Progress in Molecular Biology and Translational Science [1 ed.] 0123747600, 9780123747600

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PROGRESS IN

Molecular Biology and Translational Science Volume 87

PROGRESS IN

Molecular Biology and Translational Science Regulatory Mechanisms in Transcriptional Signaling edited by

Debabrata Chakravarti Division of Reproductive Biology Research Department of Obstetrics and Gynecology Robert H. Lurie Comprehensive Cancer Center Feinberg School of Medicine, Northwestern University Chicago, Illinois 60611, USA

Volume 87 AMSTERDAM • BOSTON • HEIDELBERG • LONDON NEW YORK • OXFORD • PARIS • SAN DIEGO SAN FRANCISCO • SINGAPORE • SYDNEY • TOKYO Academic Press is an imprint of Elsevier

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Contents

Contributors....................................................................................... Preface.............................................................................................. Introduction .......................................................................................

ix xi xv

Regulation of Metabolism by Nuclear Hormone Receptors . . . . . . . . . . . . . . . . . . . . . . . . . .

1

Huey-Jing Huang and Ira G. Schulman I. II. III. IV. V. VI. VII.

Introduction ............................................................................... The PPARs................................................................................. LXR.......................................................................................... FXR.......................................................................................... ROR ....................................................................................... ERR ....................................................................................... Summary ................................................................................... References .................................................................................

2 4 12 19 26 31 37 37

Progesterone Receptor Action in Leiomyoma and Endometrial Cancer . . . . . . . . . . . . . . . . . . . . . . . . . .

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J. Julie Kim, Elizabeth C. Sefton, and Serdar E. Bulun I. II. III. IV. V. VI. VII. VIII. IX. X.

Introduction ............................................................................... The Uterus................................................................................. Progesterone Action on the Endometrium and Myometrium............... Endometrial Cancer .................................................................... Progesterone Receptor Action in Endometrial Cancer ....................... Conclusions and Perspectives of Progesterone Action in Endometrial Cancer .................................................................... Uterine Leiomyoma ..................................................................... Progesterone Receptor Action in Leiomyoma ................................... Conclusions and Perspectives on Progesterone Action in Uterine Leiomyoma ..................................................................... Future Directions........................................................................ References .................................................................................

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54 54 56 61 62 66 66 67 73 74 74

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Nuclear Xenobiotic Receptors: Integrating Gene Regulation to Physiological Functions . . . . . . . . . . . . . .

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Jinhan He and Wen Xie I. II. III. IV.

Introduction .............................................................................. Ligands for Nuclear Receptors ...................................................... Nuclear Receptor Domain Structures............................................. Xenobiotic Receptor Functions and Their Implications in Physiology and Diseases, A Case Study ........................................... V. Pregnane X Receptor (PXR) ......................................................... VI. Constitutive Androstane Receptor (CAR) ........................................ VII. Concluding Remarks ................................................................... References ................................................................................

88 88 89 92 93 104 109 109

Emerging Roles of the Ubiquitin Proteasome System in Nuclear Hormone Receptor Signaling . . . . . . . . . . . . . 117 David M. Lonard and Bert W. O’Malley I. Introduction: Nuclear Hormone Receptors, Ubiquitin, and the Proteasome..................................................................... II. The Ubiquitin Proteasome System ................................................. III. Nuclear Receptor Interactions with the Ubiquitin Proteasome System ..................................................................... IV. Coregulators and the UPS ............................................................ V. Coregulators as UPS Targets......................................................... VI. Ubiquitin-Like Modifications in Nuclear Receptor Signaling .............. VII. Conclusion and Perspective .......................................................... References ................................................................................

118 118 120 123 125 128 129 129

Biochemical Analyses of Nuclear Receptor-Dependent Transcription with Chromatin Templates . . . . . . . . . . . . 137 Donald D. Ruhl and W. Lee Kraus I. Nuclear Receptors (NRs): Transcription Factors (TFs) with Separable Biochemical Activities.................................................................. II. Biochemical Analyses of NR Activities, Interactions, and Functions: An Historical View ...................................................................... III. Role of Chromatin in NR-Dependent Transcription .......................... IV. Biochemical Methods for the Analysis of NR-Dependent Transcription V. Biochemical Methods for the Assembly and Analysis of Chromatin...... VI. What Have We Learned About NR-Dependent Transcription from In Vitro Chromatin Assembly and Transcription Studies?...................

138 139 143 145 149 157

contents VII. Future Directions........................................................................ VIII. Summary ................................................................................... References .................................................................................

vii 174 178 178

Chromatin Remodeling and Nuclear Receptor Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Manop Buranapramest and Debabrata Chakravarti I. II. III. IV. V. VI.

Introduction ............................................................................... NR Classification and Structure ..................................................... NR Coregulators ......................................................................... Chromatin as an NR Coregulator Substrate ..................................... ATP-Dependent Chromatin Remodelers in NR Gene Regulation......... Future Directions........................................................................ References .................................................................................

194 194 197 201 212 222 225

Nuclear Receptor Repression: Regulatory Mechanisms and Physiological Implications . . . . . . . . . 235 M. David Stewart and Jiemin Wong I. II. III. IV. V. VI.

Introduction ............................................................................... Corepressors .............................................................................. Types of NR Repression ............................................................... Molecular Mechanisms of Transcriptional Repression ........................ Physiological Functions of NR-Mediated Repression ......................... Concluding Remarks.................................................................... References .................................................................................

236 238 241 244 248 252 253

The Roles and Action Mechanisms of p160/SRC Coactivators and the ANCCA Coregulator in Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Elaine Y.C. Hsia, June X. Zou, and Hong-Wu Chen I. Introduction: The Discovery of AIB1/ACTR/SRC-3 as a Nuclear Hormone Receptor Coactivator and a Gene Amplified in Cancer......... II. Aberrant Genetic Regulation of p160/SRC Expression in Cancers........ III. The p160/SRCs Functions and Their Action Mechanisms in Cancer Cells ........................................................................... IV. Functions of p160/SRCs in Tumorigenesis Revealed in Animal Models ........................................................................

262 264 275 286

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V. The Coregulator ANCCA, a Unique Target of AIB1/ACTR and a Potential Key Player in Cancer ............................................. VI. Concluding Remarks ................................................................... References ................................................................................

288 290 291

Protein Arginine Methyltransferases: Nuclear Receptor Coregulators and Beyond . . . . . . . . . . . . . . . . . . . . . . 299 Peter Kuhn and Wei Xu I. II. III. IV. V. VI. VII. VIII.

Introduction .............................................................................. Enzymatic Activity of PRMTs ....................................................... PRMTs in Transcriptional Regulation ............................................. PRMTs in Posttranscriptional Regulation ........................................ Structural Analysis of PRMTs........................................................ Small Molecule Inhibitors for PRMTs ............................................ Biological Functions of PRMTs ..................................................... Concluding Remarks ................................................................... References ................................................................................

300 301 307 318 319 323 324 332 332

Roles of Histone H3-Lysine 4 Methyltransferase Complexes in NR-Mediated Gene Transcription . . . . . . 343 Seunghee Lee, Robert G. Roeder, and Jae W. Lee I. II. III. IV. V. VI. VII.

Introduction .............................................................................. Activating Signal Cointegrator-2 (ASC-2) ........................................ Set1-Like H3K4MT Complexes..................................................... ASCOM in NR-Mediated Transactivation........................................ Cross talk of ASCOMs with Other Coactivators................................ Physiological Roles of Key Subunits of ASCOM ............................... Future Challenges ...................................................................... References ................................................................................

344 346 351 354 357 362 372 374

Index........................................................................................

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Contributors

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

Serdar E. Bulun, Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Chicago, Illinois 60611 (53) Manop Buranapramest, Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611 (193) Debabrata Chakravarti, Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611 (xv, 193) Hong‐Wu Chen, Department of Biochemistry and Molecular Medicine; and UC Davis Cancer Center/Basic Sciences, University of California at Davis, Sacramento, California 95817 (261) Ronald M. Evans, The Salk Institute for Biological Studies, La Jolla, California 92037 (xi) Jinhan He, Center for Pharmacogenetics and Department of Pharmaceutical Sciences, University of Pittsburgh, Pittsburgh, Pennsylvania 15216 (87) Elaine Y.C. Hsia, Department of Biochemistry and Molecular Medicine; and Department of Internal Medicine, School of Medicine, University of California at Davis, Sacramento, California 95817 (261) Huey‐Jing Huang, Department of Biology, Exelixis Inc., 4757 Nexus Centre Drive, San Diego, California 92121 (1) J. Julie Kim, Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Chicago, Illinois 60611 (53) W. Lee Kraus, Department of Molecular Biology and Genetics, Cornell University, Ithaca, New York 14853; and Department of Pharmacology, Weill Medical College of Cornell University, New York, New York 10021 (137) Peter Kuhn, McArdle Laboratory for Cancer Research, University of Wisconsin, Madison, Wisconsin 53706 (299) Seunghee Lee, Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas 77030 (343) Jae W. Lee, Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas 77030 (343) ix

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David M. Lonard, Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, Texas 77030 (117) Bert W. O’Malley, Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, Texas 77030 (117) Robert G. Roeder, Laboratory of Biochemistry and Molecular Biology, The Rockefeller University, New York 10021 (343) Donald D. Ruhl, Department of Molecular Biology and Genetics, Cornell University, Ithaca, New York 14853 (137) Ira G. Schulman, Center for Molecular Design and Department of Pharmacology, University of Virginia Health System, Charlottesville, Virginia 22908 (1) Elizabeth C. Sefton, Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Chicago, Illinois 60611 (53) M. David Stewart, Department of Genetics, University of Texas M.D. Anderson Cancer Center, Houston, Texas 77030 (235) Jiemin Wong, Institute of Biomedical Sciences and School of Life Sciences, East China Normal University, Shanghai 200241, China (235) Wen Xie, Center for Pharmacogenetics and Department of Pharmaceutical Sciences; and Department of Pharmacology and Chemical Biology, University of Pittsburgh, Pittsburgh, Pennsylvania 15216 (87) Wei Xu, McArdle Laboratory for Cancer Research, University of Wisconsin, Madison, Wisconsin 53706 (299) June X. Zou, Department of Internal Medicine, School of Medicine; and UC Davis Cancer Center/Basic Sciences, University of California at Davis, Sacramento, California 95817 (261)

Preface Nuclear Receptors: Past, Present, and Future

One of the major challenges in the postgenomic era is to understand the fundamental function and interplay of genes that build and maintain the organism. The availability of the complete human genome sequence, with the advent of bioinformatics tools and array technologies, has greatly accelerated this process. However, DNA sequence is inherently static and some have described the information as little more than a genomic ‘‘parts list.’’ What is more relevant is knowing how groups or networks of genes are coordinately expressed to produce unique cell function and physiology. It is important to identify the functional interactions between genes, the connections within networks, and most importantly, the regulatory code, molecules, and mechanisms that direct this complex process. Historically, the isolation and crystal structure of thyroid hormone by Kendall and Reichstein in 1914 and 1920, respectively, kicked off the first revolution in the field. This was soon followed by the isolation of vitamins A and D, bile acids, and the remaining steroid hormones between 1928 and 1952. However, how these bioactive lipids worked was clouded in mystery and the notion that they may all have a common mechanistic underpinning was never even suggested. Thus, the isolation of the glucocorticoid receptor cDNA in 1985 was critical, providing the first complete sequence of a nuclear receptor, a sequence that would prove to be a prototype of all subsequent family members.5 This helped to trigger the next revolution in our understanding by providing a commonality of signaling by bioactive nuclear hormonal lipids. We now know that nuclear hormone receptors (NRs) comprise a large family of ligand‐modulated transcription factors (TFs) that mediate responses to a wide range of lipophilic signaling molecules including lipids, steroids, retinoids, hormones, and xenobiotics.2,4,8 As sensors for these signals they provide an important link between transcriptional regulation and physiology. The NRs constitute one of the largest groups of TFs in animals (48 genes in humans, 49 in mice) and includes classic endocrine receptors that mediate the actions of the steroid hormones, thyroid hormones, and the fat‐soluble vitamins A and D mentioned earlier, as well as a large number of so‐called orphan xi

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nuclear receptors, whose ligands, target genes, and physiological functions are still largely unknown. Thanks to recent intensive research, investigation of these orphans has revealed functions that are extremely interesting, particularly as it relates to normal physiology and pathologic states such as type 2 diabetes, heart disease, and cancer. A series of key transformative advances were the isolation and characterization of the first nuclear receptor corepressors (SMRT and NCoR) and the coactivator (SRC‐1),3,7,9 respectively. This was quickly followed by the discovery that the histone acetyltransferase ‘‘CBP’’ was a nuclear receptor coactivator providing the first clear link between epigenetic modifications and hormone signaling.1,6 These were the first critical steps in deconstructing the molecular logic as to how receptors interface with the chromatin template to regulate the transcriptional process itself. Eventually, more than 300 different NR‐associated proteins have been isolated. About one third contains the prototypic ‘‘LXXLL’’ motif that suggests the potential for direct interaction with one or more NRs. Many of these proteins were identified as parts of cofactor complexes that often include between 8 and 20 proteins. If we add up the number of proteins involved in NR‐directed activation, it would most likely fall between 50 and 100 although it could easily exceed these numbers in specialized situations. Furthermore, an equal number of proteins are most likely to be involved in target gene repression. In this odd way, while the hormone itself is a relatively simple structure, it becomes much easier to see how it achieves its regulatory complexity by employing standard association/dissociation kinetics to invoke dynamic allostery upon the cognate NR, resulting in the equally dynamic recruitment and dismissal of multiple coregulator complexes. These key advances in turn lead us into our epigenomic future where receptor meets chromatin. The human and mouse genome projects opened up a great door but , as mentioned above, such projects provide a genetic parts list that is devoid of the instruction book. An understanding as to how the parts are called into action to create tissues, organs or complex physiology and disease is the next critical step in the field. The sequence helps to create a map of the chromosomes but receptors will provide a powerful tool to understand the process as to how the chromosomes are controlled. Why is this important? While there is only one genome in a person, there will be many ‘‘epigenomes’’ and this is where an understanding of complex physiologic processes will begin to emerge. It is the goal of this book to highlight some of the newer breakthroughs, techniques, and challenges to position us for an exciting future. RONALD M. EVANS The Salk Institute, La Jolla, CA

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1. Chakravarti D, LaMorte VJ, Nelson MC, Nakajima T, Schulman IG, Juguilon H, et al. Role of CBP/P300 in nuclear receptor signaling. Nature 1996;383(6595):99–103. 2. Chawla A, Repa JJ, Evans RM, Mangelsdorf DJ. Nuclear receptors and lipid physiology: opening the X‐files. Science 2001;294(5548):1866–70. 3. Chen JD, Evans RM. A transcriptional co‐repressor that interacts with nuclear hormone receptors. Nature 1995;377(6548):454–7. 4. Evans RM. The steroid and thyroid hormone receptor superfamily. Science 1988;240(4854): 889–95. 5. Hollenberg SM, Weinberger C, Ong ES, Cerelli G, Oro A, Lebo R, et al. Primary structure and expression of a functional human glucocorticoid receptor cDNA. Nature 1985;318(6047): 635–41. 6. Kamei Y, Xu L, Heinzel T, Torchia J, Kurokawa R, Gloss B, et al. A CBP integrator complex mediates transcriptional activation and AP‐1 inhibition by nuclear receptors. Cell 1996;85(3): 403–14. 7. Kurokawa R, So¨derstro¨m M, Ho¨rlein A, Halachmi S, Brown M, Rosenfeld MG, et al. Polarity‐ specific activities of retinoic acid receptors determined by a co‐repressor. Nature 1995;377 (6548):451–4. 8. Mangelsdorf DJ, Evans RM. The RXR heterodimers and orphan receptors. Cell 1995;83(6): 841–50. 9. On˜ate SA, Tsai SY, Tsai MJ, O’Malley BW. Sequence and characterization of a coactivator for the steroid hormone receptor superfamily. Science 1995;270(5240):1354–7.

Introduction: Regulatory Mechanisms in Transcriptional Signaling by Nuclear Hormone Receptors, and their Regulators: Implications in Physiology and Disease Debabrata Chakravarti Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611

This particular volume of ‘‘Progress in Molecular Biology and Translational Science’’ series provides our current state of knowledge in select areas of nuclear receptor (NR) biology. The series can also be viewed as an early celebration of the 25th anniversary of the cloning of the first nuclear hormone receptor reported by Ronald Evans and colleagues in 1985. The biochemical characterization of the estrogen receptor by Elwood Jensen, followed by the cloning of the receptors, definition of the superfamily, characterization of hormone receptor target genes, identification of endogenous and exogenous ligands, discovery of NR coregulatory proteins, and the integration of hormone signaling to chromatin function keep studies on nuclear hormone receptors at the forefront of biomedical research. Fundamental concepts deciphered using NR signaling have been used to explain and investigate mechanisms of transcriptional signaling by other transcription factors. Chapters included in this volume will reinforce this role and are expected to stimulate and help formulate novel concepts in future research investigating regulatory mechanisms in transcriptional signaling. Steroid hormones and vitamins are words that are very familiar to scientists and non-scientists alike. As a consequence, the population at large has varying levels of understanding of what these agents can do to human body. The actions of these low molecular weight lipophilic compounds are mediated by members of the NR superfamily. While hormones such as estrogen, progesterone, and thyroid hormone are synthesized by endocrine glands and organs, vitamins such as vitamin D and vitamin A come from our diet. Therefore, it was surprising that a common integrative pathway mediates the action of these different signals originating from outside and inside of our body. NRs also serve as ‘‘sensors’’ for key endogenous metabolites that regulate cholesterol and bile xv

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acid synthesis. Not surprisingly, agonists and antagonists of NRs are some of the most prescribed drugs for the treatment of a large number of human diseases. Therefore, studies on NRs span the domains of endocrinology, physiology, synthetic physiology, pathophysiology, biochemistry, and molecular biology. Very few signaling pathways influence so many diverse subspecialties and research area in the way nuclear hormone receptor signaling continues to do. To drive that point home, a simple PubMed search with the words ‘‘nuclear hormone receptors’’ identified 87,005 articles while a similar search for ‘‘nuclear factor kB (NF‐kB)’’ yielded 27,631 articles. The field is ever expanding, and new information is produced almost on daily basis. Therefore, it would be impossible to cover all important facets of nuclear hormone receptor signaling in a single volume or even multiple volumes. The purpose of this volume is therefore not to comprehensively review the field but to focus and expand on selected areas of NR biology. The choice of topics does not imply that the areas that have not been covered are any less important. So for the sake of practicality, considering a large number of reviews have been and are being published, this issue will address selected features of NR signaling. We decided to focus on a set of representative receptors, coregulators, and the playing field of NRs, the chromatin. The authors for this volume include pioneers as well as young and upcoming researchers who we believe have contributed significantly in advancing the NR field. Each chapter is organized in a similar fashion, providing current knowledge on the subject area integrated with enough background information allowing the reader to appreciate how the field evolved to its current state. The chapters are organized to progress in a logical manner giving the volume a text book-like feeling. Almost all chapters have a discussion on NRs to allow readers who want to read a single or a few selected chapters of this volume to appreciate the major functions of NRs. The members of the NR superfamily regulate hormone and vitamin action and physiology primarily by modulating transcription of receptor responsive genes. Significant insight has been obtained on the structure and function of nuclear hormone receptors. In essence, NRs are DNA‐binding transcription factors. In the absence of ligand, some hormone receptors potently repress transcription while addition of ligand not only relieves repression but also allows for robust activation of hormone responsive genes. Transcriptional repression by NRs is achieved by recruitment of SMRT/NCoR‐corepression complexes containing histone‐modifying and chromatin‐remodeling activity that helps promote a repressive chromatin state. Addition of ligand allows receptors to change conformation, causing release of corepressor complexes and recruitments of coactivator complexes with histone and chromatin‐modifying activities that promote an activated chromatin state. In this scenario, the central role of ligand is to promote exchange of cellular complexes with opposing

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enzymatic activities. This simple model is at the heart of our present day understanding of hormone receptor function. Despite the simplicity of the model, the pathways are exceedingly complex, involving hundreds of regulatory proteins that are broadly classified as NR‐coregulators. The output of this orchestration of receptors/coregulators/chromatin modification and remodeling is the regulation of genes that influence physiology. In the subsequent 10 chapters, we will review how NRs regulate physiology (Chapters 1–3), how receptors and coregulators are themselves regulated (Chapter 4), the role of chromatin and chromatin‐remodeling proteins in hormone action and transcriptional repression (Chapters 5–7), and the roles of various chromatin‐ modifying activities that transduce hormone signals to chromatin to activate transcription (Chapters 8–10). Metabolic diseases such as hypertension, obesity, diabetes, and cardiovascular disease have reached an alarming state. While it is clear that a single mechanism would not explain the complexity of each of these diseases, the role of nuclear hormone receptors as endogenous sensors of cellular metabolites are implicated in the development and progression of some of these diseases. These properties of NRs are extensively discussed in the first chapter by Huang and Schulman. NRs such as PPARs, LXRs, FXRs, and RORs play critical roles in these physiologic processes by serving as sensors of fatty acids, and cholesterol derivatives, and are discussed in detail. Finally, the intimate link between the regulation of metabolism and the control of inflammation by these members of NR superfamily are superbly described in this review. The extensive discussion on these ‘‘metabolic’’ NRs, their physiological target pathways, and how their studies might lead to development of potential future drugs should give the readers a clear idea about the importance of NR function in metabolic diseases. The roles for steroid hormones in controlling the biology of reproduction are well known. The uterus is highly responsive to estrogen and progesterone. While significant efforts have been devoted to understanding the role of estrogen in reproduction and reproductive as well as other diseases, the role of progesterone is not well studied. In Chapter 2, Kim and colleagues focus on the role of progesterone in the regulation of uterine function. Synthetic progestins or selective progesterone receptor modulators (SPRM) are used for treatment of uterine pathologies. The description of how the expression of progesterone receptors in normal endometrium and myometrium and altered progesterone signaling in endometrial cancer and leiomyoma help integrate its role in uterine biology and cancers is fascinating to read and at the same time provides a translational context in which future drugs can be modeled to treat uterine diseases. This review also highlights the role of coactivators and other signaling proteins such as Foxo1, AP1, NF‐kB, EGF, and AKT that crosstalk with progesterone receptors in impacting uterine biology. It is hoped that this

