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
Molecular Biology of B Cells
Elsevier Academic Press 84 Theobald’s Road, London WC1X 8RR, UK 525 B Street, Suite 1900, San Diego, California 92101-4495, USA Copyright © 2004, Elsevier Ltd. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Permissions may be sought directly from Elsevier’s Science & Technology Rights Department in Oxford, UK: phone: (+44) 1865 843830, fax: (+44) 1865 853333, e-mail:
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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
Preface
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 Polymorphism in VH Genes 20 Evolution 24 Genomic Considerations 24
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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 Human Immunoglobulin K Genes 27 Mouse Immunoglobulin K Genes 30 Aspects of Evolution of the K Genes 33
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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
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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) Innate Immunity 460 Infection 464
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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
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-
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
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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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Matsuda
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
TABLE 1.1
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
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
TABLE 1.2
(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.
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.
TABLE 1.3
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
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
74 psi
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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
D1 2M it4 1
Ca
D1 2M it1 9
D1 2M it1 8
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Cg2b
Cg1
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01 b
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
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3
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it2 0
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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
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.
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).
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.
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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
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 X04457 M15641
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 SYELTQPLS.VSVALGQTARITCGGN X97473,IGLV3-9 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 LPVLTQPPS.ASALLGASIKLTCTLS X57828,IGLV4-3 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-
TABLE 4.2
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).
Correspondence between serological lambda isotypes and IGLC gene and allele names Amino acid positions (1)
Serological isotype
1 (6) 112
IGLC gene and allele name
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 IMGT gene groups
IMGT gene names
IMGT functionality
List of the human IGL orphons
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
Mouse (Mus musculus, Mus spretus) IGLV germline genes
TABLE 4.7 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) IGLC
Mouse (Mus musculus, Mus spretus) IGLC genes and alleles
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
Mouse (Mus musculus, Mus spretus) IGLC RFLP Mouse (Mus musculus) TABLE 4.11
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.
C
H
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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
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.
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
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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
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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
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.
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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).
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).
Identification of Mammalian NHEJ Proteins
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
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
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
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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.
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
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).
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
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
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).
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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
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FIGURE 13.5 Three-color fluorescence of a tonsil GC to show the zonal 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
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. TABLE 14.1 Chemokine† CXCL9 (MIG) CXCL10 (IP10) CXCL11 (ITAC)
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
Chemokines involved in directing B cell movements
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
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
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
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.
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).
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;
15. Characteristics of Mucosal B Cells with Emphasis on the Human Secretory Immune System
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).
excessively aeroantigen-exposed conjunctiva, shows an IgA immunocyte density approaching that of the colon (Brandtzaeg, 1983a).
Role of Topical Antigen Exposure
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:
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
WHAT IS ACTUALLY KNOWN ABOUT HUMAN MUCOSAL B CELLS?
• 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
B
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.
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.
Survival Signals for Long-Lived 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
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
ADAPTIVE B CELL MEMORY The Interplay of Memory B and Memory Plasma Cells
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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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