Advances in Immunology Volume 89

Contents

Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Posttranscriptional Mechanisms Regulating the Inflammatory Response Georg Stoecklin and Paul Anderson 1. 2. 3. 4. 5. 6.

Abstract. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Coordinate Regulation of Pro-Inflammatory Proteins . . . . . . . . . . . Control of mRNA Stability. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Translational Inhibition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Animal Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Clinical Significance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1 1 5 16 19 23 25 26

Negative Signaling in Fc Receptor Complexes Marc Dae¨ron and Renaud Lesourne 1. 2. 3. 4. 5.

Abstract. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fc Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Positive Signaling by Activating FcRs . . . . . . . . . . . . . . . . . . . . . . . Negative Signaling by Activating FcRs . . . . . . . . . . . . . . . . . . . . . . Negative Signaling by Inhibitory FcRs . . . . . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

v

39 40 44 50 60 70 74

c on t e n ts

vi

The Surprising Diversity of Lipid Antigens for CD1-Restricted T Cells D. Branch Moody 1. 2. 3. 4. 5. 6. 7. 8.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction: From Molecules to Functions . . . . . . . . . . . . . . . . . . CD1 Protein Expression on Antigen-Presenting Cells . . . . . . . . . . Subcellular Lipid Antigen Processing Pathways. . . . . . . . . . . . . . . . 3-Dimensional Structures of CD1-b2-Microglobulin-Lipid Complexes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Microbial Antigens and Infectious Disease . . . . . . . . . . . . . . . . . . . Self Antigens, Autoreactivity, and Autoimmune Disease . . . . . . . . . Synthetic Lipid Antigens and Prospects for Immunotherapy . . . . . Conclusion: Prospects for Immunotherapy . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

87 87 90 94 102 111 119 125 126 128

Lysophospholipids as Mediators of Immunity Debby A. Lin and Joshua A. Boyce Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. LPL Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Cell Surface Receptors for LPLs and Their Signaling Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Expression of LPA and S1P Receptors by Immune Cells and Their Functions in In Vitro Studies . . . . . . . . . . . . . . . . . . . . . 5. In Vivo Functions of LPLS in Immune Responses and Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6. Clinical Applications of S1P Receptor Agonists. . . . . . . . . . . . . . . . 7. Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

141 141 143 145 148 156 159 160 160

Systemic Mastocytosis Jamie Robyn and Dean D. Metcalfe 1. 2. 3. 4.

Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mast Cell Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Biology of Kit . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mastocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

169 169 170 184 193

c o nt e n t s 5. 6. 7. 8. 9.

Prognosis and Predictive Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Supportive Care and Long-Term Management . . . . . . . . . . . . . . . . Future Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

vii 209 209 216 218 220 220

Regulation of Fibrosis by the Immune System Mark L. Lupher, Jr. and W. Michael Gallatin 1. 2. 3. 4. 5. 6. 7.

Abstract. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fibrotic Disease Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cellular Mediators of Fibrosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Inflammatory Chemokines that Regulate Fibrosis. . . . . . . . . . . . . . The Role of Integrins in Regulating the Fibrotic Response . . . . . . Other Potential Targets for Anti-Fibrotic Therapy . . . . . . . . . . . . . Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

245 245 246 249 261 264 268 270 273

Immunity and Acquired Alterations in Cognition and Emotion: Lessons from SLE Betty Diamond, Czeslawa Kowal, Patricio T. Huerta, Cynthia Aranow, Meggan Mackay, Lorraine A. DeGiorgio, Ji Lee, Antigone Triantafyllopoulou, Joel Cohen-Solal, and Bruce T. Volpe 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

Abstract. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lupus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Neuropsychiatric Lupus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mechanisms of NPSLE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mouse Models of NPSLE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Peptide Reactivity of DNA-Reactive Antibodies . . . . . . . . . . . . . . . Presence of DWEYS-Reactive Antibodies in Murine and Human SLE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Proteins Harboring the D/E W D/E Y S/G Concensus Sequence: Glutamate Receptors . . . . . . . . . . . . . . . . . . . . . . . . . . . Antibody-Mediated Neurotoxicity . . . . . . . . . . . . . . . . . . . . . . . . . . A Murine Model for Antibody-Mediated Neuronal Death . . . . . . . Antibody-Mediated Neurotoxicity in the Amygdala. . . . . . . . . . . . .

289 290 293 295 296 298 298 300 300 302 304 307

viii 12. 13. 14. 15. 16.

c on t e n ts Evidence that Antibodies Are Involved in NPSLE in Patients . . . . Anti-Peptide Antibody Activates Prolactin Secretion . . . . . . . . . . . . Anti-Peptide Antibodies and Manifestations of NP-SLE. . . . . . . . . Implications for SLE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Implications for Human Pathobiology . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

308 309 310 310 311 311

Immunodeficiencies with Autoimmune Consequences Luigi D. Notarangelo, Eleonora Gambineri, and Raffaele Badolato Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1. Autoimmune Manifestations in Primary Immune Deficiencies: Relevance and Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. AIRE, Central Tolerance, and the Pathophysiology of Autoimmune Polyendocrinopathy. . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Defective AIRE Expression Explains the Pathophysiology of Autoimmunity in Omenn Syndrome . . . . . . . . . . . . . . . . . . . . . . . . 4. CD4þ CD25þ Regulatory T Cells and the Pathophysiology of IPEX (Immunodysregulation – Polyendocrinopathy – Enteropathy – X-Linked) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Concluding Remarks. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

321 322 324 337

346 357 359

Index. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371 Contents of Recent Volumes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 385

Contributors

Numbers in parentheses indicate the pages on which the authors’ contributions begin.

Paul Anderson (1), Division of Rheumatology, Immunology, and Allergy, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts Cynthia Aranow (287), Department of Medicine, Columbia University Medical Center, New York, New York Raffaele Badolato (319), ‘‘Angelo Nocivelli’’ Institute for Molecular Medicine, Department of Pediatrics, University of Brescia, Brescia, Italy Joshua A. Boyce (141), Departments of Medicine and Pediatrics, Harvard Medical School, Division of Rheumatology, Immunology, and Allergy, Brigham and Women’s Hospital, Boston, Massachusetts Joel Cohen-Solal (287), Department of Medicine, Columbia University Medical Center, New York, New York Marc Dae¨ron (39), Unite´ d’Allergologie Mole´ culaire et Cellulaire, De´partement d’Immunologie, Institut Pasteur, Paris, France Lorraine A. DeGiorgio (287), Department of Neurology and Neuroscience, The Burke Medical Research Institute, Weill Medical College of Cornell University, White Plains, New York Betty Diamond (287), Department of Medicine, Columbia University Medical Center, New York, New York W. Michael Gallatin* (245), ICOS Corporation, Bothell, Washington Eleonora Gambineri (319), Department of Pediatrics, University of Florence, Florence, Italy Patricio T. Huerta (287), Department of Neurology and Neuroscience, The Burke Medical Research Institute, Weill Medical College of Cornell University, White Plains, New York Czeslawa Kowal (287), Department of Medicine, Columbia University Medical Center, New York, New York *Current address: Frazier Healthcare Ventures, Seattle, Washington.

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Ji Lee (287), Department of Microbiology and Immunology, Albert Einstein College of Medicine, Bronx, New York Renaud Lesourne (39), Unite´ d’Allergologie Mole´culaire et Cellulaire, De´partement d’Immunologie, Institut Pasteur, Paris, France Debby A. Lin (141), Department of Medicine, Harvard Medical School, and Division of Rheumatology, Immunology, and Allergy, Brigham and Women’s Hospital, Boston, Massachusetts Mark L. Lupher, Jr. (245), ICOS Corporation, Bothell, Washington Meggan Mackay (287), Department of Medicine, Columbia University Medical Center, New York, New York Dean D. Metcalfe (169), Laboratory of Allergic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, Maryland D. Branch Moody (87), Division of Rheumatology, Immunology, and Allergy, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts Luigi D. Notarangelo (319), ‘‘Angelo Nocivelli’’ Institute for Molecular Medicine, Department of Pediatrics, University of Brescia, Brescia, Italy Jamie Robyn (169), Laboratory of Allergic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, Maryland Georg Stoecklin (1), Division of Rheumatology, Immunology, and Allergy, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts Antigone Triantafyllopoulou (287), Department of Medicine, Montefiore Medical Center, Bronx, New York Bruce T. Volpe (287), Department of Neurology and Neuroscience, The Burke Medical Research Institute, Weill Medical College of Cornell University, White Plains, New York

Posttranscriptional Mechanisms Regulating the Inflammatory Response Georg Stoecklin and Paul Anderson Division of Rheumatology, Immunology, and Allergy; Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts

1. 2. 3. 4. 5. 6.

Abstract............................................................................................................. Coordinate Regulation of Pro‐Inflammatory Proteins ................................................. Control of mRNA Stability .................................................................................... Translational Inhibition......................................................................................... Animal Models ................................................................................................... Clinical Significance............................................................................................. Conclusions........................................................................................................ References .........................................................................................................

1 1 5 16 19 23 25 26

Abstract The inflammatory response is a complex physiologic process that requires the coordinate induction of cytokines, chemokines, angiogenic factors, effector‐ enzymes, and proteases. Although transcriptional activation is required to turn on the inflammatory response, recent studies have revealed that posttranscriptional mechanisms play an important role by determining the rate at which mRNAs encoding inflammatory effector proteins are translated and degraded. Most posttranscriptional control mechanisms function to dampen the expression of pro‐inflammatory proteins to ensure that potentially injurious proteins are not overexpressed during an inflammatory response. Here we discuss the factors that regulate the stability and translation of mRNAs encoding pro‐inflammatory proteins. 1. Coordinate Regulation of Pro‐Inflammatory Proteins In prokaryotes, the coordinate expression of genes encoding components of a metabolic pathway is often accomplished by expressing an mRNA transcript that encodes more than one protein. For example, the Lactose operon in Escherichia coli transcribes a single mRNA that encodes b‐galactosidase, permease, and transacetylase—proteins required for the utilization of lactose as a carbon source (Beckwith, 1967). Unlike their prokaryotic ancestors, processed eukaryotic transcripts generally encode a single protein. In eukaryotes, the coordinate expression of protein components of a complex functional program is accomplished by the use of regulatory nucleic acid sequences

1 advances in immunology, vol. 89 # 2006 Elsevier Inc. All rights reserved.

0065-2776/06 $35.00 DOI: 10.1016/S0065-2776(05)89001-7

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that determine rates of transcription, translation, and mRNA decay. Coordinate transcriptional control is accomplished by the selective use of promoter elements that are recognized by specific transcription factors. Activation of a transcription factor that turns on the expression of several genes involved in a common biological pathway can be sufficient to confer coordinate protein expression. One example is the heat shock response in which stress‐induced activation of heat shock factor 1 confers de novo transcription of mRNAs encoding several heat shock proteins (Voellmy, 2004). Mammalian cells have evolved posttranscriptional mechanisms that further coordinate the expression of proteins involved in a common cellular function. In some cases, regulation of mRNA stability and translation may be sufficient to coordinately regulate the expression of distinct functional classes of proteins. This appears to be the case in Saccharomyces cerevisiae in which gene array profiling has shown that mRNAs encoding subunits of complex structures such as proteasomes or ribosomes tend to have similar half‐lives (Wang et al., 2002). In many cases, posttranscriptional regulation is conferred by cis‐acting elements located in the 30 ‐untranslated region (30 UTR) of individual mRNAs. These regulatory elements recruit specific RNA‐binding proteins that, directly or indirectly, regulate mRNA translation and/or stability. A striking example of this type of regulation is provided by a family of Pumilio‐ Fem‐3‐binding factor (Puf) proteins. The Puf proteins bind to variants of a UGUR tetranucleotide motif found in the 30 UTR of selected mRNAs to regulate mRNA stability and/or translation (Wickens et al., 2002). In Saccharomyces cerevisiae, gene array profiling has revealed that Puf proteins can coordinately regulate the expression of mRNAs encoding proteins with a common function (Gerber et al., 2004). Thus, Puf1p and Puf2p were found to bind to mRNAs encoding membrane‐associated proteins, Puf3p binds to mRNAs encoding mitochondrial proteins, and Puf4p and Puf5p bind to mRNAs encoding nuclear proteins. Each Puf protein binds to a specific variant of the UGUR motif to confer specificity. These results strongly support the contention that RNA‐binding proteins coordinately regulate groups of mRNAs encoding proteins of a common function. This general mechanism has been referred to as a ‘‘post‐transcriptional operon’’ (Keene and Tenenbaum, 2002). The inflammatory response is an example of a functional program that requires the coordinate induction of proteins involved in a common function. During the inflammatory response, coordinate transcriptional control is accomplished by the selective use of promoter elements that are recognized by specific transcription factors. For example, the transcription factor Nuclear factor (NF)‐kB moves from the cytoplasm to the nucleus in T‐cells and macrophages that are exposed to inflammatory stimuli (Muller, 2001). Nuclear

P O S T T R A N S C R I P T I O N A L C O N T R O L O F I N F L A M M AT I O N

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NF‐kB binds to a nucleotide sequence element common to the promoters of several genes that encode inflammatory effector proteins (Muller, 2001). Although transcriptional activation is required for the synthesis of mRNA encoding inflammatory effector proteins, it does not determine the level of protein expression. Rather, it is the posttranscriptional regulation of mRNA stability and translation that determines levels of protein expression. Remarkably, mRNAs encoding proteins that regulate every facet of the inflammatory response are subject to these posttranscriptional control mechanisms. At the initiation phase of inflammation, Granulocyte‐macrophage colony‐stimulating factor (GM‐CSF) primes neutrophils, macrophages, and dendritic cells for subsequent pro‐inflammatory action (Hamilton, 2002). Moreover, the expression of chemokines that recruit neutrophils, eosinophils, monocytes, and dendritic cells to sites of inflammation is strongly regulated at the posttranscriptional level. Effects on cellular recruitment are potentiated by the posttranscriptional regulation of the endothelial adhesion factor Vascular cell adhesion molecule (VCAM)‐1. During the effector phase of inflammation, the expression of pro‐inflammatory cytokines such as Tumor necrosis factor (TNF)a, Interleukin (IL)‐1b, IL‐6, and Interferon (IFN)g is also subject to posttranscriptional control. Excessive production of these cytokines orchestrates tissue damage in patients with inflammatory arthritis and inflammatory bowel disease. Proteases such as Matrix metalloprotease (MMP)‐9 and MMP‐ 13 participate in the tissue damage by breaking down components of the extracellular matrix. The effector phase is further exacerbated by enzymes that synthesize pro‐inflammatory mediators such as Cyclooxygenase (COX)‐2, 15‐Lipoxygenase (LO), and inducible Nitric oxide synthase (iNOS), which are also regulated at the level of mRNA stability and translation. Taken together, it is clear that posttranscriptional control mechanisms coordinately regulate multiple aspects of the inflammatory response. In general, these mechanisms conspire to dampen inflammation. This additional level of control may be required to prevent the pathological overexpression of proteins that are potentially injurious to the host. The synchronized anti‐inflammatory effects of these regulatory programs make them ideal targets for the development of anti‐ inflammatory drugs. In this review, we will focus on two mechanisms that regulate the fate of mRNAs in the cytoplasm: control of translation efficiency and control of mRNA stability. Although earlier steps (pre‐mRNA splicing, export of the mRNA from the nucleus into the cytoplasm) and later events (protein secretion, activation by proteolytic cleavage) are extensively regulated as well, they will not be discussed here. Table 1 lists pro‐inflammatory proteins whose expression is regulated at the level of translation or mRNA stability. Most of these proteins fall into one of three categories: cytokines, chemokines, and enzymes required for the synthesis of pro‐inflammatory mediators.

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Table 1 Pro‐Inflammatory Proteins Regulated at the Posttranscriptional Level mRNA

Mechanism

Element

References

Cytokines IL‐1b

mRNA stability

AREa

IL‐1F7b IL‐2

mRNA stability mRNA stability

CRDb ARE, JREc

IL‐3

mRNA stability

ARE

IL‐6

mRNA stability

ARE

IL‐11

mRNA stability

ARE

IL‐18 TNFa

mRNA stability mRNA stability, translation

n.d. ARE, CDEd

GM‐CSF

mRNA stability, translation

ARE

G‐CSF

mRNA stability

ARE, SLDEe

IFNb

mRNA stability

ARE, CRD

IFNg

mRNA stability

ARE

VEGF

mRNA stability

ARE, 50 UTR, CRD

MCP‐1 (CCL2) Eotaxin (CCL11) GROa (KC, CXCL1) GROb (MIP‐2, CXCL2) GROg (CXCL3) IL‐8 (CXCL8)

mRNA stability mRNA stability

n.d. ARE

(Pastore et al., 2005) (Atasoy et al., 2003)

mRNA stability

ARE

mRNA stability

ARE

mRNA stability mRNA stability

ARE ARE

IP‐10 (CXCL10)

mRNA stability

n.d.

(Biswas et al., 2003; Sirenko et al., 1997; Stoeckle, 1991) (Rousseau et al., 2002; Stoeckle, 1991) (Stoeckle, 1991) (Holtmann et al., 1999; Stoeckle, 1991; Winzen et al., 1999) (Vockerodt et al., 2005)

(Fenton et al., 1988; Sirenko et al., 1997) (Bufler et al., 2004) (Chen et al., 1998; Lindsten et al., 1989; Ogilvie et al., 2005) (Nair et al., 1994; Wodnar‐Filipowicz and Moroni, 1990) (Akashi et al., 1990; Neininger et al., 2002; Stoecklin et al., 2001; Winzen et al., 1999) (Bamba et al., 2003; Yang and Yang, 1994) (Bufler et al., 2004) (Brook et al., 2000; Carballo et al., 1998; Han et al., 1990; Stoecklin et al., 2003) (Carballo et al., 2000; Grosset et al., 2004; Koeffler et al., 1988; Shaw and Kamen, 1986) (Brown et al., 1996a; Koeffler et al., 1988; Putland et al., 2002) (Raj and Pitha, 1983; Whittemore and Maniatis, 1990) (Hodge et al., 2002; Lindsten et al., 1989; Mavropoulos et al., 2005) (Coles et al., 2004; Dibbens et al., 1999; Ikeda et al., 1995; Levy et al., 1998)

Chemokines

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Table 1 (Continued) mRNA

Mechanism

Element

References

Enzymes and other pro‐inflammatory proteins VCAM‐1

mRNA stability

n.d.

MMP‐9 MMP‐13 15‐LO iNOS

mRNA stability translation translation mRNA stability

ARE ARE DICE f ARE

COX‐2

mRNA stability, translation

ARE

(Croft et al., 1999; Iademarco et al., 1995) (Huwiler et al., 2003) (Yu et al., 2003) (Ostareck et al., 1997) (Di Macro et al., 2005; Rodriguez‐Pascual et al., 2000) (Dixon et al., 2003; Lasa et al., 2000; Mukhopadhyay et al., 2003; Ridley et al., 1998; Ristimaki et al., 1994)

a

ARE: AU‐rich element. CRD: coding region determinant of instability. c JRE: JNK response element. d CDE: constitutive decay element. e SLDE: stem‐loop destabilizing element. f DICE: differentiation control element. b

2. Control of mRNA Stability 2.1. mRNA Decay Mediated by the AU‐Rich Element Many of the pro‐inflammatory transcripts that are regulated at the posttranscriptional level have an adenine/uridine‐rich element (ARE) in their 30 UTR, which targets individual transcripts for rapid cytoplasmic degradation. AREs were initially discovered by their characteristic nucleotide pattern, and the high degree of sequence conservation between different mammalian species was an early indication of their important regulatory role (Caput et al., 1986). ARE‐mediated mRNA decay (AMD) was first demonstrated by Shaw and Kamen (1986), who inserted the ARE of GM‐CSF into the 30 UTR of a normally stable reporter mRNA (in this case a b‐globin transcript), and observed that it strongly reduced reporter gene expression by destabilizing the mRNA (Shaw and Kamen, 1986). Conversely, deletion of the ARE from the 30 UTR of IL‐3 or TNFa enhances gene expression by stabilizing the mRNA (Kontoyiannis et al., 1999; Stoecklin et al., 1994). Besides inducing AMD, quantitative analysis of TNFa reporter gene expression showed that the ARE also inhibits translation of the mRNA (Han et al., 1990). AREs are U‐rich sequences that contain several copies of a canonical AUUUA pentamer.

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According to a classification that takes into account deadenylation kinetics, the typical class II AREs of cytokine mRNAs (e.g., GM‐CSF, IL‐3, and TNFa) contain a cluster of 4–7 partially overlapping AUUUA pentamers within a U‐rich context (Chen and Shyu, 1995). Class I AREs (e.g., c‐myc, c‐fos) have fewer and more scattered AUUUA pentamers, while class III AREs (e.g., c‐jun) lack this motif. Class II AREs trigger a very rapid, asynchronous shortening of the poly‐A tail (processive deadenylation activity) prior to decay of the mRNA body, whereas class I and III AREs are associated with a less rapid, synchronous shortening of the poly‐A tail (distributive deadenylation activity) (Chen et al., 1995). For class II AREs, the AUUUA pentamers are of crucial importance, since point mutations within the pentamers efficiently abrogate AMD (Stoecklin et al., 1994). The extended UUAUUUAUU nonamer is the minimal sequence that can induce mRNA degradation (Lagnado et al., 1994; Zubiaga et al., 1995). This sequence is also the minimal binding site for tristetraprolin (TTP), an ARE‐binding protein required for AMD (see below). Compared to this minimal sequence, typical AREs are much longer (50–100 nucleotides), indicating that AREs of different mRNAs may vary with respect to their destabilizing activity, their ability to inhibit translation, and the regulatory mechanisms that control these activities. Since AREs were first discovered in cytokine transcripts, the biological importance of controlling mRNA stability has primarily been established in cells of the immune system. Nevertheless, AMD is a highly conserved regulatory mechanism that functions in yeast (Vasudevan and Peltz, 2001), trypanosomes (Quijada et al., 2002), Drosophila (Jing et al., 2005), and virtually all mammalian cell lines. Based on the frequency of AU‐rich sequences in the human genome, it has been estimated that 5–10% of all mRNAs may contain an ARE (Bakheet et al., 2003). Although experimental data is still needed to confirm this high estimate of functional AREs, it is clear that AMD is a general mechanism that regulates gene expression in most, if not all, eukaryotic cells. 2.2. The Different Roles of ARE‐Binding Proteins More than 20 different proteins that bind to AREs have so far been identified, but only a subset of them has been shown to influence the stability or translation efficiency of their target mRNAs. Table 2 gives an overview of ARE‐binding proteins (ARE‐BPs) that are known to have a regulatory function: TIA‐1, TIAR, FXR1P, and CUGBP2 inhibit the translation of ARE‐ mRNAs; HuR and YB1 stabilize ARE‐mRNAs; whereas AUF1, KSRP, RHAU, and the TTP/BRF family of proteins are involved in destabilizing ARE‐mRNAs.