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review will spur interests of scientists and clinicians to study the mechanisms regulating the development and growth of these tumors that disproportionally affect women. A vast majority of NRs are classified as ‘‘orphans’’ since they do not have known ligands when the receptors were identified. Tremendous efforts from a large number of laboratories were invested in identifying ligands for these orphan receptors which led to the identification of endo and xenobiotics as potential ligands for NRs such as PXR/SXR and CAR. It became immediately clear that these receptors are major regulators of drug and endobiotic metabolism, highlighting their significant clinical and therapeutic applications in the metabolism/detoxification of drugs and removal of endogenous as well as xenobiotic compounds that are critical for the maintenance of physiologic homeostasis. This detoxification process is carried out by concerted actions of phase I cytochrome P450 enzymes, phase II conjugating enzymes, and phase III drug transporters. He and Xie (Chapter 3) review what is known to date about the roles of NRs in the transcriptional regulation of some of the key drug‐ metabolizing enzymes. A major advancement of the field has been the development of humanized transgenic animal models in which the role of human receptors in drug metabolism can be evaluated. A comprehensive understanding of the NR function in drug metabolism and the use of the humanized mouse model will significantly advance our understanding of drug–drug interactions and drug toxicity in humans which has been a major problem in drug therapeutics. This is a fascinating chapter that should be of significant interest to basic scientists, and clinicians, as well as FDA and pharmaceutical companies. In Chapter 4, Lonard and O’Malley address how the activity of the NRs themselves is regulated. This is a critical issue since the overall level of NRs and their coregulators are major determinants of hormone signaling. In a way it is surprising that while tremendous knowledge has been gained in our understanding of gene regulation by NRs, little is known about how the NRs and their coregulators are regulated and whether the proteins involved in regulating NR and coregulator levels in cells also in turn regulates transcription of hormone responsive genes. This chapter clearly establishes that the ubiquitin proteosomal system is involved in determining NR and coregulator levels as well as in modulating NR‐mediated gene expression. Some of the components of the proteosomes such as TRIP1/Sug1 and E6AP not only regulate NR function but also alter chromatin ubiquitination, thereby providing an integrative mechanism of hormone action in chromatin. Interestingly, NR ligands play an apparently paradoxical role in this process. This chapter reviews in a fascinating manner how this paradoxical role of ligand can be accounted for. Finally, this chapter also discusses the role of deubiquitylating activities in NR regulation. Questions such as how all these activities are integrated during

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hormone responsiveness, how ‘‘a unique transcription‐based Ubiquitin clock’’ is established and propagated, and whether additional proteins/modifications play roles in the above processes will keep this field active for a long time to come. The major platform on which NRs perform their act is chromatin: If you are a biochemist and want to know every step by which the whole receptor repression and activation complexes are established and how these steps are modulated by hormones and vitamins, you must turn to in vitro studies using chromatin as a template. In this review (chapter 5), Ruhl and Kraus provide a historical view about how the field of in vitro studies using chromatin templates was initiated and how researchers effectively utilized these techniques to study the molecular and biochemical aspects of NR biology. This is followed by a detailed description of how each component should be purified and reconstituted, and methods to analyze transcriptional output. Subsequent sections describe how in vitro systems helped decipher/understand the role of ligands in mediating productive interactions with coregulators. Detailed descriptions are also provided for analyzing the roles of SRC proteins, CBP/p300 coactivators, and mediators in such an in vitro system. While in vitro assays have been very useful in dissecting the role of coactivators, the system has not been as effective to study transcriptional repression by NRs. The review discusses the associated problems and how they can be overcome. Finally, these in vitro studies are integrated with in vivo genome‐wide association studies. This chapter beautifully highlights the strengths and tremendous potential of in vitro studies in understanding NR function. While significant insight has been gained in our understanding of chromatin posttranslational modifications and NR function, little is known about the role that chromatin‐remodeling proteins play in NR function. Additionally, how regulation of histone acetylation and deacetylation impact NR function has not been extensively reviewed. In Chapter 6, Buranapramest and Chakravarti review the role of chromatin remodelers and histone chaperone proteins in NR function. As evident from the review, chromatin‐remodeling proteins belonging to SWI/SNF and ISWI complexes play critical roles in both NR‐ mediated gene activation and repression. One major theme that comes out of this review is that a role of hormone in gene regulation by NRs is to alter chromatin architecture. For example, hormone treatment causes release of histones from target gene promoters most likely to prepare the template for transcriptional response. Support for this view is the observation that vitamin D3 treatment or knockdown of the chromatin assembly and remodeling protein Acf1 promotes histone H3 and H4 eviction and activation of RANKL gene. Whether hormone treatment causes histone eviction or histone replacement or promotes nucleosome sliding or exchange are some of the questions that should be addressed experimentally in the future and are discussed in great

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detail in this review. This review also summarizes the role of the INHAT (inhibitors of acetyltransferases) proteins in NR regulation. This review explains in great detail, plausible mechanisms by which INHAT activity functions in the context of NR and other transcription factors. The goal of this chapter has been to highlight the progress that has been made but more importantly to present a case for more vigorous investigation on how hormone signaling alters chromatin structure on NR target genes. The theme of transcriptional regulation by NRs is further highlighted in Chapter 7 in which Stewart and Wong discuss in more detail our present day knowledge of NR repression. The pioneering discovery of SMRT/NCoR as NR corepressors stimulated significant follow‐up research, leading to the identification of subunits of the corepression complexes. Stewart and Wong discuss the role of chromatin‐modifying activities such as histone deacetylases, methyltransferases, and demethylases in NR repression. The fact that transcriptional repression is as important as transcriptional activation by NRs in physiology is very well covered when the authors discuss the physiologic functions of NR‐ mediated repression. It becomes clear that NR repression is important in resistance to thyroid hormone syndrome, in amphibian metamorphosis, and regulation of adipogenesis. In talking about adipogenesis, the recent discovery that the mammalian Sirt1 protein is involved in PPAR‐mediated adipogenic pathway further demonstrates the power of repression. In addition to CBP/p300, the members of the p160 coactivator family play critical roles in NR signaling. Extending these observations to a clinical/translational setting, Chen and his colleagues in Chapter 8 discuss the roles of the p160/SRC/ACTR coactivators and a new coregulator termed ANCCA in human cancers. They review the genetic dysregulation of these factors including gene amplifications and translocations in various forms of cancer. One interesting theme that has developed from these studies is that these factors are also dysregulated in nonhormone responsive cancers, suggesting that these coregulators in addition to NRs also target other transcription factors to exert their effect under normal and pathophysiologic contexts. The demonstration that ACTR might be involved in E2F‐mediated transcriptional regulation of cell cycle and proliferation genes further demonstrates that a single coactivator might participate in multiple transcriptional signaling pathways, regulating critical aspects of cell growth and differentiation. Finally, the role of ACTR in modulating the IGF‐1/Akt pathway again reinforces the idea that various cellular signaling pathways are integrated in a manner that is both critical to normal physiology and in human diseases including cancer. These studies should also encourage scientists to look beyond their own favorite transcription factors. Equally fascinating is the discussion on a recently discovered coregulator ANCCA which has ATPase activity and is strongly regulated by androgen and estrogen in human cancers. Although ANCCA is regulated by hormones,

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it appears to play a predominant role in E2F‐mediated gene regulation, thereby integrating hormone signaling with other transcriptional regulators of cell cycle and proliferation. This in depth review on p160/ACTR highlights the critical importance of NR regulators functioning with NRs and beyond in human cancers. In addition to acetylation, methylation of specific arginine and lysine residues on histones also contribute to transcriptional activation and repression. In Chapter 9, Kuhn and Xu provide us with a present day understanding of the role of protein arginine methyltransferases (PRMTs) in NR function and beyond. As you read the chapter, you will encounter background information as to how the PRMTs were discovered and how they regulate NR function. The discussion on catalytic activity of PRMTs is fascinating, while the review on the role of PRMTs in the function of NRs and other transcription factors is in depth. The authors discuss the roles of PRMTs in glucose metabolism and adipogenesis, and in cancer among others. While the role of PRMTs in NR function is clear from this review, in the future it will be critically important to determine how these various histone‐modifying activities crosstalk and crossregulate histone modifications on NR target genes. Such an understanding will allow us to test whether and how hormone dictates establishment and propagation of a histone code for NR target genes. In the final chapter (Chapter 10), Lee and colleagues review the role of the ASCOM complex as a histone H3 lysine 4 methyltransferase in NR‐mediated gene activation. In general, methylation of lysine 4 plays a critical role in transcriptional activation. Therefore, hormone‐induced recruitment of the ASCOM complex by NRs provides a mechanism for hormone induction of chromatin modifications and NR gene expression. Following a theme similar to NR interactions with HATs, the ASC-2 subunit of the ASCOM complex has NR interaction domains and directly interacts with NRs. Finally, an extensive description of the phenotypes of mutant mice defective in various subunits of the ASCOM complex present a strong case for the involvement of ASCOM in critical physiologic processes including regulation of adipogenesis, cholesterol and lipid metabolism, and placental development. While most of these effects can be linked to alterations in NR signaling, it remains a possibility that ASCOM is also involved in other signaling pathways. Consistent with this view, the authors discuss additional roles of the ASCOM complex in p53 regulation. Therefore, it appears (see Chapter 8 also) that NR coregulators can also moonlight as critical components of other transcriptional signaling pathways to manifest their effects on cellular and organ physiology. In summary, as we prepare to celebrate the 25th anniversary of the cloning of the first NR, I hope the readers will agree that this review series provides a compelling case for the dominant role that the NR field played in not only deciphering the molecular mechanisms for activated transcription but also in

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describing how intimately NRs and their coregulators are involved in controlling cellular processes and human diseases. I have summarized the major talking points of the subsequent chapters to guide readers. The field of NR signaling has been extremely fortunate to have scientists of the highest caliber spending their time and effort in advancing our knowledge and it is hoped that in the next 25 years research in NR will surpass our current achievements and will provide key tools and the molecular basis for future drug development for the treatment of human diseases. To end I thank all the contributors for their love for NRs and time and effort in putting together their respective chapters. Thanks are also due to Michael Conn, who approached me to take on this enormous responsibility, and all members of the serials publication team at Elsevier including Lisa Tickner, Delsy Retchagar and Ramesh Guru Subramanian for their time and effort in keeping us on time. Lastly, I thank Ron Evans, a pioneer of NR signaling field for writing the preface for this volume. Constructive input from the scientific community is expected with the hope that in the future this thematic volume will be updated and will cover new, additional, and emerging areas of research in NR biology. Acknowledgment Research in the DC laboratory was supported in part by NIH grant R01DK65148.

Regulation of Metabolism by Nuclear Hormone Receptors Huey‐Jing Huang* and Ira G. Schulman{ *Department of Biology, Exelixis Inc., 4757 Nexus Centre Drive, San Diego, California 92121 {

Center for Molecular Design and Department of Pharmacology, University of Virginia Health System, Charlottesville, Virginia 22908

I. Introduction.................................................................................. II. The PPARs ................................................................................... A. PPARa .................................................................................... B. PPARg .................................................................................... C. PPARd .................................................................................... D. PPARs and Atherosclerosis........................................................... E. PPARs and Inflammation............................................................. III. LXR ............................................................................................ A. Regulation of Hepatic Lipid Metabolism by LXR .............................. B. Regulation of Reverse Cholesterol Transport by LXR ......................... C. LXR and Atherosclerosis ............................................................. D. LXR and Inflammation ............................................................... E. LXR and Diabetes ..................................................................... F. Therapeutic Potential of LXR Ligands ............................................ IV. FXR ............................................................................................ A. FXR and the Control of Bile Metabolism......................................... B. Gallstones, Cholestasis, and Bacterial Growth................................... C. FXR and Lipid Metabolism.......................................................... D. FXR and Atherosclerosis ............................................................. E. FXR and Diabetes ..................................................................... F. Control of Liver Regeneration and Tumorigenesis by FXR .................. G. Therapeutic Potential of FXR Ligands ............................................ V. RORa.......................................................................................... A. Regulation of Lipid Metabolism by RORs........................................ B. Role of RORs in Circadian Rhythm Control and Links to Metabolism.... C. RORs and Inflammation.............................................................. D. Therapeutic Potential of RORa Ligands .......................................... VI. ERRa .......................................................................................... A. ERRa and the PGC‐1a Pathway.................................................... B. ERRa and Diabetes ................................................................... C. ERRa in Adipose and Intestine .....................................................

Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87001-4

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Copyright 2009, Elsevier Inc. All rights reserved. 1877-1173/09 $35.00

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D. ERRa and Cancer...................................................................... E. Therapeutic Potential of ERRa Ligands .......................................... VII. Summary...................................................................................... References....................................................................................

35 36 37 37

The worldwide epidemic of metabolic disease indicates that a better understanding of the pathways contributing to the pathogenesis of this constellation of diseases need to be determined. Nuclear hormone receptors comprise a superfamily of ligand‐activated transcription factors that control development, differentiation, and metabolism. Over the last 15 years a growing number of nuclear receptors have been identified that coordinate genetic networks regulating lipid metabolism and energy utilization. Several of these receptors directly sample the levels of metabolic intermediates and use this information to regulate the synthesis, transport, and breakdown of the metabolite of interest. In contrast, other family members sense metabolic activity via the presence or absence of interacting proteins. The ability of these nuclear receptors to impact metabolism and inflammation will be discussed and the potential of each receptor subfamily to serve as drug targets for metabolic disease will be highlighted.

I. Introduction There is currently a worldwide epidemic of metabolic disease characterized by obesity, type II diabetes, hypertension, and cardiovascular disease. The factors behind this epidemic appear to be combination of genetic predisposition, high caloric diets, and our increasingly sedentary lifestyles. Indeed recent statistics from the American Heart Association indicate that almost 50% of American adults are at risk for cardiovascular disease and the Center for Disease Control reports that approximately 16 million have type II diabetes. Although a number of drugs are currently available to treat the constellation of metabolic ailments, the growing epidemic indicates that we still require a better understanding of the genetic networks and signal transduction systems that underlie the pathogenesis of these conditions. Further definition of the factors responsible for metabolic control may pave the way toward new drug targets with novel mechanisms of action for the treatment of human disease. Nuclear receptors comprise a superfamily of ligand‐dependent transcription factors that regulate genetic networks controlling cell growth, development, and metabolism. Consisting of 48 members in the human genome the superfamily includes the well‐known receptors for steroids, thyroid hormones,

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

3

and vitamins.1 Members of the nuclear receptor superfamily are characterized by a conserved structural and functional organization consisting of a heterogeneous amino terminal domain, a highly conserved central DNA‐binding domain (DBD), and a functionally complex carboxy terminal ligand‐ binding domain (LBD). The LBD mediates ligand binding, receptor homo‐ and heterodimerization, repression of transcription in the absence of ligand, and ligand‐dependent activation of transcription when agonist ligands are bound.2 Crystal structures of several LBDs support molecular and biochemical studies indicating that ligand binding promotes a conformational change in receptor structure. What appears to be a relatively flexible conserved helix near the carboxy terminus (helix 12) occupies unique positions when structures of unliganded, agonist‐occupied, and antagonist‐occupied LBDs are compared.3,4 Importantly, mutagenesis experiments indicate that helix 12, referred to as activation function 2 (AF‐2), is necessary for ligand‐dependent transactivation by nuclear receptors. The AF‐2 helix contributes an essential surface to the formation of an agonist‐dependent hydrophobic pocket that serves as a binding site for coactivators. The alternative positions occupied by the helix 12 in the unliganded or antagonist‐occupied conformations preclude the formation of this binding pocket.5,6 Classic experiments that defined the effects of glucocorticoids and thyroid hormone on metabolic control provided the foundation for the endocrine regulation of metabolism.7,8 Similar to these classical endocrine receptors, a number of orphan receptors first cloned based on homology to the well conserved receptor DBD have subsequently been shown to regulate genetic networks that control important metabolic pathways. In many cases these same pathways are deranged in instances of metabolic disease and it is this class of metabolic sensing receptors that will be the focus of this review. Several of the receptors that will be discussed including the peroxisome proliferator activated receptors (PPARs), the liver X receptors (LXRs), and the farnesoid X receptor (FXR) and perhaps the retinoid‐related orphan receptors (RORs) appear to function by directly sampling the levels of fatty acids and cholesterol derivatives via the receptor LBD and regulating genetic networks that control the synthesis, transport, and breakdown of the cognate ligand.9–14 Importantly, these fatty acid‐ and cholesterol‐derived natural ligands bind to receptors with affinities close to the physiological concentrations know to exist for these metabolites.11,12,15–18 Thus, these receptors are poised to sense and respond to small changes in the flux through the metabolic pathways that they control. The estrogen receptor related receptors (ERRs) comprise an additional subfamily of nuclear receptors that also appear to play important roles in the regulation of metabolism. In contrast to the ligand‐activated receptors mentioned earlier, the activity of the ERRa in particular appears to be controlled by the presence or

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absence of interacting proteins instead of lipid‐derived ligands. The focus of this review will be on the function of these ‘‘metabolic’’ nuclear receptors, the physiological pathways they regulate, and their potential as drug targets. There are several reoccurring themes that appear throughout this review. First, the ability of individual nuclear receptors to regulate multiple genetic networks in different tissues has made drug discovery a challenging process for this class of potential drug targets. Second, in many cases the phenotype of a genetic knockout of a particular nuclear receptor does not accurately predict the physiological activity of a receptor‐specific small molecule agonist or antagonist. Finally, there is an intimate connection between the regulation of metabolism and the control of inflammation.

II. The PPARs Three distinct members of the PPAR subfamily each encoded by a distinct gene have been identified and well characterized. PPARa (NR1C1) is highly expressed in liver, kidney, and muscle. PPARg (NR1C3) is enriched in adipose tissue and PPARb/d (NR1C2; referred to PPARd in this review) appears to be ubiquitously expressed. All three PPARs bind to DNA as heterodimers with retinoid X receptors (RXR; NR2B subgroup) and prefer to bind to direct repeats of the nuclear receptor half‐site AGGTCA separated by 1 nucleotide (DR1). Each subtype appears to have unique functions and PPARa and PPARg are the targets of the fibrate and thiazolidinedione (TZD) classes of drugs, respectively.19

A. PPARa PPARa is the molecular target of the fibrate class of drugs used for the treatment of hypertriglyceridemia. Studies in vitro and in vivo demonstrate that PPARa directly regulates a network of genes encoding the proteins required for the uptake of fatty acids, enzymes required for the oxidation of fatty acids (b oxidation), and enzymes required for ketone body utilization by binding to control regions in the promoter of these genes.19 Thus, activation of PPARa promotes the utilization of fat as an energy source. Activation of PPARa also directly induces the genes encoding the apolipoproteins apoAI and apoII, which contribute to the protein core of high density lipoprotein (HDL) particles. Thus, fibrates have the added benefit of slightly raising HDL, the ‘‘good cholesterol.’’20,21 Given its role in controlling the utilization of fatty acids for energy production it is not surprising that PPARa is required for the normal response to fasting and starvation. Mice lacking PPARa accumulate triglycerides in the liver and become hypoketonic and hypoglycemic during fasting or starvation.22,23

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Recent studies indicate the fibroblast growth factor 21 (FGF21) functions as an endocrine hormone that mediates many of the effects of PPARa. The gene encoding FGF21 is directly induced by PPARa in response to fasting via a binding site in the promoter. FGF21 in turn stimulates lipolysis in adipose tissue and ketogenesis in the liver.24 Taken together PPARa appears to function as a sensor of the fed/starved state. The increase in fatty acids derived from adipose during fasting provides PPARa agonists that promote the utilization of fat as energy and further stimulates the release of fatty acids from the adipose by the endocrine action of FGF2124 (Fig. 1).

B. PPARg PPARg is the master transcriptional regulator of adipogenesis and plays an important role in the process of lipid storage.25 Thus, PPARa and PPARg have contrasting roles in the regulation of fat metabolism; PPARa promotes the utilization of fat in the liver and muscle while activation of PPARg promotes storage in adipose. A number of naturally occurring fatty acids and prostanoids have been shown to act as PPARg agonists; however, perhaps most importantly

Adipose Fatty acids

Lypolysis PPARa

b-oxidation ketogenesis

FGF21

Liver

FIG. 1. Regulation of fatty acid utilization by PPARa. Fatty acids directly bind to PPARa and act as agonists that increase transcriptional activity. Activated PPARa directly induces genes that encode the enzymes required for b‐oxidation and ketogenesis. PPARa also increases expression of the gene encoding FGF21. FGF21 acts in an autocrine and paracine fashion to further enhance b‐oxidation and ketogenesis and FGF21 also stimulates lipolysis in adipose tissue to promote the release of fatty acids. See text for further details.

