, 195 class switch recombination (CSR) and, 290 innate immunity and, 464 mucosal B cells and, 229–231 Lyn, B cell receptor (BCR) and, 162, 163, 174, 177
M ms2LC2 complexes and, 266 mH chain gene, 107, 114 m heavy chain gene mutations, agammaglobulinemias and, 409 M cells (See memory and memory B cells) Mab231 murine IgG2a monoclonal antibody, 492, 492 MacLennan, Ian C.M., 187 macrophage inflammatory protein (MIP), class switch recombination (CSR) and, 290 MAD-2, autoimmune disorders and, 389–390 MAdCAM 1 migration of B cells and, 205, 208, 213, 215 mucosal B cells and, 234–235, 236 MAIDS syndrome, mouse B cell lineage lymphomas and, 371 Maizels, Nancy, 327 major histocompatibility class (MHC) in antibody production and response, 195 B cell development and common lymphoid precursors (CLP), 106 memory and memory B cells in, 249 mucosal B cells and, 237, 238 zebrafish (Danio rerio) and, 460 mannose-binding lectin (MBL), innate immunity and, 460 mannosidase, ER degradation enhancing mannosidase-like molecule (EDEM) in, 268 mantle cell lymphoma/myeloma, 355–356, 366 MAP kinases antigen receptor signaling regulation and, 171, 175, 178, 182 Fc (FcgRIIB) and complement receptors in, 278 MAPK/JNK pathways and class switch recombination (CSR), 290
579 mapping of antibody–antigen interactions, mutagenesis and, 496–499, 497, 498 marginal cell lymphomas, 352, 353 marginal zone (MZ) B cells, of spleen, 155, 172–176 in antibody production and response, 188–189 antibody–secreting cells (ASCs) and, homing of, 213–215 migration of B cells and, 209–211 marginal zone lymphoma (MZL), 369–370, 374 Mariuzza, Roy A., 491 Marks, James, 511 matrix attachment regions (MARs), transcription of immunoglobulin genes and, 84, 86, 90, 92–93 Matsuda, Fumihiko, 1 mBL-associated serine proteases (MASPs), innate immunity and, 462 MCTD, 393 MDC, migration of B cells and, 210 MECA79, migration of B cells and, 205 megakaryocyte development, B cell development and, 102 Melchers, Fritz, 101 membrane (M) cells, 238 mucosal B cells and, 224, 229, 230 membrane co-factor (MCP) regulator, Fc (FcgRIIB) and complement receptors in, 281 membrane receptor expression and secreted Ig, in bony fish, 419–422, 419 memory and memory B cells, 238, 247–259 adaptive memory in, 254–255 affinity maturation and, 248 antibody production and response, 188, 203 antibody–secreting cells (ASCs) and, homing of, 213–215 antigen dependency vs. lifespan in, 251 autoimmune disorders and, 248 B cell receptors (BCR) and, 248–249, 251–252 B220 and, 248 Bcl and, 251 bone marrow and, 249, 250, 251 CD8 and, 252 CD19 and, 248, 253 CD27 and, 248–249, 253 CD40 and, 255 CD70 and, 249 chemokines and, 253–254 differentiation of, 249, 251, 255 Fc (FcgRIIB) and complement receptors in, 280, 284 follicular dendritic cells (FDC) and, 251 generation of, in T cell-dependent antibody responses, 247–252 germinal center (GC) formation and, 248, 249 gut-associated lymphatic tissue (GALT) and, 249 high affinity antibody and, 339–341, 345 IgA, IgD, IgE, IgM and, 248–249
580 memory and memory B cells (continued) immune response of B cells and, 340–341, 345 interleukins and, 251–252 lifespan and homeostasis of, 249–253 localization and recirculation of memory B cells in, 249, 250 major histocompatibility class (MHC) II in, 249 markers of memory B cells in, 248 memory B cells and plasma cells in, interplay of, 254–255 migration of B cells and, 203, 211 mouse B cell lineage lymphomas and, 365 nasopharynx-associated lymphoid tissue (NALT) in, 249 Pax5 and, 255 plasma cells and, 249–254 antibody presence and persistence of, 252–253 bone marrow and, 253 CD19, CD27, CD40 and, 253 chemokines and, 253–254 generation of, 252–253 humoral immunity and, 252 interleukins and, 254, 255 lifespan of, 252–253 Pax5 and, 255 plasmablasts and, 253–254 recruitment of, to memory pool, 253–254 survival signals for, 254 TLR9 and, 255 X-box binding protein (XBP), 255 predetermination of primary vs. secondary response in B cells and, 248 “readout” system in, 247 sites of localization for, 249, 250 spleen, tonsils, lymphoid organs and, 249, 250 subsets and properties of B cells in, 248–249 survival factors, nerve growth factor (NGF) and, 251 TLR9 and, 255 Toll-like receptors (TLR9) and, 251 X-box binding protein (XBP), 255 metalloproteases, migration of B cells and, 207–208 methylation of DNA, in V(D)J recombination, 132 MIG, migration of B cells and, 215 migration of pluripotent hematopoietic stem cells (pHSC), 104 migration of B cells, 117, 156, 203–221 a4b1 and, 208, 212, 214 a4b7 and, 212, 215 a5b1 and, 214 antibody–secreting cells (ASCs) and, homing of, 213–215 autoimmune response and, 213 B cell receptors (BCR) in, 210 B1 cells and, 212 B2 cells and, 212 BLC and, 209 body cavity B cells and, 211–212 bone marrow and, 212–213
Index Burkitt’s lymphoma and BLR1 in, 215–216 CCL3 and, 210 CCL4 and, 210 CCL19 and, 206–207, 206, 208 CCL20 and, 211 CCL21 and, 206–207, 206, 208, 213 CCL22 and, 210 CCL25 and, 215 CCR7 and, 206–207, 206, 209, 210, 214 CCR9 and, 215 CD11 and, 207 CD18 and, 207 CD22 and, 212, 214–215 CD31 and, 207 CD34 and, 205 CD62L and, 204–205 CD99 and, 207 chemokine triggering of integrin activation in, 205–207, 214, 215 compartmentalization of mature B cells and, 209–213 CXCL9 and, 215 CXCL10 and, 215 CXCL11 and, 215 CXCL12 and, 206–207, 206, 212–215 CXCL13 and, 206–210, 206, 212, 213, 215 CXCR3 and, 215 CXCR4 and, 206–207, 206, 212, 214 CXCR5 and, 206–210, 206, 212, 214 cytokines and, 213 defensins and, 211 diabetes and, 213 DOCK2 and, 216 Epstein-Barr virus (EBV) and, 211 fibronectin and, 214 firm adhesion in, integrin mediation of, 207 follicular dendritic cell (FDC) and, 209 FTY720 and stopping of, 209–210, 216 fucosyltransferase T-VII in, 211 G protein coupled receptor (GPCR) and, 204 glycam 1 and, 205 helper T cells and, 203 high endothelial cell (HEC) deficiency and, 205 high endothelial venules (HEVs) and, 203–208, 205, 209–210 ICAM 1 and ICAM 2 in, 207, 208, 210, 213 IgG and IgA in, 215 immune response of B cells and, 340 immunodeficiency diseases and, 213 inflammation sites and B cells at, 213 intracellular regulation of, 216 IP10 and, 215 ITAC and, 215 junctional adhesion molecules (JAMs) in, 207 L selectins in, 204–205 LFA-1 and, 205, 207, 208, 210, 212, 214 LTa1b2 and, 213, 215 lupus and, 213 lymph nodes and, 203, 204, 205 lymphoid follicles and, 209–210 MAdCAM1 and, 205, 208, 213, 215 marginal zone (MZ) B cells and, 209, 210–211