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Table 2 ARE‐Binding Proteins ARE‐BP TIA‐1 TIAR FXRIP CUGBP2 HuR

Domains

Function

References

RRM (3)*

Inhibits translation

(Piecyk et al., 2000)

RRM (3) KH (2), RGG (1) RRM (3) RRM (3)

Inhibits translation Inhibits translation Inhibits translation Stabilizes mRNA

(Yu et al., 2003) (Garnon et al., 2005) (Mukhopadhyay et al., 2003) (Fan and Steitz, 1998; Lal et al., 2004; Peng et al., 1998; Wang et al., 2000) (Lal et al., 2004; Mazan‐Mamczarz et al., 2003) (Kullmann et al., 2002) (Capowski et al., 2001; Chen et al., 2000; Coles et al., 2004) (Lal et al., 2004; Loflin et al., 1999; Sarkar et al., 2003) (Xu et al., 2001) (Dean et al., 2002) (Chen et al., 2001; Gherzi et al., 2004) (Tran et al., 2004) (Carballo et al., 1998; Chen et al., 2001; Lai et al., 1999; Ming et al., 2001) (Lai et al., 2000; Stoecklin et al., 2000, 2002) (Lai et al., 2000)

Activates translation

YB1

CSD (1)

Inhibits translation Stabilizes mRNA

AUF1 (hnRNP D0)

RRM (2)

Destabilizes mRNA

AUF2 (CBF‐A) KSRP

RRM (2) KH (4)

Stabilizes mRNA Stabilizes mRNA Destabilizes mRNA

RHAU TTP (TIS11)

Helicase (1) C3H (2)

Destabilizes mRNA Destabilizes mRNA

BRF1 (TIS11b)

C3H (2)

Destabilizes mRNA

BRF2 (TIS11d)

C3H (2)

Destabilizes mRNA

*Designates the number of domains.

Genetic studies have provided evidence that the zinc finger proteins TTP, BRF1, and BRF2 play a central role in the degradation of ARE‐mRNAs. The TTP/BRF proteins bind to AREs with high specificity through their characteristic tandem C3H zinc finger domains (Blackshear, 2002; Varnum et al., 1991). The function of TTP was discovered by the study of TTP‐deficient mice, which develop generalized inflammatory symptoms that arise from increased levels of TNFa and GM‐CSF (Taylor et al., 1996). Cytokine overproduction in TTP
/
mice is due to an increased stability of TNFa, GM‐CSF, and IL‐2 mRNAs (Carballo et al., 1998, 2000; Ogilvie et al., 2005). TTP normally binds to the ARE of these mRNAs and promotes their rapid degradation by targeting them to the cellular RNA degradation machinery (see below). Since TTP is mainly expressed in activated macrophages and T‐cells (Cao et al., 2004; Raghavan

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et al., 2001), its principal function appears to be the control of cytokine expression in the immune system. Yet TTP is also expressed in a variety of other tissues such as lung, liver, and intestine (Cao et al., 2004; Lu and Schneider, 2004), and may thus be involved in regulating other processes. BRF1 and BRF2 are homologues of TTP which share about 70% sequence identity in the highly conserved zinc finger domain, but differ substantially in the N‐ and C‐terminal domains (Varnum et al., 1991). BRF1‐deficient mice are not viable due to embryonic lethality at day 11 post‐fertilization, probably as a result of severe placental dysfunction (Stumpo et al., 2004). A mutant cell line generated by chemical mutagenesis and selected for a defect in AMD has provided evidence for the important role of BRF1 (Stoecklin et al., 2000, 2002). This mutant cell line lacks expression of BRF1 due to point mutations in both alleles, and is unable to rapidly degrade reporter transcripts containing the AREs of TNFa, GM‐CSF, IL‐2, IL‐3, and IL‐6 (Stoecklin et al., 2001). The results obtained with TTP
/
mice and BRF1
/
cells indicate that cytokine transcripts with class II AREs are subject to a common degradation pathway. Overexpression studies have shown that BRF2, the second homologue of TTP, can also induce AMD (Lai et al., 2000). Mice expressing an N‐terminally deleted form of the BRF2 protein are viable, but exhibit complete female infertility due to block of early embryonic development at the two‐cell stage (Ramos et al., 2004). Taken together, the data from the knock‐out mice reveal that TTP plays a critical and non‐redundant role in the immune system, whereas both BRF1 and BRF2 appear to be important regulators of embryonic development. It is noteworthy that prolonged overexpression of TTP and BRF1 causes apoptotic cell death in a variety of cell lines (Johnson and Blackwell, 2002; Johnson et al., 2000), indicating that the TTP/BRF family of proteins may target yet unidentified mRNAs. Characterizing the entire spectrum of mRNAs that are regulated by TTP/BRF will help to uncover biological processes for which posttranscriptional control of mRNA stability is relevant. Although enforced tethering of TTP to an mRNA that does not contain an ARE (through a heterologous RNA‐protein interaction) is sufficient to induce rapid degradation of the mRNA (Lykke‐Andersen and Wagner, 2005), it is clear that other ARE‐BPs are also required for destabilizing ARE‐mRNAs. KSRP was initially identified as an RNA‐binding protein that enhances splicing (Min et al., 1997), and later found to interact with the 30 UTR of IL‐2 transcripts (Chen et al., 2001). KSRP binds to different AREs (e.g., IL‐2, TNFa, c‐fos) through its KH domains, and simultaneously interacts with the exosome, a large complex of about ten 30 ‐50 exonucleases and associated helicases (Mitchell and Tollervey, 2000; Raijmakers et al., 2004). Importantly, KSRP was shown to be required for AMD both in vitro and in live cells (Chen et al., 2001; Gherzi et al., 2004). The helicase RHAU was identified as a protein that

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binds to the ARE of urokinase plasminogen activator (uPA) and also interacts with the exosome (Tran et al., 2004). RHAU accelerates degradation of uPA mRNA, but it is not clear whether RHAU facilitates the degradation of other ARE‐mRNAs as well. The ARE‐BP AUF1 was identified as an activity that destabilizes an ARE‐ containing RNA in vitro (Brewer, 1991; Zhang et al., 1993). The destabilizing role of AUF1 has been confirmed by studies using siRNA to suppress AUF1 expression (Lal et al., 2004; Raineri et al., 2004), as well as by overexpression studies (Loflin et al., 1999; Sarkar et al., 2003). Its ability to interact with the exosome may provide the mechanistic basis for its activity (Chen et al., 2001). At least one study, however, has indicated that overexpression of AUF1 can also cause stabilization of ARE‐containing reporter mRNAs (Xu et al., 2001). This apparent contradiction may arise from the fact that AUF1 is expressed as four different isoforms (Wagner et al., 1998), which can differ in their activity (Raineri et al., 2004). Interestingly, overexpression of AUF2, a close homologue of AUF1, also stabilizes an ARE‐mRNA (Dean et al., 2002). HuR and its neural‐specific homologues HuB, HuC, and HuD form the embryonic lethal abnormal visual (ELAV) family of RNA‐binding proteins, all of which have high affinity for AU‐ and U‐rich sequences (Antic and Keene, 1997; Ma et al., 1996). Various studies have shown that HuR, the ubiquitously expressed member of the ELAV family, stabilizes a large number of ARE‐ mRNAs, including transcripts containing the ARE of GM‐CSF, IL‐3, and VEGF (Fan and Steitz, 1998; Ford et al., 1999; Levy et al., 1998; Ming et al., 2001; Peng et al., 1998; Raineri et al., 2004). HuR is required for the induced expression of iNOS (Di Macro et al., 2005; Rodriguez‐Pascual et al., 2000), and has also been implicated in the stabilization of IL‐8, eotaxin, COX‐2, and MMP‐9 mRNA (Atasoy et al., 2003; Cok et al., 2003; Huwiler et al., 2003; Winzen et al., 2004). Class II AREs typically contain 4 to 7 AUUUA pentamers within a U‐rich sequence of 50 or more nucleotides. This contrasts with the fact that TTP requires only the nonameric sequence UUAUUUAUU for efficient binding (Blackshear et al., 2003a; Worthington et al., 2002), and that a single C3H zinc finger of BRF2 makes direct contact with just four nucleotides: UAUU (Hudson et al., 2004). This would indicate that physiological AREs are large enough to accommodate more than one, and perhaps several ARE‐BPs. A simple model would postulate that stabilizing proteins such as HuR compete with destabilizing proteins such as TTP or AUF1 for binding to the ARE, and thereby prevent decay of the mRNA. It appears, however, that HuR does not bind within the core region of class II AREs (which consists of clustered AUUUA pentamers), but to an auxiliary, mainly U‐rich region upstream of the core region (Chen et al., 2002; Winzen et al., 2004). Paradoxically, this

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auxiliary region both enhances the destabilizing activity of the ARE core region, and allows for stabilization of the mRNA by HuR. Such an upstream auxiliary region has been described for the AREs of c‐fos, TNFa, GM‐CSF, and IL‐8 (Chen et al., 2002; Stoecklin et al., 2001; Winzen et al., 2004). Very few studies have so far directly addressed the question whether ARE‐BPs bind individually to the ARE and compete for binding, or whether they bind cooperatively to form multi‐protein ARE‐mRNPs. One study found that KSRP and TTP together compete with HuR for binding to the Pitx2 mRNA (Briata et al., 2003). Using a genome‐wide approach, another study was able to distinguish between mRNAs that are simultaneously bound by both AUF1 and HuR, and mRNAs bound by the two proteins individually (Lal et al., 2004). Given the number of proteins which can bind to AREs (at least 20), and the fact that individual AREs differ quite considerably in their sequence, it is likely that each ARE recruits a different subset of ARE‐BPs, which together determine the stability and translation efficiency of that particular transcript. At present, we lack a broader understanding of the composition of specific ARE‐ mRNP complexes, and the changes these complexes may undergo when the degradation of the mRNA is activated or inhibited. 2.3. The Pathway of ARE‐mRNA Degradation In general, degradation of mRNAs in the cytoplasm is initiated by removal of the poly‐A tail, followed by exonucleolytic decay in the 30 ‐50 or the 50 ‐30 direction. At the 30 end, the mRNA is degraded through the exosome. At the 50 end, the 7‐methyl guanosine cap is removed by the decapping complex Dcp1/Dcp2, and the mRNA body is subsequently degraded by the 50 ‐30 exonuclease Xrn1 (Cougot et al., 2004b; Fillman and Lykke‐Andersen, 2005; Parker and Song, 2004). Dcp1/Dcp2 and Xrn1 form a larger complex with the Lsm1‐7 proteins, all of which are concentrated in small cytoplasmic foci termed processing bodies (Cougot et al., 2004a; Ingelfinger et al., 2002; van Dijk et al., 2002). Processing bodies are considered to be sites of mRNA degradation in the cytoplasm. In vitro decay studies have indicated that ARE‐mRNAs are degraded in the 30 ‐50 direction by the exosome (Chen et al., 2001), and one of the exosome components, Pm‐Scl‐75, was shown to directly bind to the ARE (Mukherjee et al., 2002). On the other hand, the ARE also stimulates decapping of the RNA in vitro (Gao et al., 2001). As illustrated in Fig. 1, the TTP/BRF family of proteins plays a central role in the degradation of ARE‐mRNAs. Both in vitro and when overexpressed in live cells, TTP enhances deadenylation of ARE‐ mRNA (Fig. 1A) (Lai et al., 1999, 2003). TTP and BRF1 further interact with both the exosome (Fig. 1B) and with components of the decapping/Xrn1 complex (Fig. 1C) (Chen et al., 2001; Gherzi et al., 2004; Lykke‐Andersen

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Figure 1 The TTP/BRF family of proteins plays a central role in the degradation of ARE‐mRNAs. (A) Cytoplasmic mRNA degradation is initiated by deadenylation. (B) 30 ‐50 decay is mediated by the exosome (green), a complex of exonucleases and helicases. (C) At the 50 end, the mRNA is decapped by Dcp1/2 (yellow), followed by Xrn1‐mediated 50 ‐30 degradation. These proteins form a larger complex with Lsm1‐7, and are concentrated in cytoplasmic processing bodies. (D) The RNA‐induced silencing complex (RISC, in blue) and microRNA 16 are also required for ARE‐ mRNA degradation. The zinc finger proteins TTP/BRF1 (red) bind with high affinity to the ARE and induce degradation of the mRNA by interacting with the exosome, the decapping/Xrn1 complex, and the RISC complex.

and Wagner, 2005). This suggests that TTP/BRF1 are capable of targeting ARE‐mRNAs to both decay pathways. We have recently shown that the 50 ‐30 pathway is indeed important for degrading ARE‐mRNAs (Stoecklin et al., 2005). Using siRNA to target individual components of both pathways, Xrn1 and Lsm1 were found to be essential for AMD, whereas exosome components are less important. TTP and BRF1 also colocalize with processing bodies, a further indication that AMD uses the 50 ‐30 pathway (Kedersha et al., 2005). A recent study by Jing and coworkers (2005) revealed that AMD is dependent on the RNA‐induced silencing complex (RISC) and the microRNA miR16 (Fig. 1D). TTP interacts with two proteins of the RISC complex, Argonaute 2 and 4, which in turn allows the microRNA miR16 to base‐pair with the ARE. Interestingly, Argonaute proteins also colocalize with processing bodies and mRNAs targeted by miRNAs become concentrated in these structures (Liu et al., 2005). These novel findings indicate that the ARE actually consist of a dual code: one part of the ARE serves to recruit RNA‐binding proteins which interact with the cellular degradation machinery, while another part of the ARE (or adjacent sequences) base‐pair with regulatory microRNAs, which may contribute to the sequence specificity of the destabilizing element. A challenge of future studies will be to break this code in a way that will allow

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us to predict for individual AREs which proteins bind to it, which microRNAs are associated, and what the functional consequences are for mRNA stability and translation efficiency. 2.4. Regulation of ARE‐mRNA Decay The ARE not only targets the mRNA for rapid degradation, but also allows the tight regulation of mRNA stability in response to extracellular cues. In a variety of cell types, many cytokine mRNAs have a very short half‐life (in the range of 10–30 minutes) in resting cells, effectively preventing cytokine protein production. Upon cell stimulation, the rapid induction of cytokine expression requires both transcriptional activation and stabilization of the mRNAs. In activated cells, cytokine mRNA half‐lives are in the range of several hours, which can easily account for a 10‐fold increase in protein production. In the post‐induction phase, ARE‐mRNAs are no longer stabilized and their degradation ensures rapid return to the low, usually undetectable levels of basal cytokine production. Thus, the dynamic regulation of mRNA stability is an important means by which cytokine levels are controlled in a time‐dependent manner. The following examples (listed in Table 1) document the general importance of this mode of posttranscriptional regulation. In T‐cells stimulated with anti‐CD3 and anti‐CD28 antibodies, or phorbol ester, induction of IL‐2, IFNg, TNFa, and GM‐CSF coincides with a marked increase in the stability of the corresponding mRNAs (Bickel et al., 1990; Lindsten et al., 1989). Stabilization of IL‐3 and GM‐CSF mRNA occurs in mast cells after treatment with calcium ionophores (Wodnar‐Filipowicz and Moroni, 1990), and GM‐CSF mRNA is likewise stabilized in stimulated eosinophils (Esnault and Malter, 1999). The production of IFNg in activated NK cells involves stabilization of the mRNA (Hodge et al., 2002; Mavropoulos et al., 2005). Macrophages produce a variety of cytokines and chemokines in response to activation by lipopolysaccharide (LPS), and mRNA stabilization contributes to the induction of IL‐1b, IL‐18, TNFa, GM‐CSF, Growth regulated oncogene (GRO)a and GROb expression (Biswas et al., 2003; Brook et al., 2000; Bufler et al., 2004; Carballo et al., 2000; Rousseau et al., 2002). In monocytes activated through adhesion, mRNAs encoding IL‐1b and GROa are stabilized (Sirenko et al., 1997). Regulation of ARE‐mRNA stability is not restricted to the cells of the immune system. Virally infected fibroblasts produce IFNb in the early phase of infection. During this phase, IFNb mRNA is stable, whereas rapid degradation of IFNb mRNA during the late phase accounts for the post‐induction turnoff of IFNb expression (Raj and Pitha, 1983; Whittemore and Maniatis, 1990). Stimulation of fibroblasts and other cell types with pro‐inflammatory

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cytokines such as TNFa or IL‐1 induces the production of various cytokines and chemokines. In these cells, mRNA stabilization contributes to the induced expression of IL‐6, IL‐11, G‐CSF, GM‐CSF, Macrophage chemoattractant protein (MCP)‐1, eotaxin, GROa, GROb, GROg, and IL‐8 (Akashi et al., 1990; Atasoy et al., 2003; Bamba et al., 2003; Koeffler et al., 1988; Pastore et al., 2005; Stoeckle, 1991; Yang and Yang, 1994). Expression of the adhesion molecule VCAM‐1 on endothelial cells and fibroblast‐like synoviocytes is induced by TNFa and IL‐4, and involves stabilization of the mRNA (Croft et al., 1999; Iademarco et al., 1995). Since VCAM‐1 is overexpressed in rheumatoid arthritis synovium (Morales‐Ducret et al., 1992), it may promote the inflammation by recruiting monocytes and lymphocytes to the affected joint. The inducible nitric oxide synthase (iNOS) enzyme is upregulated in many cell types in response to bacterial products or pro‐inflammatory cytokines, leading to the sustained production of nitric oxide (NO). NO is a diffusible molecule that has beneficial antimicrobial effects in the defense against pathogens, and acts as a potent messenger in regulating tissue perfusion, epithelial permeability, and the immune response. On the other hand, overproduction of NO can be detrimental as it is associated with tissue damage and chronic inflammation (Kolios et al., 2004; MacMicking et al., 1997). The induction of iNOS occurs both at the transcriptional level, and at the posttranscriptional level through stabilization of the mRNA (Carpenter et al., 2001; Di Macro et al., 2005; Lahti et al., 2003; Perez‐Sala et al., 2001; Rodriguez‐Pascual et al., 2000). Another important effector‐enzyme of the inflammatory response, COX‐2, is also regulated at the posttranscriptional level. In a variety of cell types, IL‐1 induces the expression of COX‐2 by a mechanism that involves stabilization of the mRNA (Ridley et al., 1998; Ristimaki et al., 1994), and the same is observed in LPS‐treated monocytes (Dean et al., 1999). Taken together, these examples demonstrate that mRNA stabilization is a general mechanism by which different cell types achieve proper expression of a large group of pro‐inflammatory proteins. In most cases, an ARE in the 30 UTR plays a pivotal role in regulating the stability of the mRNA. For this reason, a major effort in the field is to identify the molecular mechanisms by which ARE‐mRNAs are stabilized. Stabilization of ARE‐mRNAs is dependent upon the activation of different signal transduction pathways. In T‐cells and mast cells, activation of the c‐jun N‐terminal kinase (JNK) pathway is required for the stabilization of IL‐2 and IL‐3 mRNA (Chen et al., 1998; Ming et al., 1998). In NIH3T3 cells, the phosphatidylinositol 3‐kinase (PI3K) pathway contributes to ARE‐mRNA stabilization (Ming et al., 2001). Induction of iNOS was linked to protein kinase Cd activation and may also require the JNK pathway (Carpenter et al., 2001;

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Lahti et al., 2003). In most cases, however, the mitogen‐activated protein kinase p38 (p38‐MAPK) pathway plays a central role in stabilizing ARE‐ mRNAs. As shown in a variety of cell types, p38‐MAPK and the downstream MAPK‐activated protein kinase 2 (MK2) are required for the posttranscriptional induction of IL‐1b, IL‐3, IL‐6, IL‐11, TNFa, IFNg, GROa, GROb, IL‐8, IP‐10, and COX‐2 (Bamba et al., 2003; Brook et al., 2000; Dean et al., 1999; Holtmann et al., 1999; Kotlyarov et al., 1999; Lasa et al., 2000; Mavropoulos et al., 2005; Ming et al., 2001; Neininger et al., 2002; Ridley et al., 1998; Rousseau et al., 2002; Sirenko et al., 1997; Vockerodt et al., 2005; Winzen et al., 1999). Given that such a large number of pro‐inflammatory proteins are induced posttranscriptionally through the p38‐MAPK–MK2 axis, developing inhibitors of this pathway has been a major focus in the search for novel anti‐inflammatory drugs. Recent findings provide first insights into the molecular mechanism that links the p38‐MAPK–MK2 pathway to regulators of AMD. Again, the TTP/ BRF proteins appear to play a central role in this process. Both p38‐MAPK and MK2 phosphorylate TTP (Carballo et al., 2001; Mahtani et al., 2001; Zhu et al., 2001), and the direct phosphorylation of TTP by MK2 at serine 52 and serine 178 leads to complex formation between TTP and the adaptor protein 14‐3‐3 (Chrestensen et al., 2003; Johnson et al., 2002; Stoecklin et al., 2004). As a consequence of 14‐3‐3 binding, TTP activity is reduced and ARE‐mRNAs thereby stabilized (Stoecklin et al., 2004). Interestingly, the activity of BRF1 is also inhibited by phosphorylation and consecutive binding of 14‐3‐3 (Schmidlin et al., 2004). Although one study questions this model (Rigby et al., 2005), phosphorylation‐induced complex formation of the TTP/BRF proteins with 14‐3‐3 may be a general mechanism by which ARE‐mRNAs are stabilized. Another target of the p38‐MAPK pathway is the ARE‐BP hnRNP‐A0 (Rousseau et al., 2002). In LPS‐stimulated macrophages, MK2 phosphorylates hnRNP‐A0 at serine 84, but it is not clear whether this contributes to stabilization of ARE‐mRNAs. The anti‐inflammatory cytokine IL‐10 is a potent inhibitor of macrophage activation that reduces the expression of many cytokines (e.g., TNFa, IL‐1, IL‐6, GM‐CSF), several chemokines, and COX‐2 (Moore et al., 2001). Although most of the inhibitory effect of IL‐10 on the expression of cytokines appears to occur at the transcriptional level in a STAT3‐dependent manner (Takeda et al., 1999; Williams et al., 2004a), there is evidence that posttranscriptional mechanisms are also involved. In LPS‐treated macrophages, IL‐10 was found to reduce TNFa, IL‐1a, and IL‐1b mRNA levels without affecting their rates of transcription, indicating that IL‐10 accelerates the decay of these mRNAs (Bogdan et al., 1992). The suppressive effect of IL‐10 on TNFa expression requires the TNFa 30 UTR (Denys et al., 2002), and another study