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was the identification that the TZD class of insulin sensitizing drugs including rosiglitazone (Avandia) and pioglitazone (Actos) are PPARg agonists.26 Although PPARg was identified as the therapeutic target of TZDs in 199526 it is still fair to say that the mechanism underlying the insulin sensitizing activity of this class of drugs is still not completely defined. Fatty acid accumulation in insulin sensitive tissues such as liver and skeletal muscle has been shown to promote insulin resistance.27 Activation of PPARg in adipose has been proposed to increase the number of adipocytes and promote the relocalization and storage of fat in adipose, protecting peripheral tissues from lipotoxicity.28 Consistent with this idea is the observation that selective knockout of PPARg in adipose eliminates the therapeutic activity of TZDs in mice29 and that a common side effect of TZD treatment in humans is weight gain due to an increase in adipose mass.30 PPARg is also expressed at relatively low levels in other tissues and selective knockout in skeletal muscle reduces the therapeutic activity of TZDs,31 suggesting that there are adipose‐independent sites of PPARg activity. The TZDs have proven to be effective drugs for improving insulin sensitivity and treating type II diabetes. However, they are not without problems. Troglitazone, the first TZD in the clinic, was taken off the market because of cases of drug‐induced liver damage.32 Additionally, recent meta‐analyses have indicated that treatment with rosiglitazone is associated with increased risk of myocardial infarction and deaths due to cardiovascular events.33 Pioglitazone treatment was also shown to be associated with an increase in serious heart failure although a significantly lower risk of myocardial infarction and death was observed in this patient population.34 Finally, muraglitazar an investigational drug that is a dual agonist of PPARa and PPARg was found to be associated with an increase in major cardiovascular events and increased incidence of death.35 The increases in cardiovascular events and mortality seen with these drugs are relatively small. Nevertheless, the wide scale use of PPARg agonists in the type II diabetic population has raised serious concerns about the safety of these drugs for long term therapy. The molecular basis underlying the increase in cardiovascular events is not clear. Changes in energy metabolism mediated by PPARs could significantly influence cardiac function. Additionally, several studies have identified edema as a side effect of TZD treatment that could impact heart function.36,37 Regulation of the epithelial sodium channel (ENaCg) in the kidney by PPARg has been suggested as potential mechanism behind the TZD‐dependent edema38 but it remains to be seen if inhibiting this channel will decrease the cardiovascular events associated with TZD treatment. Based on the recent clinical data it is unlikely that additional PPARg agonists will make it to the clinic unless a better understanding of the tissue‐specific responses of this receptor is obtained.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

7

C. PPARd Not surprisingly based on its ubiquitous expression pattern, genetic knockout of PPARd results in a number abnormalities including embryonic lethality secondary to placental defects, decreased adipose mass, mylination deficiencies, altered inflammatory responses, and impaired wound healing.39 More recent studies exploiting additional genetic models and synthetic agonists, however, have uncovered important functions for this receptor in the control of metabolism and inflammation.39,40 X‐ray crystallography, indicates that PPARd has a relatively large ligand binding pocket and in vitro studies indicate that fatty acids as well as eicosanoids including prostaglandin A1 and carbaprostacyclin function as agonists.39 Very low density lipoprotein (VLDL) particle associated fatty acids have also been demonstrated to induce PPARd target genes in a receptor‐dependent manner41 raising the possibility that PPARd regulates the synthesis, transport, and catabolism of triglyceride‐rich lipoprotein particles. Further support for a role of PPARd in lipoprotein metabolism results from studies exploring the activity of the PPARd‐specific synthetic agonist GW501516. Treatment of animals, including, nonhuman primates with GW501516 significantly increases HDL particles, lowers triglycerides and low density lipoprotein (LDL) particles, and decreases fasting insulin levels.39,40 Mechanistic studies point to regulation of the gene encoding the ATP binding cassette transporter ABCA1 by PPARd as an important step in the control of HDL levels.42 ABCA1 functions as a cholesterol transporter to transfer cholesterol out of cells to HDL particles43 and its function will be discussed further in Section III. PPARd mediated downregulation of intestinal cholesterol absorption via regulation of the gene encoding Niemann‐Pick C1‐like protein 1 (NPCL‐1), a cholesterol transporter that is the target of the cholesterol lowering drug ezetemide (Zetia), has also been suggested to play a role in the effect of PPARd on lipid levels.44 To examine the role of PPARd in specific tissues, Wang et al.45,46 fused the strong transcriptional activation domain of the viral transcription factor VP16 to the amino‐terminus of PPARd to create a ‘‘hyperactive’’ receptor that activates transcription even in the absence of agonists. Transgenic approaches were then used to express VP16–PPARd in adipose (white and brown) and skeletal muscle. In both tissues VP16–PPARd expression produced a dramatic increase in the b‐oxidation of fatty acids. In adipose, the increase fat oxidation led to a decrease in adipose mass and protection from diet‐induced obesity and insulin resistance. The protection against diet‐induced obesity results, at least in part, from increased thermogenesis in brown fat secondary to the induction of genes involved in b‐oxidation and the uncoupling of oxidative phosphorylation from ATP production.46 Uncoupling oxidative phosphorylation from ATP production by expression of uncoupling protein 1 (UCP‐1) leads to a futile cycle that

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generates heat when fat is metabolized. Weight loss was also observed in obese mice treated with the synthetic agonist GW501516,45,47 suggesting that the same PPARd‐dependent pathways can be activated pharmacologically. Nevertheless, a significant change in body weight was not detected when obese Rhesus monkeys were treated for 4 months with GW501516.42 As discussed earlier, however, significant agonist‐dependent effects on lipid metabolism and insulin level was observed in these animals indicating that GW501516 is active in nonhuman primates. Perhaps species‐dependent differences in the bioavailability, tissue distribution, and/or efficacy of GW501516 account the differences on adipose mass between rodents and primates. In skeletal muscle expression of VP16–PPARd induces genes involved in b‐oxidation, mitochondrial respiration, and increases the proportion of slow twitch oxidative muscle fibers.46 In short the muscle of these animals become fat burning machines and interestingly these transgenic mice can run on a treadmill for significantly longer times than mice without VP16–PPARd. Given the interest in ‘‘performance‐enhancing’’ drugs, Narkar et al.48 tested the ability of GW501516 to increase endurance in mice (defined as running on a treadmill until exhaustion). In contrast to the result observed with the super‐active VP16–PPARd construct, activation of endogenous PPARd with the synthetic agonist did not increase endurance. When agonist treatment was coupled with a minimal exercise regimen, however, the combination of drug with exercise produced a significantly larger increase in running time compared to exercise alone. Although pharmacological activation of PPARd alone does not improve endurance, pharmacologic activation of AMP kinase, a kinase that is activated when energy levels are low,49 is sufficient by itself to improve endurance in sedentary mice.48,50 AMP kinase, like PPARd, is known to play an important role in muscle fiber type specification50 and the endurance promoting activity of an AMP kinase activator is lost in Ppard
/
mice.48 Thus, activation of PPARd is necessary to improve endurance. Interestingly, activated AMP kinase increases the transcriptional activity of PPARd at least in part by phosphorylation of the peroxisome proliferator activated receptor g coactivator 1a (PGC‐1a), a transcriptional coactivator that directly interacts with PPARd.48 AMP kinase activity is also induced by exercise50 suggesting a simple model that exercise activated AMP kinase increases the transcriptional activity of PPARd leading to expression of a genetic network involved in the specification slow twitch oxidative muscle fibers and improved endurance (Fig. 2A). If this linear pathway is correct, one must ask why neither exercise alone nor simply activating PPARd with a synthetic agonist (bypassing AMP kinase) is sufficient to improve endurance? We would argue that there is a threshold level of PPARd activity that must be achieved in order to increase endurance (shown schematically in Fig. 2B) and that the exercise regimen used and GW501516

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A

Exercise

B 100 PPAR activity

AMP Kinase

PO4 PGC-1 RXR

PPAR

80

Threshold for improved endurance

60 40 20 AMP agonist

Exercise + PPAR agonist

Exercise

PPAR agonist

Endurance gene

VP16–PPAR

mRNA

PPAR

0

FIG. 2. PPARd‐dependent endurance pathway. (A) Exercise leads to activation of AMP kinase which phosphorylates PGC‐1a. Phosphorylated PGC‐1a acts as a transcriptional coactivator to increase the activity of PPARd leading to the induction of endurance‐promoting genes. (B) Graph illustrates the hypothetical activity of PPARd under different conditions explored by Nakar et al.48 The black line denotes a hypothesized threshold level of PPARd transcriptional activity needed for improved endurance.

individually do not achieve this level. Perhaps increasing the duration of the exercise or improving the efficacy and bioavailability of the synthetic agonist would allow these agents to function alone. Synthetic PPARd ligands have proven to be very effective in preclinical models of diabetes and GW501516 was taken into clinic for the treatment of dyslipidemia in 2006. It remains to be seen if synthetic ligands for PPARd will prove to be effective for the treatment of human disease.

D. PPARs and Atherosclerosis The important roles for the PPARs in the control of lipid metabolism prompted a number of studies investigating the activity of subtype selective agonists in mouse models of atherosclerosis.39,51 Generally, LDL receptor knockout (Ldlr
/
) mice or apoE knockout (apoE
/
) mice treated with synthetic ligand have been used as model systems. Based on these studies, one can conclude that activation of any of the 3 PPARs reduces atherosclerosis. One, however, could also conclude the opposite; that activation of the PPARs has little or no benefit for the treatment of atherosclerosis.51 It is clear that

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differences in the genetic backgrounds of the mice, their sex, the particular agonist, the diet, and the environment can influence the outcomes of these experiments in ways that are not well understood. Interestingly, treatment of hyperlipidemic Ldlr
/
and apoE
/
mice with PPAR ligands has relatively small effects on plasma lipid levels and little or no change in total plasma cholesterol is detected even in the studies that reported agonist‐dependent decreases in atherosclerosis.51 Therefore, the predominant effect of the PPARs on atherosclerosis does appear to result from changes in plasma lipid levels. A critical event in the development of atherosclerosis is the recruitment of macrophages to the underlying epithelial layer of vessel walls and the uncontrolled uptake of oxidized cholesterol.52 Continued accumulation of oxidized cholesterol by macrophages and an associated inflammatory response leads to foam cell formation and the initiation of atherosclerosis.52 Importantly, all 3 PPAR subtypes are expressed in macrophages and in vitro studies indicate that that they can modulate cholesterol homeostasis in these cells.39,51 The ability of the PPARs to regulate macrophage cholesterol metabolism lies, at least in part, through their ability to control the expression of LXRs and subsequent induction of reverse cholesterol transport,53 the process of transporting cholesterol out of cells to HDL (discussed more detail in Section III). PPAR‐dependent/LXR‐independent pathways that modulate macrophage cholesterol levels have also been identified.54

E. PPARs and Inflammation Along with the accumulation of oxidized LDL, inflammation plays a critical role in the development and progression of atherosclerosis.52 Additionally, recent findings indicate a close link between inflammation and insulin resistance.55 Pharmacologic or genetic inhibition of pathways that underlie inflammatory responses protect experimental animals from diet‐induced insulin resistance as well as atherosclerosis,52,56,57 suggesting a direct role of inflammation in the pathology of these diseases. In both diseases, inflammatory responses mediated by macrophages appear to be crucial for disease progression. In atherosclerosis, cytokines secreted by macrophages at the vessel wall promote the recruitment of additional immune cells and the proliferation of smooth muscle cells that contribute atherosclerotic lesion development.52 Macrophages also accumulate in the adipose tissue of obese animals and humans where they produce inflammatory mediators that may contribute to the development of insulin resistance.58,59 Anti‐inflammatory activity is a property shared by many members of the nuclear hormone receptor superfamily including the 3 PPAR subtypes and occurs by inhibition of the transcriptional activity of the proinflammatory transcription factors activator protein 1 (AP‐1) and nuclear factor kappa B (NFkB).60 A number of mechanisms have been proposed for this process,

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termed transrepression, including direct interactions between PPARs and the p65 subunit of NFkB, induction of the inhibitor of kappa B alpha (IkBa), regulation of c‐Jun N‐terminal kinase (JNK) activity, competition for limiting transcriptional coactivators and corepressors, and inhibition of corepressor clearance from NFkB regulated promoters.60 The contribution of chronic inflammation to metabolic disease has led to the idea that the anti‐inflammatory activity of the PPARs, particularly in macrophages, may contribute to the beneficial effects of PPAR ligands in animal models of atherosclerosis and insulin resistance.60 To our knowledge this hypothesis has not yet been tested with either PPAR ligands or PPAR mutants that dissociate the process of activation of transcription from the process transrepression. Such dissociated ligands and receptor mutants have been identified for the glucocorticoid receptor,61,62 suggesting that such reagents could be identified for the PPARs. Nevertheless, recent studies discussed below have indicated that macrophages may be critical sites of action for the activity of PPARs.63–66 Resident macrophages in tissues display significant heterogeneity.67 In obesity classically activated macrophages, also called M1 macrophages, accumulate in adipose and play role in mediating an inflammatory response that contributes to insulin resistance.68 In lean animals and people, most adipose associated macrophages display an alternatively activated or M2a phenotype. Alternatively, activated macrophages are less inflammatory and appear to play roles in tissue repair.68,69 Energy utilization also differs between these two populations of macrophages. Classically activated macrophages predominantly use glucose while a switch to oxidative metabolism is an integral component of alternative activation; linking metabolic control to macrophage phenotype and inflammation.70 Alternative activation is induced by IL‐4 and IL‐1369 and studies indicating that both PPARg and PPARd are induced in IL‐4/ IL‐13 treated macrophages promoted a number of investigators to examine the role of macrophage PPARs in models of diet‐induced obesity.51,63–66,71 Odegaard et al.65 and Hevener et al.63 used selective knockouts and bone marrow transplantations to delete PPARg in macrophages and observed glucose intolerance and increased insulin resistance in mice exposed to a high fat diet. The Odegaard et al.65 study specifically explored macrophage phenotype and determined that alternative activation was impaired, suggesting that insulin resistance observed in the absence of PPARg results from increased inflammation from M1 type macrophages. Hevener et al.63 additionally demonstrated that the therapeutic activity of rosiglitazone was compromised when macrophage PPARg was selectively eliminated, indicating that the antiinflammatory activity of PPARg contributes to the therapeutic activity of TZDs. Using macrophage selective knockouts similar to those described for PPARg, Kang et al.64 and Odegaard et al.66 demonstrated that PPARd is required for turning on the gene expression program corresponding to

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alternative activated (M2a) macrophages. The consequences of macrophage PPARd deletion are impaired glucose tolerance and an exacerbation of insulin resistance in response to a high fat diet; once again supporting a role for macrophage inflammation in the pathology of insulin resistance.64,66 Based on these studies it appears that both PPARg and PPARd play important roles in establishing the alternative activated phenotype. It has been suggested that both receptors must have distinct roles since knockout of either subtype alone is sufficient to impair alternative activation.66 Nevertheless, the exact function of each receptor in the alternative activation pathway remains to be determined. The macrophage‐selective knockout experiments described earlier suggest that the anti‐inflammatory activity of the PPARs may contribute to their therapeutic activity. Several investigators have gone a step further and suggested that specifically targeting PPARs in macrophages with tissue‐selective small molecules may be a novel and effective method for treating metabolic disease.63–66,72 The enthusiasm for such approaches, however, must be tempered with the realization that other factors including genetic background, diet, and environment may contribute to the knockout phenotypes. In a separate study, Marathe et al.73 used bone marrow transplantation approaches to selectively knockout PPARg and PPARd in hematopoietic cells either individually or together. These authors concluded that in the C58BL/6 mice the two receptors have little or no impact on the development of diet‐induced obesity and insulin resistance and that rosiglitazone is effective in the absence of macrophage PPARg.73 The roles for PPARs as important regulators of metabolism are well documented and agonists for PPARa (fibrates) and PPARg (TZDs) have been validated in humans for the treatment of metabolic disease. Nevertheless, the contribution of macrophage PPARs to the pathology of metabolic disease and the beneficial activity of PPAR agonists remains to be determined.

III. LXR The LXR subgroup of the nuclear receptor superfamily is comprised of two subtypes, LXRa (NR1H3) and LXRb (NR1H2) that are encoded by separate genes. The founding member of the subgroup LXRa, was originally cloned from a liver cDNA library, hence the name liver X receptor, and found to be highly expressed in the liver, kidney, and intestine.74 In contrast, LXRb is more ubiquitously expressed.75 Both LXRs bind to DNA and regulate transcription as heterodimers with RXRs with preferred binding to direct repeats of the nuclear receptor half‐site AGGTCA separated by four nucleotides (DR4).74,75

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A. Regulation of Hepatic Lipid Metabolism by LXR The first link between LXR and lipid metabolism came from the identification of cholesterol derivatives including 22(R)‐hydroxycholesterol, 24(S)‐ hydroxycholesterol, and 24(S),25‐epoxycholesterol as ligands that directly bind to both LXRa and LXRb and increase their transcriptional activity.16–18 More recent studies have also demonstrated that 27‐hydroxycholesterol and cholestenoic acid are LXR ligands.76,77 The identification of hydroxycholesterols as natural LXR ligands dovetailed nicely with the characterization of LXRa knockout mice. Apparently, normal under standard laboratory conditions, when challenged with a diet rich in cholesterol Lxra
/
mice accumulate massive amounts of cholesterol in the liver. Molecular analysis uncovered aberrant regulation of several genes involved in lipid and cholesterol metabolism including Cyp7a1, which encodes cholesterol 7a hydroxylase, the rate‐ limiting enzyme in the conversion of cholesterol to bile acids.78 Subsequently, the ATP binding cassette transporters ABCG5 and ABCG8 which move cholesterol out of the liver and into the intestine were identified as LXR target genes.79,80 Thus, an increase in cholesterol levels is predicted to lead to an elevation in the concentration of cholesterol‐derived LXR ligands resulting in the catabolism of cholesterol to bile acid and the excretion of cholesterol out of the liver (Fig. 3). Importantly, Cyp7a1, ABCG5, and ABCG8 all appear to be directly regulated by LXR18,79 although the binding site for LXR present in the murine Cyp7a1 gene is not conserved in the human gene.18 Along with effects on cholesterol metabolism activation of LXR agonists also increases expression of genes involved in fatty acid metabolism including the master transcriptional regulator of fatty acid synthesis, the sterol response element binding protein 1c (SREBP1c)81,82 (Fig. 3). Additionally, several of the genes encoding the enzymes involved in fatty acid metabolism including fatty acid synthase (FAS) and stearoyl CoA desaturase 1 (SCD‐1) are regulated directly or indirectly by LXR.83–85 The coordinate upregulation of fatty acid synthesis with reverse cholesterol transport is most likely to provide lipids for the transport and storage of cholesterol.

B. Regulation of Reverse Cholesterol Transport by LXR Based on the defined role for LXR in hepatic cholesterol catabolism and excretion one might have expected that synthetic LXR agonists would lower plasma cholesterol levels. Quite surprisingly, however, treatment of animals with LXR agonists significantly elevates HDL cholesterol.82 Gene expression analysis in the livers and intestines of LXR agonist‐treated mice identified ABCA1 as a direct LXR gene and this discovery stimulated great interest in the therapeutic potential of LXR agonists given the links between ABCA1, HDL metabolism, and atherosclerosis.86–88 ABCA1 is required for the process

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Macrophage

LXR

ABCA1 ABCG1

Triglyceride SREBP1c

HDL Bile acids

Cholesterol LXR

Cyp7A

ABCG5 ABCG8 LXR

FIG. 3. Regulation of cholesterol metabolism by LXR. Activation of LXR in macrophages results in the upregulation of genes encoding proteins that participate in reverse cholesterol transport including ABCA1, ABCG1, ABCG4, and apoE. In the liver, LXR activation results in upregulation of the genes encoding cholesterol 7 hydroxylase (Cyp7a), the rate limiting enzyme in the catabolism of cholesterol to bile acids, and the half ABC transporter ABCG5 and ABCG8 that function together to excrete cholesterol from the liver into the intestine. Thus, activation of LXR increases the net flux of cholesterol out of the body. LXR also regulates expression of SREBP1c, a master transcriptional regulator of genes involved in fatty acid and triglyceride synthesis. See text for details.

of reverse cholesterol transport whereby cells efflux internal cholesterol to acceptor proteins on pre‐b‐HDL particles. Loss of functional ABCA1 results in Tangier disease, a condition in which patients have extremely low levels of circulating HDL and an increased risk for developing atherosclerosis. Examination of fibroblasts isolated from subjects with Tangier disease reveals that ABCA1 defective cells are unable to efflux cholesterol, suggesting that the low HDL levels and increased risk of atherosclerosis results from a loss of reverse cholesterol transport. Historically, Tangier disease patients present with large accumulations of cholesterol‐laden macrophages in their lymph tissues, highlighting the role of ABCA1 and reverse cholesterol transport in macrophage cholesterol homeostasis.43,89,90 As described in Section II.D, accumulation of oxidized LDL cholesterol by macrophages in the arterial wall is an initiating step in the development of atherosclerotic lesions52 and recent studies with mouse knockouts of ABCA1 further support a link between reverse cholesterol transport and atherosclerosis.91–93 In support of the role of LXR as a direct regulator of ABCA1 expression and activity, treatment of primary macrophages

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or cell lines with LXR agonists results in induction of the ABCA1 gene, increase levels of ABCA1 protein, and an increase in cholesterol efflux.86,88,94 A binding site for LXR–RXR heterodimers in the ABCA1 promoter has also been described.86 Subsequent studies identified other proteins involved in the reverse cholesterol transport including ABCG1, ABCG4, and apoE as direct LXR target genes.94–97 Thus, activation of LXR results in the mobilization of cholesterol in the periphery and stimulates the catabolism and excretion of cholesterol when it arrives in the liver (Fig. 3). Interestingly, genetic deletion of LXR activity in mice (Lxra
/
/Lxrb
/
) results in the accumulation of cholesterol‐laden macrophages and splenomegaly similar to that observed in Tangier disease patients.98,99

C. LXR and Atherosclerosis The accumulation of oxidized LDL cholesterol by macrophages in blood vessel walls is an early event in the pathogenesis of atherosclerosis and it had long been suggested that reversing this process by pumping cholesterol out of macrophage foam cells would have an inhibitory effect on the progression of atherosclerosis.52 The ability of LXR to directly regulate reverse cholesterol transport in macrophages allowed two experiments to be carried out to test this hypothesis. First, transplantation of lethally irradiated apoE
/
and Ldlr
/
mice with bone marrow from wild type or Lxra
/
/Lxrb
/
mice demonstrated that genetic deletion of LXR leads to an increase in atherosclerosis is these well established mouse models.99 Second, treatment of apoE
/
and Ldlr
/
mice with synthetic LXR agonists leads to a reduction in atherosclerosis.100,101 Together the combination of genetic analysis and pharmacology clearly demonstrated the antiatherogenic activity of LXR. Not surprisingly, the mRNA levels for LXR target genes including ABCA1 and apoE are elevated in the atherosclerotic lesions of mice treated with LXR agonists.101 Subsequent studies combining bone marrow transplantation with the administration of synthetic LXR agonists have demonstrated that LXR activity in macrophages is necessary for the antiatherogenic effect of LXR ligands.102

D. LXR and Inflammation It is easy to assume that the ability of LXR to regulate lipid metabolism and reverse cholesterol transport provides the mechanistic basis for the antiatherogenic activity of LXR.99–101 Experiments using cultured macrophages, however, also demonstrate that LXR agonists can inhibit the expression of several proinflammatory genes including iNOS, COX‐2, and MMP‐9 and these compounds are effective in a murine model of irritant contact dermatitis.103,104

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Molecular studies indicate that activation of LXR decreases the transcriptional activity of NFkB using many of the same mechanisms described for the PPARs (see Section II.E).104 Since atherosclerosis is considered an inflammatory disease52 the question remains whether LXR mediates its antiatherogenic activity via control of reverse cholesterol transport, by limiting the inflammatory response, or both. Future studies that combine genetically altered macrophages (i.e., Abca1
/
) introduced by bone marrow transplantation along with the administration of LXR agonists can be used to define the individual contributions of reverse cholesterol transport and anti‐inflammatory activity to therapeutic effects of LXR ligands. Additionally, studies with the glucocorticoid receptor have shown that it is possible to identify nuclear receptor ligands that repress inflammatory genes but do not activate positively regulated glucocorticoid receptor target genes.61,62 One expects that such dissociated ligands will also be identified for LXR. A number of studies have suggested a link between viral or bacterial infections and atherosclerosis.105 In support of this hypothesis enhanced expression of Toll‐like receptors (TLR), which mediate the innate immune response to invading pathogens via stimulation of proinflammatory pathways, has been detected in human atherosclerotic lesions.106 Interestingly, studies by Castrillo et al.107 indicate that activation of TLR4 inhibits the transcriptional activity of LXR and the ability of macrophages to efflux cholesterol. The cross talk between TLR4 signaling and LXR activity suggests one potential mechanistic basis for the impact of infectious agents on cardiovascular disease. Along with effects in macrophages, recent studies have identified an important and specific role for LXRb in T cell proliferation.108 T cell activation triggers induction of the oxysterol‐metabolizing enzyme SULT2B1 and suppression of reverse cholesterol transport by decreasing the availability of endogenous LXR agonists. Consistent with a role for LXRs in T cell proliferation is the observation that knockout of LXRb confers a proliferative advantage while binding of agonists to LXRb during T cell activation inhibits mitogen‐ driven expansion. LXRa is not expressed in lymphocytes indicating that this activity is subtype selective.108 In a coordinate fashion, the SREBP2 dependent pathway for cholesterol synthesis is activated108 (Fig. 4). SREBP2 is the major transcriptional regulator of proteins required for cholesterol uptake and synthesis including the LDL receptor and HMG‐CoA reductase, the rate limiting enzyme in cholesterol biosynthesis.109 This coupling of decreased LXRb activity with activation SREBP2 insures that intracellular cholesterol levels are sufficient to support cell growth and the rapid expansion of the T cell population upon stimulation. Interestingly, inactivation of the cholesterol transporter ABCG1 attenuates the ability of LXR agonists to inhibit proliferation, directly linking cholesterol homeostasis to the antiproliferative action

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T cell receptor

SULT2B1

SREBP2

Oxysterols

Cholesterol synthesis HMG-CoA reductase

Cholesterol

LXRb Reverse cholesterol transport ABCG1

Cholesterol

FIG. 4. Regulation of T cell proliferation by LXRb. Stimulation of T cell proliferation increases the expression of SULT2B1 an enzyme that catabolizes oxysterols, the endogenous ligands for LXR. The decrease in LXR ligands reduces the transcription activity of LXRb resulting in a downregulation of cholesterol leaving cells via the reverse cholesterol transport pathway. In a coordinate fashion, SREBP2 and cholesterol biosynthetic enzymes are upregulated providing cholesterol for cell growth. See text for details.

of LXRb.108 Mice lacking LXRb also exhibit lymphoid hyperplasia and enhanced responses to antigenic challenge, indicating that proper regulation of LXR‐dependent sterol metabolism is important for immune responses.108 These results implicate LXRb signaling in a metabolic checkpoint that modulates cell proliferation and immunity.