MDC and, 210 MECA79 and, 205 metalloproteases in, 207–208 MIG and, 215 MIP1a and b, 210 MIP3a and, 211 mucosal B cells and, 223, 224 myasthenia gravis (MG) and, 213 naive B cells in, 203 omentum and “milk spot,” 211–212, 212 P selectin glycoprotein ligand (PSGL) and, 214 pathways of, 214 PECAM 1 and, 207 peripheral lymph node addressin (PNAd) in, 205 pertussis toxin (PTX) and, 205–206, 208 Peyer’s patch and, 203, 204, 205, 209, 211, 215 phosphoinositide 3 kinase and, 216 plasmablasts and, 213 PNAd and, 213 podocalyxin and, 205 relocalization following activation in, 210 rheumatoid arthritis (RA) and, 213 rolling interactions in, 204–205 rotavirus infection and, 211 secondary lymphoid tissues and, 209–210 selectins in, 214 sgp200 and, 205 sialic acid and, 212–213 Sjogren’s syndrome and, 213 spleen entry and, 208–209, 208 stopping of, enlargement of lymph tissue and, 209 stromal cell-derived factor (SDF) and, 214 subsets of B cells in, 203 T-dependent/-independent antigens and, 210 TECK and, 215 thyroiditis and, 213 tonsillar tissue and, 211 transendothelial, 207–208 tumor necrosis factor (TNF) and, 213 ulcerative colitis and, 213 VCAM-1 and, 205, 207, 208, 210, 212–214 Waldeyer’s ring of oral cavity and, 211 miniature inverted repeat elements (MITE), 480 MIP1a and b, migration of B cells and, 210 MIP3a, migration of B cells and, 211 mismatch repair (MMR) activation-induced deaminase (AID) and, 314 class switch recombination (CSR) and, 316–319 somatic hypermutation (SHM) and, 319, 332–334, 335 MLL, 11q23 abnormalities in, 350–351 MMSET in multiple myeloma, 356 MoAbs, autoimmune disorders and, 389–390 Mohr-Tranebjaerg syndrome, X-linked (Bruton’s) agammaglobulinemia (XLA) and, 406 molecular mimicry, antibody production and, 497, 501 monoclonal antibodies, 411–531, 492, 547–561 alemtuzumab and, 541
Index allergic reactions/serum sickness, 511 antibody-dependent cellular cytotoxicity (ADCC) and, 542 anti-Tac (CD25), 540–541 autoimmune disorders and, 541 CDR grafting technique in, 534–536 characteristics of display technologies in, 512–513 chimerization of, in production of, 511, 533–534, 540 clinical use of, humanized antibodies approved for, 540–541, 541 complement-dependent cytotoxicity (CDC) and, 542 complementarity-determining regions (CDR) in, 511, 523, 534–539 computer-guided design of humanized V regions in, 534–537 daclizumab and, 541 display libraries and technologies for, 411–531 expression of, E. coli for, 511, 514 gemtuzumab ozogamicin and, 541 genetic engineering and, 533–534 herpes simplex virus antibody (HSV863/HuFd79D) and, 540, 541 human anti-chimeric antibody (HACA) response in, 534 human anti-humanized antibody (HAHA) response and, 540 human anti-mouse antibodies (HAMA) response in, 511, 533, 534 humanization of, in production of, 511, 533–545 hybridoma technology in, 511, 512, 519–520, 533 immune vs. nonimmune libraries in, 521 immunogenicity and, 511 library types and applications for, 521–522 Muromonab-CD3, 533 palivizumab and, 541 phage display production of, 513–514, 516–519, 517 affinity increases using, 522–524 alternative technologies to, 524–525, 524 complementarity-determining regions (CDR) in, 523 Fab vs. scFv libraries in, 521–522 high affinity antibody selection in, 523–524 hybridoma technology bypassed by use of, 519–520 immune vs. nonimmune libraries in, 521 immunization bypassed by use of, 520–521 library types and applications in, 521–522 mutations and, introduction of, 523 selection strategies for antibodies in, 522 vector systems in, 517–519, 518 polymerase chain reaction (PCR) generation in, 511, 515, 516, 525 prokaryotic expression of antibody fragments in, 514–515 RACE techniques in, 511 ribosome display technology in, 525, 525
somatic hypermutation (SHM) in, 511 translocus mice and human Mab production in, 547–561 affinity maturation in, 557–558, 557 antibody expression in, 553–557, 556, 553 bacterial artificial chromosome (BAC) studies in, 547, 551 CD8A in, 552 embryonic stem (ES) cells in, 551 human artificial chromosome (HAC) studies in, 552, 553 human Ig transloci in, 541–542 human IgH C genes and transcriptional enhancers in, 549–550, 550 immune response of, 557–558, 557 locus stability of mouse strains in, 553 lox and LIF loci in, 552 minigene constructs in, assembly of, 547–549 mouse strains used in, 552–558 replacement of mouse C genes in, 552 size, V gene content, serum titre of mice in, 548–549, 548 transfer of chromosome fragments in, 551–552, 551 V, D, J, C regions in, 547–558 yeast artificial chromosome (YAC) studies in, 547, 549–553 trastuzumab and, 541 V heavy and light chain genes in, 511, 513, 534, 535 yeast display technology in, 524–525, 524 morpholinos, 459–460 Morse, Herbert C. III, 365 Morton, H. Craig, 223 motifs, of recombination activating genes (RAG1/RAG2), 477–479 Mott cell myelomas, 268 mouse human anti-mouse antibodies (HAMA) response in, 511, 533, 534 hypermutated Ig genes in, 327–328 immunoglobin k genes of (See immunoglobin k genes of human and mouse) immunoglobin l genes of (See immunoglobin l genes of human and mouse) monoclonal antibodies and, translocus mice and human Mab, 547–561 murine models of B cell deficiency in, 411–412, 411 nonhomologous end joining (NHEJ) and deficiency, 75–77 S region organization and, 308–310, 310 strains (translocus) used in production of human Mab, 552–558 mouse B cell lineage lymphomas (See also B cell leukemias and lymphomas), 365–379 anaplastic plasmacytoma (PCT-A) in, 372–373 B cell receptors (BCR) in, 365 B natural killer cell lymphomas in, 373 BCL in, 366, 371 BTK deficiency and, 368
581 Burkitt-like lymphoma (BLL) and, 366, 371–374 Burkitt lymphoma (BL) in, 366, 371, 374, 375 causative factors in, 365 CCND1 and, 373 CD5 and, 366 cell lines in, repertoire of genes in, 365 centroblastic DLBCL and, 370 characteristics of, 368–373 chronic lymphocytic leukemia (CLL) and, 366, 367 CIS in, 373 class switch recombination (CSR) in, 365 classification of, 365–368, 367 comparative classification of, vs. human neoplasms, 366–368, 367 comparative genomic hybridization (CGH) in, 366 diffuse large B cell lymphoma (DLBCL) in, 366, 367, 368, 370–372, 373, 374 Dunn’s classification of, 365 extraosseous PCT (PCT-E) in, 372–373 follicular B cell lymphoma (FBL) in, 370, 374 gene expression profiling in, 374–375, 374, 375 germinal centers (GC) in, 365 histiocyte-associated (HA) DLBCL and, 370–371 human disease and, 365–368, 367, 374–375 IgH and IgL loci in, 366 immunoblastic DLBCL and, 370 inverse PCR (IPCR) studies in, 373 loss of imprinting (LOI) phenomenon in, 375 MAIDS syndrome in, 371 mantle cell lymphoma in, 366 memory B cells and, 365 Mouse Models of Human Cancers Consortium and, 366–367 mouse strains used in research for, 367–368, 368 murine leukemia virus (MuLV) in, 365, 368, 373 MYC in, 366, 368, 371, 373, 375 non-Hodgkin lymphoma and, 373 pathogenesis in, 373–375 PAX5 and, 373 PIM in, 373–374 plasma cells, plasma cell neoplasms in, 365, 372–373 plasmactyomas (PCT) in, 365, 372–373, 375 pre-B cell lymphomas and, 365 precursor B cell lymphoblastic lymphoma/leukemia (Pre-B LBL) in, 369 proviral insertional mutagenesis in, 373–374 sarcomas in, 365 sites of origin in, vs. humans, 368 SJL reticular neoplasms in, 365 small B cell lymphoma (SBL) in, 369, 374 spectral karyotyping (SKY) in, 366, 375 splenic marginal zone lymphoma (MZL) in, 369–370, 374 v-abl oncogene in, 365