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indicates that IL‐10 reduces the translation of TNFa through inhibition of the p38‐MAPK–MK2 pathway (Kontoyiannis et al., 2001). Moreover, IL‐10 was found to destabilize GROa (murine KC), GM‐CSF, and G‐CSF, as well as its own mRNA (Biswas et al., 2003; Brown et al., 1996b; Kishore et al., 1999). Given that some of these results are contradictory, the contribution of posttranscriptional mechanisms to the suppressive effect of IL‐10 remains a controversial issue (Williams et al., 2004b). 2.5. Non‐ARE Decay Elements The ARE is probably the most common element that regulates mRNA stability, yet other regulatory elements have been described in a number of mRNAs encoding pro‐inflammatory proteins. For example, TNFa contains a second, constitutive decay element (CDE) in the 30 UTR, located downstream of the ARE. As opposed to AMD, CDE‐mediated mRNA decay is not inhibited by stimulation of macrophages with LPS, nor by activation of the PI3K or p38‐MAPK pathways (Stoecklin et al., 2003). The CDE may thus serve as a safeguard element that maintains stringent control over TNFa production, thereby reducing the risk of accumulating potentially injurious TNFa levels. G‐CSF is another cytokine that contains in its 30 UTR two independent destabilizing elements: an ARE and the stem‐loop destabilizing element (SLDE). The SLDE, similar to the CDE of TNFa, prevents stabilization of G‐CSF mRNA under conditions where AMD is inhibited (Brown et al., 1996a; Putland et al., 2002). The induction of IL‐2 expression in stimulated T‐cells results from both transcriptional activation and posttranscriptional mRNA stabilization (Lindsten et al., 1989; Musgrave et al., 2004). In unstimulated cells, the ARE in the 30 UTR mediates rapid decay of IL‐2 mRNA. Stabilization of IL‐2 mRNA occurs in response to JNK activation and requires, besides the ARE, a JNK‐response element (JRE) in the 50 UTR of the transcript (Chen et al., 1998). Two proteins bind to the JRE, nucleolin and the cold shock domain (CSD) protein YB1, and both are required for stabilizing IL‐2 mRNA (Chen et al., 2000). Hypoxia‐induced expression of VEGF, a major angiogenic factor, is regulated at both transcriptional and posttranscriptional levels (Ikeda et al., 1995; Levy et al., 1996; Stein et al., 1995). Under normoxic conditions, VEGF mRNA is labile. Three distinct destabilizing elements mediate rapid decay of the VEGF transcript: an ARE in the 30 UTR, a coding region determinant of instability (CDR), and an element in the 50 UTR (Dibbens et al., 1999). Interestingly, the three elements can independently mediate rapid decay of a reporter transcript, yet all three elements together are required to allow the mRNA to be stabilized in response to hypoxia (Dibbens et al., 1999). The

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ARE‐BP HuR binds to the 30 UTR of VEGF mRNA and contributes to mRNA stabilization (Levy et al., 1998). A complex that contains YB1 and polypyrimidine tract binding protein (PTB) binds to two sites in the 50 UTR and one site in the 30 UTR, and participates in the stabilization of VEGF transcripts (Coles et al., 2004). These binding sites are very similar to the binding site of YB1 in the 50 UTR of IL‐2, indicating that YB1‐containing complexes may play a general role in stabilizing different mRNAs (Coles et al., 2004). YB1 was also found to enhance the stability of GM‐CSF mRNA, although in this case, YB1 appears to directly interact with the ARE of GM‐CSF (Capowski et al., 2001). The examples of TNFa, IL‐2, and VEGF illustrate that the ARE does not operate as an isolated element, but cooperates with other elements to achieve complex control over mRNA stability and translation. By recruitment of ARE‐ BPs together with other RNA‐binding proteins, multiple transcripts can be regulated specifically and expressed differentially according to cellular requirements and extracellular cues. We are still far away from understanding the complex interplay between the factors that contribute to the stability and translation efficiency of individual mRNAs. Future studies may reveal that combining non‐ARE regulatory elements with an ARE is a common mechanism of mRNAs that is regulated at the posttranscriptional level. 3. Translational Inhibition 3.1. Translational Silencing of ARE‐Transcripts Several RNA‐binding proteins repress the translation of mRNAs encoding proteins that regulate the inflammatory response. TIA‐1 and TIAR are closely related members of the RNA‐recognition motif (RRM) family of RNA‐binding proteins that inhibit the translation of TNFa transcripts in macrophages (Anderson and Kedersha, 2002a,b; Piecyk et al., 2000), but not in T lymphocytes (Saito et al., 2001). Although LPS‐activated macrophages derived from wild‐type and TIA‐1
/
mice express similar amounts of TNFa transcripts, macrophages lacking TIA‐1 produce significantly more TNFa protein than wild‐type controls (Piecyk et al., 2000; Saito et al., 2001). In macrophages lacking TIA‐1, the percentage of TNFa transcripts found in polysomes is increased, suggesting that TIA‐1 functions as a translational silencer (Piecyk et al., 2000). The overexpression of TNFa protein in macrophages lacking TIA‐1 is strain dependent. TIA‐1
/
macrophages derived from BALB/c mice produce 3–5 times more TNFa than wild‐type controls, whereas TIA‐1
/
macrophages derived from C57BL/6 mice produce nearly 10 times more TNFa than wild‐type controls (Saito et al., 2001). Moreover, C57BL/6 mice

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spontaneously develop mild arthritis (Phillips et al., 2004). Thus, unidentified genetic modifiers determine whether TIA‐1
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mice develop arthritis. TIA‐1 similarly represses the translation of COX‐2 (Dixon et al., 2003), an enzyme that converts arachidonic acid into pro‐inflammatory prostaglandins (Sundy, 2001; Schnitzer and Hochberg, 2002). Pharmacologic inhibitors of COX‐2 are potent anti‐inflammatory agents that significantly reduce the severity of inflammatory arthritis (Schnitzer and Hochberg, 2002). The 30 UTR of COX‐2 transcripts contains an ARE that recruits a multimeric protein complex that includes TIA‐1, TIAR, hnRNP U, and HuR (Cok et al., 2003). TIA‐1 null fibroblasts express 2–3 times more COX‐2 and 2 times more prostaglandin E2 than wild‐type controls (Dixon et al., 2003). Moreover, the expression of COX‐ 2 in colon cancer‐derived cell lines is inversely correlated with the expression of TIA‐1 (Dixon et al., 2003). In colon cancer cell lines expressing abundant TIA‐1, sucrose gradient analysis shows that COX‐2 mRNA is concentrated in low‐density fractions that contain untranslated mRNPs (Dixon et al., 2003). These results suggest that TIA‐1 can coordinately repress the expression of TNFa and COX‐2 to dampen the inflammatory response. TIAR represses the translation of MMP‐13 (Yu et al., 2003), a TNFa‐ induced collagenase that has been implicated in pro‐inflammatory, angiogenic, and destructive processes within the joints of patients with rheumatoid arthritis (Konttinen et al., 1999; Moore et al., 2000; Vincenti and Brinckerhoff, 2002; Wernicke et al., 2002). Although the mechanism of TIAR‐induced translational silencing has not been investigated, the fact that TIAR
/
macrophages, like TIA‐1
/
macrophages, overexpress TNFa (Piecyk et al., 2000) suggests that TIA‐1 and TIAR use similar mechanisms to inhibit protein translation. Interestingly, a 17 amino acid peptide derived from an alternatively spliced TIAR exon enhances the expression of MMP‐13 (Yu et al., 2003), suggesting that this exon plays an important role in TIAR function. The ability of TIA‐1 and TIAR to coordinately regulate the expression of several pro‐inflammatory proteins supports the concept that posttranscriptional mechanisms play a major role in regulating the inflammatory response. CUGBP2, an RNA‐binding protein that is structurally related to HuR, has also been implicated in the translational silencing of ARE‐containing transcripts (Mukhopadhyay et al., 2003). CUGBP2 binds to two distinct AREs found in the 30 UTR of COX‐2 mRNA. Paradoxically, expression of recombinant CUGBP2 stabilizes luciferase reporter transcripts bearing the COX‐2 ARE, but reduces expression of the luciferase protein. Differential centrifugation analysis revealed that luciferase transcripts are excluded from polysomes in cells transfected with CUGBP2, but not CUGBP1 or HuR. Thus CUGBP2, like TIA‐1 and TIAR, appears to function by preventing its associated transcripts from

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moving to polysomes. It is not known whether CUGBP2, TIA‐1, and TIAR cooperate to repress the translation of ARE‐containing transcripts. The Fragile X‐related protein FXR1P has also been identified as a translational silencer that targets ARE‐containing transcripts (Garnon et al., 2005). The FXR family of RNA‐binding proteins (FMRP, FXR1P, and FXR2P) possesses two hnRNP K‐homology (KH) domains and an RGG box (Kaufmann et al., 2002). These proteins bind to both U‐rich and G‐rich sequences, and the RGG box allows FMRP to bind to the G‐quartet RNA motif (Darnell et al., 2001, 2004). Thus, the RNA specificity of the FXR proteins could be quite broad. FXR1P binds specifically to the ARE in the 30 UTR of TNFa mRNA, and FXR1P
/
macrophages express approximately 2 times more TNFa than wild‐type macrophages (Garnon et al., 2005). Polysome profiles suggest that FXR1P, like TIA‐1 and TIAR, can exclude TNFa mRNA from polysomes. These results suggest that FXR1P, TIA‐1, and TIAR may work together to bring about translational silencing of TNFa transcripts. It remains to be determined whether FXR1P also represses the translation of additional pro‐inflammatory proteins. 3.2. Mechanisms of Translational Silencing Much of what we know about the mechanism of TIA‐1‐induced translational repression comes from studies of the general translational arrest triggered by environmental stress (e.g., heat, oxidative conditions, and energy deprivation) (Anderson and Kedersha, 2002a,b; Kedersha and Anderson, 2001). Stress‐ induced translational arrest is characterized by the activation of one or more members of a family of serine/threonine kinases (e.g., double‐stranded RNA‐ dependent protein kinase R, PKR‐like endoplasmic reticulum kinase, GCN2, and heme‐regulated inhibitor kinase) (Harding et al., 2000a,b; Sood et al., 2000; Williams, 1999). These kinases phosphorylate eIF2a, a component of the ternary complex that loads tRNAiMet onto the small ribosomal subunit to initiate protein synthesis (Dever, 2002). Phosphorylation of eIF2a inhibits protein translation by reducing the availability of active ternary complex (Krishnamoorthy et al., 2001). Under these conditions, TIA‐1 promotes the assembly of a non‐canonical translation‐initiation complex that is directed to discrete cytoplasmic foci known as stress granules (Anderson and Kedersha, 2002b; Kedersha et al., 1999, 2000, 2002). Like TIA‐1, the RNA‐binding proteins TIAR, FMRP, and FXR1P are concentrated at SGs, suggesting that these proteins might work together to bring about stress‐induced translational silencing. A non‐canonical translation initiation complex assembled during stress has reduced amounts of eIF2 and eIF5 (Kedersha et al., 2002). It is possible that the assembly of these non‐canonical translation initiation complexes contribute to

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the reduced translation of ARE‐containing transcripts mediated by TIA‐1, TIAR, and FXR1P. These complexes may be unable to recruit the large ribosomal subunit, preventing the assembly of polysomes. Although stress granules are visible only when large numbers of transcripts are simultaneously subjected to translational arrest, the underlying translational control mechanism (i.e., assembly of stalled initiation complexes) regulates protein expression in both stressed and unstressed cells (Anderson and Kedersha, 2002a,b). 3.3. Translational Silencing of Non‐ARE Transcripts The translational silencing of ARE‐containing transcripts has the potential to coordinately regulate the expression of multiple pro‐inflammatory proteins. Translational silencing also regulates the expression of non‐ARE‐containing transcripts encoding proteins that control the inflammatory response. A wellcharacterized example is the translational regulation of 15‐LO, an enzyme that participates in the conversion of arachidonic acid into either leukotriene A4 (LTA4) or lipoxin A4 (LXA4) (Kuhn et al., 2002). Whereas LTA4 is converted into LTB4, a potent pro‐inflammatory lipid, LXA4 is an anti‐inflammatory lipid that promotes the resolution of inflammation (Serhan, 2001). Because of this functional dichotomy, 15‐LO has both pro‐ and anti‐inflammatory effects in a variety of experimental systems (Kuhn et al., 2002). During erythrocyte maturation, the mRNA encoding 15‐LO is expressed in the early stages of erythropoiesis, but 15‐LO protein is only expressed at a late stage of maturation (van Leyen et al., 1998). The developmental stage‐specific expression of 15‐LO is achieved by translational silencing in early erythroid progenitors. Translational silencing of 15‐LO mRNA is dependent upon a differentiation control element (DICE) in the 30 UTR (Ostareck et al., 2001). mRNP K/E1 binds hnRNP K and hnRNP E1, which prevent the recruitment of 60S ribosomal subunits to the translation initiation complex (Ostareck et al., 2001). Thus, translational silencing by TIA‐1, TIAR, and hnRNP K/E1 may all involve the exclusion of the 60S ribosomal subunit from the initiation complex. The effect of translational regulation of 15‐LO on the production of pro‐ and anti‐inflammatory lipids remains to be determined. 4. Animal Models 4.1. TNFDARE Knock‐In Mice The importance of posttranscriptional pathways in the regulation of inflammation has been dramatically demonstrated in mutant mice lacking these regulatory controls. In general, posttranscriptional control mechanisms dampen

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the expression of pro‐inflammatory proteins, and thereby prevent the pathological overexpression of these potentially dangerous products. This is certainly true in the case of TNFa, in which transgene‐derived overexpression is sufficient to induce inflammatory arthritis (Keffer et al., 1991). The contribution of posttranscriptional mechanisms to the normal regulation of TNFa production was elegantly revealed by the analysis of knock‐in mice that lack the ARE in the 3’UTR of one of the TNFa alleles (TNFDARE). Macrophages and synovial fibroblasts from TNFDARE mice overexpress TNFa mRNA and protein, and these animals spontaneously develop inflammatory arthritis and inflammatory bowel disease (Kontoyiannis et al., 1999). Because the ARE has not been implicated in the transcriptional regulation of TNFa production, these results imply that posttranscriptional controls are essential for the prevention of spontaneous inflammatory disease. Importantly, the production of TNFa from TNFDARE macrophages is not affected by inhibitors of p38‐MAPK, confirming that the ARE is required for the ability of p38‐MAPK to stabilize and promote the translation of TNFa transcripts (Kontoyiannis et al., 1999). The development of inflammatory bowel disease in TNFDARE mice requires the function of CD8þ, but not CD4þ T lymphocytes (Kontoyiannis et al., 2002). The full spectrum of intestinal inflammation requires the Th1 cytokines IL‐12 and IFN‐g, but not the Th2 cytokine IL‐4 (Kontoyiannis et al., 2002). Restricted expression of TNFa by either macrophages or T lymphocytes is sufficient to induce intestinal inflammation (Kontoyiannis et al., 2002). Moreover, studies of bone marrow chimeras revealed that the presence of the TNFDARE mutation in either bone marrow‐derived or tissue stroma‐derived cells is sufficient to induce intestinal inflammation. It is therefore likely that redundant cellular pathways act downstream of TNFa to bring about intestinal inflammation. 4.2. TTP Knock‐Out Mice Studies of TNFDARE macrophages reveal that the ARE is a destabilizing element that promotes the degradation of TNFa transcripts. The decay of TNFa mRNA is dependent upon TTP, a zinc‐finger protein that targets ARE‐ containing transcripts to the mRNA degradation machinery (see Fig. 1). Mutant mice lacking TTP develop a syndrome of cachexia, arthritis, dermatitis, and autoimmunity that results from the pathological overexpression of TNFa (Taylor et al., 1996). The administration of neutralizing antibodies reactive with TNFa prevents all aspects of this syndrome, indicating that TNFa plays a central role in disease pathogenesis. The importance of TNFa was confirmed by breeding TTP‐deficient mice with mice lacking TNF‐Receptor (R)1 or TNFR2. Whereas TTP
/
TNFR1
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mice did not develop cachexia or

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arthritis, TTP
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TNFR2
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developed more severe arthritis than mice lacking TTP alone (Carballo and Blackshear, 2001). Thus, TNFa binds to TNFR1 to bring about the pathological features of this syndrome. In contrast, interactions with TNFR2 may prevent pathology. TTP
/
mice also develop bone marrow and peripheral blood granulopoiesis, suggesting that TTP may regulate the production of granulocyte growth and/or survival factors. Interestingly, this phenotype was preserved in TTP
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mice lacking TNFR1, TNFR2, or both TNF receptors, suggesting that granulopoiesis is not mediated by overexpressed TNFa. The mechanism of granulopoiesis remains to be determined. Unlike TNFDARE mice, TTP
/
mice do not develop inflammatory bowel disease (Taylor et al., 1996). It is not surprising that the phenotypes of TNFDARE and TTP
/
mice are different. TNFDARE macrophages lack the ability to bind to several ARE‐binding proteins, including the TTP homologues BRF1 and BRF2, or translational silencers such as TIA‐1 and FXR1P. Thus, the phenotype of TNFDARE mice is likely to be more severe than that of TTP
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mice. Because TNFa transgenic mice develop arthritis, but not inflammatory bowel disease (Keffer et al., 1991; Kollias, 2004), the level of TNFa expression may not be sufficient to explain why bowel inflammation is observed in the TNFDARE strain. It is possible that tissue‐specific regulation of TNFa expression determines whether there is inflammation in the joints, the intestines, or both. 4.3. TIA‐1 and TIAR Knock‐Out Mice On the Balb/c background, mice lacking TIA‐1 do not develop spontaneous inflammatory disease. When bred onto the C57Bl/6 background, however, TIA‐1
/
mice develop mild non‐erosive arthritis (Phillips et al., 2004). Thus, genetic modifiers are important in determining the disease phenotype. Mutant mice lacking both TTP and TIA‐1 develop spontaneous inflammatory arthritis that is significantly more severe than the arthritis observed in mice lacking either TIA‐1 or TTP alone (Phillips et al., 2004). Although macrophages derived from TIA‐1
/
TTP
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mice overexpress TNFa mRNA, they express less TNFa protein than TIA‐1
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or TTP
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macrophages (Phillips et al., 2004). Whether this results from defective nuclear export of TNFa transcripts, defective translation, or defective processing of TNFa protein remains to be determined. The source of arthritigenic cytokine in mice lacking both TIA‐1 and TTP appears to be an expanded population of neutrophils found in the bone marrow and peripheral blood. The marked increase in neutrophils in TTP
/
bone marrow and peripheral blood is potentiated in mice that also lack TIA‐1. The mechanism whereby TIA‐1 and TTP cooperate to increase the maturation and/or survival of neutrophils is not known. Whereas wild‐type neutrophils produce little or no

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TNFa in response to LPS stimulation, neutrophils lacking TTP secrete significant amounts of TNFa in response to LPS (Phillips et al., 2004). It is therefore possible that neutrophils are an important source of this pro‐arthritic cytokine in mice lacking both TIA‐1 and TTP. Although neutrophils clearly contribute to the pathogenesis of arthritis (Edwards and Hallett, 1997; Pillinger and Abramson, 1995), our understanding of their precise contribution to the inflammatory process is incomplete. Neutrophils are the predominant cell type found in inflammatory synovial fluid (but not synovial pannus) derived from patients with rheumatoid arthritis. They are an important source of arachadonic acid‐derived inflammatory mediators (e.g., prostaglandins, leukotrienes, and lipoxins). In animal models of inflammatory arthritis, neutrophils are essential components of the inflammatory process (Jonsson et al., 2005; Lawlor et al., 2004; Wipke and Allen, 2001). The cooperative regulation of both neutrophil maturation and function by TIA‐1 and TTP suggests that these cells are a major source of inflammatory cytokine production in a subset of patients with rheumatoid arthritis. 4.4. G‐CSF and GM‐CSF Transgenic Mice The importance of neutrophils in inflammatory arthritis was dramatically demonstrated in mice lacking G‐CSF, an important regulator of neutrophil production. These mice exhibit impaired granulopoiesis and are protected from collagen‐induced arthritis (Lawlor et al., 2004). Moreover, administration of anti‐G‐CSF antibodies protects wild‐type mice from collagen‐induced arthritis (Lawlor et al., 2004). These results reveal that G‐CSF is required to produce the mature neutrophils which are essential effectors of synovitis. The 30 UTR of the G‐CSF mRNA encodes two regulatory elements that dampen the production of G‐CSF at the posttranscriptional level. Thus, G‐CSF may be one of the key cytokines whose posttranscriptional regulation can determine susceptibility to arthritis. Like G‐CSF, GM‐CSF contributes to the growth, differentiation, and function of granulocytes and macrophages. In both in vitro and in vivo studies, GM‐CSF has been shown to ‘‘prime’’ neutrophils and macrophages for their effector functions (Hamilton, 2002). In this capacity, GM‐CSF may play an important role in the inflammatory response. Consistent with this prediction, GM‐CSF
/
mice are resistant to methylated BSA/IL‐1‐induced arthritis (Yang and Hamilton, 2001). Moreover, anti‐GM‐CSF antibodies reduce the severity of both collagen‐ and BSA/IL‐1‐induced arthritis (Cook et al., 2001; Yang and Hamilton, 2001). These results place GM‐CSF in the category of a pro‐inflammatory protein whose expression is regulated at the posttranscriptional level. The importance of posttranscriptional regulation of GM‐CSF

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production was demonstrated in transgenic mice expressing recombinant GM‐CSF with or without the 30 ARE under control of a CMV promoter. At embryonic day 14, the ARE‐deleted transcript, but not the ARE‐containing transcript, was expressed in all tissues in which the CMV promoter was active (Houzet et al., 2001). Mice with the ARE‐deleted construct exhibit increased proliferation of granulocytes and macrophages. Taken together, these results implicate GM‐CSF as an important pro‐inflammatory cytokine subject to posttranscriptional control. 5. Clinical Significance The coordinate regulation of pro‐inflammatory proteins by posttranscriptional control pathways suggests that components of these pathways may be attractive targets for drug development. Indeed, chemical inhibitors of p38‐MAPK potently reduce the expression of pro‐inflammatory proteins in activated macrophages, and strongly support a role for posttranscriptional control in the regulation of inflammation (Pargellis and Regan, 2003). p38‐MAPK inhibitors reduce the expression of pro‐inflammatory cytokines and chemokines such as TNFa, IL‐1b, IL‐6, IL‐8, and MCP‐1. These drugs also reduce the expression of pro‐inflammatory enzymes such as COX‐2, iNOS, MMP‐1, MMP‐3, MMP‐9, and MMP‐13, as well as endothelial adhesion molecules such as E‐selectin, intracellular adhesion molecule (ICAM)‐1, and VCAM‐1 that recruit inflammatory cells to the rheumatoid synovium (Westra et al., 2004a,b, 2005). These drugs also inhibit the development of inflammatory arthritis in different animal models of arthritis (Pargellis and Regan, 2003). Several pharmaceutical companies are testing p38‐MAPK inhibitors in clinical trials involving patients with rheumatoid arthritis. In phase I clinical trials, single dose administration of these agents has been well tolerated (Parasrampuria et al., 2003). If the safety profile of these drugs is acceptable, they could become important treatments for a variety of immune‐mediated inflammatory diseases. MK2, a specific substrate for p38‐MAPK, has also been implicated in the posttranscriptional regulation of pro‐inflammatory protein expression. Macrophages derived from mutant mice lacking MK2 overexpress TNFa, IFNg, IL‐1, IL‐6, and nitric oxide following LPS activation (Kotlyarov et al., 1999). Thus, MK2 inhibitors, similar to p38‐MAPK inhibitors, have the potential to coordinately reduce the expression of several classes of proteins with pro‐inflammatory activity. The effects of MK2 on mRNA stability and translation are complex and poorly understood. In macrophages lacking MK2, the half‐life of IL‐6 mRNA is reduced more than 10 fold, whereas the half‐ life of TNFa mRNA is essentially unchanged (Neininger et al., 2002).