E. LXR and Diabetes Treatment of experimental animals with LXR agonists leads to increases in hepatic fatty acid synthesis and plasma triglyceride levels.81,82 Since elevations in fatty acids have been linked to insulin resistance and type II diabetes several investigators have examined the cross talk between LXR activity and glucose metabolism. Interestingly, along with upregulating fatty acid synthesis, activation of LXR also represses expression of the genes encoding the enzymes of gluconeogenesis in the liver including phosphoenolpyruvate carboxy kinase (PEPCK) and glucose 6‐phosphatase110,111 and induces expression of GLUT4 in adipose tissue.110 Thus, in many ways activation of LXR mimics treatment with insulin. Perhaps not surprisingly in light of this ‘‘insulin‐like’’ activity, LXR ligands decrease hepatic glucose output and lower blood glucose levels in animal models of type II diabetes.110,111 The observation, however, that LXR

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ligands can behave as insulin sensitizers even in face of relatively large increases in plasma triglyceride levels suggests the possibility of a broader role for LXR in regulating glucose homeostasis. Indeed, Mitro et al.112 have demonstrated that glucose can directly bind to LXRs with relatively weak affinity (millimolar) and function as an agonist. Thus, LXRs may directly sense glucose and regulate carbohydrate levels in manner similar to the control of cholesterol metabolism. The observation that LXRs are also active in skeletal muscle113 and that Lxra
/
/Lxrb
/
mice are resistant to diet‐induced obesity114 further supports a role for the LXRs as important coordinators of energy metabolism.

F. Therapeutic Potential of LXR Ligands The antiatherogenic, anti‐inflammatory, and antidiabetic activities of LXR agonists in animal models have highlighted the therapeutic potential of LXRa and LXRb. Nevertheless, the link between LXR activity and triglyceride metabolism has clearly dampened the enthusiasm surrounding this target class. Treatment of mice and hamsters with synthetic LXR agonists results in a significant increase in plasma triglyceride levels82 and elevations in LDL cholesterol have been observed in nonhuman primates.115 Treatment of patients with a drug that raises lipids is not a viable option for the treatment of metabolic diseases and approaches to separate the beneficial activities of LXR ligands from unwanted side effects need to be explored. Furthermore, studies in human cells have shown that LXR agonists also increase expression of the gene encoding the cholesterol ester transfer protein (CETP).116 CETP functions to transfer cholesterol esters from HDL to apolipoprotein B containing lipoprotein particles and CETP activity has been shown to inversely correlate with atherosclerosis.117–119 Indeed, CETP inhibitors are currently being explored for the treatment of atherosclerosis.118 Interestingly, defects in hepatic cholesterol metabolism are detected in Lxra
/
single knockout mice indicating that LXRb is not functionally redundant with LXRa.78 In contrast, cholesterol and triglyceride levels appear normal in Lxrb
/
mice suggesting that LXRa mediates most, if not all, of the effects of LXR ligands on triglyceride metabolism.120 The relatively low level of LXRb in the liver most likely accounts for lack of functional redundancy in this tissue. Nevertheless, in macrophages either LXRa or LXRb alone appears to be sufficient to mediate the effects of LXR ligands on reverse cholesterol transport and inflammatory gene expression. Additionally, a recent study by Bradley et al.121 indicates that the antiatherogenic activity of LXR agonists are maintained in apoE/
/Lxra
/
double knockout mice, suggesting that LXRb alone is sufficient to limit atherosclerosis. Taken together these observations have led several investigators to suggest that LXRb‐selective ligands may provide a

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mechanistic basis for identification of LXR ligands with improved therapeutic profiles.122 The enthusiasm for LXRb‐selective ligands must be tempered with the realization that the spectrum of activities measured in the complete absence of LXRa activity may differ when a subtype selective synthetic ligand is used. Additionally, the observation that the ligand binding pockets of LXRa and LXRb defined by crystallography123–126 differ by only one amino acid suggests that identification of selective ligands may not be simple. While the therapeutic activity of LXRb‐selective ligands is still an open question, it has been possible to identify ligands for other nuclear receptors that exhibit a restricted set of activities and therefore allow the separation of beneficial therapeutic activities from unwanted side effects. Perhaps the best examples of such compounds are the selective estrogen receptor modulators such as roloxifene that function as estrogen receptor agonists in some tissues and estrogen receptor antagonists in others.127 More recently, synthetic ligands for PPARg have been identified that appear to separate the insulin sensitizing activity of PPARg from unwanted effects on weight gain.128,129 A common feature of all these selective receptor modulators is that they appear to function as partial or weak agonists when characterized in vitro. When bound to receptors selective modulators produce unique conformational changes that cannot be achieved by more typical agonists.3,4 The outcome of these unique conformations is an alteration in interactions between receptors and the down‐stream coregulator proteins that mediate the transcriptional response leading to ligand‐specific effects on gene expression.127,130 Since the LXRs function in multiple tissues to mediate effects on lipid metabolism, glucose homeostasis, and inflammation we expect that the identification of selective LXR modulators will yield compounds with beneficial therapeutic activities.

IV. FXR FXRa (NR1H4), like the PPARs and LXRs, was originally identified as an orphan member of the nuclear hormone receptor superfamily and subsequently shown be activated by the direct binding of farnesol.131 FXR binds to DNA as an obligate heterodimer with RXR and prefers to bind to inverted repeats of the nuclear receptor half‐site AGGTCA separated by four basepairs (IR4).132,133 A second FXR subtype, FXRb (NR1H5), has been identified in rodents, rabbits, and dogs but is a pseudogene in humans and primates.133 FXRb will not be discussed in this review and we will refer to FXRa as simply FXR. The expression pattern of FXR is relatively restricted to the liver, intestine, kidney, and adrenal gland. Low levels of expression of the FXR mRNA have also been reported in adipose tissue, heart, and smooth muscle cells, however, it is not clear if FXR is functionally active in the latter three

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locations.133 Hepatic mRNA levels of FXR are elevated by prolonged fasting and by overexpression of the transcriptional coactivator PGC‐1a.134 PGC‐1a is an important transcriptional regulator of hepatic gluconeogenesis and energy metabolism135 suggesting that FXR may play a role in the response to nutritional status.

A. FXR and the Control of Bile Metabolism A number of studies have demonstrated that bile acids such as chenodeoxycholic acid (CDCA), deoxycholic acid (DCA), and cholic acid (CA) bind directly to the FXR and function as activators of FXR‐regulated genes.136–138 Importantly, FXR activation by bile acids leads to downregulation of Cyp7a1, the gene encoding the rate limiting enzyme (cholesterol 7a‐hydroxylase) in the conversion of cholesterol to bile acids.136–138 Inhibition of Cyp7a1 by FXR occurs indirectly via the FXR‐dependent upregulation of the small heterodimeric partner (SHP, NR0B2) a transcriptional repressor139 (Fig. 5). SHP lacks the canonical DBD found in most other members of the nuclear receptor superfamily but can dimerize with other receptors via a LBD. SHP interacts strongly with corepressors proteins so dimerization with SHP generally leads to

Liver FXR

FXR

Bile acid resin

Cholesterol

SHP

FGF19

CYP7A1

FGF19

7a-hydroxycholesterol

CYP8B1

AKR1D1 Intestine

CA

MCA CDCA

FIG. 5. Regulation of bile acid synthesis by FXR. In the liver, bile acids act as FXR agonists leading to the induction of SHP, a transcriptional repressor. SHP strongly represses the transcription of CYP8B1 and relatively weakly represses CYP7A1. The net result of activating FXR in the liver is a change in bile acid composition toward the more hydrophobic bile acids MCA (mice) or CDCA (humans). MCA and CDCA poorly promote the absorption of cholesterol in the intestine relative to CA. In the intestine, activation of FXR increases the expression of FGF19 which acts in a paracine fashion to strongly inhibit CYP7A1 in the liver and an overall decrease in bile acid synthesis. Bile acids resins sequester bile acids in the intestine, decreasing FXR activity, reducing the levels of FGF19, and consequently upregulating the levels of CYP7A1. See text for details.

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an inhibition of transcription.140 In the context of Cyp7a1, SHP dimerizes with and inhibits the liver receptor homolog 1 (LRH1, NR5A2), a third nuclear hormone receptor that functions as a constitutive activator of Cyp7a1.139 SHP also inhibits expression of the Cyp8b1 gene encoding sterol 12a‐hydroxylase.141,142 Sterol 12a‐hydroxylase sits at a branch point in the bile acid synthetic pathway that determines the polarity of primary bile acids (Fig. 5). When Cyp8b1 is active CA is the primary product; when Cyp8b1 is inactive CDCA (humans) and muricholic acid (MCA) (mice) are the predominant species.143 Along with regulation of bile acid synthetic enzymes via control of SHP expression, FXR induces the gene encoding FGF19 (FGF15 is the mouse ortholog) which also functions to inhibit expression of Cyp7a1144,145 (Fig. 5). FXR19 is expressed in the intestine, not the liver, suggesting that intestinal FXR senses the level of bile acids at this site and regulates hepatic bile acid synthesis in an endocrine fashion via the production of FGF19.144,145 FGF19 binds to FGF receptor 4 on the surface of hepatocytes and appears to inhibit Cyp7a1 by activating a JNK‐dependent145 or mitogen activated kinase (MAP)‐ dependent146 phosphorylation cascade. FGF19 also stimulates the normal refilling of the gallbladder with bile acids after cholecystokinin‐dependent emptying.147 Taken together these results suggest that when bile acid levels are high in the intestine that FXR functions to inhibit further synthesis and stimulate gallbladder filling. Interestingly, tissue‐specific knockouts of FXR indicate that the intestine is the dominant site for FXR‐dependent regulation of Cyp7a1 while the liver is the dominant site for regulation of Cyp8b1.148 Thus, activation of FXR in the intestine leads to an inhibition of bile acid synthesis while in the liver activation of FXR would primarily impact the composition of bile acids produced (Fig. 5). The consequence of altering bile acid composition on lipid metabolism will be discussed further below.

B. Gallstones, Cholestasis, and Bacterial Growth Bile comprised of bile acids, phospholipids, and cholesterol is stored in the gallbladder and released upon feeding. An excess of cholesterol to bile acids in the bile leads to the precipitation of cholesterol and formation of cholesterol gallstones. According to the American Gastroenterological Association, almost 24 million Americans suffer from gallstones and the condition can be quite painful. Along with controlling bile acid synthesis, FXR also regulates genes required for the hepatic uptake, conjugation, and excretion of bile acids.149–152 The bile salt export pump (BSEP), multidrug resistant‐associated protein 2 (MRP2, ABCC2), and the multidrug resistance P‐glycoportein 3 (MDR3, ABCB4) are localized on the canalicular membrane of hepatocytes and secrete bile acids from hepatocytes into the bile canaliculi.133 These three transporters are regulated directly by FXR and induced by FXR agonists.132,133 Moschetta et al.153 demonstrated that treatment with the synthetic FXR agonist

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GW4064 reduces the number of gallstones in a gallstone susceptible strain of mice. The beneficial effect of the FXR agonist most likely arises from increased expression of the canalicular bile acid transporters and increased transport of bile acids into the gallbladder. In contrast, FXR knockout mice are more susceptible to developing gallstones than FXRþ/þ mice when placed on a lithogenic diet.153 The current treatment for gallstones is surgical removal of the gallbladder (cholecystectomy) raising the possibility that FXR agonists could provide a novel nonsurgical treatment for this disease. Cholestasis is defined as any condition that impairs the flow of bile acids out of the liver and is many times associated with liver damage. Generally, cholestatic diseases are defined as obstructive (the flow of bile is physically blocked) or nonobstructive.154,155 Not surprisingly, GW4064 and 6‐ethyl CDCA, a derivative of CDCA that is a potent FXR agonist, has proven effective in several models of drug induced nonobstructive cholestasis.156–158 The activity of FXR agonists in these models most likely arises from their ability to induce the transport of bile acids out of the liver and to reduce further bile acid synthesis. Inflammation and fibrosis are commonly associated with liver damage and FXR agonists also reduce the expression of markers of fibrosis and inflammation perhaps via induction of the transcriptional repressor SHP as described for Cyp7a1.159–161 As we have seen with the PPARs and LXRs, regulation of metabolism by FXR appears to be intimately linked with the control of inflammation. Obstruction of bile flow is also associated with intestinal bacterial growth and injury to the intestinal mucosa. Fxr
/
mice have bacterial overgrowth in the ileum and compromised epithelial barrier most likely resulting from impaired bile flow. On the other hand, FXR activation induces a number of genes in the intestine involved in enteroprotection including angiogenin, nitric oxide synthase, and IL‐18.162

C. FXR and Lipid Metabolism Clinical trials examining the utility of bile acids for the treatment of cholesterol gallstones first demonstrated that increasing bile acid levels leads to a corresponding decrease in plasma triglycerides.163–165 The observation that Fxr
/
mice have elevated triglyceride levels166 and that a synthetic FXR agonist lowers triglycerides in rats167 supports an essential role for FXR in mediating the effects of bile acids on triglyceride metabolism. Indeed, FXR has been identified as a direct regulator of a number of genes encoding proteins involved in triglyceride synthesis and catabolism including apoCII, apoCIII, apoAV, FGF19, SREBP1c, syndecan 1, the VLDL receptor, complement C3/acylation stimulating protein, Insig‐2, and PPARa.144,150,151,168–172 The contribution of these different mechanisms to FXR‐dependent control of triglycerides remains to be determined.

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While a role for FXR in the control of triglyceride metabolism appears to be well established, the effect of FXR on cholesterol homeostasis is less clear. Repression of Cyp7a1 by FXR should inhibit the conversion of cholesterol to bile acids resulting in an increase in cholesterol levels (Fig. 5). In support of this prediction, humans with a genetic deficiency in CYP7A1 present with elevated cholesterol levels.173 Nevertheless, no evidence for increased cholesterol was observed in humans treated with bile acids165 and genetic knockout of FXR results in an increase in cholesterol levels174,175; not the expected decrease predicted if feedback inhibition of Cyp7a1 is eliminated. Similarly, studies in animals and humans have shown that bile acids repress the production of VLDL.176,177 As described earlier, activation of FXR in the liver will strongly inhibit Cyp8b1 expression.141,142 Sterol 12a‐hydroxylase, the enzyme encoded by Cyp8b1, sits at a branch point in the bile acid synthetic pathway and its enzymatic activity is required for the synthesis of CA. The parallel arm in the pathway leads to synthesis of MCA in mice and CDCA in humans.143 Thus, modulating expression of Cyp8b1 expression alters bile composition (Fig. 5). Importantly, individual bile acids differ in their ability to promote intestinal cholesterol absorption and MCA, of all bile acids tested, promotes the lowest amount of cholesterol absorption while CA promotes the greatest amount.178 CDCA is in between CA and MCA 178. Thus, activating FXR in the liver should decrease cholesterol absorption in the intestine. On the other hand, eliminating FXR activity in the liver with antagonists or by genetic knockout should increase cholesterol absorption by the intestine (Fig. 5). Thus, we would argue that many of the effects of modulating FXR activity on lipid levels may result from influencing the absorption of fat and cholesterol in the intestine.

D. FXR and Atherosclerosis The ability of FXR to control triglyceride and cholesterol metabolism suggests that activating or inhibiting this receptor should have significant effects on the development and progression of atherosclerosis. Indeed, treatment of apoE
/
mice with 6‐ethyl CDCA, a derivative of CDCA that functions as an FXR agonist, reduces atherosclerosis.179 Based on this data and on studies demonstrating elevated triglyceride and cholesterol levels in Fxr
/
mice, one would predict that eliminating FXR activity in a proatherosclerotic mouse background would increase atherosclerosis. The predicted result of increased atherosclerosis in apoE
/
/Fxr
/
mice was observed in one published study.174 Two other studies, however, one using apoE
/
/Fxr
/
180 mice and the other using Ldlr
/
/Fxr
/
181 mice detected decreased atherosclerosis compared to apoE
/
and Ldlr
/
controls. Interestingly, both of the studies that measured decreased atherosclerosis detected changes in plasma lipid levels that one would expect to actually promote atherosclerosis.180,181 Both studies also detected decreased expression of CD36 in macrophages isolated

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from double knockout mice. CD36 is a scavenger receptor responsible for the uptake of oxidized cholesterol by macrophages and decreased expression would be expected to reduce atherosclerosis by limiting macrophage foam cell development.182 FXR is not expressed in macrophages and both studies suggest that alterations in lipid profiles that result from deleting FXR may influence the expression of CD36 in peripheral cells.180,181 In the study using the Ldlr
/
background, the authors also detected a significant increase in VLDL cholesterol and decreases in LDL and HDL.181 This change in lipoprotein particle profile, particularly the decrease in LDL, may also contribute to the decrease in atherosclerosis observed in this study. The different results seen among the various atherosclerosis studies again highlight the care that needs to be taken in interpreting such experiments. Furthermore, these studies clearly illustrate that it can be difficult to predict the activity of a nuclear receptor agonist or antagonist from the phenotype of a knockout.

E. FXR and Diabetes Analysis of Fxr
/
mice has demonstrated impaired glucose tolerance and insulin resistance in the liver and in peripheral tissues (muscle and fat).183,184 The effects on insulin resistance most likely arise from the elevated fatty acid levels observed in these mice. Similarly, the synthetic FXR agonist GW4064 improves insulin sensitivity in several diabetic models including db/db, ob/ob, and KK‐A(y) mice.185 Infection of these mice with an adenovirus expressing a ‘‘super‐active’’ VP16–FXR construct also improves insulin sensitivity suggesting that the liver is the major site of FXR activity with regards to diabetes.185 Examination of these diabetic models suggests that activation of FXR reduces hepatic gluconeogenesis and increases glycogen synthesis thus reducing plasma glucose levels.183–185 The effects of FXR on gluconeogenesis and glycogen synthesis are somewhat surprising given the report that FXR is induced by prolonged fasting,134 a condition when gluconeogenesis would be expected to be high and glycogen synthesis low. The mechanistic basis for these effects remains to be determined.

F. Control of Liver Regeneration and Tumorigenesis by FXR Recent studies indicate that deletion of FXR in mice impairs the process the liver regeneration after hepatectomy186 and increases the incidence of hepatocellular adenoma, carcinoma, and hepatocholangiocellular carcinoma in older (12 months) animals.187,188 Opposite results are observed when wild‐ type mice are fed diets enriched in bile acids; 187,188 however, to our knowledge

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synthetic agonists have not been examined in these models. It appears that the ability of FXR to control genes involved in inflammation and cell cycle control accounts for these observations.186–188

G. Therapeutic Potential of FXR Ligands Studies in humans with bile acid resins such as cholestyramine that absorb bile acids indicate that depletion of bile acids decreases cholesterol levels.189 The activity of bile acid resins most likely results from the upregulation of CYP7A1 and increased catabolism of cholesterol to restore the bile acids that have been removed (Fig. 5). While these resins are an effective means to lower cholesterol compliance is generally poor. Bile acid resin powders must be mixed with water or fruit juice and taken once or twice daily with meals. Tablets must be taken with large amounts of fluids to avoid gastrointestinal symptoms. Resin therapy may also produce a variety of side effects including constipation, bloating, nausea, and gas. The observation that FXR controls the bile acid‐ dependent repression of CYP7A1 suggested to many that an FXR antagonist would block bile acid‐dependent repression of CYP7A1 and produce the benefits of a bile acid resin without unwanted side effects.132 Guggulsterone, the active ingredient of a naturally occurring medicinal cholesterol lowering agent isolated from the guggul tree, has been reported to be an FXR antagonist.190 The putative cholesterol‐lowering activity of guggulsterone is consistent with the predicted activity of an FXR antagonist. Guggulsterone, however, appears to have multiple biological targets and it is not clear if any of the physiological effects of this compound are derived from inhibiting FXR.190–192 In contrast, all the activities associated with FXR agonists including lowering lipid levels, decreasing atherosclerosis, improving insulin sensitivity, and hepatoprotection appear to be beneficial. We would suggest that the apparent disconnect between the activities of bile acid resins (upregulate CYP7A1, lower cholesterol) and FXR agonists (downregulate CYP7A1, lower cholesterol) lies in the site of the action of these two agents. Bile acid resins function in the intestine and remain there for extended periods. Depleting bile acids in the intestine will decrease the expression of FGF19, relieve the feedback inhibition on CYP7A1, and promote the breakdown of cholesterol (Fig. 5). In contrast, the synthetic agonists tested are relatively rapidly absorbed into the blood stream and accumulate in the liver. In the liver, activation of FXR will induce SHP and strongly inhibit expression of CYP8B1 shifting bile acid composition toward MCA/CDCA and limiting cholesterol absorption in the intestine (Fig. 5). Activation of FXR in the liver has the added benefits of promoting bile flow, and protecting against tissue damage. Thus, all the preclinical studies suggest that FXR agonists have the ideal therapeutic profile for the treatment of human disease. Currently, 6‐ethyl CDCA is in phase II clinical trial for the

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treatment of cholestasis and Wyeth has completed a Phase I clinical study of a synthetic FXR agonist (information on ClinicalTrials.gov). The data from these studies are not currently available.