582 Mouse Genome Sequencing Consortium (MGSC), immunoglobin l genes (IGL), mouse and, 51 Mouse Models of Human Cancers Consortium, 366–367 MRL, 275 MSH2/MSH6, somatic hypermutation (SHM) and, 332–335 mucosa-associated chemokine (MEC), mucosal B cells and, 236, 237 mucosa-associated lymphoid tissue (MALT), mucosal B cells and, 223, 225, 227–232, 238 mucosal B cells, 223–246, 224 adhesion molecules in, 234–237 a4b7 and, 236, 237 antigen–presenting cells (APCs) and, 223–224, 226 B cell receptors (BCR) and, 230, 231 B cell retention, proliferation, terminal differentiation and, 237–238 BCA 1 and, 231 BLC and, 231 bronchus-associated lymphoid tissue (BALT) in, 224–225, 236–237 CCL19 and, 235 CCL21 and, 235 CCL25 and, 236 CCL28 and, 236 CCR10 and, 236, 237 CCR7 and, 232, 235 CCR9 and, 236 CD3 and, 227 CD4 and, 231 CD5 and, 227 CD11 and, 235 CD18 and, 235 CD19 and, 227, 231 CD27 and, 227, 232 CD29 and, 235, 237 CD38 and, 232, 236 CD40 and, 227, 231 CD44 and, 236, 237 CD49 and, 237 CD54 and, 235 CD57 and, 231 CD70 and, 232 CD120 and, 235 characteristics of B cells in, 226–229 chemokines and, 231, 234–236, 237 class switch and IgA isotype promotion in, 232–234 compartmentalization of mature B cells and, 239 CR1 and CR2 in, 231 Crohn’s disease and, 228 CXCL12 and, 231, 235 CXCL13 and, 231, 235 CXCR4 and, 231, 235 CXCR5 and, 232, 235 differentiation and dispersion of GC B cells in, 231–232 disparate isotype distribution of other immunocytes in, 228
Index Epstein-Barr virus-induced chemokine (ELC) and, 235 exit of B cells from GC, 232 exocrine epithelia and, 238 follicle-associated epithelium (FAE) and, 224, 226, 229 follicular dendritic cell (FDC) and, 229–231, 230 germinal center (GC) formation and, molecular interactions in, 229–231 germinal center dendritic cells (GCDC) and, 231 gut-associated lymphoid tissues (GALT) and, 224, 225, 226–227, 227, 231, 233–238, 249 gut lamina propria and, 236 high endothelial venules (HEVs), 234–235 homing and retention mechanisms in, 234–238 ICAM and, 235 iccosomes and, 231 IgA and germinal centers in, 223, 224, 226–227, 226, 232–234 cell cycle in, 224 promoting stimuli in, 233–234 subclass distribution in, 227 IgD and, 238 IgG and, 228, 238 IgM and, 223, 224, 228 immune inductive tissue compartments and, 223–226 immunodeficiency diseases and, 237 inflammatory bowel disease (IBD) and, 228 isolated lymphoid follicles (ILFs) in, 224, 227 J region or J chain in, 223, 224, 228–229 leukocyte function-associated (LFA) and, 235 lipopolysaccharide (LPS) and, 227 LTa and LTb in, 230 LTa1b2 and, 231 lymph nodes in, 225–226 lymphotoxins and, 229–231 MAdCAM 1 and, 234–236 major histocompatibility class (MHC) II, 237, 238 membrane (M) cells in, 224, 229, 230, 238 migration of, 223, 224 mucosa-associated chemokine (MEC) and, 236, 237 mucosa-associated lymphoid tissue (MALT) and, 223, 225, 227–232, 238 mucosal homing molecules beyond the gut, 236–237 naive and primed B cells traffic from GALT, 235–236, 236 nasopharynx-associated lymphoid tissue (NALT) in, 224–225, 236–237, 249 peritoneal cavity and, 226 Peyer’s patches and, 223, 224, 226–227, 229, 230, 235, 238 PNAd and, 237 positive selection and plasma cell induction in, 231–232
questions still remaining about, 238–239 secretory antibodies (SIgA and SIgM) in, 223, 224, 227, 239 severe combined immunodeficiency disease (SCID) and, 237 SLC and, 235 stromal cell-derived factor (SDF) and, 231 T cell signaling and, 237, 238 TECK and, 236, 237 topical antigen exposure and, 238 tumor necrosis factor (TNF) and, 230 ulcerative colitis and, 228 vascular cell adhesion molecule (VCAM-1) and, 237 multiple myeloma, 356 multiple sclerosis, 382 Muramatsu, Masamichi, 307 murine leukemia virus (MuLV), 365, 368, 373 murine models of B cell deficiency, 411–412, 411 muromonab/Muromonab-CD3, 533, 541 mutation mutation (See also somatic hypermutation), 327 activation-induced deaminase (AID) and, 314 antibody production and, 496–499, 503–505 antibody–antigen interface, accommodation of, 499–500 autoimmune disorders and, 381–382 B cell development and, 103, 117, 148 class switch recombination (CSR) and, 289, 311 high affinity antibody and, 339 immunodeficiency diseases and, 148, 403, 404 immunoglobulin assembly and secretion, 267–268 mouse B cell lineage lymphomas and, proviral insertional mutagenesis in, 373–374 phage display and, introduction of, 523 recombination activating genes (RAG1/RAG2), 69 somatic hypermutation (SHM) and, 328–329 zebrafish (Danio rerio) and, genetic screening and, 457–459, 465 MutSa, somatic hypermutation (SHM) and, 333, 334 myasthenia gravis (MG), 213, 385, 392–393 MYC Burkitt lymphoma/B cell leukemia, activation of, 354–355 mouse B cell lineage lymphomas and, 366, 368, 371, 373, 375 myeloid cell development, with B cell development, 102 myeloid Ig-like receptor (MIR), 182 myeloid progenitor (pM) cells, B cell development and, 107 myeloma, 355–356
N NADPH-oxidase, B cell receptor (BCR) and, 163, 163
Index naive B cells antibody production and response, 188, 203 migration of B cells and, 203 nasopharynx-associated lymphoid tissue (NALT), 224–225, 236–237, 249 National Center for Biotechnology Information (NCBI), 38 natural killer cells, 141, 142 B cell development and, 108 B natural killer cell lymphomas in, 373 class switch recombination (CSR) and, 290–291 innate immunity and, 460, 462 killer-cell immunoglobulin-like receptors (KIRs) in, 428, 462 pluripotent hematopoietic stem cells (pHSC) and, 104 subset development in B cells, 155 transcription factors and, 108 natural, polyspecific autoantibodies, 382–383 NEMO and class switch recombination (CSR), 290 neonatal lupus (See also systemic lupus erythematosus), 393 nerve growth factor (NGF), memory and memory B cells in, 251 neutropenia, X-linked (Bruton’s) agammaglobulinemia (XLA) and, 404 new antigen receptors (NAR), somatic hypermutation (SHM) and, 485 NFATc1/NFATc2, 255 NF-kB/rel proteins, 93–96 class switch recombination (CSR) and, 290–292, 297, 312 signaling to activate, 95–96 NF-kB pathway, B cell leukemias and lymphomas and, and translocations associated with, 354 nick cleavage, class switch recombination (CSR) and, staggered, 314–315 Nijmegen breakage syndrome 1 (Nbs-1) gene, class switch recombination (CSR) and, 319 Nitschke, Lars, 171 Noggin, pluripotent hematopoietic stem cells (pHSC) and, 103 non-Hodgkin lymphoma, 373 nonhomologous end joining (NHEJ), 71–78, 71 ataxia telangiectasia mutated (ATM) protein and, 74 class switch recombination (CSR) and, 297, 310, 311, 315–316, 319 coding joint (CJ) in, 72, 72 DNA polymerases and, 75 DNA-PKcs and Artemis, 74–76 double strand breaks (DSBs) and, 77 functions of, 73–75 H2AX histone and, 77 hairpin end opening in, 75 immunodeficiency diseases and, 73–77 Ku70 and Ku80, 73–74, 76 mammalian, identification of, 73 mice deficient in, 75–77 repair of DNA and, 77
somatic hypermutation (SHM) and, 319 suppression of RAG-initiated translocations and, 77–78 V(D)J recombination and, 61, 67, 70, 71–78, 71, 72 XRCC4 and DNA ligase IV, 74, 76 nonimmunoglobulin genes, Vh region, human IgH, 6–7 non-T cell activation linker (NTAL), B cell receptor (BCR) and, 165 nontemplated somatic hypermutation (SHM), 328, 329 NOTCH/NOTCH1 pluripotent hematopoietic stem cells (pHSC) and, 103 T cell development and, 108–109 novel immune type receptors (NITR), innate immunity and, 462 nuclear localization/sublocalization transcription of immunoglobulin genes and, 92 V(D)J recombination and, 132 numbering in IMGT system, 39–40 NZB, 275