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This could be due to the presence of a second decay element, the CDE, in the TNFa 30 UTR (Stoecklin et al., 2003). It remains to be determined whether differences in the spectrum of pro‐inflammatory proteins regulated by p38‐MAPK and MK2 will affect the clinical efficacy of these drugs. Comparative pre‐clinical and clinical evaluation of p38‐MAPK and MK2 inhibitors will be required to determine the potential for these drugs in the treatment of inflammation. A number of other compounds have also been shown to interfere with cytokine production at the posttranscriptional level. In macrophages, the radicicol analogue A inhibits IL‐1b, IL‐6, and TNFa production by accelerating mRNA decay in an ARE‐dependent manner (Kastelic et al., 1996). In monocytes, the cannabinoid ajulemic acid reduces LPS‐induced IL‐1b production, which may involve accelerated IL‐1b mRNA decay (Bidinger et al., 2003). In osteoblasts, a tetracycline derivative was shown to inhibit IL‐1b‐ induced IL‐6 production by accelerating decay of IL‐6 mRNA (Kirkwood et al., 2003). Thalidomide, once in use as a sedative but withdrawn from the market after its severe teratogenic effects were discovered, reduces TNFa and COX‐2 expression by accelerating the degradation of the corresponding mRNAs. Due to its anti‐inflammatory activity, thalidomide has been reintroduced for the treatment of erythema nodosum leprosum, aphtous stomatitis, Behcet syndrome, chronic cutaneous systemic lupus erythematosus, and graft‐versus‐host disease (Calabrese and Fleischer, 2000). Studies have also shown efficacy against aphthous ulcers and other HIV‐associated conditions including Kaposi sarcoma, as well as in the treatment of multiple myeloma. By interfering with posttranscriptional control mechanisms, these drugs have the capacity to dampen the expression of several pro‐inflammatory proteins simultaneously, and may therefore prove useful for the treatment of a variety of inflammatory conditions. The importance of posttranscriptional control mechanisms in the regulation of inflammation suggests that individual components of these regulatory pathways may influence disease susceptibility. Patients expressing single nucleotide polymorphisms that reduce the expression and/or function of TTP, TIA‐1, TIAR, and FXR1P could have an increased risk of developing inflammatory arthritis or inflammatory bowel disease. Sequence analysis has identified 13 polymorphisms in the coding regions of TTP and its two homologues BRF1 and BRF2 (Blackshear et al., 2003b). Six of these mutations would result in amino acid changes that could alter protein function. Another mutation was a dinucleotide substitution that would prevent splicing of the single intron in ZFP36L1, the gene encoding for BRF1. Analysis of lymphoblasts from this individual confirmed that the expression of ZFP36L1 mRNA was reduced by 50%. Case control association studies will be required to determine whether

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any of these gene polymorphisms are associated with increased susceptibility to immune‐mediated inflammatory disease. 6. Conclusions Posttranscriptional regulation of mRNA stability and translation are major determinants of pro‐inflammatory protein expression. An ARE in the 30 UTR of a large number of mRNAs encoding pro‐inflammatory proteins plays a central role by recruiting a specific set of ARE‐BPs, which exert their functions by (i) repressing translation of the bound mRNA, (ii) targeting the mRNA for rapid degradation, or (iii) stabilizing the mRNA in response to extracellular cues. In addition, non‐ARE elements participate in regulating stability and translation of certain mRNAs, and provide a further level of complexity. A molecular understanding of the underlying mechanisms will help to characterize new targets for the development of a next generation of anti‐inflammatory drugs. Genetic polymorphisms within the genes encoding posttranscriptional regulatory proteins are likely to identify individuals with enhanced susceptibility to immune‐mediated inflammatory diseases. As our understanding of posttranscriptional regulatory pathways is still rudimentary, this growing area of research will continue to provide important insights into the fundamental principles that determine gene expression. Much remains to be learned. It is not clear how posttranscriptional control pathways discriminate between the many transcripts that contain regulatory elements in their 30 UTRs. Gene array analysis of mRNAs that co‐precipitate with TIA‐1 or TIAR has revealed that these RNA‐binding proteins can associate with thousands of transcripts (Lopez de Silanes et al., 2005; M. Gorospe, personal communication). Yet functional studies suggest that the mRNAs regulated by these proteins may be much more restricted. It is likely that multiple RNA‐binding proteins expressed in individual cells work together to select mRNAs that are subject to post‐ transcriptional regulation. We know very little about how different ARE‐PBs cooperatively regulate mRNA stability and translation. The potential participation of microRNAs provides further opportunities for fine‐tuning the mRNA selection process. Defining the complex interplay of all factors associated with ARE and non‐ARE elements will help us to fully understand the posttranscriptional operon that governs the expression of pro‐inflammatory proteins. Acknowledgments We would like to thank Nancy Kedersha (Brigham and Women’s Hospital, Boston) for helpful comments on the manuscript. PA was supported by National Institute of Health grants AI‐33600 and AI‐50167.

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Negative Signaling in Fc Receptor Complexes Marc Dae¨ron and Renaud Lesourne Unite´ d’Allergologie Mole´culaire et Cellulaire, De´partement d’Immunologie, Institut Pasteur, Paris, France

1. 2. 3. 4. 5.

Abstract............................................................................................................. Fc Receptors ...................................................................................................... Positive Signaling by Activating FcRs ...................................................................... Negative Signaling by Activating FcRs..................................................................... Negative Signaling by Inhibitory FcRs..................................................................... Conclusion ......................................................................................................... References .........................................................................................................

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Abstract Cell activation results from the transient displacement of an active balance between positive and negative signaling. This displacement depends in part on the engagement of cell surface receptors by extracellular ligands. Among these are receptors for the Fc portion of immunoglobulins (FcRs). FcRs are widely expressed by cells of hematopoietic origin. When binding antibodies, FcRs provide these cells with immunoreceptors capable of triggering numerous biological responses in response to a specific antigen. FcR‐dependent cell activation is regulated by negative signals which are generated together with positive signals within signalosomes that form upon FcR engagement. Many molecules involved in positive signaling, including the FcRb subunit, the src kinase lyn, the cytosolic adapter Grb2, and the transmembrane adapters LAT and NTAL, are indeed also involved in negative signaling. A major player in negative regulation of FcR signaling is the inositol 5‐phosphatase SHIP1. Several layers of negative regulation operate sequentially as FcRs are engaged by extracellular ligands with an increasing valency. A background protein tyrosine phosphatase‐dependent negative regulation maintains cells in a «resting» state. SHIP1‐dependent negative regulation can be detected as soon as high‐affinity FcRs are occupied by antibodies in the absence of antigen. It increases when activating FcRs are engaged by multivalent ligands and, further, when FcR aggregation increases, accounting for the bell‐shaped dose‐ response curve observed in excess of ligand. Finally, F‐actin skeleton‐associated high‐molecular weight SHIP1, recruited to phosphorylated ITIMs, concentrates in signaling complexes when activating FcRs are coengaged with inhibitory FcRs by immune complexes. Based on these data, activating and inhibitory FcRs could be used for new therapeutic approaches to immune disorders.

39 advances in immunology, vol. 89 # 2006 Elsevier Inc. All rights reserved.

0065-2776/06 $35.00 DOI: 10.1016/S0065-2776(05)89002-9

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When sensitized with IgE antibodies, mouse mast cells and human basophils release granular mediators and secrete pro‐inflammatory cytokines and chemokines in response to stimulation by specific antigen. These biological responses depend on high‐affinity receptors for the Fc portion of IgE antibodies (FceRI) that are expressed by the two cell types (Ishizaka et al., 1970; Metzger et al., 1986; Prouvost‐Danon and Binaghi, 1970). For a given concentration of IgE used for sensitization, mediator release increases with the concentration of antigen used for challenge up to a maximum. Release then decreases as the concentration of antigen further increases (Dembo et al., 1978). Peritoneal mouse mast cells also degranulate when challenged by preformed IgG immune complexes (Prouvost‐Danon et al., 1966). IgG‐ induced responses depend on low‐affinity receptors for the Fc portion of IgG (FcgRIIIA) (Dae¨ron et al., 1992; Hazenbos et al., 1996) that bind immune complexes with a high avidity. Bone Marrow‐derived Mast Cells (BMMC) do not respond or respond very poorly to IgG immune complexes, although they express FcgRIIIA (Benhamou et al., 1990). Likewise, human blood basophils release no or little histamine in response to immune complexes (Van Toorenenbergen and Aalberse, 1981), although they express another type of low‐affinity IgG receptor (FcgRIIA) which can activate mast cells (Dae¨ron et al., 1995a). These observations have long been interpreted as resulting from an inefficient engagement of activating receptors by high concentrations of antigen or by IgG immune complexes. Actually, these experiments show that negative regulation occurs in Fc Receptor (FcR) complexes. One is an example of autonomous negative regulation of activating FcRs; others are examples of negative regulation by inhibitory FcRs. These examples were selected from studies of FcRs in mast cells and basophils. FcR‐ dependent negative signaling is not peculiar to these cells. Mast cells are, however, convenient models to study FcR signaling, and they will often be used as examples throughout this review. 1. Fc Receptors 1.1. FcRs, the Third Type of Immunoreceptors Receptors for the Fc portion of immunoglobulins are immunoreceptors of the third type. They ‘‘recognize’’ neither native antigens as B Cell Receptors (BCRs) do, nor the association of antigen‐derived peptides with Major Histocompatibility Complex molecules, as T Cell Receptors (TCRs) do, but antigen‐ antibody complexes. Even though they do not themselves bind to antigen, they enable cells to respond specifically to antigen. Antibodies indeed function as extracellular adapter molecules when their Fab and Fc portions bind

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simultaneously to specific epitopes on antigen and to FcRs on cell membrane, respectively. BCRs, TCRs, and FcRs are receptors for the three forms under which any given antigen can interact with and deliver signals to cells of the immune system. BCRs and TCRs are assigned specificity at an early stage during B and T cell differentiation through somatic DNA rearrangements. Combinations of variable gene segments determine the clonally restricted specificity of lymphocytes. Specificity persists over cell divisions, as it is transmitted to the progeny within a given clone. These unique features of lymphocytes have several consequences. Altogether, the lymphocytes of an individual can recognize virtually all antigens this individual can be exposed to. Their number being finite, only a small number of naı¨ve lymphocytes can respond to a given antigen. Lymphocytes therefore need first to undergo clonal expansion for significant numbers of cells expressing antigen receptors with any given specificity to be generated and to mount an adaptive immune response. In addition, B and T lymphocytes are not ready‐to‐work effector cells. They need to differentiate into antibody‐producing plasma cells and into helper, regulatory, or cytoxic T cells, respectively, before they can act on antigen. Unlike lymphocytes, large numbers of differentiated cells of hematopoietic origin are capable of exerting a variety of biological activities without needing to proliferate and/or to differentiate. These mostly myeloid cells are the primary effectors of innate immunity. They are equipped with pattern‐ recognition receptors which enable them to interact with structures borne or secreted by microorganisms, but they lack antigen receptors. Most myeloid cells, however, express FcRs. FcRs provide these cells with immunoreceptors and a bona fide immunological specificity. Antigen specificity is provided by antibodies that happen to be present in the environment and bind to FcRs. As these antibodies, polyclonal in nature, have different specificities, one FcR‐ expressing cell can respond specifically to a wide repertoire of different antigens. This repertoire can, theoretically, be as wide as that of the whole population of B cells. In the presence of specific antibodies, FcRs enroll in adaptive immunity, the many cells involved in innate immunity. Besides endowing them with specificity, FcRs can indeed generate intracellular signals which modulate their biological activities. Some FcRs activate, whereas others inhibit cellular responses. 1.2. Activating FcRs Most FcRs are activating receptors (Dae¨ron, 1997; Ravetch and Bolland, 2001; Ravetch and Kinet, 1991). Activating FcRs comprise receptors for IgA (FcaRI), IgE (FceRI), and IgG (FcgRI, FcgRIIA/C, FcgRIIIA, and FcgRIV).

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They include high‐affinity receptors (FcaRI, FceRI, FcgR, and FcgIV) which can bind monomeric immunoglobulins, and low‐affinity receptors (FcgRIIA/C and FcgRIIIA) which cannot, but which can bind multivalent antigen‐antibody complexes and immunoglobulin aggregates with a high avidity (Hulett and Hogarth, 1994). As a consequence, a proportion of high‐affinity FcRs are occupied in vivo, whereas low‐affinity FcRs remain free in spite of the high concentrations of immunoglobulins present in the extracellular milieu. With one exception in humans (FcgRIIA/C), activating FcRs are multi‐chain receptors composed by one immunoglobulin‐binding FcRa subunit and one (FcRg) or two (FcRg and FcRb) common transduction subunits. As for other immunoreceptors, the cell‐activating properties of FcRs depend on the presence of Immunoreceptor Tyrosine‐based Activation Motif(s) (ITAMs) in the intracytoplasmic domains of their transduction subunits (Reth, 1989). Activating FcRs are expressed by myeloid cells and by lymphoid cells with no classical antigen receptor (i.e., NK cells [Perussia et al., 1989] and intraepithelial g/d T cells of the intestine [Deusch et al., 1991; Sandor et al., 1992; Woodward and Jenkinson, 2001]). They are not expressed by mature T and B lymphocytes. Lymphocytes therefore do not express more than one type of antigen receptor, and activating FcRs do not interfere with lymphocyte activation triggered by clonally expressed antigen receptors. Interestingly, however, activating FcRs are transiently expressed by pre‐B and pre‐T cells, before they express a functional BCR or TCR, respectively (Sandor and Lynch, 1992). Low levels of FcgRIIIA were recently reported to be expressed on a subset of self‐specific murine CD8 T cells and to efficiently trigger antibody‐dependent cell‐ mediated cytotoxicity (Dhanji et al., 2005). Differing from other immunoreceptors, which induce both cell activation and proliferation, FcRs induce cell activation only. Activating FcRs do not induce unique biological responses, but biological activities that can be induced by other receptors in the same cell. 1.3. Inhibitory FcRs Inhibitory FcRs consist of one family of low‐affinity receptors for IgG, referred to as FcgRIIB (Dae¨ron, 1997; Ravetch and Bolland, 2001). FcgRIIB are single‐chain receptors, encoded by one gene named fcgr2b, which generates two (FcgRIIB1 and FcgRIIB2 in humans) or three (FcgRIIB1, FcgRIIB1’, and FcgRIIB2 in mice) isoforms of membrane receptors, by alternative splicing of sequences encoded by the first intracytoplasmic exon (Hibbs et al., 1986; Latour et al., 1996; Lewis et al., 1986; Ravetch et al., 1986). One distinctive feature of the fcgr2b gene is indeed that one exon encodes the transmembrane domain and three others the intracytoplasmic domain of FcgRIIB (in other FcR genes, a single exon encodes both the transmembrane

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and the intracytoplasmic domains) (Brooks et al., 1989; Hibbs et al., 1988). The inhibitory properties of FcgRIIB depend on an Immunoreceptor Tyrosine‐ based Inhibition Motif (ITIM) (Dae¨ron et al., 1995a), encoded by the third intracytoplasmic exon of the fcgr2b gene, and located in the intracytoplasmic domain of all murine and human FcgRIIB isoforms. FcgRIIB are expressed by myeloid and, with two exceptions, by lymphoid cells. The two exceptions are NK cells and resting T cells which express a variety of other inhibitory receptors involved in cell–cell interactions (Long, 1999). FcgRIIB can negatively regulate cell activation triggered by all ITAM‐containing receptors (Dae¨ron et al., 1995a) as well as cell proliferation triggered by growth factor receptors with an intrinsic kinase activity (Malbec et al., 1999). In order to exert their inhibitory properties, FcgRIIB must be co‐engaged with activating receptors by a common extracellular ligand at the surface of the same cell (Dae¨ron et al., 1995b). The specificity of negative regulation is therefore under the control of two antigen‐specific recognition processes: that of IgG antibodies which engage FcgRIIB and that of immunoreceptors with which FcgRIIB are coaggregated. 1.4. Activating and Inhibitory FcRs in Physiology and Pathology The aggregation of identical FcRs only (homo‐aggregation) is a rare situation in physiology. Even when cells express one type of FcR only (e.g., FcgRIIB in murine B cells, or FcgRIIIA in murine NK cells), immune complexes can co‐ engage FcRs with other immunoreceptors (BCRs in B cells, or NK Receptors on NK cells). Several FcRs are coaggregated when IgG immune complexes interact with cells that co‐express several FcgRs (FcgRI, FcgRIIB, and FcgRIIIA on macrophages or dendritic cells, for instance, or FcgRIIA and FcgRIIB on human B cells) or with cells that co‐express FcRs for several classes of antibodies (mouse mast cells, for instance, where IgG immune complexes can coaggregate FcgRs and FceRI‐bound IgE). Hetero‐aggregation, that is, the coaggregation of different types of FcRs or the coaggregation of FcRs with other immunoreceptors, is actually a rule, rather than an exception, under physiological conditions. Because there are FcRs for all antibody classes, because immune complexes contain more than one class of antibody, and because most cells express more than one type of FcRs, various combinations of FcRs can be engaged at the cell surface to form hetero‐aggregates with a non‐predetermined composition. FcRs can thus generate a variety of signaling complexes, depending on the relative proportion of receptors of the various types that are co‐engaged by immune complexes. The in vivo biological significance of FcgRIIB‐dependent negative regulation of activating FcR‐dependent physio‐pathological processes has been

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established using mice rendered deficient for FcgRIIB by homologous recombination (Ravetch and Bolland, 2001). Compared to wt mice, FcgRIIB‐ deficient mice were found to produce more antibodies (Takai et al., 1996), to exhibit exaggerated anaphylactic reactions and Arthus reactions of a higher intensity (Takai et al., 1996; Ujike et al., 1999), to be more susceptible to collagen‐induced arthritis (Kleinau et al., 2000; Yuasa et al., 1999), and, in the C57BL/6 background, to spontaneously develop Lupus‐like syndromes (Bolland and Ravetch, 2000). FcgRIIB were also shown to critically determine the protective effects of anti‐tumor therapeutic antibodies in a murine model of melanoma (Clynes et al., 1998), and of IVIG in a model of idiopathic thrombopenic purpura (Samuelsson et al., 2001). 2. Positive Signaling by Activating FcRs 2.1. Positive Signaling in Resting Cells Positive signals are generated even before immunoreceptors are engaged by extracellular ligands. This can be readily unraveled by treating cells with the tyrosine phosphatase inhibitor pervanadate. Pervanadate‐treated cells display an array of tyrosyl‐phosphorylated molecules, including immunoreceptors, indicating that protein tyrosine kinases are active in resting cells but that their substrates are constantly dephosphorylated by tyrosine phosphatases. It follows that cell activation results from a transient displacement of a physiological balance between positive and negative signals that controls cellular responses. Interestingly, the expression of multi‐subunit immunoreceptors such as BCRs was found to be required (and sufficient?) for intracellular signaling molecules to be phosphorylated in pervanadate‐treated cells (Wossning and Reth, 2004), suggesting that signaling complexes can be organized by immunoreceptors even in the absence of known extracellular ligands, but that positive signals emanating from such complexes are either insufficient to lead to cell activation or are dampened by an autonomous type of negative regulation of immunoreceptor signaling. The displacement of the constitutive balance between positive and negative signals that leads to biological responses primarily depends on extracellular ligands which engage surface receptors. 2.2. FcR Engagement and the Constitution of Signalosomes Activating FcRs trigger signals when aggregated by antibody and multivalent antigen. Dimerization was, long ago, shown to be the minimal degree of FceRI aggregation capable of generating activation signals sufficient for triggering

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mediator release by mast cells (Segal et al., 1977; Siraganian et al., 1975). Intracellular signals are generated within juxta‐membrane signaling complexes that assemble under FcR aggregates and form signalosomes. Signalosomes are transient structures which contain the signaling complexes generated at a given time and at a given location, in which signaling molecules can meet and interact with each other. These comprise receptors that are co‐engaged by common extracellular ligands, molecules that are recruited underneath, and molecules that are contained in subcellular compartments into which receptor aggregates translocate. Signalosomes are dynamic structures which evolve with time and with their intracellular location. Molecules are sequentially recruited first, as complexes build up and get organized around transmembrane adapters. Recruitment depends in part on inducible molecular changes, such as phosphorylation, on the generation of specific molecules, and on location or relocation of molecules into subcellular compartments. It is stabilized by cooperative interactions between molecules with several binding sites and by cytosolic adapters. The composition of signalosomes then rapidly changes as recruited enzymes meet substrates and act on them. Finally signalosomes are dismantled as signaling molecules are ubiquitinated and degraded by the proteasome. 2.3. Generation of Positive Signals by Activating FcRs An initial event in signal transduction by activating FcRs is the activation of src‐family protein tyrosine kinases. In resting cells, these kinases are maintained in an inactive state as a result of the phosphorylation of a regulatory C‐terminal tyrosine by the C‐terminal tyrosine Src kinase Csk (Okada et al., 1991). This confers the molecule a closed conformation that prevents substrates to have access to the catalytic site of the kinase (Cole et al., 2003). The regulatory tyrosine is dephosphorylated by the transmembrane protein tyrosine phosphatase CD45 (Burns et al., 1994; Thomas and Brown, 1999). Supporting a role of CD45 in FceRI signaling, CD45‐deficient mast cells displayed reduced IgE‐induced mediator release and CD45‐deficient mice were refractory to IgE‐induced systemic anaphylaxis (Berger et al., 1994). How CD45 becomes involved upon FcR receptor engagement is unclear. Whatever the mechanism, src kinases are activated and they can phosphorylate tyrosine residues in the ITAMs of FcR transduction subunits. In most cases, the responsible kinase is Lyn. Whether src kinases are constitutively associated with FcR subunits and transphosphorylate ITAMs upon FcR aggregation (Pribluda et al., 1994), or whether ITAMs are phosphorylated in lipid rafts, where src kinases are concentrated (Brown and London, 2000), upon translocation of FcR aggregates into these microdomains (Field et al., 1997) still

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needs to be clarified. In any case, phosphorylated ITAMs provide docking sites that mediate the recruitment of SH2 domain‐containing molecules, among which is the two‐SH2 domain‐containing protein tyrosine kinase Syk (Benhamou et al., 1993). Once recruited, Syk is tyrosyl‐phosphorylated by src kinases and it further auto‐phosphorylates (Kimura et al., 1996). This activates its catalytic activity. Syk then phosphorylates tyrosines in multiple molecules (Costello et al., 1996). Among these are the cytosolic adapter molecule SH2 domain‐containing Leukocyte Protein of 76 kDa (SLP‐76) (Hendricks‐Taylor et al., 1997; Kettner et al., 2003) and the raft‐associated transmembrane adapter Linker for Activation of T cells (LAT) (Wonerow and Watson, 2001). A parallel series of src kinase‐initiated events was described, following FceRI aggregation in mouse mast cells. Fyn was indeed found to tyrosyl‐ phosphorylate the cytosolic adapter Gab2, thus enabling its association with the p85 subunit of Phosphatidylinositol 3‐kinase (PI3K) via its SH2 domain, and the subsequent activation of the p110 catalytic subunit of this enzyme (Parravincini et al., 2002). PI3K generates phosphatidyl (3,4,5)tris‐phosphate [PI(3,4,5)P3] by adding a phosphate group at position 3 in phosphatidyl (4,5) bis‐phosphate. Several molecules that contain a Pleckstrin Homology (PH) domain are recruited to the membrane by PI(3,4,5)P3. 2.4. Organization of FcR Signaling Complexes by Adapter Proteins The many molecular interactions that occur in signalosomes generate signals that are organized by tyrosine‐rich adapter molecules which, when phosphorylated, function as scaffold proteins. These include cytosolic and transmembrane adapters. SLP‐76 is one such cytosolic adapter. Besides its N‐terminal SH2 domain, SLP‐76 contains a central proline‐rich region and multiple C‐terminal tyrosines (Jackman et al., 1995). Once phosphorylated, it binds to a variety of molecules including the exchange factor Vav (Tuosto et al., 1996) and other adapters such as Gads, Nck, and SLAP‐130 (Boerth et al., 2000). Based on studies of cells from SLP‐76‐deficient mice, SLP‐76 was shown to contribute to the activation of phospholipase C‐g (PLC‐g) and to the activation of Mitogen‐Activated protein (MAP) kinases (Pivniouk et al., 1999). Transmembrane adapters consist of a short extracellular domain, unlikely to bind extracellular ligands; a single transmembrane domain; and a long intracellular domain devoid of molecular interaction domains, but rich in tyrosine residues. When phosphorylated upon FcR engagement, these tyrosines function as inducible docking sites for cytosolic molecules having SH2 domains.