V. RORa The RAR‐related orphan receptor (ROR) family consists of three members (RORa, NR1F1; RORb, NR1F2; and RORg, NR1F3) that bind to DNA as monomers to regulate transcription.193–195 ROR response elements (ROREs) consist of a six basepair A/T rich region immediately preceding a half site AGGTCA motif. RORs are constitutively active transcription factors which recruit coactivators in the absence of exogenous ligand.196–199 Nevertheless, crystal structures suggest that these orphan receptors can bind and be regulated by ligands. All‐trans retinoic acid was crystallized with LBD of RORb and can inhibit RORb transcriptional activity in cells, suggesting a possible role of RORb in regulating retinoid action.200 Interestingly, a structure of RORa obtained using baculovirus‐expressed protein unexpectedly identified cholesterol as a ligand for RORa.201 This suggests that cholesterol or derivatives of cholesterol might function as physiological ligands for RORa. Based on this structure, point mutations of RORa that prevent cholesterol binding were generated and the ability of these mutants to activate a reporter gene with an RORE was impaired.201 Further confirmation of cholesterol as a RORa ligand was provided by decreased RORa activity in cells treated with lovastatin or hydroxypropyl‐B‐cyclodextrin to lower intracellular cholesterol. Under these cholesterol‐depleted conditions, adding back cholesterol restores RORa activity.201 Although it cannot be ruled out that the effect of cholesterol is indirect, these results demonstrate that RORa activity can be modulated by cholesterol status in cells. Thus, RORa may act as a lipid sensor of the intracellular cholesterol levels and play an important role in the regulation of cholesterol homeostasis.

A. Regulation of Lipid Metabolism by RORs RORa is highly expressed not only in blood, brain, skin, intestine but also in metabolically important tissues such as liver, muscle, and adipose tissue.202 The physiological function of RORa in vivo has been revealed using staggerer mice (RORasg/sg). These mice contain a 122‐bp deletion in the coding sequence of RORa that prevents translation of a functional LBD.203,204 The most obvious defect in RORasg/sg mice is their balance deficit due to massive neurodegeneration in the cerebellum, caused by a developmental defect in Purkinje cells.

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This cerebellar atrophic phenotype was also found in RORa knockout (RORa
/
) mice, providing validation that the RORasg/sg defects are due to the functional disruption of RORa gene.205 The role of RORa in lipid homeostasis and atherosclerosis/obesity was uncovered by the observation of metabolic abnormalities in RORasg/sg mice. RORasg/sg mice are smaller than wild type despite hyperphagia and have lower total and HDL cholesterol levels. The decrease in HDL is thought to arise from reduced plasma ApoA‐I and ApoA‐II, both major apolipoproteins of HDL, in these mice and from decreased expression of ApoA‐I in the intestine, previously shown to be a direct target gene for RORa.206 The reverse cholesterol transporters ABCA1 and ABCG1 genes are also downregulated in the liver from RORasg/sg mice.207 Not surprisingly given the low levels of HDL, when fed with an atherogenic diet RORasg/sg mice are more susceptible to develop atherosclerosis compared to wild‐type mice.208 Plasma triglycerides levels from RORasg/sg mice are reduced and this is correlated with decreased hepatic and intestinal expression of ApoC‐III, a component of HDL and VLDL that regulates triglyceride levels and is also a RORa target gene.209 Interestingly, a recent genome‐wide association study indicated that individuals who are heterozygous for a null mutation in ApoC‐III have low plasma triglycerides, a favorable cholesterol profile and decreased risk for cardiovascular disease.210 Consistent with the lowering of plasma triglycerides, liver triglycerides are also found to be decreased in RORasg/sg mice.207 Examination of genes involved in lipogenesis in the liver show that SREBP‐1c and FAS expression are reduced with little change in LXR expression, suggesting factors other than LXR may be involved in regulating expression of these genes. Using transfection and chromatin immunoprecipitation assays, RORa was shown to bind to the SREBP‐1c promoter and modulate its activity. Thus, the decreased triglyceride levels detected in staggerer mice may be a direct result of RORa deficiency. Perhaps related to the defect in triglyceride metabolism, RORasg/sg mice have reduced adiposity associated with decreased fat pad mass and adipocyte size207 and are resistant to diet‐induced weight obesity. Compared to wild‐type mice, there is a lack of fat accumulation in the gonadal and inguinal white adipose and interscapular brown adipose of RORasg/sg mice on high fat diet.207 These results further demonstrate the importance of RORa in lipogenesis in the adipose tissue and liver. In muscle, overexpression of a dominant negative RORa that lacks the entire LBD and part of the hinge region results in a reduced expression of SREBP‐1c and other lipogenic genes (FAS, SCD‐1, SCD‐2). Similarly, genes involved in lipid and cholesterol efflux and homeostasis (ABCA1, apoE, CAV3) are repressed in these cells versus the wild‐type cells. Other genes involved in lipid absorption (CD36, FABP3), lipid catabolism (LPL, CPT1, ACS4, adipoR1,

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adipoR2, PGC‐1), lipid storage (ADRP) are also downregulated.211 Collectively, these in vivo and in vitro studies demonstrate that RORa is an important modulator of lipid homeostasis in the intestine, liver, adipose tissue, and muscle. The involvement of the other two ROR family members, RORb and RORg, in lipid metabolism is less clear. RORb expression is restricted mostly to brain and retina and expressed poorly in other tissues such as liver, muscle, and adipose tissue.194,202,212 RORg is highly expressed in muscle and its function in lipid and glucose metabolism has been examined using the in vitro muscle cell model. In these studies, although expression of a dominant negative RORg that lacks AF‐2 domain significantly represses RORg‐dependent gene expression, genes involved in lipid metabolism are not affected.213 Similarly, RORg‐deficient mice have normal triglycerides and cholesterol levels.214 Interestingly, serum glucose levels are significantly lower in RORg‐deficient mice but normal in staggerer mice, suggesting that RORs have different roles in lipid and glucose homeostasis.214 Rev‐erba is an orphan nuclear receptor which can bind ROREs and act as a dominant‐negative repressor of RORs by competing for the same DNA binding sites.215 Consistent with the idea that RORs and Rev‐erba compete for binding sites is the observation that Rev‐erba can repress ApoA‐I and ApoC‐III gene expression in HepG2 cells and primary hepatocytes.216–218 In Rev‐erba‐ deficient mice, serum triglycerides are elevated and this is associated with the increases in serum apoC‐III concentration and liver apoC‐III RNA levels.217 Since RORs and Rev‐erbs are coexpressed in many tissues, the effects of these receptors on lipid metabolism are likely to be determined by the ratio of their expression levels, which can be altered by diet or circadian rhythm (see below).

B. Role of RORs in Circadian Rhythm Control and Links to Metabolism There is increasing evidence suggesting that regulation of the biological clock is linked to energy metabolism. Mutation of the CLOCK (Circadian locomotor output cycles kaput) gene, a key transcription factor of the molecular circadian clock, results in a phenotype of metabolic disease in mice.219 Homozygous Clock mutant mice lose the diurnal rhythm in food intake and their metabolic rates are decreased. They gain more weight than wild‐type mice and have higher triglycerides, cholesterol, and glucose in the serum and accumulation of fat in the liver, all hallmarks of metabolic syndrome. Conversely, a high fat diet which induces the same metabolic phenotypes also alters the cyclic expression of genes important for maintaining circadian rhythm,202 emphasizing the interrelationship of metabolic homeostasis and molecular circadian clock. Genomic profiling of suprachiasmatic nuclei (SCN) and liver has identified an important role of the RORE in gene expression during circadian cycle. This implies that RORs and Rev‐erba may be important transcriptional circadian

29

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Relative RNA levels

regulators.220 Indeed, RORs and Rev‐erba display circadian expression in the SCN and peripheral tissues important for metabolism (liver, muscle, WAT, BAT).202,220 In these tissues, Rev‐erba and Rev‐erbb show similar circadian pattern with highest level at CT (circadian time) 4–8 and lowest at CT16–20. Expression of different ROR isoforms is more tissue‐specific. In the SCN, RORg is not detected and RORb expression is high but shows little oscillation whereas RORa expression is cyclic with highest level at CT4–12. In the WAT, RORa expression peaks at CT12, right after the peaks of Rev‐erbs expression. In the liver and BAT, RORg expression reaches its maximal level at CT16 and minimal level at CT4, showing an antiphase pattern to Rev‐erbs expression. Interestingly, expression of Bmal1 (Brain and muscle ARNT like protein 1), another important gene involved in regulating circadian clock,221 generally reaches the lowest level and highest level in both SCN and metabolically important tissues (liver, muscle, fat) at the peak and trough of Rev‐erbs expression, respectively (Fig. 6).202,220 Bmal1 gene contains ROREs in its promoter region and is likely be regulated by RORs and Rev‐erbs. Indeed, Bmal1 has been shown to be a target gene of RORa and RORg and its transcription is activated by RORs and repressed by Rev‐erbs through competing for the same ROREs (Fig. 6).222–224 The importance of Rev‐erba and RORa in Bmal1 transcription and circadian rhythm is further demonstrated in mice deficient in these orphan nuclear receptors. In wild‐type mice the amplitudes of Bmal1 RNA oscillations in the

Bmal1

Rev-erbs

0

4

RORs

8

12

16

20

24 CT

Rev-erbs

ROREs

Bmal1

FIG. 6. Schematic representation of circadian Bmal1 expression and its regulation by RORs and Rev‐erbs. RORs and Rev‐erbs bind to the ROREs in the promoter of Bmal1 gene and activate or repress its transcription depending on the relative expression levels of these two classes of orphan nuclear receptors. Both Rev‐erba and Rev‐erbb cycle in an antiphase to Bmal1 in all tissues (SCN, liver, muscle, adipose) whereas three ROR isoforms exhibit different circadian patterns in different tissues. Decrease of Rev‐erbs expression in cells may allow the RORs to function as an activator which is inhibited when Rev‐erbs levels are high.

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liver are at least 20‐fold with lowest point at CT8. This cyclic change in Bmal1 expression is reduced to less than 2‐fold in Rev‐erba
/
mice.225 Similarly, the dramatic decrease of Bmal1 expression in the SCN at CT4 compared to that at CT16 is also lost in these mice. The Rev‐erba
/
mice have shortened period lengths of their locomotor activity rhythm in constant darkness or light, and the distribution of period lengths is much more variable than in wild‐type mice. Interestingly, the more variable and shortened period of locomotor activity rhythm is also seen in the RORasg/sg mice.222,223 In the SCN of the RORasg/sg mice, the peak Bmal1 expression at CT15–18 is much reduced compared to wild‐type mice. These studies suggest that RORa and Rev‐erba are required for maintaining normal cyclic Bmal1 expression and circadian clock function. The RORasg/sg and Rev‐erba
/
mice, as discussed earlier, have abnormal triglycerides levels. Plasma triglycerides in mice show circadian rhythm with highest level at CT4 and lowest level at CT16.226 This cyclic change in triglycerides is lost in Bmal1
/
mice, with similar levels at CT4 and CT16. It is possible that Bmal1 expression, regulated by RORs and Rev‐erbs, may be required for lipid homeostasis. Mouse embryonic fibroblasts (MEFs) isolated from Bmal1
/
mice and Bmal1‐knockdown 3T3‐L1 cells are resistant to adipocyte differentiation and accumulate less lipids.227 This is correlated with a significantly reduced induction of adipocyte‐related genes such as PPARg and aP2. Reexpression of Bmal1 in Bmal1
/
MEFs can restore the adipocyte gene expression and lipid accumulation. In mature 3T3‐L1 adipocytes, overexpression of Bmal1 increases lipid synthesis activity which is associated with an increase in the RNA levels of genes involved in lipid metabolism such as SREBP‐1a, FAS, and aP2. Interestingly, the RORasg/sg mice have reduced adiposity and decreased expression of lipogenic genes.207 The similarity of these lipogenic deficiency phenotypes in Bmal1
/
and ROR
/
cells provides another link between circadian rhythm and lipid metabolism.

C. RORs and Inflammation As we have observed with the other metabolic sensing nuclear receptors, RORa also appears to play an important role in regulating inflammation. In the immune system, RORa is expressed in both lymphoid and myeloid cells with highest expression in macrophages.228 Peritoneal macrophages isolated from the RORasg/sg mice express higher RNA levels of IL‐1b, TNFa, and IL‐1a than wild‐ type mice upon LPS stimulation.229 In Rora
/
mice, bone marrow derived macrophages also produce more IL‐6 and TNFa than those obtained from wild‐type mice.228 Since inflammation is also a key element in atherosclerosis, the activity of RORa was examined in primary human aortic smooth muscle cells. RORa is expressed in these cells and overexpression of RORa inhibits TNFa‐ induced IL‐6, IL‐8, and COX‐2 expression.230 In addition, RORa upregulates

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gene transcription of IkBa, the major inhibitory protein of the NF‐kB signaling pathway, through a RORE in the IkBa promoter in these primary muscle cells. The role for RORa in regulating IkBa was confirmed in vivo by the significantly lower basal levels of IkBa mRNA in aortas from the RORasg/sg mice than the wild‐ type mice. These studies suggest that RORa is anti‐inflammatory and may have a protective role during inflammation and metabolic disease.

D. Therapeutic Potential of RORa Ligands Studies in vivo and in vitro have demonstrated the importance of RORa in regulating expression of genes that control cholesterol and triglyceride levels as well as circadian rhythm. Recent studies linking the circadian rhythm to energy metabolism and demonstrating the essential role of ROREs in circadian gene expression suggest that RORs and their dominant negative regulators Rev‐erbs are likely to be important players in integrating the circadian clock with lipid metabolism. Identification of cholesterol and ATRA as RORa and RORb ligands suggests that it may be possible to develop small molecules that can bind RORs and modulate their activities. Based on the atherosclerotic and inflammatory phenotypes of RORasg/sg mice, a RORa agonist will likely to be beneficial in improving cardiovascular disease. Nevertheless, an RORa agonist may also potentially increase lipid synthesis in the adipose and liver. As we have observed with the other receptors discussed in the review, the challenge for drug discovery targeting RORa will be to identify small molecules that function in a tissue‐selective manner in order to maximize the therapeutic benefit relative to unwanted side effects.

VI. ERRa Estrogen receptor related receptora (ERRa; NR1B1) is one of the three members of the ERR family (ERRa, ERRb, NR1B2; ERRg, NR1B3) that bind to the ERR responses element (ERRE) composed of a nuclear receptor half‐site preceded by three nucleotides with the consensus sequence TNAAGGTCA.231 ERRa is most related to estrogen receptor a (ERa), sharing a high degree of homology within their DBD (68% identity). Not surprisingly, ERRa also bind estrogen receptor response elements (ERE) consisting of an inverted repeat of the core AGGTCA separated by three nucleotide.232 Indeed, several ERE‐containing gene promoters can be regulated by both ERa and ERRa,232–234 suggesting potential cross talk between the signaling pathways mediated by these two receptors. ERRa is constitutively active, assuming a LBD conformation that resembles that of agonist‐bound nuclear receptors in the absence of ligand.235,236 A crystal structure of ERRa complexed with an LXXLL peptide from PGC‐1a

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indicates that the LBD of ERRa is almost completely filled with hydrophobic side chains, resulting in a small ligand binding pocket which can only be occupied by compounds with no more than the equivalent of four or five carbon atoms. Based on the sequence similarity between ERRa and ERa, a series of estrogen‐like compounds were screened for their ability to modulate ERRa transcriptional activity. The phytoestrogens geneistein, daidzein, and biochanin as well as 6,30 ,40 ‐trihydroxyflavone were identified as agonists of ERRa by a virtual ligand screening using the agonist‐bound ERa as the template.237 The activity of these phytoestrogens on ERRa was confirmed in cell transfection assays but they also activate ERs and other ERRs. Using a yeast one‐hybrid assay with an ERRa‐binding DNA region from the human aromatase gene, two organocholorine (pesticides toxaphene and chlordane) were identified to interact with ERRa.238 These compounds exhibit weak agonist activity on ERa but inhibit transcriptional activity of ERRa in mammalian cells. Diethylstilbesterol (DES), a potent ER agonist, was also identified as an antagonist of ERRa with an IC50 at 1 mM using an in vitro coactivator interaction assay.239 In cells, DES disrupts the ERRa‐coactivator interaction and inhibits the constitutive transcriptional activity of ERRa. The first synthetic ERRa ligand XCT790, and inverse agonist, was derived from high throughput screening using an in vitro coactivator interaction assay.240,241 This compound is a potent inhibitor of ERRa transcriptional activity in cells (IC50: 300–500 nM) with no significant activity on related nuclear receptors such as ERRg and ERa at concentrations below 10 mM, providing a pharmacological tool for understanding the functions of ERRa. Recently, another ERRa inverse agonist was derived from high throughput screening and crystallized with ERRa LBD.242 This compound inhibits the interaction of ERRa and PGC‐1a peptide in a biochemical assay with an IC50 of 190 nM. Binding of this antagonist positions Helix 12 in the coactivator binding groove where the PGC‐1a peptide is located in the apo‐ERRa crystal structure,236 providing an explanation for the molecular mechanism of an ERRa inverse agonist. The identification of numerous ERRa inverse antagonists and few agonists suggest that the small ligand binding pocket is likely to accommodate only tightly packed molecules and most compounds that bind will result in a disruption of the ERRa transcriptional activity.

A. ERRa and the PGC‐1a Pathway ERRa is ubiquitously expressed with higher levels in the tissues (muscle, kidney, heart, and BAT) that have high metabolic needs.243,244 This expression pattern of ERRa is very similar to that of PGC‐1a (which has been shown to induce ERRa gene expression through the ERRE in the promoter region and also function as an ERRa coactivator).245,246 Physiological stimuli such as

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starvation, exposure to cold temperatures, and exercise increase both PGC‐1a and ERRa gene expression,244,246,247 suggesting that these two proteins may mediate similar cellular functions. PGC‐1a plays an important role in regulating energy metabolism and mitochondrial biogenesis in muscle and BAT.248,249 In the course of identifying transcription factors and their cis‐regulatory elements that are important in mediating the activities of PGC‐1a, ERR binding sites (ERREs) were found to be one of the key motifs enriched in the genes upregulated by PGC‐1a in muscle cells.250 This suggests that an important function of PGC‐1a in muscle is to modulate transcriptional activity of ERRs. The ERRE motif is highly enriched in the promoters of genes involved in oxidative phosphorylation which are also targeted by PGC‐1a.249 The role of ERRa in the regulation of PGC‐1a‐mediated mitochondrial functions was demonstrated by using the ERRa‐specific antagonist XCT790.250 In C2C12 myotube cells, PGC‐1a increases expression of ERRa and genes involved in oxidative phosphorylation but XCT790 diminishes this induction. XCT790 also significantly reduces the PGC‐1a‐increased total mitochondrial respiration. The requirement of ERRa in the PGC‐1a mediated mitochondrial biogenesis is demonstrated in an osteosarcoma cell line SAOS2.251 Overexpression of PGC‐1a in these cells induces genes encoding mitochondrial proteins such as mitochondrial DNA replication and transcription (mtTFA), cytochrome c, somatic (Cyt c), and ATP synthase b (ATPsynb), and mitochondrial DNA contents are also increased. Inhibition of ERRa expression using an siRNA approach results in the decrease in expression of these mitochondrial proteins and functions mediated by PGC‐1a. In these studies, ERRa is also shown to bind to the ERREs in the promoters of ATPsynb and Cyt c genes. PGC‐1a also plays a role in regulating glucose and fatty acid metabolism in an ERRa‐dependent manner. Overexpression of PGC‐1 in C2C12 myotube cells induces gene expression of pyruvate dehydrogesnase kinase 4 (PDK4), which contains an ERRE in the promoter region.252 PGC‐1a‐mediated activation of the PDK4 promoter requires ERRa since this activity is not seen when using MEFs from Erra
/
mice compared to those from the wild‐type mice. Stimulation of PDK expression suggests that glucose oxidation is inhibited and energy utilization is switched from glucose to fatty acids. Indeed, PGC‐1a expression in C2C12 myotube cells decreases the glucose oxidation rate and is accompanied by an increase in the expression of median‐chain acylcoenyme A dehydrogenase (MCAD), an enzyme that mediates the first step in the mitochondrial b‐oxidation of fatty acids. MCAD is a target gene of ERRa and coexpressed with ERRa and PGC‐1 in tissues that preferentially utilize fatty acids as energy substrates.243 The PGC‐1‐induced MCAD expression can be inhibited using the ERRa‐specific inhibitor XCT790,250 further demonstrating the requirement of ERRa in PGC‐1‐mediated fatty acid oxidation.

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Another metabolic function of PGC‐1a affected by ERRa is its activity on gluconeogenesis in the liver. Overexpression of PGC‐1a in primary hepatocytes increases expression of gluconeogenic enzymes phosphoenolpyruvate carboxykinase (PEPCK) and glucose‐6‐phosphatase (G6Pase) and results in increased glucose production.253 Injection of an adenovirus expressing PGC‐1a into rats also increases blood glucose levels, associated with increases in PEPCK and G6Pase expression. Similarly, PGC‐1a also increases PEPCK gene expression in HepG2 cells and this is accompanied by an increase in ERRa expression as predicted.254 However, coexpression of ERRa decreases this PGC‐1a‐ mediated induction of PEPCK which contains an ERRE in its promoter region, suggesting ERRa functions as a transcriptional repressor of PGC‐1a in this context. Indeed, inhibition of ERRa expression using a siRNA increases PGC‐1a‐induced PEPCK expression in HepG2 cells. The inhibitory activity of ERRa on PGC‐1‐mediated gluconeogenesis is further confirmed by the significantly increased expression of PEPCK and glycerol kinase genes in livers from the Erra
/
mice compared to the wild‐type mice. Interestingly, ERRa still functions as an activator of PGC‐1a genes involved in the oxidative phosphorylation in the liver since the PGC‐1a‐induced mitochondrial gene expression in HepG2 cells is not activated but inhibited by ERRa siRNA. This suggests that ERRa represses gene expression in a promoter‐specific manner and is not a general inhibitor in the liver.

B. ERRa and Diabetes PGC‐1a and the genes involved in the oxidative phosphorylation are decreased in human diabetic muscles, suggesting that muscle mitochondrial dysfunction is linked to diabetes.255 PGC‐1a is also increased in the livers of diabetic mouse models, supporting the hypothesis that increased glucose production stimulated by PGC‐1a has a negative impact on insulin sensitivity.253 The ability of PGC‐1a to increase expression of genes involved in both mitochondrial oxidative phosphorylation in muscle and gluconeogenesis in liver suggest that tissue‐specific modulation of PGC‐1a will be needed to achieve a net beneficial effect for the treatment of diabetes. However, the ability of ERRa to activate PGC‐1a‐induced oxidative phosphorylation in muscle and repress PGC‐1a‐mediated gluconeogenesis in liver (as discussed earlier) presents an opportunity to increase mitochondrial function without the detrimental effect on glucose production by targeting ERRa.