O OBF, B cell development and, 109 Oca-B, V(D)J recombination and, 130 Oct1 and Oct2, 130, 298 class switch recombination (CSR) and, 298 OCT-1 and OCT-2, 109 octamer proteins, transcription of immunoglobulin genes and, regulation by, 88 Oettinger, Marjorie, 61 Omenn syndrome, recombination activating genes (RAG1/RAG2), 69 omentum “milk spot,” B cell development and, 211–212, 212 open and shut joint recombination activating genes (RAG1/RAG2), 70 V(D)J recombination and, 61, 70 open reading frame (ORF) human IgH and, VH region, 3–4 immunoglobin l genes (IGL), human/mouse, 37 orphons immunoglobin l genes (IGL), human and, 49, 49 VH region, human IgH, 10–11 Vk genes of human/mouse and, 28, 30, 33 Orthoclone OKT3, 533 Osborne, Barbara A., 433 OX40L, 255
P P selectin glycoprotein ligand (PSGL) in migration of B cells, 214 P1/PDL-1, subset development in B cells, 158 p97/cdc48, immunoglobulin assembly and secretion, 268
583 paired complex (PC), V(D)J recombination and, 67 paired immunoglobulin-like receptors (PIR AB), 181–182 palivizumab, 541, 541 pathogenic autoantibody production (See autoimmune disorders) pattern recognition receptors, innate immunity and, 463 Pax5, 255 B cell development and, 109 common lymphoid precursors (CLP), 106, 107 deficiency in pre-B cells and, plasticity of, 109–110 class switch recombination (CSR) and, 298–299 marginal cell lymphomas and, 352 memory and memory B cells in, 255 mouse B cell lineage lymphomas and, 373 pluripotent hematopoietic stem cells (pHSC) and, 104 transcription of immunoglobulin genes and, lineage and stage-specific Ig enhancers in, 88, 93 V(D)J recombination and, 130 PDb1, V(D)J recombination and, 129 PECAM 1, migration of B cells and, 207 pediatric early B-lineage acute lymphoblastic leukemia, 349–352, 350 pemphigus, 385 periarteriolar lymphocyte sheath (PALS), B cell development and, 117 peripheral lymph node addressin (PNAd), 205 peritoneal cavity, B cell development in, 155, 211–212, 226 PERK, immunoglobulin assembly and secretion, 270 pertussis toxin (PTX), in studies of migration of B cells, 205–206, 208 Peyer’s patch in antibody production and response, 189, 203 ileal (IPP), 443 memory and memory B cells in, 249, 250 in migration of B cells, 203, 204, 205, 209, 211, 215 mucosal B cells and, 223, 224–230, 235, 238 phage display of monoclonal antibodies, 511, 512 affinity increases using, 522–524 alternative technologies to, 524–525, 524 complementarity-determining regions (CDR) in, 523 ELISA analysis in, 518–519 Fab vs. scFv libraries in, 521–522 high affinity antibody selection in, 523–524 hybridoma technology bypassed by, 519–520 immune vs. nonimmune libraries in, 521 immunization bypassed by use of, 520–521 library types and applications in, 521–522 monoclonal antibody display libraries and, 513–514, 516–519, 517 mutations and, introduction of, 523
584 phage display of monoclonal antibodies (continued) selection strategies for antibodies in, 522 vector systems in, 517–519, 518 phagocytic (myeloid) cells, innate immunity and, in zebrafish, 461–462 phosphoinositide 3 kinase (PI3K) antigen receptor signaling regulation and, 177 B cell receptor (BCR) and, 164 migration of B cells and, 216 PhyloGene tree map, Immunogenetics Database (IMGT), 55, 57 phylogenic relationships of immunoglobulin classes in vertebrates, 420 PIM, mouse B cell lineage lymphomas and, 373–374 pL cell compartments, B cell development and common lymphoid precursors (CLP), 106 plasma cells (See also plasmablasts) adaptive B cell memory and, 254–255 anaplastic plasmacytoma (PCT-A) in, 372–373 in antibody production and response, 188 bone marrow and, 253 CD19, CD27, CD40 and, 253 chemokines and, 253–254 differentiation of, 255, 268–270, 269 extraosseous PCT (PCT-E) in, 372–373 Fc (FcgRIIB) and complement receptors in, 284 generation of, 252–253 humoral immunity and, 252 immunoglobulin assembly and secretion, differentiation to, 268–270, 269 induction of, positive selection and, 231–232 interleukins and, 255 lifespan of, 252–253 memory and memory B cells in, 249, 251, 252–254 mouse B cell lineage lymphomas and, 365, 372–373 Pax5 and, 255 plasmactyomas (PCT) in, 365, 372–373, 375 survival signals for, 254 TLR9 and, 255 X-box binding protein (XBP), 255 plasmablasts, 340 in antibody production and response, 187, 189, 190–191, 197, 253–254 antibody–secreting cells (ASCs) and, homing of, 213–215 memory and memory B cells in, 253–254 migration of B cells and, 213 plasmacytomas (PCT), 365, 372–373, 375 plasticity of pluripotent hematopoietic stem cells (pHSC), 105–106, 105 platelet development (See also hematopoiesis), 102 PLCg2, antigen receptor signaling regulation and, 175, 177, 182 pleural cavity, B cell development in, 155, 211–212 PLNK, B cell development and, 113
Index pluripotent hematopoietic stem cells (pHSC), 103–106 cytokines and, 104 hemangioblasts as early progenitors of, 104–105 long-term reconstitution potential of, 104 migration and homing in, 104 natural killer cells and, 104 PAX5 and, 104 plasticity vs. stability of, 105–106, 105 pluripotency of, 104 RUNX-1 and, 104 self-renewal ability in, 103–104 short-term (ST), 104 stem cell factor (SCF) and, 104 TRANCE cytokine and, 104 transdifferentiation to and from nonhematopoietic cells, 106 vascular endothelium development and, 104–105 PNAd migration of B cells and, 213 mucosal B cells and, 237 podocalyxin and migration of B cells, 205 poliomyelitis, X-linked (Bruton’s) agammaglobulinemia (XLA) and, 403–404 polychondritis, 385 polymerase chain reaction (PCR) generation monoclonal antibody display libraries and, 511, 515, 516, 525 transcription of immunoglobulin genes and, 91 polymerases, error-prone, somatic hypermutation (SHM) and, 335 polymorphisms Ch region, human IgH, 14–15, 14 immunoglobin l genes (IGL), human and, 49 lagomorphs (rabbits and hares) and, VHa in, 437 Vk genes, human and, 30 VH region, human IgH, 4–5 polymyositis, 383 post-cleavage complex, RAG, in V(D)J recombination, 67 PRDM1, 255 pre-B cell lymphomas, mouse B cell lineage, 365 pre-B cell receptors (pre-BCRs), 101, 127, 142, 144–145 immunoglobulin assembly and secretion, 265–266 pre-B cells, immunoglobulin assembly and secretion, 262 pre-B cell lymphoblastic leukemias (pre-B LBL), 351, 369 pre-B-1 cells, 112 primary biliary cirrhosis (PBC), 390–391, 391 pro-B cell lymphoblastic leukemias of adolescence, 353–354 prokaryotic expression of antibody fragments, in monoclonal antibody display, 514–515 promoters of Ig genes, 83–85 transcription of immunoglobulin genes and, 83–85
lower vertebrate immunoglobulin genes and, 425–427 protein kinase C (PKC), B cell receptor (BCR) and, 164 protein tyrosine kinases (PTK), B cell receptor (BCR) and, 162 protein tyrosine phosphatases (PTP), B cell receptor (BCR) and, 162 proteins binding in Ig transcriptional regulatory elements, 86–89, 87 protochordates, V region diversification in, 428–429 PSA testing, 533 pseudogenes immunoglobin l genes (IGL), human/mouse, 37 Vk genes, mouse and, 31–32, 32 VH region, human IgH, 11, 12 Vk genes of human/mouse and, 28 pTa, B cell development and common lymphoid precursors (CLP), 106 PU.1 B cell development and, 107–109 class switch recombination (CSR) and, 292 transcription of immunoglobulin genes and, lineage and stage-specific Ig enhancers in, 88, 90, 93 V(D)J recombination and, 130 pulse field gel electrophoresis (PFGE) human IgH and, 2 immunoglobin l genes (IGL), mouse and, 50, 51 Vk genes, mouse and, 31 pyk-2, 156–157