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Transmembrane adapters are of two types, depending on the presence, in their intracytoplasmic domain, of a juxtamembrane CxxC motif which targets them to lipid rafts. The Protein Associated with GEMs/Csk‐binding protein (PAG/Cbp), LAT, Non‐T cell Activation Linker/Linker of Activation for B cells (NTAL/LAB), and Lck‐Interacting Membrane protein (LIME) have such a palmitoylation site. The T cell Receptor‐Interacting Molecule (TRIM), SHP‐2‐Interacting Transmembrane adapter (SIT), and Linker of Activation for X cells (LAX) do not, and they are excluded from lipid rafts (Kliche et al., 2004; Togni et al., 2004). LAT was shown to support positive signaling triggered not only by TCR, but also by FceRI, and the mechanisms by which it concurs to mast cell activation and to T cell activation are thought to be similar. FceRI aggregation in BMMC from LAT
/
mice triggered a reduced phosphorylation of SLP‐76 and of PLC‐g, resulting in a decreased Ca2þ mobilization and MAP Kinase activation and, ultimately, in a decreased release of preformed mediators and secretion of cytokines (Saitoh et al., 2000). FcRb/FcRg ITAMs and Syk were phosphorylated as in wt cells. These observations suggested that LAT primarily serves as a coupling molecule between immunoreceptors and intracellular signaling pathways leading to cellular responses (Sommers et al., 2004). LAT contains many tyrosines (9 in mice, 10 in humans) in its intracytoplasmic domain (Weber et al., 1998; Zhang et al., 1998a). It is tyrosyl‐phosphorylated by Syk following FceRI engagement, and serves as a scaffold molecule by providing multiple docking sites for additional SH2 domain‐containing cytosolic enzymes and adapters to be recruited. These include PLC‐g, protein tyrosine kinases of the Tec family, the p85 subunit of PI3K, exchange factors of the Vav family, and the adapters Gads, Grap, and Grb2 (Weber et al., 1998; Zhang et al., 1998a, 2000). Works based on mutational analysis of LAT identified critical tyrosine residues involved in the recruitment of these molecules in T cells (Zhang et al., 2000; Zhu et al., 2003). These were the four distal tyrosines (Y132, Y171, Y191, and Y226 in humans, and their homologues in mice Y136, Y175, Y195, and Y235). Specifically, Y132/136 was demonstrated as being the major binding site for PLC‐g, and the three distal tyrosines (Y171/175, Y191/195, and Y226/235) binding sites for Gads, Grap, and Grb2 (Zhang et al., 2000). The two sets of binding sites also contribute to the recruitment of other molecules such as SLP‐76 via Gads and they cooperate to stabilize the binding of molecules recruited by each other. A mutational analysis of the four distal tyrosines of LAT, in LAT
/
BMMC reconstituted in vitro with wt or mutant LAT (Saitoh et al., 2003), confirmed that, once phosphorylated upon FceRI engagement, these residues play critical roles for FceRI signaling by recruiting the same set of signaling molecules in mast cells as in T cells.

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2.5. Intracellular Propagation of FcR Signals Molecules recruited and activated in signalosomes concur to the activation of metabolic pathways which propagate signals intracellularly up to the nucleus and back to the plasma membrane. Several pathways are used by activating FcRs. They are, with variations, the same as pathways used by other immunoreceptors. Some lead to the calcium response, while others lead to the activation of transcription factors. These pathways are not linear, but tightly interconnected. We will briefly underline only critical steps that either contribute to or are targets of negative regulation. The calcium response results from the recruitment and activation of PLC‐g1 and/or 2, depending on the cell type (Wang et al., 2000; Wen et al., 2002). The recruitment of PLC‐g involves the interaction of one of its SH2 domain with phosphorylated Y136 on LAT, and the interaction with the adapter Gads which binds to phosphorylated LAT terminal tyrosines. PLC‐g is also recruited to the membrane through the interaction of its PH domain with newly formed PI (3,4,5)P3. PLC‐g is subsequently activated as a result of the phosphorylation of specific tyrosine residues by Syk and by the Tec kinase Btk (Humphries et al., 2004), respectively. PLC‐g generates inositol (1,4,5)tris‐phosphate [IP(1,4,5) P3 or IP3] and Diacyl Glycerol (DAG). IP3 triggers an efflux of intracellular Ca2þ from the endoplasmic reticulum and, secondarily, an influx of extracellular Ca2þ. The result is a markedly increased intracellular Ca2þ concentration. Intracellular Ca2þ is critical for exocytosis in mast cells. It also activates calcineurin. This phosphatase dephosphorylates the Nuclear Factor of Activation for T cells (NF‐AT), which enables its translocation from the cytosol to the nucleus (Stankunas et al., 1999). DAG upregulates the catalytic activity of several among the many serine‐ threonine kinases of the Protein Kinase C (PKC) family. Following further activation as a result of the phosphorylation of several serine/threonine and tyrosine residues, PKCs phosphorylate a variety of substrates involved in the activation of MAP kinases (Kawakami et al., 1998) and of transcription factors (Turner and Cantrell, 1997), and in mast cell degranulation (Buccione et al., 1994). PKCs can also threonyl‐phosphorylate FcRg (Pribluda et al., 1997), which contributes to the activation of Syk (Swann et al., 1999). Another substrate of DAG‐activated PKCs is the serine Protein Kinase D (PKD) (Valverde et al., 1994). PKD is abundant in mast cells and, when activated upon FceRI engagement, it contributes to the regulation of transcriptional activity of NF‐kB (Johannes et al., 1998). NFkB activation was observed upon FceRI aggregation in mast cells (Hundley, Blood, 2004) and dendritic cells (Kraft et al., 2002), preceded by the seryl‐phosphorylation and degradation of IkB, and it was reported to be

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involved in the generation of several cytokines (Marquardt and Walker, 2000). NF‐kB was also activated in human monocytes (Drechsler et al., 2002) and mesangial cells (Duque et al., 1997), as a consequence of FcgRs and FcaRI aggregation, respectively. Three sets of MAP kinases are activated upon FcR engagement: Erk1/2, JNK, and p38 (Dong et al., 2002). Erk1/2 are the terminal effector kinases of the Ras pathway, JNK and p38, effector kinases of the rac pathway. Ras and Rac are small G proteins which are in an inactive form when associated with GDP, and in an active form when associated with GTP. The replacement of GDP by GTP on Ras and Rac depends on the exchange factors Sos and Vav, respectively (Cantrell, 1998; Downward, 1996). It initiates a cascade of serine/ threonine phosphorylations, the ultimate substrates of which are MAP kinases. Phosphorylated MAP kinases are translocated into the nucleus where they can phosphorylate transcription factors. These associate with NF‐AT to form a complex which can bind to specific sites in the promoter of cytokine genes and initiate their transcription. 2.6. Ligand Valency Influences FceRI‐Dependent Mast Cell Secretory Responses Several observations recently challenged the widely accepted concept that the binding of monomeric IgE to FceRI generates no detectable signal and no detectable response. An exposure of mast cells to IgE in the absence of antigen was indeed reported (1) to up‐regulate the expression of membrane FceRI (Hsu and MacGlashan, 1996; MacGlashan et al., 1997), (2) to increase the survival of mast cells in the absence of growth factors (Asai et al., 2001; Kawakami and Galli, 2002), and (3) to induce cytokine secretion (Kalesnikoff et al., 2001; Kohno et al., 2005; Pandey et al., 2004). The effect of monomeric IgE on mast cell survival and cytokine secretion was found to depend on the FcRg ITAM (Kohno et al., 2005), but not the upregulation of FceRI expression. The effect on receptor expression was shown to result from slowing down the removal of FceRI from the membrane and its subsequent degradation without affecting the rate of FceRI synthesis (Borkowski et al., 2001). As a consequence, FceRI accumulate on the mast cell membrane without requiring detectable intracellular signals. By contrast, the effects on mast cell survival and cytokine secretion were found to be restricted to anti‐DNP/TNP IgE, to vary markedly from one mAb IgE to another (Kitaura et al., 2005), and most importantly, to be inhibited by a monovalent hapten such as DNP‐lysine (Tanaka et al., 2005). These effects, therefore, must be understood as resulting from FceRI aggregation, whatever the mechanism, that is, to obey the general rule. Interestingly, however, quantitative variations of receptor aggregation were found to result in qualitative

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variation in cellular responses. A low level of FceRI aggregation, induced by incubating mast cells with IgE in the absence of (known) antigen, triggered intracellular signals leading to the secretion of IL‐3 or MCP‐1, but not to degranulation, whereas a high level of receptor aggregation, induced by incubating with multivalent antigen mast cells sensitized with the same IgE, triggered both degranulation and cytokine secretion (Gonzalez‐Espinosa et al., 2003; Kohno et al., 2005; Yamasaki et al., 2004). Molecular mechanisms that enable quantitative differences in receptor aggregation to produce qualitatively different responses remain to be elucidated. 3. Negative Signaling by Activating FcRs FcR‐dependent cell activation is negatively regulated by several inhibitory mechanisms generated by activating FcRs themselves. Some are triggered together with activation mechanisms by ITAM‐containing FcRs and contribute to their own, autonomous control. Others can be triggered by activating FcRs in the absence of detectable positive signals, although they depend on ITAMs, and can negatively regulate signaling triggered by other activating receptors expressed by the same cell. 3.1. Autonomous Negative Regulation of Activating FcRs When engaged by antibody and antigen, activating FcRs not only generate positive signals, but also negative signals. This autonomous negative regulation controls the intensity and duration of positive signals. Negative signaling depends on several mechanisms involving a variety of molecules. Interestingly, many among the proteins which contribute to negative regulation are the same as those which contribute to positive regulation. These include receptor subunits, protein tyrosine kinases, adapter molecules, and phosphatases. 3.1.1. FcRb The mast cell‐specific FcRb subunit was first understood to function as an amplifier of signals generated by FcRg upon FceRI aggregation (Adamczewski et al., 1995; Dombrowicz et al., 1998; Lin et al., 1996). Differing from mouse or rat FceRIa, which need to associate with both FcRg and FcRb in order to be expressed at the mast cell membrane, human FceRIa need to associate with FcRg only. As a consequence, FceRI can be expressed in human mast cells with or without FcRb. They can also be expressed by human monocytes, macrophages, and eosinophils, which do not express FcRb (Maurer et al., 1994), but not in corresponding murine cells. Signals triggered by FcRb‐

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associated FceRI were found to be of a higher intensity than signals triggered by FceRI associated with FcRg only (Dombrowicz et al., 1998). FcRb also enhances IgE‐induced allergic responses by up‐regulating the surface expression of FceRI (Donnadieu et al., 2003). Recently, however, FcRb was found to generate ITAM‐dependent negative signals. The FcRb ITAM has a unique feature. Compared to other ITAMs, the FcRb ITAM contains an additional tyrosine residue, in the 6‐residue sequence that separates the two canonical YxxL motifs:

Human Mouse

FcRg

FcRb

YTGLSTRNQETYETL YTGLNTRSQETYETL

YEELNIYSATYSEL YEELNVYSPIYSEL

Based on a mutational analysis, this additional tyrosine was shown to be involved in the negative regulation of IgE‐induced signals. The activation of the MAP kinases Erk and p38, the activation of NF‐kB, and, ultimately, the secretion of IL‐6, IL‐13, and TNF‐a were indeed enhanced when this residue was mutated into phenylalanine (Furumoto et al., 2004). No marked effect was observed on the activation of PLC‐g, the Ca2þ response, the generation of leukotrienes, and the release of b‐hexosaminidase, suggesting that this tyrosine is not critical in signal amplification. These altered responses were reminiscent of the phenotype of mast cells derived from Lyn‐deficient mice (Odom et al., 2004). Indeed, pull‐down experiments using beads coated with phosphopeptides corresponding to a wt or an altered FcRb ITAM showed that the additional tyrosine could mediate the binding of Lyn, and also of the SH2 domain‐containing inositol phosphatase SHIP1. Supporting an in vivo significance of this in vitro analysis, slightly less FcRb coprecipitated with Lyn, and SHIP1 was less phosphorylated following FceRI engagement, when the additional tyrosine was mutated in FcRb. FcRb may therefore contribute to the involvement of SHIP1 and to the recruitment of Lyn in FceRI signaling complexes. Increasing evidence supports the idea that this src family protein tyrosine kinase contributes to negative regulation of immunoreceptor signaling and, possibly, more than to positive regulation as originally thought. 3.1.2. Lyn The src‐family protein tyrosine kinase Lyn was shown to play a critical role in the initiation of IgE‐induced signal transduction in mast cells. Lyn was indeed demonstrated to be responsible for the phosphorylation of both FcRb and FcRg ITAMs upon FceRI aggregation and for the initial phosphorylation of

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Syk, when the latter has been recruited to the phosphorylated FcRg ITAM (Jouvin et al., 1994; Kihara and Siraganian, 1994; Scharenberg et al., 1995). Lyn was therefore considered first as a major player in positive signaling. When Lyn‐deficient mice became available, it became apparent that Lyn is involved in a variety of negative regulatory processes. B cells from Lyn
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mice were found to be hyper‐responsive to BCR engagement (Chan et al., 1997; Hibbs et al., 2002), and to IL‐4 stimulation (Janas et al., 1999). Lyn
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mast cells were also more responsive to proliferative signals delivered by IL‐3 or Stem Cell Factor (Hernandez‐Hansen et al., 2004). Importantly, Lyn
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mast cells were more responsive to FceRI‐dependent activation signals (Kawakami et al., 2000; Nishizumi and Yamamoto, 1997). As expected, IgE‐induced phosphorylation of FceRI ITAMs was reduced in Lyn
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BMMC (Kawakami et al., 2000; Kovarova et al., 2001). The phosphorylation of Csk‐binding protein (Cbp) was abrogated and, as a consequence, the coprecipitation of Csk with this scaffold adapter protein observed in wt mast cells was lost in Lyn
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mast cells. Noticeably, the catalytic activity of Fyn was increased in these cells, and hyperactive Fyn was phosphorylated on tyrosine 417, in the activation loop of the kinase. The hyper‐responsiveness of Lyn
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mast cells to IgE could be ascribed to this kinase as this phenotype was abrogated in BMMC derived from doubly deficient Lyn
/
/Fyn
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mice (Odom et al., 2004). Altogether these data provided the following explanation to the negative role of Lyn in mast cell activation. In wt cells, Lyn phosphorylates Cbp which recruits Csk. Csk phosphorylates the regulatory tyrosines 508 and 528 of Fyn and thereby inhibits its catalytic activity (Odom et al., 2004). Interestingly, the phenotype of Lyn
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mice was reminiscent of an ‘‘allergic’’ phenotype which could not be accounted for by the hyper‐reactivity of mast cells only. As these mice grew older, they displayed an increased serum IgE concentration, an upregulation of FceRI expression on mast cells, increased numbers of peritoneal mast cells and eosinophils, and elevated levels of plasma histamine (Odom et al., 2004). Most of these allergy‐associated traits could be ascribed to a screwed isotypic switch toward IgE during B cell differentiation due to the hyper‐responsiveness of Lyn
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B cells to IL‐4, and to the consequences of an increased IgE serum concentration. Finally, besides its first recognized role in positive signaling by immunoreceptors, a critical role of Lyn kinase in negative signaling that dampens cell activation by these receptors must be considered. Whether Lyn primarily contributes to positive or to negative signaling may depend on the cell type and on engaged receptors. 3.1.3. LAT LAT has been first understood to organize signalosomes generated by activating receptors and to couple them with downstream signaling pathways leading

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to cellular responses (Sommers et al., 2004). LAT is critical for TCR signals involved in early T cell differentiation and, indeed, LAT‐deficient mice display an arrest in thymocyte development with a block in both TCRab and gd T cell differentiation (Zhang et al., 1999). Unexpectedly, knock‐in mice, expressing LAT with a single point mutation of the PLC‐g‐binding site (Y136F), displayed an aberrant T cell development characterized by a partial block in early T cell differentiation and polyclonal lymphoproliferative disorder, resulting in abnormally high numbers of CD4þ TCRab T cells that secreted abnormally high levels of TH2 cytokines in the periphery. As a consequence of this exaggerated TH2 polarization, serum IgG1 concentrations were 5000‐fold higher than in wild‐type mice, serum IgE concentrations were in the range of several mg/ml, instead of a few mg/ml, and peripheral tissues were massively infiltrated with eosinophils. The differentiation of TCRgd T cells was unaffected (Aguado et al., 2002; Sommers et al., 2002). Likewise, knock‐in mice bearing point mutations of the adapter‐binding three distal tyrosines of LAT (Y175F, Y195F, and Y235F) displayed a complete block in the differentiation of TCRab T cells and an abnormal differentiation of TCRgd T cells, also resulting in an exaggerated TH2 polarization and massive proliferation. As a result, IL‐4 secretion was increased, and the serum concentrations of IgG1 and IgE were 500‐ and 1000‐fold higher than in normal mice, respectively (Nun˜ez‐Cruz et al., 2003). Although they affect two distinct T cell lineages, respectively, the two types of LAT tyrosine mutations therefore seemed to inhibit a negative regulation that normally controls terminal T cell differentiation. This suggested that, besides positive signals, LAT might support negative signals that normally regulate terminal T cell differentiation and proliferation, and that this regulation, which differentially affects TCRab and TCRgd signaling, depends on distinct tyrosine residues. Our analysis of IgE‐induced biological responses of cultured mast cells derived from the same knock‐in mice led to the same conclusion for FceRI signaling. A systematic comparison of biologic responses observed in pairs of mutants enabled us to dissect the respective roles played by LAT tyrosines in mast cells (Malbec et al., 2004). As expected, Y136 and the three distal tyrosines differentially contributed to exocytosis and the secretion of cytokines, on the one hand, and to the generation or the activation of major cytosolic effectors such as intracellular Ca2þ and the terminal MAP kinases of the ras pathway, Erk1/2, on the other hand. Interestingly, mutations unraveled the existence of negative signals, generated by distinct LAT tyrosines. Thus Y136 had a negative effect on mediator release when Y175, 195, and 235 were mutated and, conversely, Y175, 195, and 235 had a negative effect when Y136 was mutated. Positive and negative signals generated by different segments of the LAT molecule are apparently additive. Thus, sequences containing the five proximal tyrosines

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could abrogate the negative effects of Y136 in the absence of the three distal tyrosines or the negative effects of the three distal tyrosines in the absence of Y136, observed on b‐hexosaminidase release in BMMC. Importantly, LAT can integrate positive and negative signals even when in a wt configuration. Thus, the four distal tyrosines together had a positive effect on b‐hexosaminidase release in BMMC, but of a lower magnitude than the intense positive effects of either Y136 alone or of the three distal tyrosines alone. These observations would be best explained if LAT could promote the assembly of a signaling complex composed of a mixture of intracellular molecules with antagonistic properties. 3.1.4. NTAL Another transmembrane adapter was recently cloned as a result of a search for the B cell homologue of LAT and was named Linker for activation of B cells (LAB) (Janssen et al., 2003). Because it is expressed not only by B cells, but also by monocytes, NK cells, mast cells, and platelets, this molecule was also named Non‐T cell Activation Linker (NTAL) (Brdicka et al., 2002). NTAL is encoded in humans by the WBSCR5 gene, on chromosome 7 (Brdicka et al., 2002; Martindale et al., 2000). It consists of a single polypeptide resembling LAT, with a short extracellular domain, a transmembrane domain with a potential palmitoylation CxxC motif, and a long intracytoplasmic domain containing nine tyrosine residues that are phosphorylated upon immunoreceptor engagement and provide multiple binding sites for SH2 domain‐containing molecules. Grb2, Sos, Gab1, and c‐Cbl indeed coprecipated with phosphorylated NTAL in monocytes and B cells (Brdicka et al., 2002). Differing from LAT, NTAL contains no PLC‐g binding site. When expressed in LAT‐deficient T cells, NTAL could partially restore TCR signaling (Koonpaew et al., 2004), and LAT
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mice expressing an NTAL transgene under the control of the CD2 promoter had a phenotype resembling that of LAT Y136F knock‐in mice (Janssen et al., 2004). Based on these observations, NTAL was proposed to play, in B cells, a similar role as the one LAT plays in T cells (Brdicka et al., 2002). Because mast cells co‐express LAT and NTAL and because FceRI signaling was reduced, but not abrogated in BMMC derived from LAT‐ deficient mice, the two adapters were thought to play complementary roles in mast cell activation. Surprisingly, the genetic deletion of NTAL resulted in increased, rather than diminished, IgE‐induced release of granular mediators and secretion of cytokines by mast cells (Volna et al., 2004; Zhu et al., 2004). The tyrosyl‐phosphorylation of Syk, LAT, and PLC‐g1 and 2 were increased, as well as the phosphorylation of the Erk, p38, and JNK MAP kinases in BMMC from NTAL‐deficient mice. The activity of PI3K, the concentration of PI