C. ERRa in Adipose and Intestine Erra
/
mice have a 50–60% reduction in the weight of the inguinal, epididymal, and peritoneal fat pads with decreased adipocytes size.256 These mice are also resistant to high fat diet‐induced obesity. Microarray analysis of

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the adipose tissues from ERRa
/
mice show downregulation of the genes involved in fatty acid synthesis such as FAS, and SREBP1. Lipogenesis in the Erra
/
mice assessed by treating with 3H2O and measuring the amount of radioactive label incorporated into triacylglycerol show that there is a 30–50% decrease in 3H incorporation into lipids in the inguinal, epidymal, and perirenal fat. These results indicate that ERRa is important for normal lipid metabolism in the white adipose tissue. In the brown adipose tissue (BAT) of Erra
/
mice, expression of mitochondrial genes important for oxidative phosphorylation and fatty acid oxidation is decreased, demonstrating the importance of ERRa in mediating mitochondrial energy metabolism in this tissue.257 Mitochondrial density and DNA contents are also decreased in the BAT of Erra
/
mice compared to the wild‐type mice, confirming the in vivo function of ERRa in mitochondrial biogenesis. The physiological significance of the decreased mitochondrial function and mass is shown by the inability of Erra
/
mice to maintain proper body temperature when exposed to cold, suggesting a defect in adaptive thermogenesis. The lean phenotype of Erra
/
mice arises without decreased food consumption or increased energy expenditure.256 One possible explanation for the phenotype of these mice is a defect in intestinal adsorption of dietary nutrients including fat. ERRa is expressed in epithelia cells throughout the small intestine244,258 and fat malabsorption is observed in the Erra
/
pups.256 Microarray analysis of intestinal RNA from Erra
/
mice show that genes involved in oxidative phosphorylation are downregulated and intestinal b‐oxidation activity is also reduced, confirming the in vivo role of ERRa in mitochondrial metabolism in this tissue.258 Additionally, expression of genes involved in dietary lipid digestion and absorption such as pancreatic lipase‐related protein 2 (PLRP2), fatty acid‐binding protein 1 and 2 (L‐FABP and I‐FABP), and apolipoprotein A‐IV (apoA‐IV) are also downregulated in the intestine of Erra
/
mice.256

D. ERRa and Cancer The observation that ERRs and ERs can bind to the same DNA sequences suggests that ERRs may influence well characterized estrogen‐dependent pathways. Interestingly, ERRa has been identified as a negative prognostic factor for breast cancers. ERRa is expressed in many breast cancer cell lines including the ER‐positive MCF‐7 and ER‐negative MDA‐MB‐231 cells. Treatment of these cell lines with high concentrations of DES, which antagonizes ERR activity, inhibits cell proliferation.234 Knockdown of ERRa expression in the MDA‐MB‐231 cells using siRNA also results in a reduction in cell migration in vitro and tumor growth when implanted as xenografts into athymic

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nude mice.259 Recently, the angiogenic vascular endothelial growth factor (VEGF) was identified as a direct target gene for ERRa, providing another evidence linking ERRa to tumor progression and metastasis.260 These results suggest that pharmacological manipulation of ERRa activity may be of therapeutic use in treating breast cancer patients.

E. Therapeutic Potential of ERRa Ligands Although Erra
/
mice exhibit a relatively mild metabolic phenotype this could be due to the upregulation of compensatory pathways. Indeed, expression of PGC‐1a is increased in the heart, liver, and muscle of Erra
/
mice and ERRg is also upregulated in the heart of these mice.254,261 Several studies indicate mitochondrial oxidative phosphorylation is reduced in patients with type II diabetes255,262 and ability of ERRa to induce mitochondrial biogenesis and oxidative phosphorylation in conjunction with PGC‐1a suggests the straight forward hypothesis that an ERRa agonist would be useful for the treatment of this disease (Fig. 7). Nevertheless, this simple hypothesis is confounded by the recent observation that over expression of PGC‐1a in muscle actually promotes insulin resistance.263 Additionally, to our knowledge none of the compounds reported to have ERR agonist activity in vitro and in cell culture systems have been shown to act as ERR agonists in animals. Hydrophobic amino acid residues in the LBD of ERRa appear to function as an intramolecular agonist favoring a transcriptionally active conformation and we feel it is unlikely that synthetic agonists with the efficacy to significantly influence gene expression in animals will be identified. On the contrary, ERRa antagonists that disrupt the active confirmation have been identified240–242,250 and we anticipate that the further development of such inhibitors with sufficient systemic exposure will help elucidate the in vivo function of ERRa. Clearly, such inhibitors may have therapeutic value for the treatment of cancer and one wonders about the metabolic impact of such ligands. Will inhibiting ERRa function to decrease

Gluconeogenesis (e.g., PEPCK) Fatty acid oxidation (e.g., MCAD) Glucose oxidation (e.g., PDK4) Lipid transport (e.g., ApoA-IV)

ERRa

Tumor growth (e.g., pS2) Angiogenesis (e.g., VEGF)

Oxidative phosphorylation (e.g., Cyt c)

FIG. 7. ERRa activity and its target genes. ERRa binds to the ERREs in the promoters of its target genes and activates or represses their expression. The mitochondrial functions and other activities of ERRa have been characterized using a combination of the Erra
/
mice, siRNA, and ERRa‐specific antagonists.

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lipid absorption perhaps having a beneficial effect on plasma lipids or will decreases in oxidative phosphorylation speed the progression of diabetes (Fig. 7)?

VII. Summary Over the last 15 years a growing number of nuclear hormone receptors have been demonstrated to play important roles in the control of metabolism at the level of gene expression. Nature has seized on the ability of nuclear receptors to function as ligand‐controlled transcription factors and exploited this ability to evolve a family of metabolic sensors that coordinate metabolism in response to changes in the levels of important end‐products and intermediates. Recent studies have also defined an intimate link between the regulation of metabolism and the control of inflammation. Excitingly, many of the receptors highlighted in this review appear to function at this interface. Although the biological necessity for linking metabolism with inflammation is not yet clear, the best evidence suggests that control of energy utilization plays an important role in determining inflammatory status. Since many of the pathways regulated by the PPARs, LXRs, FXRs, RORs, and ERRs are deranged in states of metabolic disease these receptors have become important drug targets for development of novel drugs for human disease. Indeed, preclinical and clinical studies have uncovered many beneficial activities for small molecules that regulate these receptors. Nevertheless, in many cases there also appear to be unwanted activities of many of these small molecules that can or may limit their therapeutic use. Future studies that better define the tissue‐specific activities of the metabolic nuclear receptors and the trans‐acting factors that interact with receptors to impart tissue‐selective gene expression patterns will pave the way toward the identification of effective drugs for metabolic disease. References 1. Maglich JM, Sluder A, Guan X, Shi Y, McKee DD, Carrick K, et al. Comparison of complete nuclear receptor sets from the human, Caenorhabditis elegans and Drosophila genomes. Genome Biol 2001;2:RESEARCH0029. 2. Mangelsdorf DJ, Thummel C, Beato M, Herrlich P, Schutz G, Umesono K, et al. The nuclear receptor superfamily: the second decade. Cell 1995;83:835–9. 3. Steinmetz AC, Renaud JP, Moras D. Binding of ligands and activation of transcription by nuclear receptors. Annu Rev Biophys Biomol Struct 2001;30:329–59. 4. Greschik H, Moras D. Structure‐activity relationship of nuclear receptor‐ligand interactions. Curr Top Med Chem 2003;3:1573–99. 5. McKenna NJ, O’Malley BW. Combinatorial control of gene expression by nuclear receptors and coregulators. Cell 2002;108:465–74.

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6. Westin S, Rosenfeld MG, Glass CK. Nuclear receptor coactivators. Adv Pharmacol 2000;47:89–112. 7. Yen PM. Physiological and molecular basis of thyroid hormone action. Physiol Rev 2001;81:1097–142. 8. Schacke H, Docke WD, Asadullah K. Mechanisms involved in the side effects of glucocorticoids. Pharmacol Ther 2002;96:23–43. 9. Chawla A, Repa JJ, Evans RM, Mangelsdorf DJ. Nuclear receptors and lipid physiology: opening the X‐files. Science 2001;294:1866–70. 10. Edwards PA, Kast HR, Anisfeld AM. BAREing it all. The adoption of lxr and fxr and their roles in lipid homeostasis. J Lipid Res 2002;43:2–12. 11. Forman BM, Chen J, Evans RM. Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator‐activated receptors alpha and delta. Proc Natl Acad Sci USA 1997;94:4312–7. 12. Kliewer SA, Sundseth SS, Jones SA, Brown PJ, Wisely GB, Koble CS, et al. Fatty acids and eicosanoids regulate gene expression through direct interactions with peroxisome proliferator‐activated receptors alpha and gamma. Proc Natl Acad Sci USA 1997;94:4318–23. 13. Lee CH, Olson P, Evans RM. Minireview: lipid metabolism, metabolic diseases, and peroxisome proliferator‐activated receptors. Endocrinology 2003;144:2201–7. 14. Repa JJ, Mangelsdorf DJ. Nuclear receptor regulation of cholesterol and bile acid metabolism. Curr Opin Biotechnol 1999;10:557–63. 15. Forman BM, Ruan B, Chen J, Schroepfer GJ, Jr., Evans RM. The orphan nuclear receptor LXRalpha is positively and negatively regulated by distinct products of mevalonate metabolism. Proc Natl Acad Sci USA 1997;94:10588–93. 16. Janowski BA, Grogan MJ, Jones SA, Wisely GB, Kliewer SA, Corey EJ, et al. Structural requirements of ligands for the oxysterol liver X receptors LXRalpha and LXRbeta. Proc Natl Acad Sci USA 1999;96:266–71. 17. Janowski BA, Willy PJ, Devi TR, Falck JR, Mangelsdorf DJ. An oxysterol signalling pathway mediated by the nuclear receptor LXRa. Nature 1996;383:728–31. 18. Lehmann JM, Kliewer SA, Moore LB, Smith‐Oliver TA, Oliver BB, Su JL, et al. Activation of the nuclear receptor LXR by oxysterols defines a new hormone response pathway. J Biol Chem 1997;272:3137–40. 19. Bensinger SJ, Tontonoz P. Integration of metabolism and inflammation by lipid‐activated nuclear receptors. Nature 2008;454:470–7. 20. Kersten S, Desvergne B, Wahli W. Roles of PPARs in health and disease. Nature 2000;405:421–4. 21. Lefebvre P, Chinetti G, Fruchart JC, Staels B. Sorting out the roles of PPAR alpha in energy metabolism and vascular homeostasis. J Clin Invest 2006;116:571–80. 22. Kersten S, Seydoux J, Peters JM, Gonzalez FJ, Desvergne B, Wahli W. Peroxisome proliferator‐activated receptor alpha mediates the adaptive response to fasting. J Clin Invest 1999;103:1489–98. 23. Leone TC, Weinheimer CJ, Kelly DP. A critical role for the peroxisome proliferator‐ activated receptor alpha (PPARalpha) in the cellular fasting response: the PPARalpha‐ null mouse as a model of fatty acid oxidation disorders. Proc Natl Acad Sci USA 1999; 96:7473–8. 24. Inagaki T, Dutchak P, Zhao G, Ding X, Gautron L, Parameswara V, et al. Endocrine regulation of the fasting response by PPARalpha‐mediated induction of fibroblast growth factor 21. Cell Metab 2007;5:415–25. 25. Beaven SW, Tontonoz P. Nuclear receptors in lipid metabolism: targeting the heart of dyslipidemia. Annu Rev Med 2006;57:313–29.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

39

26. Forman BM, Tontonoz P, Chen J, Brun RP, Spiegelman BM, Evans RM. 15‐Deoxy‐delta 12, 14‐prostaglandin J2 is a ligand for the adipocyte determination factor PPAR gamma. Cell 1995;83:803–12. 27. Kraegen EW, Cooney GJ. Free fatty acids and skeletal muscle insulin resistance. Curr Opin Lipidol 2008;19:235–41. 28. Medina‐Gomez G, Gray SL, Yetukuri L, Shimomura K, Virtue S, Campbell M, et al. PPAR gamma 2 prevents lipotoxicity by controlling adipose tissue expandability and peripheral lipid metabolism. PLoS Genet 2007;3:e64. 29. He W, Barak Y, Hevener A, Olson P, Liao D, Le J, et al. Adipose‐specific peroxisome proliferator‐activated receptor gamma knockout causes insulin resistance in fat and liver but not in muscle. Proc Natl Acad Sci USA 2003;100:15712–7. 30. Tsai YS, Maeda N. PPARgamma: a critical determinant of body fat distribution in humans and mice. Trends Cardiovasc Med 2005;15:81–5. 31. Hevener AL, He W, Barak Y, Le J, Bandyopadhyay G, Olson P, et al. Muscle‐specific Pparg deletion causes insulin resistance. Nat Med 2003;9:1491–7. 32. Colca JR, Kletzien RF. What has prevented the expansion of insulin sensitisers? Expert Opin Investig Drugs 2006;15:205–10. 33. Nissen SE, Wolski K. Effect of rosiglitazone on the risk of myocardial infarction and death from cardiovascular causes. N Engl J Med 2007;356:2457–71. 34. Lincoff AM, Wolski K, Nicholls SJ, Nissen SE. Pioglitazone and risk of cardiovascular events in patients with type 2 diabetes mellitus: a meta‐analysis of randomized trials. JAMA 2007;298:1180–8. 35. Nissen SE, Wolski K, Topol EJ. Effect of muraglitazar on death and major adverse cardiovascular events in patients with type 2 diabetes mellitus. JAMA 2005;294:2581–6. 36. Hollenberg NK. Considerations for management of fluid dynamic issues associated with thiazolidinediones. Am J Med 2003;115(Suppl 8A), 111S–115S. 37. Karalliedde J, Buckingham RE. Thiazolidinediones and their fluid‐related adverse effects: facts, fiction and putative management strategies. Drug Saf 2007;30:741–53. 38. Guan Y, Hao C, Cha DR, Rao R, Lu W, Kohan DE, et al. Thiazolidinediones expand body fluid volume through PPARgamma stimulation of ENaC‐mediated renal salt absorption. Nat Med 2005;11:861–6. 39. Barish GD, Narkar VA, Evans RM. PPAR delta: a dagger in the heart of the metabolic syndrome. J Clin Invest 2006;116:590–7. 40. Furnsinn C, Willson TM, Brunmair B. Peroxisome proliferator‐activated receptor‐delta, a regulator of oxidative capacity, fuel switching and cholesterol transport. Diabetologia 2007;50:8–17. 41. Chawla A, Lee CH, Barak Y, He W, Rosenfeld J, Liao D, et al. PPARdelta is a very low‐density lipoprotein sensor in macrophages. Proc Natl Acad Sci USA 2003;100:1268–73. 42. Oliver WR, Jr., Shenk JL, Snaith MR, Russell CS, Plunket KD, Bodkin NL, et al. A selective peroxisome proliferator‐activated receptor delta agonist promotes reverse cholesterol transport. Proc Natl Acad Sci USA 2001;98:5306–11. 43. Oram JF, Lawn RM. ABCA1. The gatekeeper for eliminating excess tissue cholesterol. J Lipid Res 2001;42:1173–9. 44. van der Veen JN, Kruit JK, Havinga R, Baller JF, Chimini G, Lestavel S, et al. Reduced cholesterol absorption upon PPARdelta activation coincides with decreased intestinal expression of NPC1L1. J Lipid Res 2005;46:526–34. 45. Wang YX, Lee CH, Tiep S, Yu RT, Ham J, Kang H, et al. Peroxisome‐proliferator‐activated receptor delta activates fat metabolism to prevent obesity. Cell 2003;113:159–70. 46. Wang YX, Zhang CL, Yu RT, Cho HK, Nelson MC, Bayuga‐Ocampo CR, et al. Regulation of muscle fiber type and running endurance by PPARdelta. PLoS Biol 2004;2:e294.

40

HUANG AND SCHULMAN

47. Tanaka T, Yamamoto J, Iwasaki S, Asaba H, Hamura H, Ikeda Y, et al. Activation of peroxisome proliferator‐activated receptor delta induces fatty acid beta‐oxidation in skeletal muscle and attenuates metabolic syndrome. Proc Natl Acad Sci USA 2003;100:15924–9. 48. Narkar VA, Downes M, Yu RT, Embler E, Wang YX, Banayo E, et al. AMPK and PPARdelta agonists are exercise mimetics. Cell 2008;134:405–15. 49. Hardie DG. AMP‐activated/SNF1 protein kinases: conserved guardians of cellular energy. Nat Rev Mol Cell Biol 2007;8:774–85. 50. Rockl KS, Hirshman MF, Brandauer J, Fujii N, Witters LA, Goodyear LJ. Skeletal muscle adaptation to exercise training: AMP‐activated protein kinase mediates muscle fiber type shift. Diabetes 2007;56:2062–9. 51. Li AC, Glass CK. PPAR‐ and LXR‐dependent pathways controlling lipid metabolism and the development of atherosclerosis. J Lipid Res 2004;45:2161–73. 52. Glass CK, Witztum JL. Atherosclerosis. the road ahead. Cell 2001;104:503–16. 53. Chawla A, Boisvert WA, Lee CH, Laffitte BA, Barak Y, Joseph SB, et al. A PPAR gamma‐LXR‐ ABCA1 pathway in macrophages is involved in cholesterol efflux and atherogenesis. Mol Cell 2001;7:161–71. 54. Li AC, Binder CJ, Gutierrez A, Brown KK, Plotkin CR, Pattison JW, et al. Differential inhibition of macrophage foam‐cell formation and atherosclerosis in mice by PPARalpha, beta/delta, and gamma. J Clin Invest 2004;114:1564–76. 55. Schenk S, Saberi M, Olefsky JM. Insulin sensitivity: modulation by nutrients and inflammation. J Clin Invest 2008;118:2992–3002. 56. Arkan MC, Hevener AL, Greten FR, Maeda S, Li ZW, Long JM, et al. IKK‐beta links inflammation to obesity‐induced insulin resistance. Nat Med 2005;11:191–8. 57. Solinas G, Vilcu C, Neels JG, Bandyopadhyay GK, Luo JL, Naugler W, et al. JNK1 in hematopoietically derived cells contributes to diet‐induced inflammation and insulin resistance without affecting obesity. Cell Metab 2007;6:386–97. 58. Weisberg SP, McCann D, Desai M, Rosenbaum M, Leibel RL, Ferrante AW, Jr. Obesity is associated with macrophage accumulation in adipose tissue. J Clin Invest 2003;112:1796–808. 59. Xu H, Barnes GT, Yang Q, Tan G, Yang D, Chou CJ, et al. Chronic inflammation in fat plays a crucial role in the development of obesity‐related insulin resistance. J Clin Invest 2003;112:1821–30. 60. Ricote M, Glass CK. PPARs and molecular mechanisms of transrepression. Biochim Biophys Acta 2007;1771:926–35. 61. Coghlan MJ, Jacobson PB, Lane B, Nakane M, Lin CW, Elmore SW, et al. A novel antiinflammatory maintains glucocorticoid efficacy with reduced side effects. Mol Endocrinol 2003;17:860–9. 62. Miner JN. Designer glucocorticoids. Biochem Pharmacol 2002;64:355–61. 63. Hevener AL, Olefsky JM, Reichart D, Nguyen MT, Bandyopadyhay G, Leung HY, et al. Macrophage PPAR gamma is required for normal skeletal muscle and hepatic insulin sensitivity and full antidiabetic effects of thiazolidinediones. J Clin Invest 2007;117:1658–69. 64. Kang K, Reilly SM, Karabacak V, Gangl MR, Fitzgerald K, Hatano B, et al. Adipocyte‐derived Th2 cytokines and myeloid PPARdelta regulate macrophage polarization and insulin sensitivity. Cell Metab 2008;7:485–95. 65. Odegaard JI, Ricardo‐Gonzalez RR, Goforth MH, Morel CR, Subramanian V, Mukundan L, et al. Macrophage‐specific PPARgamma controls alternative activation and improves insulin resistance. Nature 2007;447:1116–20. 66. Odegaard JI, Ricardo‐Gonzalez RR, Red Eagle A, Vats D, Morel CR, Goforth MH, et al. Alternative M2 activation of Kupffer cells by PPARdelta ameliorates obesity‐induced insulin resistance. Cell Metab 2008;7:496–507.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

41

67. Gordon S, Taylor PR. Monocyte and macrophage heterogeneity. Nat Rev Immunol 2005;5:953–64. 68. Lumeng CN, Bodzin JL, Saltiel AR. Obesity induces a phenotypic switch in adipose tissue macrophage polarization. J Clin Invest 2007;117:175–84. 69. Gordon S. Alternative activation of macrophages. Nat Rev Immunol 2003;3:23–35. 70. Vats D, Mukundan L, Odegaard JI, Zhang L, Smith KL, Morel CR, et al. Oxidative metabolism and PGC‐1beta attenuate macrophage‐mediated inflammation. Cell Metab 2006;4:13–24. 71. Barish GD, Atkins AR, Downes M, Olson P, Chong LW, Nelson M, et al. PPARdelta regulates multiple proinflammatory pathways to suppress atherosclerosis. Proc Natl Acad Sci USA 2008;105:4271–6. 72. Li AC, Glass CK. The macrophage foam cell as a target for therapeutic intervention. Nat Med 2002;8:1235–42. 73. Marathe C, Bradley MN, Hong C, Chao L, Wilpitz D, Salazar J, et al. Preserved glucose tolerance in high fat diet‐fed C57BL/6 mice transplanted with PPARgamma
/
, PPARdelta
/
, PPARgamma delta
/
or LXRalpha beta
/
bone marrow. J Lipid Res 2009;50:214–24. 74. Willy PJ, Umesono K, Ong ES, Evans RM, Heyman RA, Mangelsdorf DJ. LXR, a nuclear receptor that defines a distinct retinoid response pathway. Genes Dev 1995;9:1033–45. 75. Shinar DM, Endo N, Rutledge SJ, Vogel R, Rodan GA, Schmidt A. NER, a new member of the gene family encoding the human steroid hormone nuclear receptor. Gene 1994;147:273–6. 76. Song C, Liao S. Cholestenoic acid is a naturally occurring ligand for liver X receptor alpha. Endocrinology 2000;141:4180–4. 77. Fu X, Menke JG, Chen Y, Zhou G, MacNaul KL, Wright SD, et al. 27‐hydroxycholesterol is an endogenous ligand for liver X receptor in cholesterol‐loaded cells. J Biol Chem 2001;276:38378–87. 78. Peet DJ, Turley SD, Ma W, Janowski BA, Lobaccaro J‐MA, Hammer RE, et al. Cholesterol and bile acid metabolism are impaired in mice lacking the nuclear oxysterol receptor LXRa. Cell 1998;93:693–704. 79. Repa JJ, Berge KE, Pomajzl C, Richardson JA, Hobbs H, Mangelsdorf DJ. Regulation of ATP‐ binding cassette sterol transporters ABCG5 and ABCG8 by the liver X receptors alpha and beta. J Biol Chem 2002;277:18793–800. 80. Berge KE, Tian H, Graf GA, Yu L, Grishin NV, Schultz J, et al. Accumulation of dietary cholesterol in sitosterolemia caused by mutations in adjacent ABC transporters. Science 2000;290:1771–5. 81. Repa JJ, Liang G, Ou J, Bashmakov Y, Lobaccaro JM, Shimomura I, et al. Regulation of mouse sterol regulatory element‐binding protein‐1c gene (SREBP‐1c) by oxysterol receptors, LXRalpha and LXRbeta. Genes Dev 2000;14:2819–30. 82. Schultz JR, Tu H, Luk A, Repa JJ, Medina JC, Li L, et al. Role of LXRs in control of lipogenesis. Genes Dev 2000;14:2831–8. 83. Zhang Y, Repa JJ, Gauthier K, Mangelsdorf DJ. Regulation of lipoprotein lipase by the oxysterol receptors, LXRalpha and LXRbeta. J Biol Chem 2001;276:43018–24. 84. Wang Y, Kurdi‐Haidar B, Oram JF. LXR‐mediated activation of macrophage stearoyl‐CoA desaturase generates unsaturated fatty acids that destabilize ABCA1. J Lipid Res 2004;45:972–80. 85. Joseph SB, Laffitte BA, Patel PH, Watson MA, Matsukuma KE, Walczak R, et al. Direct and indirect mechanisms for regulation of fatty acid synthase gene expression by LXRs. J Biol Chem 2002;277:11019–25. 86. Costet P, Luo Y, Wang N, Tall AR. Sterol‐dependent transactivation of the ABC1 promoter by the liver X receptor/retinoid X receptor. J Biol Chem 2000;275:28240–5.