R rabbits (See lagomorphs) RACE techniques, monoclonal antibody display libraries and, 511 RAD51, somatic hypermutation (SHM) and, 335 Radbruch, Andreas, 247 rag gene, in zebrafish (Danio rerio) and, 454, 456–458, 456, 458, 465 Rajewsky, Klaus, 247 Ravetch, Jeffrey V., 275 “readout” system, memory and memory B cells in, 247 rearranged (unproductive) genes, immunoglobin l genes (IGL), human/mouse, 37 “recombinasome,” in class switch recombination (CSR), 319 recombination activating genes (RAG1/RAG2), 68–71, 307 active site identification in, 69 B cell development and, 106–109, 112, 114, 119, 127 C terminal region/domain of RAG 2 and, 68–69 conservation of, between species, 68 domains and motifs in, 477–479 double strand break (DSB) and, 72–73, 77 exons in, 68
Index full length proteins vs. core versions, differences in V(D)J recombination, 69–70 genomic structure of, sequence similiarity and, 68 hairpin end opening in, 75 hinge region in RAG 2 and, 68 hybrid joining (HJ) in, 70 identification of, 68 immunodeficiency diseases and, 69, 75–77, 148 introns in, 68 kelch repeats in, 478–479 linkage and structure in, 477 lymphoid-specific nature of, 68 lymphoid tumors and, 71 mechanisms of transposition in, 479–480, 479 miniature inverted repeat elements (MITE) and, 480 mutations in, 69 nonhomologous end joining (NHEJ) in, 70 Omenn syndrome and, 69 open and shut joints in, 70 post-cleavage complex in, V(D)J recombination, 67 recombination signal sequence (RSS) binding to, 66–67, 70–71, 71 severe combined immune deficiency (SCID) disease and, 75–77 suppression of translocation in, by NHEJ factors, 77–78 tranposition mediated by, 70–71 V(D)J recombination and, 61, 62, 62–63, 65–71, 127, 131, 133, 134, 135, 136, 141, 142, 144, 473–480 zinc fingers in RAG1, 69 recombination signal sequence (RSS) cleavage requirements of, in V(D)J recombination, 66 mutational analysis of, during V(D)J recombination, 64 RAG1 and RAG2 binding to, 66–67, 70–71, 71 V(D)J recombination and, 61–66, 63, 128, 129, 131, 145–146, 473, 474, 476, 482, 483 Vh region, human IgH, 9 red pulp of spleen, migration of B cells and, 208, 340 redox regulation of BCR signaling, 162– 163 regulatory elements, transcription of immunoglobulin genes and, in heavy and light chain genes and, 83–86, 84, 85 Rel homology domain (RHD) proteins, transcription of immunoglobulin genes and, 94 relics Vk genes, mouse and, 33 Vk genes of human/mouse and, 28 reptiles, class switching recombination (CSR) in, 422–423 Reth, Michael, 161 reticular neoplasms, SJL, 365
reverse transcription PCR (RT-PCR), monoclonal antibody display libraries and, 525 RFLP, immunoglobin l genes (IGL), mouse and, 52, 55 rheumatoid arthritis (RA), 111, 213, 393 ribosome display technology, monoclonal antibody display libraries and, 525, 525 RIIB (See Fc and complement receptors) rituximab, 541 RNA PolII/mediator, transcription of immunoglobulin genes and, 90 rolling interactions in migration of B cells, 204–205 rotavirus infection, migration of B cells and, 211 RUNX1, 349, 459 pluripotent hematopoietic stem cells (pHSC) and, 104 Russell bodies, immunoglobulin assembly and secretion, 268
S S regions, class switch recombination (CSR) and, 308–319, 309 sarcomas, mouse B cell lineage lymphomas and, 365 SATB1, MAR binding protein, 93 Sca 1, 106, 141 Scharff, Matthew D., 327 Schlissel, Mark S., 127 scl genes, zebrafish (Danio rerio) and, 451, 452 scleroderma, 383, 393 SDF (See stromal cell-derived factor) secretory antibodies (SIgA and SIgM), 223, 224, 227, 239 Sekiguchi, JoAnn, 61 selectins in migration of B cells, 214 self-antigen (See autoimmune disorders) self-reactive B cells, negative selection of Fc (FcgRIIB) and complement receptors in, 284–285 Sen, Ranjan, 83 Sequence Retrieval System (SRS), immunoglobin l genes (IGL), human/mouse, 38 serum sickness, 511 severe combined immune deficiency (SCID) disease B cell development and, 109 mucosal B cells and, 237 nonhomologous end joining (NHEJ) and, 73–77 recombination activating genes (RAG1/RAG2), 69 sgp200 and migration of B cells, 205 SH2 domain, B cell receptor (BCR) and, 162, 165 SH2 domain containing inositol 5-phosphatase (SHIP) antigen receptor signaling regulation and, 182 B cell development and, 147
585 class switch recombination (CSR) and, 291 Fc (FcgRIIB) and complement receptors in, 277–278 Shc, antigen receptor signaling regulation and, 177 sheep, somatic hypermutation (SHM) and, nontemplated mutation in, pre-immune diversification, 329 Shlomchik, Mark J., 339 short-term (ST), pluripotent hematopoietic stem cells (pHSC) and, 104 SHP-1 B cell receptor (BCR) and, 166, 177–178 subset development in B cells, 156 antigen receptor signaling regulation and, 181 B cell receptor (BCR) and, 162 sialic acid, migration of B cells and, 212–213 single nucleotide polymorphisms (SNP), human IgH and, Vh region, 4–5 single strand breaks (SSB), somatic hypermutation (SHM) and, 334, 335 Sitia, Roberto, 261 SJL reticular neoplasms in, 365 Sjogren’s syndrome (See also systemic lupus erythematosus), 213, 393, 394 SLAP-130, B cell receptor (BCR) and, 165 SLC, mucosal B cells and, 235 SLP-65 B cell development and, 113 B cell receptor (BCR) and, 163–166, 177 small B cell lymphoma (SBL), 369, 374 Smith, George, 516 somatic hypermutation (SHM), 319–320, 327–338 activation and targeting of, transcription and cis elements in, 329–330 activation-induced deaminase (AID), 307, 314, 319–320, 328–332, 335 antibody production and, 503–505, 511 APOBEC-1 and, 331 B cell development and, limited window for, 330–331 BCDX2 and, 335 BCL 6 in, 330 break and repair pathway in, 334 bursa of Fabricius and, 330, 435–436 C region and, 327 C to U deamination and base excision repair in, 332, 333 C/G base pairs and, 332, 334 CD95 in, 330 characteristics of, 327–329 chicken, templated mutation in, pre-immune diversification, 328–329 cis elements in, 485 class switch recombination (CSR) and, 289, 307, 319–320, 330, 331 complementarity-determining regions (CDR) and, 327 co-opting of targeting elements in, 330 diffuse large cell lymphomas (DLCL) and, 330 DNA breaks in, 334–335 DNA-PKcs in, 335