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(3,4,5)P3, the amount of IP3, and the Ca2þ response were also increased. NTAL therefore appears to negatively regulate FceRI signaling. The mechanism of inhibition still needs to be elucidated. Whether NTAL recruits an inhibitory molecule, such as a phosphatase, is one possibility that has not been convincingly demonstrated. Whether a competition between LAT and NTAL, which recruit a common set of adapter molecules, exists is supported by the observation that the phosphorylation of LAT is increased in the absence of NTAL (Volna et al., 2004) and that, reciprocally, the phosphorylation of NTAL is increased in the absence of LAT. The augmented phosphorylation of LAT in NTAL
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mast cells is likely to explain the increased phosphorylation of PLC‐g and its consequences on IP3 production and Ca2þ mobilization. Also, NTAL lacks a PLC‐g binding site, and the recruitment of PLC‐g by LAT requires the cooperative binding of several among the adapters that are recruited by both LAT and NTAL and that NTAL might sequester. Against the competition hypothesis, LAT and NTAL were found to reside in distinct lipid rafts on the plasma membrane (Volna et al., 2004). Whether these different microdomains could possibly merge during FceRI signaling, as it was suggested (Rivera, 2005), needs to be demonstrated. Interestingly, NTAL may not only generate negative signals, but also contribute to positive signals in mast cells. These could be observed in the absence of LAT. Inhibition of mediator release was indeed reported to be more pronounced in BMMC from LAT‐ and NTAL‐doubly deficient mice than in BMMC from LAT‐deficient mice (Volna et al., 2004; Zhu et al., 2004). A positive role of NTAL could be seen on Ca2þ responses in T and B lymphocytes (Brdicka et al., 2002; Janssen et al., 2004) and on Stem Cell factor‐ induced activation of human mast cells (Tkaczyk et al., 2004). A recent analysis performed in DT40 B cells proposed that, when recruiting Grb2, phosphorylated NTAL removes a Grb2‐dependent inhibitory effect on the BCR‐induced influx of extracellular Ca2þ (Stork et al., 2004). This inhibitory effect could be due to protein tyrosine phosphatases and inositol phosphatases which associate with Grb2 in different conditions. 3.1.5. Protein Tyrosine Phosp hatases Protein tyrosine phosphatases are thought to negatively regulate FcR signaling. Supporting evidence is, however, scarce. The SH2 domain‐containing Protein Tyrosine Phosphatase SHP‐1 has been implicated in FceRI signaling by using trapping mutants (Xie et al., 2000). SHP‐1 was reported to associate with the phosphorylated ITAM of FcgRIIA, Syk, the p85 subunit of PI3K, and Dok‐1, and to decrease the tysrosyl‐phosphorylation of intracellular proteins, upon FcgRIIA aggregation in the macrophage‐like THP‐1 cells (Ganesan

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et al., 2003). SHP‐1 also contains several consensus binding motifs for the SH2 domain of Grb2, and the inhibitory effect of Grb2‐SHP‐1 complexes was observed on cytokine receptor signaling (Minoo et al., 2004). The possible role of the second SH2 domain‐containing Protein Tyrosine Phosphatase SHP‐ 2 in the negative regulation of FcR‐dependent cell activation remains to be demonstrated. Several protein phosphatase devoid of SH2 domain were also found to be activated upon FceRI aggregation and to dephosphorylate ITAMs (Swieter et al., 1995). 3.1.6. Inositol Phosp hatases Inositol phosphatases, by contrast, play a prominent role in controlling FcR‐ dependent cell activation. The inositol 3‐phosphatase PTEN was involved in FcgR signaling as Akt and MAP kinase phosphorylation induced upon FcgRIIIA aggregation was enhanced in macrophages from PTEN
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mice, resulting in enhanced cytokine secretion (Cao et al., 2004). SHIP2 was tyrosyl‐ phosphorylated upon FcgRI engagement in THP‐1 cells or upon FcgRIIA engagement in human peripheral blood monocytes following upregulation by LPS, and it associated via its SH2 domain to the phosphorylated ITAM of this receptor (Pengal et al., 2003). Finally, SHIP1 was described to inhibit FcgRIIA‐dependent phagocytosis in THP‐1 cells (Nakamura et al., 2002), to coprecipitate with phosphorylated FcgRIIA, and to negatively regulate NFk‐ B‐mediated gene transcription during phagocytosis in human myeloid cells (Tridandapani et al., 2002). SHIP1 activity was reported to associate with the phosphorylated z subunit and to negatively regulate FcgRIIIA‐dependent ADCC in human NK cells (Galandrini et al., 2002). SHIP1 was found to bind in vitro to phosphopeptides corresponding to the FcRb ITAM (Kimura et al., 1997) and to interact with FcRb when examined by yeast triple hybrid assay (Osborne et al., 1996). The possible in vivo recruitment of this phosphatase in FceRI signaling complexes remains elusive as, so far, it was not reported to coprecipitate with FceRI, including the FcRb subunit, following receptor engagement in mast cells. SHIP1 was, however, understood to play a central regulatory role in the autonomous negative regulation of FceRI signaling. This conclusion was based on studies of SHIP1‐deficient mice. As they get older, SHIP1
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mice spontaneously develop a splenomegaly and a progressive lung infiltration by myeloid cells that leads to a waste syndrome and, ultimately, to a shortened life span. Their myeloid progenitor cells are hyper‐responsive to cytokines, such as IL‐3, and growth factors, such as Granulocyte/Macrophage Colony‐Stimulating Factor and Stem Cell Factor (Helgason et al., 1998). Interestingly, BMMC derived from SHIP1
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mice are hyper‐responsive not only to Stem Cell Factor‐, but also to IgE‐dependent

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stimulation. Such cells indeed release more b‐hexosaminidase than do BMMC derived from wt mice in response to FceRI aggregation by IgE and antigen. Supporting the conclusion that BMMC from SHIP1
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mice could respond to a lower degree of receptor aggregation, IgE anti‐DNP alone could trigger these cells, but not wt‐type cells, to release b‐hexosaminidase, as well as an array of cytokines. These antigen‐independent responses were inhibited by a monovalent hapten such as DNP‐lysine (Huber et al., 1998; Kalesnikoff et al., 2001). IgE‐induced increased degranulation was correlated with augmented and sustained Ca2þ mobilization and Erk1/2 activation. The phosphorylation of Shc, which associates constitutively to SHIP1, was reduced in the absence of SHIP1, but, surprisingly, FcRb phosphorylation was increased. Based on these data, SHIP1 was proposed to raise the threshold of FceRI aggregation needed to generate activation signals and to function as a ‘‘gatekeeper’’ of mast cell degranulation (Huber et al., 1998). SHIP1 is constitutively active. By contrast with SHPs, the phosphatase activity of SHIP1 is not up‐regulated when its SH2 domain binds to a tyrosyl‐phosphorylated motif, but when it is translocated close to the membrane (Bolland et al., 1998). The expression of a membrane‐targeted CD8‐SHIP1 chimera in COS cells constitutively induced a three‐fold higher enzymatic activity than the expression of a cytosolic form of SHIP1 (Phee et al., 2000). A simple explanation is that, under these conditions, SHIP1 is located close to its membrane substrate. SHIP1 removes 5‐phosphate groups in the inositol ring of 3‐phosphorylated inositides and phosphatidylinositides. Its substrates are inositol (1,3,4,5)tetrakis‐phosphate [I(1,3,4,5)P4] and PI(3,4,5)P3, which are hydrolyzed into inositol (1,3,4)tris‐phosphate and into phosphatidylinositol (3,4)bis‐phosphate, respectively (Damen et al., 1996). SHIP1 can therefore prevent PI(3,4,5)P3‐dependent critical upstream events leading to the Ca2þ response and, as a consequence, inhibit cell responses (Scharenberg and Kinet, 1998; Scharenberg et al., 1998). Another role of SHIP1 in autonomous negative regulation was recently unraveled. This regulation accounts for the bell‐shaped curve of mast cell activation as a function of antigen concentration. Inhibition of biological responses in excess of antigen is unique neither to FceRI nor to mast cells. It was long interpreted as resulting from a progressive decrease in receptor aggregation, due to a competition of high concentrations of antigen for efficiently crosslinking FceRI‐bound IgE (Dembo et al., 1978; Wofsy et al., 1978), although negative regulation had previously been hypothesized as an explanation, resulting from an excess of receptor aggregation (Magro and Alexander, 1974). Supporting experimental predictions deduced from a mathematical analysis (Delisi and Siraganian, 1979), recent works provided evidence that intracellular signals do not decrease, but increase, as the concentration of

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antigen increases. Thus, the tyrosyl‐phosphorylation of intracellular proteins in whole cell lysates and, more specifically, of FcRb and PLC‐g were of a higher magnitude in BMMC stimulated with supra‐optimal concentrations of antigen than in BMMC stimulated with an optimal antigen concentration. The secretory response decreases, however, because negative signals increase and become dominant over positive signals. Supporting this interpretation, the inducible tyrosyl‐phosphorylation of SHIP1 dose‐dependently increased with the concentration of antigen, even after supra‐optimal concentrations were reached. Most importantly, inhibition of secretion induced by an excess of antigen in mast cells derived from wt mice was abrogated in mast cells derived from SHIP1‐deficient mice (Gimborn et al., 2005). These data altogether indicate that SHIP1, possibly recruited by FcRb when heavily phosphorylated as a result of supra‐optimal receptor aggregation, is the effector of autonomous negative regulation of FceRI signaling that dampens mast cell activation in excess of ligand. 3.1.7. Cbl Finally, ubiquitination of receptors and signaling molecules, followed by proteasomal degradation, were shown to terminate cell activation. Thus, following FceRI engagement, FcRb and FcRg, as well as Syk, undergo rapid c‐Cbl‐ dependent E3 ligase‐mediated ubiquitination (Gimborn et al., 2005). Lyn also associates with c‐Cbl and is ubiquitinated and degraded in IgE‐activated mast cells (Kyo et al., 2003). Likewise, Syk and ZAP‐70 are ubiquitinated following FcgRIIIA engagement in human NK cells (Paolini et al., 2001). 3.2. Promiscuous Negative Regulation of Activating FcRs by FcaRI ITAM‐containing FcRs were recently demonstrated to have the ability of generating not only positive and negative signals which regulate each other, but also negative signals which can affect positive signals delivered by other activating FcRs in the same cell. FcaRI are such receptors. They bind monomeric IgA with a moderate affinity and dimeric IgA with a high avidity (Wines et al., 2001). FcaRI are encoded by genes of the Leukocyte Receptor Complex, on chromosome 19. They share with receptors encoded by this gene family a KIR‐type orientation, instead of an FcR‐type orientation of their extracellular domains (Herr et al., 2003). Although FcaRI can be expressed without, FcaRI associate with FcRg and, upon aggregation by IgA immune complexes, they trigger cell activation like other ITAM‐containing immunoreceptors. They are expressed by a variety of myeloid cells which contribute to inflammation (Monteiro and Van De Winkel, 2003).

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Surprisingly, the engagement of FcRg‐associated FcaRI by monomeric ligands—Fab fragments of mAbs against the extracellular domains of human FcaRI or human serum IgA—was found to negatively regulate the in vitro phagocytosis of IgG‐opsonized bacteria by human monocytes or IgE‐ dependent exocytosis in the rat mast cell line RBL‐2H3 transfected with cDNA encoding FcaRI. When administered intraperitoneally into human FcaRI transgenic mice, anti‐FcaRI Fab fragments also inhibited bronchial constriction and airway infiltration by inflammatory cells induced by IgE and antigen in a murine model of allergic asthma. Using chimeric molecules made of the a subunit of FcaRI the transmembrane domain of which had a point mutation preventing the association with FcRg and the intracytoplasmic domain of which was replaced by that of FcRg (FcaRI/FcRg chimeras) expressed in RBL transfectants, the authors demonstrated that inhibition depended on the FcRg ITAM, and that both tyrosines were required for inhibition. These tyrosines were phosphorylated following monovalent engagement of FcaRI/ FcRg chimeras, but to a much lower extent than following plurivalent engagement. Inhibition was a slow process, taking 6 hrs to be complete. Interestingly, inhibition induced by monovalent ligands was correlated with the coprecipitation of SHP‐1 with weakly phosphorylated FcaRI/FcRg chimeras. Indeed, SHP‐1 did not detectably coprecipitate with chimeras that were heavily phosphorylated following cell activation induced by multivalent ligands. Finally, the coprecipitation of SHP‐1 with FcaRI/FcRg chimeras was dose‐dependently inhibited by a MEK inhibitor, suggesting a positive role of Erk in SHP‐1 recruitment. Intriguingly, when engaged by monovalent Fab fragments of a mAb against the extracellular domain of FcgRIIB, FcgRIIB/FcRg chimeras failed to inhibit IgE‐induced mediator release in the same cells, suggesting that, beside the intracytoplasmic ITAM, the ligand and/or the extracellular domain of the chimera were critical for inhibition (Pasquier et al., 2005). Also, inhibition is unlikely to depend on the mere membrane recruitment of SHP‐1. IgE‐induced mediator release and intracellular signaling were indeed not impaired in RBL transfectants expressing FcgRIIB whose intracytoplasmic domain had been replaced by the catalytic domain of SHP‐1 (Hardre´‐Lie´nard and Dae¨ron, unpublished data). Whatever the mechanism of inhibition, these results have several important implications. First, they support the evidence that, although not able to fully activate cells, interactions of ITAM‐containing immunoreceptors with monovalent ligands can generate intracellular signals. Second, they indicate that FcaRI can generate either positive or negative signaling depending on extracellular ligands available (i.e., depending on whether IgA are in complexes with specific antigen or not). Whether other ITAM‐containing receptors may exert similar dual functions or whether it is a unique feature of FcaRI is not

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known. FceRI do not seem to inhibit cell activation by other ITAM‐containing receptors when occupied by monomeric IgE as they are under physiologic conditions. Third, they suggest that FcaRI may negatively regulate activation signals triggered by many other receptors. Negative regulation by ITAM‐ containing receptors apparently did not require that inhibitory and activating receptors be coaggregated at the cell surface. If this conclusion proves to be correct, one can expect that many biological responses be affected by monovalent ligand‐induced negative regulation by FcaRI. SHP‐1 can indeed inhibit most if not all activation processes triggered by receptors whose signaling depends on tyrosyl‐phosphorylation of proteins. Finally, these findings provide a possible explanation and molecular basis to the paradox that, although IgA receptors can activate inflammatory cells (Patry et al., 1995), IgA have long been known to have general anti‐inflammatory effects (Russell et al., 1997) and to the observation that selective IgA deficiencies are correlated with increased susceptibility to autoimmune and allergic diseases (Schaffer et al., 1991). 4. Negative Signaling by Inhibitory FcRs By contrast with FcaRI‐dependent negative regulation, FcgRIIB‐dependent negative regulation requires that the inhibitory receptors be coaggregated with activating receptors by a common extracellular ligand and affects cell signaling triggered by these receptors. 4.1. Inhibitory FcRs and ITIMs The inhibitory properties of FcgRIIB lie on the presence of an ITIM in their intracytoplasmic domain. First identified in FcgRIIB (Dae¨ron et al., 1995a), ITIMs were subsequently found in a large number of inhibitory receptors that control the biologic activities of hematopoietic cells (Long, 1999). Sequence alignments of these ITIMs made it possible to define ITIMs structurally. ITIMs consist of a sequence containing a single tyrosine (Y) followed by a hydrophobic residue (I, V, or L) at position Y þ 3 and preceeded by a less conserved hydrophobic residue at position Y
2 (Vivier and Dae¨ron, 1997). One consequence of the coaggregation of FcgRIIB with activating receptors is the phosphorylation of their ITIM. FcgRIIB are not tyrosyl‐ phosphorylated when aggregated at the cell surface. They become phosphorylated when they are coaggregated with activating immunoreceptors (D’Ambrosio et al., 1995) because these provide the src kinase which phosphorylates both ITAMs and ITIMs in receptor coaggregates (Malbec et al., 1998). Due to this peculiarity, FcgRIIB are not inhibitory in resting cells. They do not establish a threshold that must be overcome by activating receptors.

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They become functional ‘‘upon request’’ only, when cell activation has been launched. The phosphorylation of the FcgRIIB ITIM is indeed critical to initiate negative regulation. 4.2. The Recruitment of SHIP1 by FcgRIIB Inhibitory receptors carrying phosphorylated ITIMs (pITIMs) were shown to recruit SH2 domain‐containing cytosolic phosphatases that interfere with signals transduced by ITAM‐bearing receptors (Bolland and Ravetch, 1999). Four such phosphatases have been identified in mice and in humans: the two‐ SH2 domain‐containing Protein Tyrosine Phosphatases SHP‐1 and SHP‐2 and the single‐SH2 domain‐containing inositol 5‐phosphatases SHIP1 and SHIP2. Phosphorylated ITIMs differ from phosphorylated ITAMs by their specificity for SH2‐containing molecules. ITIMs recruit phosphatases only, whereas ITAMs recruit protein tyrosine kinases, adapter molecules, and phosphatases. FcgRIIB were found to differ from other ITIM‐containing receptors by being capable of recruiting SHIP1 and SHIP2. The FcgRIIB ITIM indeed has an affinity for the SH2 domain of SHIPs that other ITIMs lack. Our investigation of the bases of this unique specificity identified several parameters as being critical for SHIP1 to be recruited by FcgRIIB. 4.2.1 The Y þ 2 Leucine Determines the Affinity of the FcgRIIB ITIM for SHIP1/2 First of all, the affinity of FcgRIIB for SHIPs depends on a specific amino acid at position Y þ 2 in the ITIM. As expected from studies that established the molecular bases of the affinity of SH2 domains of other molecules for tyrosyl‐ phosphorylated peptides, the affinity of pITIMs for the SH2 domains of these phosphatases required the conservation of both the Y and the Y þ 3 residues. Synthetic peptides corresponding to pITIMs of all ITIM‐bearing molecules were found to bind SHP‐1 and SHP‐2 in vitro (Burshtyn et al., 1996; D’Ambrosio et al., 1995). The in vitro binding of SHP‐1 and SHP‐2 to the pITIMs of KIR2DL3 and FcgRIIB depends on the Y
2 residue (Ve´ly et al., 1997). Phosphorylated peptides corresponding to the FcgRIIB ITIM, but not phosphorylated peptides corresponding to the KIR2DL3 ITIMs, bound also SHIP1 and SHIP2 (Muraille et al., 2000; Ono et al., 1996). To identify the SHIP‐binding site in FcgRIIB, we exchanged residues between the FcgRIIB ITIM and the N‐terminal ITIM of KIR2DL3. Loss‐of‐function and gain‐of‐ function substitutions identified the Y þ 2 leucine, in the FcgRIIB ITIM, as determining the binding of both SHIP1 and SHIP2, but not the binding of SHP‐1 or SHP‐2. Conversely, the Y
2 isoleucine that determines the in vitro

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binding of SHP‐1 and SHP‐2 affected neither the in vitro binding nor the in vivo recruitment of SHIP1 or SHIP2 (Bruhns et al., 2000). One hydrophobic residue in the ITIM of FcgRIIB therefore determines the affinity for SHIPs. This residue is symmetrical to another hydrophobic residue that determines the affinity of all ITIMs for SHPs. It defines a SHIP‐binding site, distinct from a SHP‐binding site, which confers FcgRIIB their ability to recruit SHIP1 and SHIP2. 4.2.2. The Density of pITIM Determines the Selective Recruitment of SHIP1/2 by FcgRIIB Intriguingly, these two binding sites are not used in vivo. Although agarose beads coated with phosphorylated peptides corresponding to the FcgRIIB ITIM bind in vitro both SHIP1/2 and SHP‐1/2, phosphorylated FcgRIIB, recruit selectively SHIP1/2 in vivo (Fong et al., 1996; Muraille et al., 2000; Ono et al., 1996). When investigating the reasons for this discordance, we found that beads coated with low amounts of pITIM bound SHIP1, but not SHP‐1, i.e., they behaved in vitro like phosphorylated FcgRIIB in vivo. The same was found when examining the binding of pITIM‐coated beads to GST fusion proteins containing the SH2 domain of SHIP1 or the two SH2 domains of SHP‐1. The reason is that the affinity of the SH2 domain of SHIP1 is high enough for binding to pITIM‐coated beads, but not that of either the N‐ or the C‐terminal SH2 domain of SHP‐1 (Lesourne et al., 2001). SHP‐1 indeed requires its two SH2 domains to bind to two pITIMs that are close enough to enable a cooperative interaction. This condition is fulfilled in vitro when beads are coated with sufficient amounts of pITIMs or in vivo when two tandem pITIMs are present in the intracytoplasmic domain of inhibitory receptors such as KIR2DL3. The deletion (Bruhns et al., 1999) or the mutation (Burshtyn et al., 1996) of either ITIM indeed abrogated the ability of KIR2DL3 to recruit SHP‐1. This is not fulfilled by FcgRIIB when coaggregated with activating receptors. When trying to increase FcgRIIB phosphorylation in B cells and mast cells, we found that concentrations of extracellular ligands optimal for FcgRIIB phosphorylation failed to induce the recruitment of SHP‐1. SHP‐1 was, however, recruited by FcgRIIB when the receptors were hyperphosphorylated following cell treatment with pervanadate (Lesourne et al., 2001). These data suggest that, although it can be reached under non‐physiological conditions, a high enough level of FcgRIIB phosphorylation may not be reached, under physiological conditions, to enable the in vivo recruitment of SHP‐1. Whether a regulatory mechanism limits the phosphorylation of FcgRIIB and whether (pathological?) conditions that would lead to the hyperphosphorylation of FcgRIIB might enable the

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recruitment of SHP‐1 that would dephosphorylate signaling molecules are interesting possibilities that remain to be demonstrated. 4.2.3. The Recruitment of SHIP1 by FcgRIIB Requires the Cooperative Recruitment of Cytosolic Adapters Surprisingly, we found that, although sufficient for binding SHIP1 or SHIP2 in vitro, the FcgRIIB pITIM is not sufficient for the receptors to recruit these phosphatases in vivo. It is a general consensus that the FcgRIIB ITIM is both necessary and sufficient for inhibition of cell activation. The conclusion that it is necessary was based on the pioneer work by Amigorena and coworkers who showed that a 13‐amino acid deletion, which was later understood to encompass the ITIM, abrogated inhibition in B cells (Amigorena et al., 1992). A point mutation of the ITIM tyrosine also abrogated FcgRIIB‐dependent inhibition of mast cell and T cell activation (Dae¨ron et al., 1995a), and abolished (Muta et al., 1994) or reduced (Fong et al., 2000) the calcium response in B cells. The conclusion that the ITIM is sufficient was based on works by Muta and coworkers who showed that a chimeric molecule whose intracytoplasmic domain contained the murine FcgRIIB ITIM retained inhibitory properties in B cells (Muta et al., 1994). A C‐terminal deletion of the intracytoplasmic domain of murine FcgRIIB, which left the ITIM intact, however, prevented SHIP1 from being detectably coprecipitated, and reduced the inhibitory effect of FcgRIIB on BCR signaling (Fong et al., 2000). Our recent study showed that this C‐terminal sequence contains a second tyrosine‐based motif that mediates the recruitment of the cytosolic adapter proteins Grb2 and Grap via their SH2 domain and that contributes to the recruitment of SHIP1. The recruitment of the phosphatase indeed required an intact adapter‐binding motif and, conversely, the recruitment of adapters required an intact phosphatase‐binding motif. The reason is that Grb2 and Grap are constitutively associated with SHIP1 via their C‐terminal SH3 domain, and this association increases upon coaggregation of BCR with FcgRIIB. Grb2/Grap thus form a tri‐molecular complex with SHIP1 and FcgRIIB. This stabilizes the binding of the phosphatase to the ITIM and enables its recruitment by murine FcgRIIB. Supporting this conclusion, SHIP1 failed to coprecipitate with FcgRIIB, when tyrosyl‐phosphorylated upon coligation with BCR in mutant DT40 cells lacking both Grb2 and Grap (Isnardi et al., 2004). This requirement may not be peculiar to the interactions between FcgRIIB1, SHIP1, and Grb2. As discussed above, molecules that contain two SH2 domains require the cooperative binding of these two domains to two sequences containing phosphorylated tyrosines in order to be recruited in vivo. The recruitment of ZAP‐70 and Syk (Bu et al., 1995; Kurosaki et al., 1995), or SHP‐1 (Lesourne et al., 2001),