42

HUANG AND SCHULMAN

87. Repa JJ, Turley SD, Lobaccaro JA, Medina J, Li L, Lustig K, et al. Regulation of absorption and ABC1‐mediated efflux of cholesterol by RXR heterodimers. Science 2000;289:1524–9. 88. Venkateswaran A, Laffitte BA, Joseph SB, Mak PA, Wilpitz DC, Edwards PA, et al. Control of cellular cholesterol efflux by the nuclear oxysterol receptor LXR alpha. Proc Natl Acad Sci USA 2000;97:12097–102. 89. Hayden MR, Clee SM, Brooks‐Wilson A, Genest J, Jr., Attie A, Kastelein JJ. Cholesterol efflux regulatory protein, Tangier disease and familial high‐density lipoprotein deficiency. Curr Opin Lipidol 2000;11:117–22. 90. Hobbs HH, Rader DJ. ABC1: connecting yellow tonsils, neuropathy, and very low HDL. J Clin Invest 1999;104:1015–7. 91. Aiello RJ, Brees D, Francone OL. ABCA1‐deficient mice: insights into the role of monocyte lipid efflux in HDL formation and inflammation. Arterioscler Thromb Vasc Biol 2003;23:972–80. 92. Singaraja RR, Fievet C, Castro G, James ER, Hennuyer N, Clee SM, et al. Increased ABCA1 activity protects against atherosclerosis. J Clin Invest 2002;110:35–42. 93. Joyce CW, Amar MJ, Lambert G, Vaisman BL, Paigen B, Najib‐Fruchart J, et al. The ATP binding cassette transporter A1 (ABCA1) modulates the development of aortic atherosclerosis in C57BL/6 and apoE‐knockout mice. Proc Natl Acad Sci USA 2002;99:407–12. 94. Wagner BL, Valledor AF, Shao G, Daige CL, Bischoff ED, Petrowski M, et al. Promoter‐ specific roles for liver X receptor/corepressor complexes in the regulation of ABCA1 and SREBP1 gene expression. Mol Cell Biol 2003;23:5780–9. 95. Kennedy MA, Venkateswaran A, Tarr PT, Xenarios I, Kudoh J, Shimizu N, et al. Characterization of the human ABCG1 gene: liver X receptor activates an internal promoter that produces a novel transcript encoding an alternative form of the protein. J Biol Chem 2001;276:39438–47. 96. Laffitte BA, Repa JJ, Joseph SB, Wilpitz DC, Kast HR, Mangelsdorf DJ, et al. LXRs control lipid‐inducible expression of the apolipoprotein E gene in macrophages and adipocytes. Proc Natl Acad Sci USA 2001;98:507–12. 97. Wang N, Lan D, Chen W, Matsuura F, Tall AR. ATP‐binding cassette transporters G1 and G4 mediate cellular cholesterol efflux to high‐density lipoproteins. Proc Natl Acad Sci USA 2004;101:9774–9. 98. Schuster GU, Parini P, Wang L, Alberti S, Steffensen KR, Hansson GK, et al. Accumulation of foam cells in liver X receptor‐deficient mice. Circulation 2002;106:1147–53. 99. Tangirala RK, Bischoff ED, Joseph SB, Wagner BL, Walczak R, Laffitte BA, et al. Identification of macrophage liver X receptors as inhibitors of atherosclerosis. Proc Natl Acad Sci USA 2002;99:11896–901. 100. Terasaka N, Hiroshima A, Koieyama T, Ubukata N, Morikawa Y, Nakai D, et al. T‐0901317, a synthetic liver X receptor ligand, inhibits development of atherosclerosis in LDL receptor‐ deficient mice. FEBS Lett 2003;536:6–11. 101. Joseph SB, McKilligin E, Pei L, Watson MA, Collins AR, Laffitte BA, et al. Synthetic LXR ligand inhibits the development of atherosclerosis in mice. Proc Natl Acad Sci USA 2002;99:7604–9. 102. Levin N, Bischoff ED, Daige CL, Thomas D, Vu CT, Heyman RA, et al. Macrophage liver X receptor is required for antiatherogenic activity of LXR agonists. Arterioscler Thromb Vasc Biol 2005;25:135–42. 103. Fowler AJ, Sheu MY, Schmuth M, Kao J, Fluhr JW, Rhein L, et al. Liver X receptor activators display anti‐inflammatory activity in irritant and allergic contact dermatitis models: liver‐X‐ receptor‐specific inhibition of inflammation and primary cytokine production. J Invest Dermatol 2003;120:246–55.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

43

104. Joseph SB, Castrillo A, Laffitte BA, Mangelsdorf DJ, Tontonoz P. Reciprocal regulation of inflammation and lipid metabolism by liver X receptors. Nat Med 2003;9:213–9. 105. Buja LM. Does atherosclerosis have an infectious etiology? Circulation 1996;94:872–3. 106. Edfeldt K, Swedenborg J, Hansson GK, Yan ZQ. Expression of toll‐like receptors in human atherosclerotic lesions: a possible pathway for plaque activation. Circulation 2002;105:1158–61. 107. Castrillo A, Joseph SB, Vaidya SA, Haberland M, Fogelman AM, Cheng G, et al. Crosstalk between LXR and toll‐like receptor signaling mediates bacterial and viral antagonism of cholesterol metabolism. Mol Cell 2003;12:805–16. 108. Bensinger SJ, Bradley MN, Joseph SB, Zelcer N, Janssen EM, Hausner MA, et al. LXR signaling couples sterol metabolism to proliferation in the acquired immune response. Cell 2008;134:97–111. 109. Brown MS, Goldstein JL. The SREBP pathway: regulation of cholesterol metabolism by proteolysis of a membrane‐bound transcription factor. Cell 1997;89:331–40. 110. Laffitte BA, Chao LC, Li J, Walczak R, Hummasti S, Joseph SB, et al. Activation of liver X receptor improves glucose tolerance through coordinate regulation of glucose metabolism in liver and adipose tissue. Proc Natl Acad Sci USA 2003;100:5419–24. 111. Cao G, Liang Y, Broderick CL, Oldham BA, Beyer TP, Schmidt RJ, et al. Antidiabetic action of a liver x receptor agonist mediated by inhibition of hepatic gluconeogenesis. J Biol Chem 2003;278:1131–6. 112. Mitro N, Mak PA, Vargas L, Godio C, Hampton E, Molteni V, et al. The nuclear receptor LXR is a glucose sensor. Nature 2007;445:219–23. 113. Muscat GE, Wagner BL, Hou J, Tangirala RK, Bischoff ED, Rohde P, et al. Regulation of cholesterol homeostasis and lipid metabolism in skeletal muscle by liver X receptors. J Biol Chem 2002;277:40722–8. 114. Kalaany NY, Gauthier KC, Zavacki AM, Mammen PP, Kitazume T, Peterson JA, et al. LXRs regulate the balance between fat storage and oxidation. Cell Metab 2005;1:231–44. 115. Groot PH, Pearce NJ, Yates JW, Stocker C, Sauermelch C, Doe CP, et al. Synthetic LXR agonists increase LDL in CETP species. J Lipid Res 2005;46:2182–91. 116. Luo Y, Tall AR. Sterol upregulation of human CETP expression in vitro and in transgenic mice by an LXR element. J Clin Invest 2000;105:513–20. 117. de Grooth GJ, Smilde TJ, Van Wissen S, Klerkx AH, Zwinderman AH, Fruchart JC, et al. The relationship between cholesteryl ester transfer protein levels and risk factor profile in patients with familial hypercholesterolemia. Atherosclerosis 2004;173:261–7. 118. Brousseau ME, Schaefer EJ, Wolfe ML, Bloedon LT, Digenio AG, Clark RW, et al. Effects of an inhibitor of cholesteryl ester transfer protein on HDL cholesterol. N Engl J Med 2004;350:1505–15. 119. Klerkx AH, de Grooth GJ, Zwinderman AH, Jukema JW, Kuivenhoven JA, Kastelein JJ. Cholesteryl ester transfer protein concentration is associated with progression of atherosclerosis and response to pravastatin in men with coronary artery disease (REGRESS). Eur J Clin Invest 2004;34:21–8. 120. Alberti S, Schuster G, Parini P, Feltkamp D, Diczfalusy U, Rudling M, et al. Hepatic cholesterol metabolism and resistance to dietary cholesterol in LXRbeta‐deficient mice. J Clin Invest 2001;107:565–73. 121. Bradley MN, Hong C, Chen M, Joseph SB, Wilpitz DC, Wang X, et al. Ligand activation of LXR beta reverses atherosclerosis and cellular cholesterol overload in mice lacking LXR alpha and apoE. J Clin Invest 2007;117:2337–46. 122. Lund EG, Menke JG, Sparrow CP. Liver X receptor agonists as potential therapeutic agents for dyslipidemia and atherosclerosis. Arterioscler Thromb Vasc Biol 2003;23:1169–77.

44

HUANG AND SCHULMAN

123. Hoerer S, Schmid A, Heckel A, Budzinski RM, Nar H. Crystal structure of the human liver X receptor beta ligand‐binding domain in complex with a synthetic agonist. J Mol Biol 2003;334:853–61. 124. Farnegardh M, Bonn T, Sun S, Ljunggren J, Ahola H, Wilhelmsson A, et al. The three‐ dimensional structure of the liver X receptor beta reveals a flexible ligand‐binding pocket that can accommodate fundamentally different ligands. J Biol Chem 2003;278:38821–8. 125. Williams S, Bledsoe RK, Collins JL, Boggs S, Lambert MH, Miller AB, et al. X‐ray crystal structure of the liver X receptor beta ligand binding domain: regulation by a histidine‐ tryptophan switch. J Biol Chem 2003;278:27138–43. 126. Svensson S, Ostberg T, Jacobsson M, Norstrom C, Stefansson K, Hallen D, et al. Crystal structure of the heterodimeric complex of LXRalpha and RXRbeta ligand‐binding domains in a fully agonistic conformation. EMBO J 2003;22:4625–33. 127. McDonnell DP, Connor CE, Wijayaratne A, Chang CY, Norris JD. Definition of the molecular and cellular mechanisms underlying the tissue‐selective agonist/antagonist activities of selective estrogen receptor modulators. Recent Prog Horm Res 2002;57:295–316. 128. Oberfield JL, Collins JL, Holmes CP, Goreham DM, Cooper JP, Cobb JE, et al. A peroxisome proliferator‐activated receptor gamma ligand inhibits adipocyte differentiation. Proc Natl Acad Sci USA 1999;96:6102–6. 129. Rocchi S, Picard F, Vamecq J, Gelman L, Potier N, Zeyer D, et al. A unique PPARgamma ligand with potent insulin‐sensitizing yet weak adipogenic activity. Mol Cell 2002;8:737–47. 130. Schulman IG, Heyman RA. The flip side: identifying small molecule regulators of nuclear receptors. Chem Biol 2004;11:639–46. 131. Forman BM, Goode E, Chen J, Oro AE, Bradley DJ, Perlmann T, et al. Identification of a nuclear receptor that is activated by farnesol metabolites. Cell 1995;81:687–93. 132. Westin S, Heyman RA, Martin R. FXR, a therapeutic target for bile acid and lipid disorders. Mini Rev Med Chem 2005;5:719–27. 133. Zhang Y, Edwards PA. FXR signaling in metabolic disease. FEBS Lett 2008;582:10–8. 134. Zhang Y, Castellani LW, Sinal CJ, Gonzalez FJ, Edwards PA. Peroxisome proliferator‐ activated receptor‐gamma coactivator 1alpha (PGC‐1alpha) regulates triglyceride metabolism by activation of the nuclear receptor FXR. Genes Dev 2004;18:157–69. 135. Handschin C, Spiegelman BM. Peroxisome proliferator‐activated receptor gamma coactivator 1 coactivators, energy homeostasis, and metabolism. Endocr Rev 2006;27:728–35. 136. Makishima M, Okamoto AY, Repa JJ, Tu H, Learned RM, Luk A, et al. Identification of a nuclear receptor for bile acids. Science 1999;284:1362–5. 137. Parks DJ, Blanchard SG, Bledsoe RK, Chandra G, Consler TG, Kliewer SA, et al. Bile acids: natural ligands for an orphan nuclear receptor. Science 1999;284:1365–8. 138. Wang H, Chen J, Hollister K, Sowers LC, Forman BM. Endogenous bile acids are ligands for the nuclear receptor FXR/BAR. Mol Cell 1999;3:543–53. 139. Goodwin B, Jones SA, Price RR, Watson MA, McKee DD, Moore LB, et al. A regulatory cascade of the nuclear receptors FXR, SHP‐1, and LRH‐1 represses bile acid biosynthesis. Mol Cell 2000;6:517–26. 140. Lee YS, Chanda D, Sim J, Park YY, Choi HS. Structure and function of the atypical orphan nuclear receptor small heterodimer partner. Int Rev Cytol 2007;261:117–58. 141. Boulias K, Katrakili N, Bamberg K, Underhill P, Greenfield A, Talianidis I. Regulation of hepatic metabolic pathways by the orphan nuclear receptor SHP. EMBO J 2005;24:2624–33. 142. Yang Y, Zhang M, Eggertsen G, Chiang JY. On the mechanism of bile acid inhibition of rat sterol 12alpha‐hydroxylase gene (CYP8B1) transcription: roles of alpha‐fetoprotein transcription factor and hepatocyte nuclear factor 4alpha. Biochim Biophys Acta 2002;1583:63–73. 143. Russell DW. The enzymes, regulation, and genetics of bile acid synthesis. Annu Rev Biochem 2003;72:137–74.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

45

144. Holt JA, Luo G, Billin AN, Bisi J, McNeill YY, Kozarsky KF, et al. Definition of a novel growth factor‐dependent signal cascade for the suppression of bile acid biosynthesis. Genes Dev 2003;17:1581–91. 145. Inagaki T, Choi M, Moschetta A, Peng L, Cummins CL, McDonald JG, et al. Fibroblast growth factor 15 functions as an enterohepatic signal to regulate bile acid homeostasis. Cell Metab 2005;2:217–25. 146. Song KH, Li T, Owsley E, Strom S, Chiang JY. Bile acids activate fibroblast growth factor 19 signaling in human hepatocytes to inhibit cholesterol 7alpha‐hydroxylase gene expression. Hepatology 2009;49:297–05. 147. Choi M, Moschetta A, Bookout AL, Peng L, Umetani M, Holmstrom SR, et al. Identification of a hormonal basis for gallbladder filling. Nat Med 2006;12:1253–5. 148. Kim I, Ahn SH, Inagaki T, Choi M, Ito S, Guo GL, et al. Differential regulation of bile acid homeostasis by the farnesoid X receptor in liver and intestine. J Lipid Res 2007;48:2664–72. 149. Ananthanarayanan M, Balasubramanian N, Makishima M, Mangelsdorf DJ, Suchy FJ. Human bile salt export pump promoter is transactivated by the farnesoid X receptor/bile acid receptor. J Biol Chem 2001;276:28857–65. 150. Claudel T, Inoue Y, Barbier O, Duran‐Sandoval D, Kosykh V, Fruchart J, et al. Farnesoid X receptor agonists suppress hepatic apolipoprotein CIII expression. Gastroenterology 2003;125:544–55. 151. Kast HR, Nguyen CM, Sinal CJ, Jones SA, Laffitte BA, Reue K, et al. Farnesoid X‐activated receptor induces apolipoprotein C‐II transcription: a molecular mechanism linking plasma triglyceride levels to bile acids. Mol Endocrinol 2001;15:1720–8. 152. Pircher PC, Kitto JL, Petrowski ML, Tangirala RK, Bischoff ED, Schulman IG, et al. Farnesoid X receptor regulates bile acid‐amino acid conjugation. J Biol Chem 2003; 278:27703–11. 153. Moschetta A, Bookout AL, Mangelsdorf DJ. Prevention of cholesterol gallstone disease by FXR agonists in a mouse model. Nat Med 2004;10:1352–8. 154. Trauner M, Boyer JL. Cholestatic syndromes. Curr Opin Gastroenterol 2004;20:220–30. 155. Trauner M, Meier PJ, Boyer JL. Molecular pathogenesis of cholestasis. N Engl J Med 1998;339:1217–27. 156. Fiorucci S, Clerici C, Antonelli E, Orlandi S, Goodwin B, Sadeghpour BM, et al. Protective effects of 6‐ethyl chenodeoxycholic acid, a farnesoid X receptor ligand, in estrogen‐induced cholestasis. J Pharmacol Exp Ther 2005;313:604–12. 157. Liu Y, Binz J, Numerick MJ, Dennis S, Luo G, Desai B, et al. Hepatoprotection by the farnesoid X receptor agonist GW4064 in rat models of intra‐ and extrahepatic cholestasis. J Clin Invest 2003;112:1678–87. 158. Pellicciari R, Fiorucci S, Camaioni E, Clerici C, Costantino G, Maloney PR, et al. 6alpha‐ ethyl‐chenodeoxycholic acid (6‐ECDCA), a potent and selective FXR agonist endowed with anticholestatic activity. J Med Chem 2002;45:3569–72. 159. Fiorucci S, Antonelli E, Rizzo G, Renga B, Mencarelli A, Riccardi L, et al. The nuclear receptor SHP mediates inhibition of hepatic stellate cells by FXR and protects against liver fibrosis. Gastroenterology 2004;127:1497–512. 160. Fiorucci S, Rizzo G, Antonelli E, Renga B, Mencarelli A, Riccardi L, et al. Cross‐talk between farnesoid‐X‐receptor (FXR) and peroxisome proliferator‐activated receptor gamma contributes to the antifibrotic activity of FXR ligands in rodent models of liver cirrhosis. J Pharmacol Exp Ther 2005;315:58–68. 161. Fiorucci S, Rizzo G, Antonelli E, Renga B, Mencarelli A, Riccardi L, et al. A farnesoid x receptor‐small heterodimer partner regulatory cascade modulates tissue metalloproteinase inhibitor‐1 and matrix metalloprotease expression in hepatic stellate cells and promotes resolution of liver fibrosis. J Pharmacol Exp Ther 2005;314:584–95.

46

HUANG AND SCHULMAN

162. Inagaki T, Moschetta A, Lee YK, Peng L, Zhao G, Downes M, et al. Regulation of antibacterial defense in the small intestine by the nuclear bile acid receptor. Proc Natl Acad Sci USA 2006;103:3920–5. 163. Angelin B, Einarsson K, Hellstrom K, Leijd B. Effects of cholestyramine and chenodeoxycholic acid on the metabolism of endogenous triglyceride in hyperlipoproteinemia. J Lipid Res 1978;19:1017–24. 164. Bateson MC, Maclean D, Evans JR, Bouchier IA. Chenodeoxycholic acid therapy for hypertriglyceridaemia in men. Br J Clin Pharmacol 1978;5:249–54. 165. Iser JH, Sali A. Chenodeoxycholic acid: a review of its pharmacological properties and therapeutic use. Drugs 1981;21:90–119. 166. Sinal CJ, Tohkin M, Miyata M, Ward JM, Lambert G, Gonzalez FJ. Targeted disruption of the nuclear receptor FXR/BAR impairs bile acid and lipid homeostasis. Cell 2000;102:731–44. 167. Maloney PR, Parks DJ, Haffner CD, Fivush AM, Chandra G, Plunket KD, et al. Identification of a chemical tool for the orphan nuclear receptor FXR. J Med Chem 2000;43:2971–4. 168. Hubbert ML, Zhang Y, Lee FY, Edwards PA. Regulation of hepatic Insig‐2 by the farnesoid X receptor. Mol Endocrinol 2007;21:1359–69. 169. Li J, Pircher PC, Schulman IG, Westin SK. Regulation of complement C3 expression by the bile acid receptor FXR. J Biol Chem 2005;280:7427–34. 170. Pineda Torra I, Claudel T, Duval C, Kosykh V, Fruchart JC, Staels B. Bile acids induce the expression of the human peroxisome proliferator‐activated receptor alpha gene via activation of the farnesoid X receptor. Mol Endocrinol 2003;17:259–72. 171. Prieur X, Coste H, Rodriguez JC. The human apolipoprotein AV gene is regulated by peroxisome proliferator‐activated receptor‐alpha and contains a novel farnesoid X‐activated receptor response element. J Biol Chem 2003;278:25468–80. 172. Watanabe M, Houten SM, Wang L, Moschetta A, Mangelsdorf DJ, Heyman RA, et al. Bile acids lower triglyceride levels via a pathway involving FXR, SHP, and SREBP‐1c. J Clin Invest 2004;113:1408–18. 173. Pullinger CR, Eng C, Salen G, Shefer S, Batta AK, Erickson SK, et al. Human cholesterol 7alpha‐hydroxylase (CYP7A1) deficiency has a hypercholesterolemic phenotype. J Clin Invest 2002;110:109–17. 174. Hanniman EA, Lambert G, McCarthy TC, Sinal CJ. Loss of functional farnesoid X‐receptor increases atherosclerotic lesions in apolipoprotein E‐deficient mice. J Lipid Res 2005;46:2595–604. 175. Lambert G, Amar MJ, Guo G, Brewer HB, Jr., Gonzalez FJ, Sinal CJ. The farnesoid X‐receptor is an essential regulator of cholesterol homeostasis. J Biol Chem 2003;278:2563–70. 176. Elzinga BM, Havinga R, Baller JF, Wolters H, Bloks V, Mensenkamp AR, et al. The role of transhepatic bile salt flux in the control of hepatic secretion of triacylglycerol‐rich lipoproteins in vivo in rodents. Biochim Biophys Acta 2002;1573:9–20. 177. Kang S, Davis RA. Cholesterol and hepatic lipoprotein assembly and secretion. Biochim Biophys Acta 2000;1529:223–30. 178. Wang DQ, Tazuma S, Cohen DE, Carey MC. Feeding natural hydrophilic bile acids inhibits intestinal cholesterol absorption: studies in the gallstone‐susceptible mouse. Am J Physiol Gastrointest Liver Physiol 2003;285:G494–502. 179. Mencarelli A, Renga B, Distrutti E, Fiorucci S. Anti‐atherosclerotic effect of Farnesoid‐X‐ Receptor in ApoE
/
mice. Am J Physiol Heart Circ Physiol 2009;296:H272–H281. 180. Guo GL, Santamarina‐Fojo S, Akiyama TE, Amar MJ, Paigen BJ, Brewer B, Jr., et al. Effects of FXR in foam‐cell formation and atherosclerosis development. Biochim Biophys Acta 2006;1761:1401–9.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