586 somatic hypermutation (SHM) (continued) double strand breaks (DSBs) and, 319, 334–335 error-prone polymerases in, 335 evolution of, 320, 335, 484–485 g-H2AX in, 335 germinal centers (GC) and, 330 lower vertebrates, 485 lymphoid organs and, 330 mice, hypermutated Ig genes in, 327–328 mismatch repair (MMR) and, 319, 332–335 monoclonal antibody display libraries and, 511 MSH2/MSH6 in, 332–335 MutSa and, 333, 334 new antigen receptors (NAR) in, 485 nonhomologous end joining (NHEJ) in, 319 rabbits, templated and nontemplated mutation in, antigen-activated, 329 RAD51 and, 335 sequestered microenvironments for, 330–331 sheep, nontemplated mutation in, preimmune diversification, 329 single strand breaks (SSB) in, 334, 335 T cell receptors (TCR) and, 485 targeted mutagensis and, 328–329 templated vs. nontemplated mutation in, 328, 329, 335 transcription in, 329–330 transformed cell lines and, 330–331 unselected mutations patterns in, 328 uracil DNA glycosylase in, 332 ur-V genes and, 485 V region and, 327–329 V(D)J recombination and, 327, 330, 484–485 zone of hypermutation in, 327–328, 328 southern hybridization, human IgH and, 2 SOX-4, B cell development and, 109 spectral karyotyping (SKY), 366, 375 Spi-1, B cell development and, 107 spleen antibody–secreting cells (ASCs) and, homing of, 213–215 B cell development and, 117, 155, 187 immune response of B cells and, 340 marginal zone (MZ) of, 155, 172–176 memory and memory B cells in, 249, 250 migration of B cells and, 208–209, 208 red pulp and white pulp in, 208 white pulp cords in, 208 splenic marginal zone lymphoma (MZL), 369–370, 374 Src homology 2 domain containing inositol-5phosphatase (See SHIP) S-S recombination, class switch recombination (CSR) and, with looped-out CH gene deletion in, 307–308, 308 staggered nick cleavage, class switch recombination (CSR) and, 314–315 STAT and STAT1 antigen receptor signaling regulation and, 175 class switch recombination (CSR) and, 292, 293 Stavnezer, Janet, 307
Index Steiner, Lisa A., 449 stem cell development, 102 stem cell factor (SCF), 104 Stevenson, Freda K., 381 strength of antibody–antigen interaction, 499 stress response, endoplasmic reticulum (ER) and Ig production in, 269–270 stromal cell-derived factor (SDF) B cell development and, 103 class switch recombination (CSR) and, 290 migration of B cells and, 214 mucosal B cells and, 231 plasma cells, memory and, 253, 254 subcellular localization, transcription of immunoglobulin genes and, in IkB proteins and, 94–95, 132 subgroups, immunoglobin l genes (IGL), human/mouse, 38 subset development and function of B cells, 155–160, 248–249 antibody production and, 155 B 1 cells in, 187 B cell receptors (BCR) and, 156–157, 248–249 B lymphoid follicles (FO) and, 155 B1 and B2 precursors for, 158 B1 cells in, 155, 172, 212 B2 cells and, 212 BAFF in, 157 CD40 and, 158 CD5 and, 157 chemokines and, 156 compartmentalization of, 157, 209–213, 239 CXCL13 in, 157, 192, 195 factors involved in formation of, 157–158 Fas and, 157–158 G protein coupled receptors (GPCRs) in, 156, 204 ICAM-1 and VCAM-1 in, 157 immune response and, 155 localization in, 156 lymph nodes and, 155, 187, 188–189 marginal zone (MZ) B cells and, 155, 172–176, 188–189, 209, 210–211 migration of B cells and, 156, 203 natural killer cells and, 155 P1/PDL-1 and, 158 peritoneal and pleural cavities in, 155, 211–212, 226 pyk-2 and, 156–157 selection and differentiation in, 156–157 T cell development and, 155 T cell receptor (TCR) genes and, 155 Sundberg, Eric J., 491 superantigenic drive on B cells, autoimmune disorders and, 388–390 supercoiling of DNA, transcription of immunoglobulin genes and, 89 surrogate light chain (SL/SLC) B cell development and, 107, 110–111, 144–145 expression of, 111 immunoglobulin assembly and secretion, 262, 265–266 ligands for pre-B cell receptors in, 111
structure and assembly of pre-B cell receptor in, 110–111 SV40 enhancer, transcription of immunoglobulin genes and, 83 SWAP-70, class switch recombination (CSR) and, 290, 297 switch region, Ch region, human IgH, 14 switch sequences, class switch recombination (CSR) and, as transcriptional stimulatory elements in, 291 Syk B cell development and, 113 B cell receptor (BCR) and coupling, 162, 163, 177 systemic lupus erythematosus (SLE), 213, 275, 382, 385, 393–394 Fc (FcgRIIB) and complement receptors in, 284, 286
T T cell development, 127, 141, 142 antibody production and response, 187, 188–189, 190, 210 B cell development and, 108–109 class switch recombination (CSR) and, 290 germinal centers (GC) and, 195–197 germinal centers (GC) and, in secondary follicles, 194 growth and differentiatiion, 190 linker of activated T cell (LAT) in, 165 memory and memory B cell generation and, 247–252 mucosal B cells and, 237, 238 non-T cell activation linker (NTAL) in, 165 primary cognate interaction of B cells with, 189–190 subset development in B cells, 155 T cell receptor (TCR) genes and, 155 thymocytes and, 127 transcription factors and, 108 X-linked (Bruton’s) agammaglobulinemia (XLA) and, 408 zebrafish (Danio rerio) and, 456–457 T cell-independent (TI) antigens, class switch recombination (CSR) and, 289 T cell receptor (TCR) B cell development and, 127–130 evolution of loci for, 481–483, 481 lower vertebrate immunoglobulin genes and, 417, 427–428, 428 somatic hypermutation (SHM) and, 485 V(D)J recombination and, 62, 64–65, 133–136 t(11;18)(q21;q21) and API2-MLT/MALT1 fusion gene in marginal cell lymphomas, 353 t(14;18) and activation of BCL2 in follicular lymphoma, 352–353, 370 t(17;19)(q21-q22;13) and E2A-HLF fusion gene in pro-B cell lymphoblastic leukemias of adolescence, 353–354 t(21;21)(p13;q22), E2A-PBX1 fusion gene and, pre-B cell lymphoblastic leukemias, 351
Index t(21;21)(p13;q22), TEL-AML1 (ETV6RUNX1) fusion gene and, pediatric early B-lineage acute lymphoblastic leukemia, 349–352, 349, 350 t(3;14)(q27;q32) and BCL6 activation in diffuse large cell lymphoma, 351–352 t(4;14)(p16;q34) and activation of FGFR3 and MMSET in multiple myeloma, 356 t(8;14)(q24;q23) and MYC activation in Burkitt lymphoma/B cell leukemia, 354–355 t(9;14)(p13;q32) and PAX5 activation in marginal cell lymphomas, 352 t(9;22)(q34;q11) and BCR-ABL fusion gene in adult early B-lineage acute lymphoblastic leukemia, 355–356 TAL-1, B cell development and, 107 TCF-1, V(D)J recombination and, 130 TCRa/b, in zebrafish (Danio rerio) and, 454, 455–457 TdT B cell development and, 106, 107, 114, 144 Fc (FcgRIIB) and complement receptors in, 282 TEA, V(D)J recombination and, 129, 135–136 TECK migration of B cells and, 215 mucosal B cells and, 236, 237 plasma cells, memory and, 254 TEL-AML1 (ETV6-RUNX1) fusion gene and, pediatric early B-lineage acute lymphoblastic leukemia, 349–352, 350 templated somatic hypermutation (SHM) and, 328, 329, 335 TFE3, transcription of immunoglobulin genes and, 89 TGF-b, 292, 293, 312 thermodynamics of antibody–antigen interactions, 496 thoracic cavity, B cell development in, 211–212 3¢ IgH enhancers, class switch recombination (CSR) and, 293–295, 316 Thy 1 B cell development and, 106, 141 common lymphoid precursors (CLP) and, 106 thymocytes, T cell development and, 127 thymus, zebrafish (Danio rerio) and, development of, 456–459 thymus-dependent antigens, antigen receptor signaling regulation and, CD19 and, 173 thymus-independent antigens, antigen receptor signaling regulation and, CD19 in, 172–173 thyroiditis, 213, 383 TLR9, memory and memory B cells in, 255 TNFR (See also tumor necrosis factor) class switch recombination (CSR) and, 290 transcription of immunoglobulin genes and, 95 TNFSF5, hyper-IgM syndrome (HIGM) and, 410 Toll-like receptors (TLR) class switch recombination (CSR) and, 290 innate immunity and, 460, 463 memory and memory B cells in, 251