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required the conservation of their two SH2 domains and the conservation of the two tyrosines of ITAMs in immunoreceptors (Kimura et al., 1996) or of the two ITIMs in KIRs (Bruhns et al., 1999; Burshtyn et al., 1999), respectively. Moreover, molecules that contain a single SH2 domain were found to require the cooperation of other SH2 domain‐containing molecules in order to be recruited (Yamasaki et al., 2003). One can therefore propose that one SH2 domain alone may not be sufficient for enabling stable interactions between signaling molecules. 4.3. SHIP1 Accounts for FcgRIIB‐Dependent Negative Regulation 4.3.1. SHIP1 is Necessary and Sufficient for FcgRIIB‐Dependent Negative Regulation Once it has been stably recruited, SHIP1 is the effector of FcgRIIB‐ dependent negative regulation. Evidence supporting this conclusion is as follows. FcgRIIB‐dependent negative regulation was abolished in cultured mast cells derived from the bone marrow of SHIP1‐deficient mice (Malbec et al., 2001), but not in mast cells derived from the bone marrow of moth‐eaten mice which are deficient in SHP‐1 (Fong et al., 1996). FcgRIIB‐dependent inhibition of Ca2þ mobilization was abolished in SHIP1‐deficient chicken DT40 B cells, but not in SHP‐1‐deficient (Ono et al., 1997) or in SHP‐2‐ deficient (Isnardi et al., unpublished observation) DT40 cells. Noticeably, FcgRIIB‐dependent inhibition was only reduced in B cells from SHIP1‐ deficient mice (Brauweiler et al., 2000), possibly because SHIP‐2 could partially replace SHIP1. Although also present in mast cells, SHIP2 could, however, not mediate FcgRIIB‐inhibition in SHIP1‐deficient mast cells. Inhibition was also partially reduced in B cells from moth‐eaten mice (D’Ambrosio et al., 1995). One possible reason is that SHP‐1‐deficient B cells are constitutively hyper‐activated (Pani et al., 1995), which might make BCR‐dependent signaling more difficult to inhibit. These data indicate that SHIP1 is necessary for FcgRIIB‐dependent inhibition of mast cell activation and, most probably, of B cell activation. Evidence that SHIP1 is also sufficient is as follows. B cell (Ono et al., 1997) and mast cell (Malbec et al., 2001) activation were comparably inhibited when BCRs or FceRI were coaggregated with wt FcgRIIB or with FcgRIIB whose intracytoplasmic domain had been replaced by the catalytic domain of SHIP1. In an analysis of a series of FcgRIIB‐SHIP chimeras, we found that, when coaggregated with BCR in the FcgR‐deficient cell line IIA1.6, SHIP1 chimeras abolished IL‐2 secretion, Ca2þ mobilization, Akt phosphorylation, and Erk1/2 phosphorylation. Under the same conditions, SHIP2 chimeras inhibited Akt phosphorylation, but did not affect Erk1/2

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phosphorylation, Ca2þ mobilization, and IL‐2 secretion (Hardre´‐Lie´nard and Dae¨ron, unpublished data). 4.3.2. Two Effector Mechanisms Are Used By SHIP1 in FcgRIIB‐Dependent Negative Regulation SHIP1 mediates FcgRIIB‐dependent inhibition by at least two distinct mechanisms. One depends on its catalytic activity, the other does not. By dephosphorylating PI(3,4,5)P3, SHIP1 prevents the recruitment of PH domain‐containing molecules such as PKB/Akt. The serine/threonine phosphorylation of PKB/Akt observed following BCR or FceRI aggregation was indeed abrogated upon coaggregation of these immunoreceptors with FcgRIIB (Jacob et al., 1999; Malbec et al., 2001). PKB/Akt phosphorylation depends on the membrane translocation of PKB/Akt and of PDK1, the responsible kinase. Both contain one PH domain which targets both the substrate and the enzyme to PI(3,4,5)P3‐rich membrane regions. PKB/Akt phosphorylation is therefore an indirect means to estimate the amount of membrane PI(3,4,5)P3 (Carver et al., 2000). Supporting this approximation, when transfected into B cells, a GFP construct containing the PH domain of Akt that is diffusely distributed in the cytosol of resting cells, translocates to the membrane following BCR aggregation. This translocation was prevented when BCR were coaggregated with FcgRIIB (Astoul et al., 1999). PKB/Akt phosphorylation is critical for mechanisms that prevent apoptosis. Although useful to assess PI(3,4,5)P3 degradation, and although it was recently reported to promote IgG immune complex‐induced phagocytosis in murine macrophages (Ganesan et al., 2004), PKB/Akt is not known to be a major player in signaling pathways leading to cell activation. PLC‐g and Tec kinases are. Like PKB/Akt, PLC‐g and Tec kinases contain a PH domain which mediates or contributes to their membrane recruitment via PI(3,4,5)P3. When translocated to the membrane, Tec kinases are thought to be tyrosyl‐phosphorylated/activated by Lyn and, together with Syk, to phosphorylate PLC‐g. The mechanism by which SHIP1 can negatively regulate the activity of Tec kinases was recently documented. SHIP1, as well as SHIP2, were reported to bind preferentially to the Tec kinase itself, and to inhibit its activity. Binding occurs through the SH3 domain of Tec, and mutations of this domain generated a hyperactive form of Tec. Constitutively active Tec could also be generated by introducing mutations that targeted this kinase to the membrane. Since Tec activity is positively regulated by its membrane localization, mostly via its recruitment to PI(3,4,5)P3, it was proposed that, by hydrolyzing PI(3,4,5)P3, SHIP1/2 could prevent the membrane recruitment and, hence, the activation of Tec (Tomlinson et al., 2004). This explanation of the inhibition of Ca2þ responses observed upon coaggregation of FcgRIIB

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with immunoreceptors and the Fyn/Gab2/PI3K pathway that was described in mast cells (Parravincini et al., 2002) are not readily compatible. This Fyn‐ initiated pathway leads to the generation of PI(3,4,5)P3 by PI3K, whereas the Lyn/Syk/LAT/PLC‐g leads to Ca2þ mobilization. The mechanism of SHIP1‐ mediated FcgRIIB‐dependent inhibition of the Ca2þ response is more difficult to understand if the substrate of SHIP1 does not belong to the same pathway as that which leads to PLC‐g activation. These apparently conflicting data may be reconciled if one considers that bridges exist between the two pathways as suggested by the decreased phosphorylation of PLC‐g observed in Gab2
/
mice (Gu et al., 2001). PLC‐g is indeed recruited both by PI(3,4,5)P3 and by LAT, as well as Btk, via Gads and SLP76. PI3K is recruited both by Gab2 and, via Gads, by LAT (Schraven et al., 1999). The coaggregation of FcgRIIB with immunoreceptors markedly inhibits the phosphorylation/activation of MAP kinases. SHIP1‐dependent PI(3,4,5)P3 degradation may affect the recruitment of the exchange factor Vav, which is translocated to the membrane via its PH domain, and the subsequent generation of Rac‐GTP that leads to the activation of JNK and p38. Inhibition of Erk1/2 activation also depends on SHIP1. It, however, does not depend on the phosphatase activity of the enzyme. SHIP has a tyrosine‐rich C‐terminal segment which contains NPXY motifs. It is constitutively tyrosyl‐phosphorylated. It is further phosphorylated following immunoreceptor‐dependent cell activation, and even further when recruited by FcgRIIB. The responsible kinase is thought to be Lyn. The phosphorylation of SHIP1 does not affect its enzymatic activity, but it confers this phosphatase the properties of an adapter molecule which can affect positive signals, independently of its catalytic activity. This conclusion stemmed from the observation that the adapter molecule Dok‐1 becomes heavily phosphorylated following the coaggregation of BCR with FcgRIIB in murine B cells (Tamir et al., 2000). Dok‐1 is a member of a family of adapter proteins that are tyrosyl‐phosphorylated upon engagement of a variety of cytokine receptors, growth factor receptors, and immunoreceptors. Dok phosphorylation depends on its membrane recruitment, and membrane‐targeted Dok‐1 was constitutively phosphorylated. Dok‐1 can be phosphorylated by Lyn or by Tec. Stem Cell factor‐induced Dok‐1 phosphorylation was, however, prevented in mast cells derived from Lyn
/
mice, indicating that Lyn is primarily responsible for Dok‐1 phosphorylation in these cells (Liang et al., 2002). When tyrosyl‐phosphorylated, Dok‐1 recruits a variety of SH2 domain‐containing molecules including rasGAP which negatively regulates Ras activation. Dok‐1 contains an N‐terminal PH domain, a PTB domain, and a proline/tyrosine‐rich C‐terminal sequence. The role of Dok‐1 in FcgRIIB‐dependent negative regulation was analyzed using chimeric molecules made by replacing the intracytoplasmic domain of

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FcgRIIB by the PH and PTB domain‐containing N‐terminal half of Dok‐1 or the proline/tyrosine‐rich C‐terminal half of Dok‐1. SHIP1 coprecipitated with the N‐terminal Dok chimera, whereas rasGAP coprecipitated with the C‐terminal Dok chimera when chimeras were coaggregated with BCR (Tamir et al., 2000). Ras‐GAP contains an SH2, an SH3, another SH2, and a PH domain, followed by a catalytic domain which can enhance the auto‐catalytic activity of ras‐GTP. As a consequence, Ras‐GTP is converted into RasGDP, and the Ras pathway is extinguished. Indeed, Erk1/2 activation seen upon BCR aggregation was inhibited upon coaggregation of BCR with the C‐terminal Dok chimera, but not with the N‐terminal Dok chimera (Tamir et al., 2000). Based on these data, it was proposed that, when recruited by FcgRIIB and tyrosyl‐phosphorylated, SHIP1 recruits Dok‐1 via the PTB domain of the latter. Dok‐1 becomes tyrosyl‐phosphorylated and recruits rasGAP via the SH2 domain of the latter. rasGAP turns Ras off and prevents the activation of Erk1/2. Similar results were observed when FcgRIIB were coaggregated with FceRI in mast cells (Ott et al., 2002). Supporting this scenario, MAP kinase activation was enhanced in response to BCR aggregation, and inhibition of cell proliferation in response to the coaggregation of BCR with FcgRIIB was abolished in B cells from Dok‐1‐deficient mice (Yamanashi et al., 2000). 4.4. FcgRIIB Amplify the Autonomous Negative Regulation of Activating FcRs 4.4.1. FcgRIIB‐Dependent Negative Regulation of FceRI Signaling Does not Occur in Lipid Rafts Lipid rafts are cholesterol/glycosphygolipid‐rich membrane micro‐domains (Brown and London, 2000; Horejsi, 2003) that diffuse laterally within the plasma membrane (Pralle et al., 2000). They play a critical role in positive signaling by FceRI. Disruption of rafts, using cholesterol‐depleting drugs, dramatically decreases early phosphorylation events induced upon FceRI aggregation (Sheets et al., 1999). According to a current model, FceRI are excluded from rafts in resting mast cells, whereas signaling proteins that are covalently associated with saturated fatty acids, such as Lyn (Young et al., 2003) and LAT (Zhang et al., 1998b), are concentrated in these domains. Upon aggregation, a fraction of FceRI transiently translocate into rafts (Field et al., 1997), bringing FceRI and raft‐associated signaling proteins close to each other. Kono and coworkers reported that FcgRIIB can translocate into lipid rafts upon aggregation in RBL‐2H3 cells (Kono et al., 2002) and Aman and coworkers reported that, when coaggregated with BCRs in A20 lymphoma

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B cells, FcgRIIB recruited SHIP1 preferentially in low‐density detergent‐ resistant membrane compartments (Aman et al., 2000). We failed to observe a detectable translocation of FcgRIIB into lipid rafts, when coaggregated with FceRI. Actually the coaggregation of FcgRIIB with FceRI partially inhibited the translocation of FceRI into lipid rafts. The recruitment of SHIP1 by FcgRIIB is therefore not likely to take place in lipid rafts in mast cells. Because FcgRIIB are phosphorylated by the raft‐associated protein tyrosine kinase Lyn upon coaggregation with FceRI (Malbec et al., 1998), FcgRIIB may, however, transiently translocate into rafts where they are possibly phosphorylated. 4.4.2. FcgRIIB Associate with the Submembranous F‐Actin Skeleton When analyzing the contents of subcellular fractions prepared from RBL‐2H3 cells, we observed that FcgRIIB and SHIP1 were located in different subcellular compartments in resting cells. Following cell disruption in hypotonic buffer, differential centrifugation and solubilization of resulting fractions, most, if not all, FcgRIIB were indeed recovered in membrane fraction, whereas SHIP1 was recovered in the cytosolic and in the F‐actin skeleton fractions. The submembranous F‐actin skeleton, which connects F‐actin‐associated proteins with membrane proteins and phospholipids (Luna and Hitt, 1992), is another subcellular compartment. Unlike rafts, the submembranous F‐actin skeleton is not critical for FceRI‐dependent positive signaling. Rather, it seems to be involved in constitutive negative regulation of FceRI signaling. Indeed, drugs such as latrunculin, which prevent actin polymerization, enhance mast cell degranulation (Frigeri and Apgar, 1999). Interestingly, inhibition of degranulation observed in excess of antigen was markedly reduced in cells treated with latrunculin B, and actin could coprecipitate with SHIP1 in BMMC (Gimborn et al., 2005). Since FcgRIIB inhibit mast cell activation by recruiting SHIP1, the two molecules must meet somewhere. We found that, when coaggregated with FceRI, FcgRIIB heavily translocated into the F‐actin skeleton compartment. This translocation did not require that FcgRIIB be coaggregated with FceRI as FcgRIIB were similarly translocated upon aggregation by specific ligands. Surprisingly, it did not require either the intracytoplasmic domain of FcgRIIB as tail‐less FcgRIIB behaved similarly as intact receptors. Like FcgRIIB, FceRI were found in the membrane fraction in resting cells and, albeit in lower proportions, they dose‐dependently translocated into the F‐actin skeleton fraction when aggregated by IgE and antigen. The coaggregation with FcgRIIB did not increase but facilitated FceRI translocation which reached comparable levels at lower concentrations of antigen. Since tail‐less FcgRIIB

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could enhance the translocation of FceRI into the F‐actin skeleton fraction but failed to inhibit mast cell activation, when coaggregated with FceRI (Lesourne et al., 2005), this effect of FcgRIIB on FceRI cannot alone account for negative regulation. 4.4.3. FcgRIIB Concentrate SHIP1 Close to FceRI Signaling Complexes in the F‐Actin Skeleton Filamin 1 is an actin‐binding protein that was previously reported to associate with SHIP2 in platelets (Dyson et al., 2001, 2003). We found that SHIP1 and Filamin 1 were recovered in the same sub‐cellular fractions as SHIP1 and that SHIP1 coprecipitated with filamin 1 in unstimulated RBL‐2H3 cells. Noticeably, the high‐molecular weight isoform of SHIP1 was predominant in the F‐actin skeleton fraction and it preferentially coprecipitated with filamin 1, whereas the two main SHIP1 isoforms were equally distributed in the cytosolic fraction. SHIP2 was proposed to associate with filamin via its proline‐rich C‐terminal region that is conserved in high‐molecular weight isoforms of SHIP1, but is spliced out in low‐molecular weight isoforms. Interestingly, the high‐molecular weight isoform of SHIP1 also preferentially coprecipitated with phosphorylated FcgRIIB, following their coaggregation with FceRI. These data altogether suggested that FcgRIIB could recruit filamin‐bound SHIP1 in the submembranous F‐actin skeleton compartment. This possibility was examined in intact cells by confocal microscopy. Upon coaggregation, FcgRIIB and FceRI rapidly formed small FcR patches on the plasma membrane. Both SHIP1 and filamin 1, but not F‐actin, co‐patched with FcRs. As the size of patches enlarged with time, higher amounts of SHIP1 colocalized with FcR patches. Surprisingly, filamin 1, as well as F‐actin, were excluded from large FcR patches (Lesourne et al., 2005). Based on these data, we propose a dynamic model according to which the translocation of FcgRIIB into the cytoskeleton enables these receptors to meet filamin‐bound SHIP1. The high‐avidity cooperative interactions between SHIP1, Grb2, and FcgRIIB are likely to displace SHIP1 from filamin and to concentrate the phosphatase in FcR signaling complexes. Supporting this critical role of the cytoskeleton, FcgRIIB‐dependent negative regulation of IgE‐induced mediator release was markedly reduced in latrunculin B‐treated cells. As for the exclusion of filamin and F‐actin from large FcR patches, one may hypothesize that the increased local degradation of PI(3,4,5)P3 by SHIP1 might decrease the rate of actin polymerization. Actin is indeed constantly polymerized and depolymerized and actin polymerization depends on PI3K (Bhargavi et al., 1998). Finally, we propose that FcgRIIB negatively regulate FceRI signaling by two mechanisms. First, they facilitate the translocation of FceRI into the F‐actin skeleton

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compartment, thus enhancing SHIP1‐dependent constitutive negative regulation of FceRI at low antigen concentrations. Second, FcgRIIB concentrate SHIP1 in the vicinity of FceRI. Supporting this interpretation, SHIP1 readily coprecipitates with phosphorylated FcgRIIB but not with with FceRI. It follows that FcgRIIB act as amplifiers of SHIP1‐dependent constitutive negative regulation of FceRI signaling. 5. Conclusion FcRs are critical molecules of the immune system as they mediate most biological activities of the main effectors of the so‐called humoral immunity, that is, antibodies. Because they are ubiquitously expressed (mostly, but not only) by cells of hematopoietic origin, and because antibodies circulate in the blood stream, FcRs are involved in a wide array of biological activities in physiology. They also contribute to a variety of pathological processes. FcRs can trigger the release of potentially harmful—in some cases, life‐ threatening—inflammatory mediators, and induce destructive cytotoxic mechanisms, but (or therefore?) their activating properties are tightly controlled by regulatory mechanisms. As a consequence, immune responses are normally nonpathogenic. These regulatory mechanisms are primarily based on negative signaling that counterbalances positive signaling. Several levels of negative regulation can act on a given activating FcR. Negative regulation depends on different molecular mechanisms that may be used sequentially, depending on the conditions. A critical condition is the aggregation state of FcRs. Protein tyrosine phosphatase‐dependent negative regulation operates in resting cells when multi‐subunit FcRs are expressed on the plasma membrane and not yet engaged by any ligand (Fig. 1A). SHIP1‐ dependent negative regulation operates in mast cells whose FceRI are occupied by ‘‘monomeric’’ IgE (Fig. 1B). Unknown regulatory mechanisms account for the selective expression of some cytokine genes in mast cells exposed to IgE in the absence of antigen. Promiscuous SHP‐1‐dependent negative regulation is also triggered in cells whose FcaRI are occupied by monomeric IgA. Negative regulation involving multiple molecules that generate negative signals of different types operates as soon as positive signals are generated by activating FcRs. These include receptor subunits, kinases and phosphatases, and cytosolic and transmembrane adapter molecules. SHIP1 is a major player in the negative regulation that controls antigen‐induced IgE‐dependent mast cell activation (Fig. 1C). When further aggregated by supra‐optimal concentrations of ligand, FceRI associate with the F‐actin skeleton where the filamin 1‐bound high‐molecular weight isoform of SHIP1 resides. SHIP1 extinguishes positive signals and prevents mediator release. One, however, does not know

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which molecular interaction(s) enable its recruitment in FceRI signaling complexes (Fig. 1D). When they are co‐engaged by IgG immune complexes, FcgRIIB facilitate the association of FceRI with the F‐actin skeleton, and tyrosyl‐phosphorylated FcgRIIB recruit and concentrate high‐molecular weight SHIP1 in the signaling complex, where it dephosphorylates PI(3,4,5) P3, becomes C‐terminally tyrosyl‐phosphorylated, and recruits Dok‐1 (Fig. 1E). As consequences, both the Ca2þ response and the activation of MAP kinases are inhibited. Noticeably, negative signaling often uses molecules that are also involved in positive signaling. The ITAM‐containing FcR subunit FcRb generates positive signals that complement FcRg‐dependent signaling. It contributes to bring Lyn in the signalosome and, possibly SHIP1. Lyn phosphorylates not only FcR ITAMs and Syk, but also SHIP1, enabling this phosphatase to inhibit the Ras pathway via the sequential recruitment of Dok‐1 and rasGAP. Lyn phosphorylates also Cbp, enabling Csk to be recruited and to prevent Fyn from being activated and to lead to the activation of PI3K. Grb2 can be recruited via its SH2 domain by phosphorylated adapters such as LAT, NTAL, or Shc, in activating FcR signaling complexes and contribute to positive regulation, but also by FcgRIIB and contribute to negative regulation. It is constitutively associated, via its N‐terminal SH3 domain, with the exchange factor Sos which activates Ras, but also, via its C‐terminal SH3 domain, with SHIP1 which inhibits Ras. Grb2 can also interact, via its SH2 domain, with phosphorylated SHP‐1 which dephosphorylates signaling molecules. LAT is critical for positive TCR‐ and FcR‐dependent signaling but, as revealed by knock‐in mice expressing LAT with selective tyrosine mutations, it also contributes to generate negative signals. NTAL may function both as a LAT equivalent in B cells and as a LAT antagonist in mast cells and, in these cells, its overall dominant negative effect results from an integration of negative and positive signals. Noticeably, molecules involved in negative regulation such as SHP‐1 (Xie et al., 2000) and SHIP1 (Giallourakis et al., 2000) can also have positive effects when overexpressed. Finally, depending on the ligand valency—IgA alone or in complex with multivalent antigen—FcaRI, can either prevent or induce inflammatory responses. Altogether, data listed above lead to the conclusion that molecules have no biological functions, but biological properties only. What ultimately determines a ‘‘function’’ is the context in which a set of molecules interact in sequence with each other. This context depends on the organization of signaling complexes that transiently form and function in different subcellular compartments where different molecules reside or are translocated. As a consequence, and as learnt from the study of knock‐out mice, therapeutic approaches aiming at targeting any specific molecule can be expected to have

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Figure 1 Five levels of negative regulation in FcR complexes. Molecules in black are primarily involved in the generation of positive signals, molecules in red are primarily involved in the generation of negative signals, molecules in blue are involved in the generation of both positive and negative signals. (A) Positive and negative regulation in resting cells. Protein tyrosine kinases

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‘‘paradoxical’’ unwanted effects. An alternative is to act on the balance between positive and negative signaling in appropriate cells. Since most cells are constitutively equipped with both activating and inhibitory FcRs, these can be used as therapeutic tools. One way is to increase the expression of FcRs of one type or of the other. This is apparently what happens when intravenous immunoglobulins (IVIG) are administered and upregulate the expression of FcgRIIB (Bruhns et al., 2003). Another way is to bring more FcRs of one type into complexes of FcRs of the other type. In vitro and in vivo proofs of concepts were recently provided that one can favor negative regulation using bispecific synthetic molecules capable of co‐engaging FceRI and FcgRIIB on human mast cells and basophils, and reduce IgE‐dependent human mast cell activation (Tam et al., 2004), allergen‐induced systemic anaphylaxis, and airway hyper‐responsiveness in transgenic mice expressing human FceRI (Zhu et al., 2005). Similar approaches can be envisioned in other diseases requiring immune responses to be dampened. Conversely, other molecules can be tailored to favor positive regulation in pathological situations requiring immune responses to be boostered. For these approaches to develop and be mastered, further investigations are needed in order to understand what determines the ratio of activating and inhibitory FcRs expressed at the cell and protein tyrosine phosphatases constitutively phosphorylate and dephosphorylate, respectively, intracellular proteins. Possibly resulting activation signals do not lead to a detectable cellular reponse. (B) SHIP1 as a gatekeeper of mast cell activation. Positive signals triggered by IgE in the absence of antigen are constitutively negatively regulated by SHIP1. As a result, wt mast cells usually do not degranulate when sensitized with IgE and are not challenged with antigen, but SHIP1‐deficient mast cells do. (C) Negative signals generated together with positive signals by activating FcRs. Upon aggregation of activating FcRs by antibodies and multivalent antigens, both the Lyn/Syk/PLC‐g and the Fyn/Gab2/PI3K pathways are activated, leading to cell activation. These positive signals are counterbalanced by negative signals. By phosphorylating Cpb, Lyn enables Csk to be recruited and to inhibit Fyn. By phosphorylating SHIP1, Lyn enables Dok1 to be recruited and to inhibit Ras via rasGAP. SHIP1 is possibly recruited by phosphorylated FcRb. NTAL also negatively regulates FceRI signaling by not yet clear mechanisms. Biological responses of the cell results from the integration of these antagonistic signals. (D) Negative signals generated by activating FcRs in excess of ligand. When supra‐optimally engaged by an excess of ligand, FceRI aggregates associate with the F‐actin skeleton, where the high molecular isoform of SHIP1 is constitutively associated with Filamin 1. As a consequence, more SHIP1 is involved in negative regulation as indicated by its increased phosphorylation. The result is a dose‐dependent inhibition of degranulation. (E) Negative regulation by FcgRIIB. When coaggregated with FceRI, FcgRIIB are phosphorylated by Lyn, associate with the F‐actin skeleton, and recruit F‐actin‐associated SHIP1. The recruitment of SHIP1 involves the interactions of its SH2 domain with specific residues in the FcgRIIB phosphorylated ITIM and of its C‐terminal prolin‐rich region with the C‐terminal SH3 domain of Grb2 which, itself, binds to the phosphorylated C‐terminal tyrosine of FcgRIIB via its SH2 domain. FcgRIIB thus concentrate SHIP1 in FceRI signaling complex and, by inhibiting both the Ca2þ response and the activation of MAP Kinases, extinguish all cellular responses.

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surface, whether activating or inhibitory FcRs can be preferentially engaged by antibodies, how FcRs generate positive and negative signals, and how these signals are integrated within cells. Acknowledgments Our works discussed in this review were supported in part by the Institut National de la Sante´ et de la Recherche Me´dicale (INSERM), the Fondation pour la Recherche Me´dicale (FRM), the Association pour la Recherche sur le Cancer (ARC), and the Institut Pasteur. RL was the recipient of fellowships from the ARC and from the Socie´te´ Franc¸aise d’Allergologie et d’Immunologie Clinique (SFAIC).

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The Surprising Diversity of Lipid Antigens for CD1‐Restricted T Cells D. Branch Moody Division of Rheumatology, Immunology, and Allergy, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts

1. 2. 3. 4. 5. 6. 7. 8.

Abstract............................................................................................................. Introduction: From Molecules to Functions ............................................................. CD1 Protein Expression on Antigen‐Presenting Cells ................................................ Subcellular Lipid Antigen Processing Pathways......................................................... 3‐Dimensional Structures of CD1‐b2‐Microglobulin‐Lipid Complexes .......................... Microbial Antigens and Infectious Disease ............................................................... Self Antigens, Autoreactivity, and Autoimmune Disease.............................................. Synthetic Lipid Antigens and Prospects for Immunotherapy........................................ Conclusion: Prospects for Immunotherapy ............................................................... References .........................................................................................................

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Abstract CD1 proteins have been conserved throughout mammalian evolution and function to present lipid antigens to T cells. Crystal structures of CD1‐lipid complexes show that CD1 antigen‐binding grooves are composed of four pockets and two antigen entry portals. This structural information now provides a detailed understanding of how CD1‐binding grooves capture a surprisingly diverse array of lipid ligands. CD1‐expressing APCs are able to acquire lipid antigens from their own pool of lipids and from exogenous sources, including microbial pathogens, bystander cells, or even the systemic circulation. CD1 proteins bind to certain antigens using high stringency loading reactions within endosomes that involve low pH, glycosidases, and lipid transfer proteins. Other antigens can directly load onto CD1 proteins using low stringency mechanisms that are independent of cellular factors. New evidence from in vivo systems shows that CD1‐restricted T cells influence outcomes in infectious, autoimmune, and allergic diseases. These studies lead to a broader view of the natural function of ab T cells, which involves recognition of both cellular proteins and lipids. 1. Introduction: From Molecules to Functions The discovery of the CD1 antigen presentation system represents an advance in immunology because it shows that T cells scan and respond to changes in the lipid content of target cells. The central questions in the study of

87 advances in immunology, vol. 89 # 2006 Elsevier Inc. All rights reserved.

0065-2776/06 $35.00 DOI: 10.1016/S0065-2776(05)89003-0

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CD1‐restricted T cells have unfolded in a very different way from those that led to the discovery of the major histocompatibility complex (MHC). The study of the MHC started by establishing the role of T cells in controlling viral infection and rejecting transplanted tissues (McDevitt and Benacerraf, 1969; Zinkernagel and Doherty, 1974) and represented a three‐decade‐long search for the molecules that mediate T cell activation (Garboczi et al., 1996; Garcia et al., 1996; Kappler et al., 1983; McIntyre and Allison, 1983; Oldstone et al., 1988). In contrast, detailed molecular information about CD1 proteins and CD1 genes was available for many years prior to the first evidence that CD1 proteins control the activation of T cells. McMichael and Milstein discovered CD1 proteins using the monoclonal antibody, NA1/34, which bound a protein on human thymocytes with an apparent molecular weight of 45 kilodaltons. The ligand bound by this antibody showed some biochemical similarities to the MHC class I protein, but could be distinguished by its lack of recognition by an MHC class I‐specific antibody, W6/32 (McMichael et al., 1979). Initially known as human thymocyte antigen‐1 (HTA‐1), this protein was renamed as the first cluster of differentiation antigen (CD1) because its discovery represented the first use of monoclonal antibodies to define a specific cell surface marker on human lymphocytes. Cellular studies showed that the CD1 heavy chain associates with b‐2 microglobulin to form heterodimers (Cotner et al., 1981; Terhorst et al., 1981). Subsequently, the five CD1 genes (CD1A, CD1B, CD1C, CD1D, CD1E) were cloned from human thymocytes and mapped to chromosome 1 (Calabi and Milstein, 1986). The first studies of CD1 function showed that human ab and gd T cell clones were directly reactive with CD1 in the sense that their activation could be blocked with antibodies against CD1 proteins, but the activation did not require an exogenous antigen (Porcelli et al., 1989). Shortly thereafter, it was found that antigens from microbial pathogens could be taken up into endosomes for processing reactions within human dendritic cells and that these microbial antigens were lipidic in nature (Beckman et al., 1994; Porcelli et al., 1992). Twenty‐five years after the discovery of CD1 proteins, current work in the field seeks to understand the precise functions of CD1‐ restricted T cells in influencing infectious, autoimmune, allergic, vascular, and neoplastic diseases. In considering the natural functions of CD1‐restricted T cells in immune response, two populations within the CD1‐restricted T cell repertoire have been recognized and categorized, based on whether or not they express conserved or diverse TCRs (Fig. 1). In general, CD1‐restricted T cells are capable of expressing diverse TCR a and b chains and recognize structurally diverse classes of lipid antigens. Within the larger CD1‐restricted T cell repertoire, NK T cells are an abundant and functionally important subset

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Figure 1 The larger repertoire of diverse CD1‐restricted T cells is composed of clones expressing TCRs with diverse structural elements, including differing TCR a‐ and b‐variable gene segments with varied N‐region additions. NK T cells are a large subpopulation of CD1‐restricted T cells that recognize CD1d and express conserved TCRs with an invariant TCR a chain and limited diversity of TCR Vb genes.

that expresses nearly invariant TCRs composed of a canonical a chain (Va14Ja18 in mice and Va24Ja18 in humans) paired with a limited number of b‐chains. The discovery of NK T cells involved the identification of T cells that lack CD4 and CD8 coreceptors, express Vb8‐containing TCRs and could be identified by staining C‐type lectins that are also found on NK cells such as NK1.1 (CD161) (Budd et al., 1987; Fowlkes et al., 1987). Separately, knockout of MHC class II proteins revealed a set of CD4þ T cells that express TCR a chains with limited diversity (Bendelac et al., 1994; Cosgrove et al., 1991). The molecular targets of recognition of NK T cells were solved in two key studies, which showed that CD1d expression on APCs was necessary for their activation (Bendelac, 1995) and that the CD1d‐mediated activation was greatly enhanced by treating APCs with synthetic glycolipids related in structure to a‐galactosyl ceramides (Kawano et al., 1997). The use of the term NK T cell to refer to CD1d‐restricted T cells with invariant TCRs persists for these historical reasons and because NK cells and NK T cells have some limited functional similarities. For example, both NK cells and NK T cells use T‐bet transcription factors (Townsend et al., 2004), expand in the presence of IL‐15 (Matsuda et al., 2002), and have at least some reliance on NK1.1 for activation (Exley et al., 1998). However, the primary activation signals for NK T cells involve TCRs and CD1d, not NK complex‐ encoded receptors. Further, many conventional T cells express NK1.1, and many CD1d‐restricted T cells with invariant TCR a chains do not express NK1.1 (Gumperz et al., 2002). The relative lack of sensitivity and specificity of NK1.1 and other NK complex‐encoded proteins for distinguishing NK T

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cells from MHC‐restricted T cells has led to the waning use of these markers as the primary defining features of NK T cells. Instead, NK T cells are identified by the presence of invariant TCR a chains and activation by CD1d in combination with glycolipid antigens (Fig. 1). This review of CD1‐restricted T cells follows the path of evolution of CD1 research—from molecular structures to in vivo studies—with emphasis on the newest advances in each area.

2. CD1 Protein Expression on Antigen‐Presenting Cells 2.1. Three Groups of CD1 Proteins The human CD1 locus encodes five genes: CD1A, CD1B, CD1C, CD1D, and CD1E (Calabi and Milstein, 1986). Prior to accumulation of any knowledge of the functions of CD1 proteins, a grouping system was proposed based on levels of amino acid sequence homology among the five members of the human CD1 family (Calabi et al., 1989). CD1a, CD1b, and CD1c were most homologous to one another and were designated as the group 1 isoforms. CD1d demonstrated the lowest overall homology to other CD1 proteins and was designated as the group 2 isoform. CD1e was similarly homologous to both groups, so it was not originally classified, but is now known as the group 3 isoform. Although this classification system was originally devised based on amino acid sequence, it remains useful because more detailed studies of gene regulation and CD1 function have generally, but not universally, found differing functions for group 1, group 2, and group 3 CD1 proteins. For example, each of the group 1 isoforms (CD1a, CD1b, CD1c) bind and present microbial lipid antigens to diverse CD1‐restricted T cells, whereas the group 2 isoform (CD1d) is more clearly associated with a function in activating invariant NK T cells. Because NK T cells generally function to regulate other cells, whereas diverse T cells are thought to have direct effector functions against cells with altered lipids, these differences in antigen profiles may also translate into differing functions in vivo. There are, however, exceptions to these apparently differing immunoregulatory and effector functions of group 1 and 2 CD1 proteins, as NK T cells have been recently found to directly recognize microbial lipids (Kinjo et al., 2005; Mattner et al., 2005), and T cells that are directly reactive to group 1 CD1 proteins regulate DC maturation (Vincent et al., 2003). The function of the group 3 isoform, CD1e, is distinct because it is not detected at the surface of APCs and does not function in antigen display (Angenieux et al., 2000, 2005). Instead, it functions in the processing and transfer of lipids to other CD proteins (de la Salle et al., 2005).

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2.2. Evolutionary Conservation of CD1 CD1 proteins are expressed in every mammalian species that has been examined to date, including monkeys, horses, sheep, cows, rabbits, guinea pigs, rats, and mice. Because the human system was the first to be described, CD1 nomenclature designates non‐human sequences according to the human ortholog with which they share the highest level of overall sequence homology. When a species expresses more than one ortholog of a given human gene, each protein is given an arbitrary numerical designation. For example, guinea pigs express three CD1c orthologs (CD1c1, CD1c2, CD1c3), and rabbits express two orthologs of CD1a (CD1a1 and CD1a2) (Dascher et al., 1999; Hayes and Knight, 2001). This nomenclature reflects the fact that most mammalian species have larger numbers of CD1 genes, with mice and rats being the notable exceptions. Recent advances point to new insights in the evolutionary origins of the CD1 system. MHC class I and II molecules are conserved in bony and cartilaginous fish but are lacking in jawless fish. Based on fossil evidence that dates this split, MHC antigen‐presenting molecules are thought to have arisen approximately 500 million years ago, contemporaneously with recombinases that allowed production of rearranged TCRs, forming the basis of the acquired immune system (Laird et al., 2000). Sequence analyses of MHC I, MHC II, and CD1 genes show similar levels of relatedness (despite differences in domain organization), and it has been argued that these three families of antigen‐presenting molecules are derived from a single ancestral gene (Hughes, 1991; Porcelli, 1995). However, there has been controversy as to whether the CD1 system is an ancient branch of the adaptive immune system or evolved from MHC class I proteins under selective pressure over a relatively short span. For example, computer‐based homology analyses had predicted that CD1 may have diverged early in vertebrate evolution, whereas others suggested that divergence might have occurred later, perhaps even after the bird–mammal (synapsid–diapsid) split, approximately 300 million years ago (Hughes, 1991; Porcelli, 1995). Direct evidence that ancestral CD1 genes predate the bird–mammal divergence in the form of CD1 gene sequences in early vertebrates was lacking until recently. Two studies have independently isolated avian CD1 orthologs in chickens (Gallus gallus), which have sequence or structural homologies with mammalian CD1 genes (Miller et al., 2005; Salomonsen et al., 2005). Although the ability of these avian CD1 gene products to present antigens has not been analyzed, these chicken CD1 genes are predicted to encode proteins with requisitely high amino acid homology with mammalian CD1 proteins. Also, chicken CD1 proteins have hydrophobic amino acids in the predicted

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antigen‐binding domains. Importantly, both studies found that chicken CD1 genes are tightly linked with the MHC complex, providing the first example of direct linkage between CD1 and MHC genes. These studies indicate that the CD1 system predates evolution of mammalian species and more directly support prior arguments that CD1 and MHC genes diverged from a common ancestral gene. 2.3. Constitutive and Inducible Expression of CD1 on APCs With the exception of epithelial cells that line the gastrointestinal tract (Somnay‐Wadgaonkar et al., 1999; van de Wal et al., 2003), the expression of CD1 proteins is generally limited to hemopoietically derived cells with specialized functions in antigen presentation and immune response. All five human CD1 proteins are expressed on thymocytes at high levels (Calabi and Milstein, 1986). Most evidence suggests that thymocytes downregulate CD1 proteins prior to their exit to the periphery. However, there is some evidence for expression of CD1a and CD1c proteins on a small subset of T cells in the peripheral blood (Salamone et al., 2001a,b). B cells represent another effector population that constitutively expresses CD1 proteins at high levels. Although CD1c and CD1d are found on a minority of peripheral blood B cells, many studies of secondary lymphoid tissues show that these two isoforms are expressed on large numbers of B cells that localize to the interface of B cell‐ and T cell‐rich areas of secondary lymphoid tissues and co‐express markers of marginal zone B cells. In fact, formal analysis of patterns of CD1c and CD1d expression have found high correlation with localization to the marginal zone, such that CD1c and CD1d can be considered relatively specific markers of marginal zone B cells (Weller et al., 2004). Marginal zone B cells are positioned so that they can rapidly sample blood‐borne antigens, and new evidence indicates that the expression of CD1d at this site regulates Ig production in a way that can alter infection. For example, marginal zone B cells from CD1d‐ deficient mice produce lower levels of borrelia‐specific IgM than wild‐type mice, and passive transfer of IgM partially rescues the worsened infection seen in CD1d‐deficient mice infected by Borrelia burgdorferii and Borellia hermsii (Belperron et al., 2005; Kumar et al., 2000). In contrast to the apparently constitutive expression of CD1 on thymocytes, B cells, and epithelia, the patterns of expression of individual CD1 isoforms on myeloid cells are complex and change in response to cellular activation. CD1d proteins are expressed at low but consistently detectable levels on monocytes. In contrast, fresh human monocytes from peripheral blood lack expression of CD1a, CD1b, and CD1c, but can be induced to express these group 1 isoforms at high levels after treatment with GM‐CSF, IL‐4, and other factors that

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promote DC maturation (Porcelli et al., 1992; Sallusto and Lanzavecchia, 1994). Indeed, group 1 CD1 protein expression is increasingly used as a lineage marker that allows myeloid DCs to be distinguished from activated macrophages. However, modulation of CD1 expression on DCs has only recently come under study as a possible physiological means of regulating the functions of CD1‐restricted T cells, leading to the identification of molecular signals that turn off or on CD1 antigen presentation in DCs. One study has shown that infection of CD1‐expressing dendritic cells by Mycobacterium tuberculosis, a pathogen that makes several classes of CD1‐presented lipids, resulted in the complete loss of detectable CD1b proteins over a period of several days. Such downregulation of CD1 expression in vitro led to the speculation that this could represent a mechanism of immune evasion in vivo (Stenger et al., 1998b). However, similar studies of in vitro‐derived dendritic cells have found only partial or no downregulation of CD1 proteins in response to mycobacterial infection (Giuliani et al., 2001; Henderson et al., 1997). Also, studies testing the effect of Mycobacterium tuberculosis infection of fresh monocytes have found upregulation of CD1a, CD1b, and CD1c after infection. The mechanism of CD1 upregulation in response to mycobacteria involves lipid agonists of Toll‐like receptor 2 (TLR‐2), such as lipoarabinomannan, which promotes transcription and translation of group 1 CD1 proteins (Roura‐Mir et al., 2005b). A similar process likely occurs in vivo, as Mycobacterium leprae infection increases group 1 CD1 protein expression on DCs within the skin of patients with tuberculoid leprosy (Krutzik et al., 2005; Sieling et al., 1999). Also, group 1 CD1 proteins are expressed in peribronchial tissues and brochioalveolar lavage samples of tuberculosis patients (Buettner et al., 2005; Uehira et al., 2002). Thus, the dominant effect of mycobacteria is to generate a local inflammatory reaction that leads to increased expression of group 1 CD1 proteins on myeloid cells at sites of infection. New insights into the differential timing of expression of CD1a, CD1b, CD1c, CD1d, and CD1e on maturing myeloid DCs point to potentially distinct biological functions of each of the three groups of CD1 proteins. Kinetic measurements of group 1 CD1 induction at the cell surface in response to cytokines, TLR agonists, or mycobacterial lipids show that cell surface expression is first detected 2 days after activation, peaking at day 3. This delayed expression of group 1 CD1 proteins relates to the cellular mechanism of group 1 CD1 induction, which requires synthesis of new proteins, rather than redirected trafficking of pre‐formed proteins, as is the case with MHC class II (Roura‐Mir et al., 2005b). Certain stimuli that promote group 1 CD1 expression lead to the concurrent downregulation of CD1d and vice versa. For example, the upregulation of CD1d expression by agonists of

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the peroxisome proliferator activated receptor g (PPAR‐g) pathway is accompanied by decreased expression of CD1a (Szatmari et al., 2004), and mycobacterial cell wall products that increase group 1 CD1 expression decrease CD1d expression. The differing timing and opposite responses of group 1 and group 2 CD1 proteins imply distinct functions. Rapid activation of NK T cells over minutes to hours by constitutively expressed CD1d proteins is one criterion supporting the description of NK T cells as ‘‘lymphocytes of innate immunity’’ (Benlagha and Bendelac, 2000). In contrast, the delayed expression of group 1 CD1 proteins on DCs, as well as their regulation by antecedent activation of pattern recognition receptors of the innate immune system, are more in keeping with a slightly delayed pathogen recognition system that becomes effective during the transition from innate to adaptive immune responses. These new insights derived from in vitro studies of the agonists, receptors, and kinetics of expression of CD1 now form a rationale for further investigation of the dynamic patterns of expression of CD1 proteins in vivo. Given the clear evidence for cell‐type and activation‐dependent patterns of expression of CD1 proteins, there currently exists remarkably little information regarding signaling molecules, promoter elements, or transcriptional mechanisms that control CD1 expressions, at the surface of myeloid APCs. Agonists of TLRs and PPAR‐g both signal through nuclear factor kappa B (NFkB), implicating this signaling pathway in the control of CD1 expression. There has been some preliminary mapping of CD1 promoters, and new evidence shows that E26 transformation‐specific (ets) transcription factors control CD1d expression (Calabi et al., 1989; Geng et al., 2005). 3. Subcellular Lipid Antigen Processing Pathways New studies indicate that the loading of lipid antigens onto cellular CD1 proteins is regulated by vesicular ATPases, cell surface lectins, lipid transfer proteins, and other cellular cofactors with distinct patterns of expression within subcellular compartments of APCs. These studies are beginning to coalesce into models of coordinated subcellular pathways of lipid antigen processing. Whereas protein antigen processing centrally involves cleaving antigens to generate smaller peptides, lipid antigen processing is coming to be understood as a multi‐stage transfer process for moving relatively insoluble antigens through aqueous solutions for loading into the hydrophobic grooves of CD1 proteins. Viewed from this perspective, the trafficking of CD1 proteins through the secretory, cell surface, and endosomal pathways is not merely a means to disperse CD1 proteins throughout diverse lumenal compartments of APCs. Instead, the complex subcellular trafficking patterns offer a means to

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regulate the loading of differing subclasses of antigens into CD1 proteins and thereby influence the repertoire of lipids that are ultimately displayed to T cells. In particular, the routing of CD1 trafficking through late endosomes and lysosomes renders CD1 proteins able to capture antigens that cannot be captured within other cellular subcompartments. 3.1. CD1 Translation and Egress to the Cell Surface Leader sequences in the CD1 genes result in cotranslational insertion into the endoplasmic reticulum (ER), so that the a1, a2, and a3 portions of the heavy chain are inserted in the lumen, leaving 6 or more amino acids extending into the cytoplasm (cytoplasmic tails). The folding of CD1 heavy chains is regulated by calnexin and calreticulin and appears to be followed by rapid association with b‐2 microglobulin in the ER (Bauer et al., 1997; Huttinger et al., 1999; Kang and Cresswell, 2002a; Sugita and Brenner, 1994). Whether or not ER‐resident lipids associate with CD1 proteins during the initial folding process has not been directly assessed in cells, but there is indirect evidence that lipids promote CD1 folding. For example, addition of lipids to unfolded recombinant