tonsillar B cells in antibody production and response, 189 memory and memory B cells in, 249, 250 migration of B cells and, 211 TRAF in antibody production and response, 189, 196, 197 class switch recombination (CSR) and, 290 TRANCE B cell development and, 109 pluripotent hematopoietic stem cells (pHSC) and, 104 tranposition, recombination activating genes (RAG1/RAG2)-mediated, 70 trans-acting factors, V(D)J recombination and, 130 transcription factors class switch recombination (CSR) and, 3¢ IgH regulatory elements control, 297–299 control of, 107–109 PU-1 and, 107–108 transcription of immunoglobulin genes, 83–100, 131 activation of Ig locus for rearrangement and, 90–91, 90 alleles of IgH in, 92 B cell lineage and, commitment to, 93 B cell lineage specific activator protein (BSAP) in, 88 B cell specificity in, 83 base unpairing regions (BURs) in, 93 C regions and, 83 chromatin structure and, 90, 130–132 class switch recombination (CSR) and, 91, 147 current research in, 89–93 D region in, 91 Dh-Cu region activation in, 91, 92 different proteins binding to one site in, regulation by, 88 dimerizing proteins in, regulation by, 87–88 discoveries resulting from studies in, 93–96 DNA binding proteins and regulation in, cooperative interactions in, 88–89 DNA-dependent protein kinase (DNA-PK) in, 95 DQ52 promoter in, 129 E2-A proteins in, 93 enhancers 3¢ of Ca in, 86 ets family proteins and partners in, regulation by, 88–89, 93 Em intronic enhancer in, 83–91, 84, 129 gene rearrangement and, 91 hematopoeitic cell development and, 93 Ig enhancer activity regulation, multi-site, multi-mechanism, 86–88 Ig promoters in, 83–85 IRF4, C/EBPb, TFE3 and, 89 IkB kinase (IKK) complex in, 95–96 J regions and, 83, 91 k intronic and 3¢ enhancers in, 85, 86, 88– 89 l enhancers in, 85, 86, 88–89 looping in, 89–90
587 lower vertebrate immunoglobulin genes and, 425–427 matrix attachment regions (MARs) in, 84, 86, 90, 92–93 mechanisms of enhancer action for, 89–93 NF-kB/rel proteins and, 93–96 nuclear sublocalization in, 92 octamer proteins in, regulation by, 88 Pax5 protein and, lineage and stage-specific Ig enhancers in, 88, 93 polymerase chain reaction (PCR) studies in, 91 probablity of initiation in, 89 proteins binding regulatory elements in, 86–89, 87 PU.1 protein and, lineage and stage-specific Ig enhancers in, 88, 90, 93 regulatory elements in heavy and light chain genes and, 83–86, 84, 85 Rel homology domain (RHD) proteins and, 94 RNA PolII/mediator in, 90 signaling to activate NF-kB/rel proteins, 95–96 somatic hypermutation (SHM) and, 329– 330 subcellular localization in IkB proteins and, 94–95, 132 SV40 enhancer in, 83 tracking of, 89 V region in, 84, 91 V(D)J recombination and, 83 VH gene activation in, 92 transcriptional enhancers, 128–130, 129 transcriptional promoters, 129 transcytosis, immunoglobulin assembly and secretion, 267 transformed cell lines, somatic hypermutation (SHM) and, 330–331 transgenesis in zebrafish, 459 transient receptor potential (TRP), B cell receptor (BCR) and, 164 translocus mice and human monoclonal antibodies (See also monoclonal antibodies), 547–561 affinity maturation in, 557–558, 557 antibody expression in, 553–557, 556 bacterial artificial chromosome (BAC) studies in, 547, 551 CD8A in, 552 embryonic stem (ES) cells in, 551 human artificial chromosome (HAC) studies in, 552, 553 human Ig transloci in, 541–542 immune response of, 557–558, 557 locus stability of mouse strains in, 553 lox and LIF loci in, 552 minigene constructs in, assembly of, 547–549 replacement of mouse C genes in, 552 size, V gene content, serum titre of mice in, 548–549, 548 transfer of chromosome fragments in, 551–552, 551 V, D, J, C regions in, 547–558, 547
588
Index
translocus mice and human monoclonal antibodies (continued) yeast artificial chromosome (YAC) studies in, 547, 549–552, 553 transposons, V(D)J recombination and, 473–480, 474, 475 trastuzumab, 541, 541 Tsurushita, Naoya, 533 tumor necrosis factor (TNF), 255 B cell development and, 118, 157 innate immunity and, 464 migration of B cells and, 213 mucosal B cells and, 230 12/23 rule in, 62, 63, 64–65, 128
U ulcerative colitis, 213, 228, 541 unfolded protein response (UPR), immunoglobulin assembly and secretion, 269–270, 270 UNG, activation-induced deaminase (AID) and, 314 unique numbering in IMGT system, 39–40 Université Montpellier II, 37 uracil DNA glycosylase, somatic hypermutation (SHM) and, 332 ur-V gene, V(D)J recombination and, 483–486 uveitis, 382, 541
V V chain bifunctional genes (VCBP), 429 V (variable) genes/regions, 1 antibody production and, 491, 493–495 autoimmune disorders and, 381–388, 385, 386, 387, 394, 395, 395 B cell development and, 115 beyond 12/23 (B12/23) rule in, 64–65 class switch recombination (CSR) and, 307 fibrinogen-related proteins (FREP) and, 486 high affinity antibody and, 339 immunoglobin l genes (IGL), human, 38, 38, 39, 41, 41, 46, 48, 50, 51–52, 53, 54 immunoglobin l genes (IGL), mouse, 38, 39, 39, 41, 50, 51–52, 53, 54 immunoglobulin assembly and secretion, 261, 262 immunoglobulin genes in higher vertebrates and, polymorphisms in, 437 monoclonal antibodies, 511, 513, 534 in protochordates, V region diversification in, 428–429 somatic hypermutation (SHM) and, 327–329 transcription of immunoglobulin genes and, 84, 91 translocus mice and human Mab and, 547–558 12/23 rule in, 62, 63, 64–65, 128 V chain bifunctional genes (VCBP) in, 429 V(D)J recombination and, 62 zebrafish (Danio rerio) and, 455 Vk genes, human, 27–30 bacterial artificial chromosome (BAC) studies in, 27
elucidation of, 27–28 evolution of, 33–34 k protein classification in, 29 mapping of, 28, 28 orphons in, 28, 30 polymorphisms, 30 pseudogenes, relics, repetitive elements in, 28–29 rearranged and expressed genes in, 29–30 sequencing of, 34 V-J rearrangement in, 29–30 yeast artificial chromosone (YAC) studies in, 27 Vk genes, mouse, 30–33 bacterial artificial chromosome (BAC) studies in, 31, 33 evolution of, 33–34 genes and pseudogenes in, 31–32, 32 relics and orphons in, 33 sequencing of, 34 yeast artificial chromosome (YAC) studies in, 31 V(D)J recombination, 61–82, 473–489 accessibility hypothesis in, 128 activation of Ig locus for transcription and, 90–91, 90 “alien seed” hypothesis in, 473–480 allelic exclusion in IgH loci in, 133–134 Igk loci and, 135, 145 isotypic exclusion and, 127–140, 128 TCRb loci in, 134–135 12/23 rule in, 62, 63, 64–65, 128 AMuLv in, 134 antigen receptor genes and, 61, 62–64, 63, 64 in artiodactyls (hoofed, grazing animals), 441 assay of, in vitro, 63, 64 B cell development and, 101, 109, 110, 112, 113, 114, 119, 141 B cell receptor (BCR) loci in, evolution of, 481–483, 481 beyond 12/23 (B12/23) rule in, 64–65 biochemistry of, 65–66 cell cyle checkpoint machinery and, 77 chromatin dynamics and, 130–132 class switch recombination (CSR) and, 289, 292, 307, 312, 315 coding flanks in, 65 coding joint (CJ) in, 61, 72 complementarity-determining regions (CDRs) and, 61, 146, 483 coordinated regulation of transcription, recombination, replication in, 300 CpG methylation and, 132 CREB and, 130 D segment creation, germline signal joint formation in, 482 deletional, 63, 63 diversification in, 473–489 DNA generated in, 61, 131 DNA polymerases during, 75 double strand break (DSB) in, 61, 62, 71, 71, 72–73, 77 DQ52 promoter in, 129
E2A protein and, 130 Ea and, 130 enhancer and promoter control of, 128–130, 129 Em intronic enhancer and, 83, 129, 292 evolutionary origins of, 473 “exaptation” in, 473, 474, 480 fibrinogen-related proteins (FREP) and, 486 germline coding joint formation and gene segment fusion in, 481–482, 481 germline transcriptions in, 128, 131 H2AX histone and, 77 hairpin end opening in, 75 HEB and, 130 HIN and HIX sites in, 474, 476, 476 histone acetyltransferases (HATs) in, 131 histone deacetylases (HDACs) in, 131 HMG1/HMG2 DNA bending protein in, 65–68, 128, 131 horizontal or lateral transfer in, 473–474 human IgH and, 1 hybrid joint (HJ) in, 61, 70 Ig light chain isotypic exclusion in, 136 IgH and, ordered rearrangement of, 132–133 IL7 and, 133 immune type receptors (ITRs) and, 476 immunoglobulin (Ig) genes and, 62 impairment of, 129–130 innate defense V gene theory and, 485–486 inversional, 63, 63 inverted terminal repeats (ITRs) in, 473, 474 isotypic exclusion in, 136 kelch repeats in, 478–479 LEF-1 and, 130 lymphoid tumors and, 71 mechanisms of, 479–480, 479 methylation of DNA in, 132 mutational analysis of RSS in, 64 nonhomologous end joining (NHEJ) in, 61, 67, 70, 71–78, 71, 72 nuclear localization and, 132 Oca-B and, 130 Oct1 and Oct 2 in, 130 open and shut joint in, 61, 70 ordered rearrangement with Ig and TCR loci in, 132–133 paired complex (PC) in, 67 Pax5, 130 PDb1 and, 129 PU.1 and, 130 recombination activating genes (RAG1/RAG2) and, 61–63, 62, 65–71, 127, 131, 133, 134, 135, 136, 141, 142, 144, 307, 473–474, 477–480 domains and motifs in, 477–479 “exaptation” in, 480 kelch repeats in, 478–479 linkage and structure in, 477 mechanisms of transposition in, 479–480, 479 miniature inverted repeat elements (MITE) and, 480 post-cleavage complex in, 67 RSS binding in, 66–67
589
Index recombination signal sequences (RSS) in, 61–66, 63, 128, 129, 131, 145–146, 473, 474, 476, 482, 483 repair of DNA and, 77 RSS swaps and locus speciation in, germline hybrid joint formation, 482 sequence length diversity and, 483–484, 483 site similarities in recombination, 474–477 somatic hypermutation (SHM) and, 327, 330, 484–485 spacer length in, 62, 66 suppression of RAG translocation in, by NHEJ factors, 77–78 T cell receptor (TCR) genes and, 62, 64–65, 127, 128, 129, 130, 133, 134–136 evolution of, 481–483, 481 TCF-1 and, 130 TEA in, 129, 135–136 trans-acting factors in, 130 transcription of immunoglobulin genes and, 83, 129 transcriptional enhancers in, 128–130, 129 transcriptional promoters in, 129 transposons and, 473–480, 474, 475 unpairing of DNA and distortion in, 66 ur-V gene in, 483–486 V, D, and J regions in, 62 in zebrafish (Danio rerio) and, 453–454, 457 v-abl oncogene, mouse B cell lineage lymphomas and, 365 vascular endothelium, pluripotent hematopoietic stem cells (pHSC) and, development and, 104–105 vasointestinal protein (VIP), class switch recombination (CSR) and, 293 Vasquez, Maximiliano, 533 VCAM-1 in antibody production and response, 195 migration of B cells and, 205, 207, 208, 210–214 mucosal B cells and, 237 subset development in B cells, 157 VCAM-4, pluripotent hematopoietic stem cells (pHSC) and, 105 vector systems, in phage display, 517–519, 518 VEGF, 105, 505 vesicular stomatitis virus (VSV), 253 Vh region, human IgH, 1, 2–7 5¢ regulatory regions in, 8–9, 8 analysis of, 7–10 B cell and immune response in, 9–10 chromosome 14 and, evolution of, 10 chromosome 15 and 16 segments of, 5–6, 6 classification of segments in, 4, 4 evolution of, 10–12 nonimmunoglobulin genes in, 6–7 open reading frames (ORF) in, 3–4 orphon, evolution of, 10–11 overexpression of, 10 polymorphism of, 4–5 primary repertoire of, 9–10 pseudogenes and gene conversion in, 11, 12 recombination signal sequence in, 9
segment usage and repertoire formation in, 10 subgroups and families of, 7–8, 7 total number of segments, 3–4 transcription of immunoglobulin genes and, 92 V-J rearrangement, V k genes, human and, 29–30 Vk region (See immunoglobin k genes of human and mouse) VLA4, in antibody production and response, 195 VpreB, 109–113, 114, 127, 142, 143–144 B cell development and common lymphoid precursors (CLP), 106 V-QUEST alignment tool, Immunogenetics Database (IMGT) and, 40
W Waldeyer’s ring of oral cavity, migration of B cells and, 211 warm hemolytic anemia, 385 water molecues at antibody–antigen interface, 493 Weizmann Institute, 38 white pulp cords of spleen, 208, 340 Wienands, Jurgen, 161 Willett, Catherine E., 449 Wnts, pluripotent hematopoietic stem cells (pHSC) and, 103 Wu, Gililan E., 473
X X-box binding protein (XBP), 255 X-linked (Bruton’s) agammaglobulinemia (XLA), 403–409, 404 atypical, 405 autoimmune disorders associated with, 404 B cell development and function in, 407–408 BTK mutations and, 405–409, 406, 407 calcium flux/transport in, 408 carrier detection in, 408–400 CD19 in, 408 CD20 in, 408 CD21 in, 408 CD34 in, 408 CD38 in, 408 deafness-dystonia protein gene and, 406 diagnosis of, 408–409 encephalomyelitis syndrome and, 403–404 growth hormone deficiency and, 407 HLA-DR, 408 IgM in, 407–408 malignancies associated with, 404–405, 404 Mohr-Tranebjaerg syndrome and, 406 neutropenia and, 404 poliomyelitis and, 403–404 T cell function in XLA, 408 XBP 1 in antibody production and response, 190 immunoglobulin assembly and secretion, 269, 270
XRCC4, nonhomologous end joining (NHEJ) and, 74, 76
Y yeast artificial chromosome (YAC) human IgH and, 2 translocus mice and human Mab and, 547, 549–552, 553 Vk genes, human and, 27 Vk genes, mouse and, 31 yeast display technology, monoclonal antibody display libraries and, 524–525, 524
Z Zachau, Hans G., 27 zebrafish (Danio rerio) immune system, 449–472 adaptive immunity in, 450, 452–457 B cell development in, 457 C regions in, 453–454 drug research and, 465 GATA-1 genes in, 459 GATA-2 genes in, 451–452, 459 genetic analysis and genome of, 449 genetic mapping in, 450 genetic screens and mutations in, 457–459, 465 hematopoiesis in, 450–452, 450, 451 human disease and, 459 IgM in, 453, 454–455 Ikaros gene in, 454, 456 immune system gene expression during development in, 455–457 immune system genes in, 453–454 infections and adaptive immunity in, 464 innate immunity in, 460–464 cellular effectors of, 461 complement system in, 462 cytokines in, 463–464 cytotoxic cells in, 462 pattern recognition receptors in, 463 phagocytic (myeloid) cells in, 461–462 intermediate cell mass (ICM) stage in, 451–452, 460 light chain gene encoding in, 455 lymphoid organs in, 452–453 major histocompatibility complex (MHC) I and II in, 460 markers used in research with, 449–450 morphants in, 459–460 patterns of gene expression in, 460 RAG gene in, 454, 456, 457, 456, 458, 458, 465 scl genes in, 451, 452 TCRa/b in, 454, 455, 456–457 thymus and T cell developoment in, 456–459 transgenesis in, 459 V regions in, 455 V(D)J recombination and, 453–454, 457 Zhang, Zhixin, 141 zinc fingers, recombination activating genes (RAG1/RAG2), 69