47

181. Zhang Y, Wang X, Vales C, Lee FY, Lee H, Lusis AJ, et al. FXR deficiency causes reduced atherosclerosis in Ldlr
/
mice. Arterioscler Thromb Vasc Biol 2006;26:2316–21. 182. Nicholson AC, Han J, Febbraio M, Silversterin RL, Hajjar DP. Role of CD36, the macrophage class B scavenger receptor, in atherosclerosis. Ann N Y Acad Sci 2001; 947:224–8. 183. Cariou B, van Harmelen K, Duran‐Sandoval D, van Dijk TH, Grefhorst A, Abdelkarim M, et al. The farnesoid X receptor modulates adiposity and peripheral insulin sensitivity in mice. J Biol Chem 2006;281:11039–49. 184. Ma K, Saha PK, Chan L, Moore DD. Farnesoid X receptor is essential for normal glucose homeostasis. J Clin Invest 2006;116:1102–9. 185. Zhang Y, Lee FY, Barrera G, Lee H, Vales C, Gonzalez FJ, et al. Activation of the nuclear receptor FXR improves hyperglycemia and hyperlipidemia in diabetic mice. Proc Natl Acad Sci USA 2006;103:1006–11. 186. Huang W, Ma K, Zhang J, Qatanani M, Cuvillier J, Liu J, et al. Nuclear receptor‐dependent bile acid signaling is required for normal liver regeneration. Science 2006;312:233–6. 187. Kim I, Morimura K, Shah Y, Yang Q, Ward JM, Gonzalez FJ. Spontaneous hepatocarcinogenesis in farnesoid X receptor‐null mice. Carcinogenesis 2007;28:940–6. 188. Yang F, Huang X, Yi T, Yen Y, Moore DD, Huang W. Spontaneous development of liver tumors in the absence of the bile acid receptor farnesoid X receptor. Cancer Res 2007;67:863–7. 189. Bays HE, Goldberg RB. The ‘forgotten’ bile acid sequestrants: is now a good time to remember? Am J Ther 2007;14:567–80. 190. Urizar NL, Liverman AB, Dodds DT, Silva FV, Ordentlich P, Yan Y, et al. A natural product that lowers cholesterol as an antagonist ligand for FXR. Science 2002;296:1703–6. 191. Burris TP, Montrose C, Houck KA, Osborne HE, Bocchinfuso WP, Yaden BC, et al. The hypolipidemic natural product guggulsterone is a promiscuous steroid receptor ligand. Mol Pharmacol 2005;67:948–54. 192. Owsley E, Chiang JY. Guggulsterone antagonizes farnesoid X receptor induction of bile salt export pump but activates pregnane X receptor to inhibit cholesterol 7alpha‐hydroxylase gene. Biochem Biophys Res Commun 2003;304:191–5. 193. Giguere V, Tini M, Flock G, Ong E, Evans RM, Otulakowski G. Isoform‐specific amino‐ terminal domains dictate DNA‐binding properties of ROR alpha, a novel family of orphan hormone nuclear receptors. Genes Dev 1994;8:538–53. 194. Carlberg C, Hooft van Huijsduijnen R, Staple JK, DeLamarter JF, Becker‐Andre M. RZRs, a new family of retinoid‐related orphan receptors that function as both monomers and homodimers. Mol Endocrinol 1994;8:757–70. 195. Medvedev A, Yan ZH, Hirose T, Giguere V, Jetten AM. Cloning of a cDNA encoding the murine orphan receptor RZR/ROR gamma and characterization of its response element. Gene 1996;181:199–206. 196. Atkins GB, Hu X, Guenther MG, Rachez C, Freedman LP, Lazar MA. Coactivators for the orphan nuclear receptor RORalpha. Mol Endocrinol 1999;13:1550–7. 197. Xie H, Sadim MS, Sun Z. RORgammat recruits steroid receptor coactivators to ensure thymocyte survival. J Immunol 2005;175:3800–9. 198. Moraitis AN, Giguere V, Thompson CC. Novel mechanism of nuclear receptor corepressor interaction dictated by activation function 2 helix determinants. Mol Cell Biol 2002;22:6831–41. 199. Johnson DR, Lovett JM, Hirsch M, Xia F, Chen JD. NuRD complex component Mi‐2beta binds to and represses RORgamma‐mediated transcriptional activation. Biochem Biophys Res Commun 2004;318:714–8.

48

HUANG AND SCHULMAN

200. Stehlin‐Gaon C, Willmann D, Zeyer D, Sanglier S, Van Dorsselaer A, Renaud JP, et al. All‐ trans retinoic acid is a ligand for the orphan nuclear receptor ROR beta. Nat Struct Biol 2003;10:820–5. 201. Kallen JA, Schlaeppi JM, Bitsch F, Geisse S, Geiser M, Delhon I, et al. X‐ray structure of the hRORalpha LBD at 1.63 A: structural and functional data that cholesterol or a cholesterol derivative is the natural ligand of RORalpha. Structure 2002;10:1697–707. 202. Yang X, Downes M, Yu RT, Bookout AL, He W, Straume M, et al. Nuclear receptor expression links the circadian clock to metabolism. Cell 2006;126:801–10. 203. Hamilton BA, Frankel WN, Kerrebrock AW, Hawkins TL, FitzHugh W, Kusumi K, et al. Disruption of the nuclear hormone receptor RORalpha in staggerer mice. Nature 1996;379:736–9. 204. Matysiak‐Scholze U, Nehls M. The structural integrity of ROR alpha isoforms is mutated in staggerer mice: cerebellar coexpression of ROR alpha1 and ROR alpha4. Genomics 1997;43:78–84. 205. Steinmayr M, Andre E, Conquet F, Rondi‐Reig L, Delhaye‐Bouchaud N, Auclair N, et al. staggerer phenotype in retinoid‐related orphan receptor alpha‐deficient mice. Proc Natl Acad Sci USA 1998;95:3960–5. 206. Vu‐Dac N, Gervois P, Grotzinger T, De Vos P, Schoonjans K, Fruchart JC, et al. Transcriptional regulation of apolipoprotein A‐I gene expression by the nuclear receptor RORalpha. J Biol Chem 1997;272:22401–4. 207. Lau P, Fitzsimmons RL, Raichur S, Wang SC, Lechtken A, Muscat GE. The orphan nuclear receptor, RORalpha, regulates gene expression that controls lipid metabolism: staggerer (SG/ SG) mice are resistant to diet‐induced obesity. J Biol Chem 2008;283:18411–21. 208. Mamontova A, Seguret‐Mace S, Esposito B, Chaniale C, Bouly M, Delhaye‐Bouchaud N, et al. Severe atherosclerosis and hypoalphalipoproteinemia in the staggerer mouse, a mutant of the nuclear receptor RORalpha. Circulation 1998;98:2738–43. 209. Raspe E, Duez H, Gervois P, Fievet C, Fruchart JC, Besnard S, et al. Transcriptional regulation of apolipoprotein C‐III gene expression by the orphan nuclear receptor RORalpha. J Biol Chem 2001;276:2865–71. 210. Pollin TI, Damcott CM, Shen H, Ott SH, Shelton J, Horenstein RB, et al. A null mutation in human APOC3 confers a favorable plasma lipid profile and apparent cardioprotection. Science 2008;322:1702–5. 211. Lau P, Nixon SJ, Parton RG, Muscat GE. RORalpha regulates the expression of genes involved in lipid homeostasis in skeletal muscle cells: caveolin‐3 and CPT‐1 are direct targets of ROR. J Biol Chem 2004;279:36828–40. 212. Andre E, Conquet F, Steinmayr M, Stratton SC, Porciatti V, Becker‐Andre M. Disruption of retinoid‐related orphan receptor beta changes circadian behavior, causes retinal degeneration and leads to vacillans phenotype in mice. EMBO J 1998;17:3867–77. 213. Raichur S, Lau P, Staels B, Muscat GE. Retinoid‐related orphan receptor gamma regulates several genes that control metabolism in skeletal muscle cells: links to modulation of reactive oxygen species production. J Mol Endocrinol 2007;39:29–44. 214. Kang HS, Angers M, Beak JY, Wu X, Gimble JM, Wada T, et al. Gene expression profiling reveals a regulatory role for ROR alpha and ROR gamma in phase I and phase II metabolism. Physiol Genomics 2007;31:281–94. 215. Forman BM, Chen J, Blumberg B, Kliewer SA, Henshaw R, Ong ES, et al. Cross‐talk among ROR alpha 1 and the Rev‐erb family of orphan nuclear receptors. Mol Endocrinol 1994;8:1253–61. 216. Vu‐Dac N, Chopin‐Delannoy S, Gervois P, Bonnelye E, Martin G, Fruchart JC, et al. The nuclear receptors peroxisome proliferator‐activated receptor alpha and Rev‐erbalpha mediate the species‐specific regulation of apolipoprotein A‐I expression by fibrates. J Biol Chem 1998;273:25713–20.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

49

217. Raspe E, Duez H, Mansen A, Fontaine C, Fievet C, Fruchart JC, et al. Identification of Rev‐erbalpha as a physiological repressor of apoC‐III gene transcription. J Lipid Res 2002;43:2172–9. 218. Coste H, Rodriguez JC. Orphan nuclear hormone receptor Rev‐erbalpha regulates the human apolipoprotein CIII promoter. J Biol Chem 2002;277:27120–9. 219. Turek FW, Joshu C, Kohsaka A, Lin E, Ivanova G, McDearmon E, et al. Obesity and metabolic syndrome in circadian Clock mutant mice. Science 2005;308:1043–5. 220. Ueda HR, Chen W, Adachi A, Wakamatsu H, Hayashi S, Takasugi T, et al. A transcription factor response element for gene expression during circadian night. Nature 2002;418:534–9. 221. Bunger MK, Wilsbacher LD, Moran SM, Clendenin C, Radcliffe LA, Hogenesch JB, et al. Mop3 is an essential component of the master circadian pacemaker in mammals. Cell 2000;103:1009–17. 222. Sato TK, Panda S, Miraglia LJ, Reyes TM, Rudic RD, McNamara P, et al. A functional genomics strategy reveals Rora as a component of the mammalian circadian clock. Neuron 2004;43:527–37. 223. Akashi M, Takumi T. The orphan nuclear receptor RORalpha regulates circadian transcription of the mammalian core‐clock Bmal1. Nat Struct Mol Biol 2005;12:441–8. 224. Guillaumond F, Dardente H, Giguere V, Cermakian N. Differential control of Bmal1 circadian transcription by REV‐ERB and ROR nuclear receptors. J Biol Rhythms 2005;20:391–403. 225. Preitner N, Damiola F, Lopez‐Molina L, Zakany J, Duboule D, Albrecht U, et al. The orphan nuclear receptor REV‐ERBalpha controls circadian transcription within the positive limb of the mammalian circadian oscillator. Cell 2002;110:251–60. 226. Rudic RD, McNamara P, Curtis AM, Boston RC, Panda S, Hogenesch JB, et al. BMAL1 and CLOCK, two essential components of the circadian clock, are involved in glucose homeostasis. PLoS Biol 2004;2:e377. 227. Shimba S, Ishii N, Ohta Y, Ohno T, Watabe Y, Hayashi M, et al. Brain and muscle Arnt‐like protein‐1 (BMAL1), a component of the molecular clock, regulates adipogenesis. Proc Natl Acad Sci USA 2005;102:12071–6. 228. Dzhagalov I, Giguere V, He YW. Lymphocyte development and function in the absence of retinoic acid‐related orphan receptor alpha. J Immunol 2004;173:2952–9. 229. Kopmels B, Mariani J, Delhaye‐Bouchaud N, Audibert F, Fradelizi D, Wollman EE. Evidence for a hyperexcitability state of staggerer mutant mice macrophages. J Neurochem 1992;58:192–9. 230. Delerive P, Monte D, Dubois G, Trottein F, Fruchart‐Najib J, Mariani J, et al. The orphan nuclear receptor ROR alpha is a negative regulator of the inflammatory response. EMBO Rep 2001;2:42–8. 231. Vanacker JM, Bonnelye E, Chopin‐Delannoy S, Delmarre C, Cavailles V, Laudet V. Transcriptional activities of the orphan nuclear receptor ERR alpha (estrogen receptor‐related receptor‐alpha). Mol Endocrinol 1999;13:764–73. 232. Vanacker JM, Pettersson K, Gustafsson JA, Laudet V. Transcriptional targets shared by estrogen receptor‐ related receptors (ERRs) and estrogen receptor (ER) alpha, but not by ERbeta. EMBO J 1999;18:4270–9. 233. Yang N, Shigeta H, Shi H, Teng CT. Estrogen‐related receptor, hERR1, modulates estrogen receptor‐mediated response of human lactoferrin gene promoter. J Biol Chem 1996;271:5795–804. 234. Lu D, Kiriyama Y, Lee KY, Giguere V. Transcriptional regulation of the estrogen‐inducible pS2 breast cancer marker gene by the ERR family of orphan nuclear receptors. Cancer Res 2001;61:6755–61. 235. Chen S, Zhou D, Yang C, Sherman M. Molecular basis for the constitutive activity of estrogen‐ related receptor alpha‐1. J Biol Chem 2001;276:28465–70.

50

HUANG AND SCHULMAN

236. Kallen J, Schlaeppi JM, Bitsch F, Filipuzzi I, Schilb A, Riou V, et al. Evidence for ligand‐ independent transcriptional activation of the human estrogen‐related receptor alpha (ERRalpha): crystal structure of ERRalpha ligand binding domain in complex with peroxisome proliferator‐activated receptor coactivator‐1alpha. J Biol Chem 2004;279:49330–7. 237. Suetsugi M, Su L, Karlsberg K, Yuan YC, Chen S. Flavone and isoflavone phytoestrogens are agonists of estrogen‐related receptors. Mol Cancer Res 2003;1:981–91. 238. Yang C, Chen S. Two organochlorine pesticides, toxaphene and chlordane, are antagonists for estrogen‐related receptor alpha‐1 orphan receptor. Cancer Res 1999;59:4519–24. 239. Tremblay GB, Kunath T, Bergeron D, Lapointe L, Champigny C, Bader JA, et al. Diethylstilbestrol regulates trophoblast stem cell differentiation as a ligand of orphan nuclear receptor ERR beta. Genes Dev 2001;15:833–8. 240. Busch BB, Stevens WC, Jr., Martin R, Ordentlich P, Zhou S, Sapp DW, et al. Identification of a selective inverse agonist for the orphan nuclear receptor estrogen‐related receptor alpha. J Med Chem 2004;47:5593–6. 241. Willy PJ, Murray IR, Qian J, Busch BB, Stevens WC, Jr., Martin R, et al. Regulation of PPARgamma coactivator 1alpha (PGC‐1alpha) signaling by an estrogen‐related receptor alpha (ERRalpha) ligand. Proc Natl Acad Sci USA 2004;101:8912–7. 242. Kallen J, Lattmann R, Beerli R, Blechschmidt A, Blommers MJ, Geiser M, et al. Crystal structure of human estrogen‐related receptor alpha in complex with a synthetic inverse agonist reveals its novel molecular mechanism. J Biol Chem 2007;282:23231–9. 243. Sladek R, Bader JA, Giguere V. The orphan nuclear receptor estrogen‐related receptor alpha is a transcriptional regulator of the human medium‐chain acyl coenzyme A dehydrogenase gene. Mol Cell Biol 1997;17:5400–9. 244. Ichida M, Nemoto S, Finkel T. Identification of a specific molecular repressor of the peroxisome proliferator‐activated receptor gamma Coactivator‐1 alpha (PGC‐1alpha). J Biol Chem 2002;277:50991–5. 245. Huss JM, Kopp RP, Kelly DP. Peroxisome proliferator‐activated receptor coactivator‐1alpha (PGC‐1alpha) coactivates the cardiac‐enriched nuclear receptors estrogen‐related receptor‐ alpha and ‐gamma. Identification of novel leucine‐rich interaction motif within PGC‐1alpha. J Biol Chem 2002;277:40265–74. 246. Schreiber SN, Knutti D, Brogli K, Uhlmann T, Kralli A. The transcriptional coactivator PGC‐1 regulates the expression and activity of the orphan nuclear receptor estrogen‐related receptor alpha (ERRalpha). J Biol Chem 2003;278:9013–8. 247. Cartoni R, Leger B, Hock MB, Praz M, Crettenand A, Pich S, et al. Mitofusins 1/2 and ERRalpha expression are increased in human skeletal muscle after physical exercise. J Physiol 2005;567:349–58. 248. Puigserver P, Wu Z, Park CW, Graves R, Wright M, Spiegelman BM. A cold‐inducible coactivator of nuclear receptors linked to adaptive thermogenesis. Cell 1998;92:829–39. 249. Wu Z, Puigserver P, Andersson U, Zhang C, Adelmant G, Mootha V, et al. Mechanisms controlling mitochondrial biogenesis and respiration through the thermogenic coactivator PGC‐1. Cell 1999;98:115–24. 250. Mootha VK, Handschin C, Arlow D, Xie X, St Pierre J, Sihag S, et al. Erralpha and Gabpa/b specify PGC‐1alpha‐dependent oxidative phosphorylation gene expression that is altered in diabetic muscle. Proc Natl Acad Sci USA 2004;101:6570–5. 251. Schreiber SN, Emter R, Hock MB, Knutti D, Cardenas J, Podvinec M, et al. The estrogen‐ related receptor alpha (ERRalpha) functions in PPARgamma coactivator 1alpha (PGC‐ 1alpha)‐induced mitochondrial biogenesis. Proc Natl Acad Sci USA 2004;101:6472–7. 252. Wende AR, Huss JM, Schaeffer PJ, Giguere V, Kelly DP. PGC‐1alpha coactivates PDK4 gene expression via the orphan nuclear receptor ERRalpha: a mechanism for transcriptional control of muscle glucose metabolism. Mol Cell Biol 2005;25:10684–94.

REGULATION OF METABOLISM BY NUCLEAR RECEPTORS

51

253. Yoon JC, Puigserver P, Chen G, Donovan J, Wu Z, Rhee J, et al. Control of hepatic gluconeogenesis through the transcriptional coactivator PGC‐1. Nature 2001;413:131–8. 254. Herzog B, Cardenas J, Hall RK, Villena JA, Budge PJ, Giguere V, et al. Estrogen‐related receptor alpha is a repressor of phosphoenolpyruvate carboxykinase gene transcription. J Biol Chem 2006;281:99–106. 255. Mootha VK, Lindgren CM, Eriksson KF, Subramanian A, Sihag S, Lehar J, et al. PGC‐1alpha‐ responsive genes involved in oxidative phosphorylation are coordinately downregulated in human diabetes. Nat Genet 2003;34:267–73. 256. Luo J, Sladek R, Carrier J, Bader JA, Richard D, Giguere V. Reduced fat mass in mice lacking orphan nuclear receptor estrogen‐related receptor alpha. Mol Cell Biol 2003;23:7947–56. 257. Villena JA, Hock MB, Chang WY, Barcas JE, Giguere V, Kralli A. Orphan nuclear receptor estrogen‐related receptor alpha is essential for adaptive thermogenesis. Proc Natl Acad Sci USA 2007;104:1418–23. 258. Carrier JC, Deblois G, Champigny C, Levy E, Giguere V. Estrogen‐related receptor alpha (ERRalpha) is a transcriptional regulator of apolipoprotein A‐IV and controls lipid handling in the intestine. J Biol Chem 2004;279:52052–8. 259. Stein RA, Chang CY, Kazmin DA, Way J, Schroeder T, Wergin M, et al. Estrogen‐related receptor alpha is critical for the growth of estrogen receptor‐negative breast cancer. Cancer Res 2008;68:8805–12. 260. Arany Z, Foo SY, Ma Y, Ruas JL, Bommi‐Reddy A, Girnun G, et al. HIF‐independent regulation of VEGF and angiogenesis by the transcriptional coactivator PGC‐1alpha. Nature 2008;451:1008–12. 261. Huss JM, Torra IP, Staels B, Giguere V, Kelly DP. Estrogen‐related receptor alpha directs peroxisome proliferator‐activated receptor alpha signaling in the transcriptional control of energy metabolism in cardiac and skeletal muscle. Mol Cell Biol 2004;24:9079–91. 262. Patti ME, Butte AJ, Crunkhorn S, Cusi K, Berria R, Kashyap S, et al. Coordinated reduction of genes of oxidative metabolism in humans with insulin resistance and diabetes: potential role of PGC1 and NRF1. Proc Natl Acad Sci USA 2003;100:8466–71. 263. Choi CS, Befroy DE, Codella R, Kim S, Reznick RM, Hwang YJ, et al. Paradoxical effects of increased expression of PGC‐1alpha on muscle mitochondrial function and insulin‐stimulated muscle glucose metabolism. Proc Natl Acad Sci USA 2008;105:19926–31.

Progesterone Receptor Action in Leiomyoma and Endometrial Cancer J. Julie Kim, Elizabeth C. Sefton, and Serdar E. Bulun Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Chicago, Illinois 60611

I. Introduction ................................................................................ II. The Uterus .................................................................................. III. Progesterone Action on the Endometrium and Myometrium ................... A. Physiological Response to Progesterone ......................................... B. Progesterone Receptors ............................................................. C. Coregulators of PR ................................................................... D. Progesterone Receptors in the Endometrium and Myometrium........... IV. Endometrial Cancer ...................................................................... V. Progesterone Receptor Action in Endometrial Cancer ........................... A. Progestin Therapy in Women ...................................................... B. Progesterone Receptors in Endometrial Cancer............................... C. Genes Regulated by Progestins in Endometrial Cancer...................... D. Transcriptional Activity of Progesterone Receptors in Endometrial Cancer.................................................................. VI. Conclusions and Perspectives of Progesterone Action in Endometrial Cancer ................................................................ VII. Uterine Leiomyoma....................................................................... VIII. Progesterone Receptor Action in Leiomyoma....................................... A. Relevance of Progesterone in Uterine Leiomyomas .......................... B. Progesterone Action on Genes Associated with Proliferation, Apoptosis, and ECM Deposition ................................................................ C. Growth Factor Regulation in Leiomyoma by Progesterone ................. D. Activation of Signaling Pathways in Leiomyoma by Progesterone and Estrogen ................................................................................ IX. Conclusions and Perspectives on Progesterone Action in Uterine Leiomyoma.................................................................................. X. Future Directions ......................................................................... References ..................................................................................

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Progesterone is a key hormone in the regulation of uterine function. In the normal physiological context, progesterone is primarily involved in remodeling of the endometrium and maintaining a quiescent myometrium. When pathologies of the uterus develop, specifically, endometrial cancer and uterine Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87002-6

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leiomyoma, response to progesterone is usually altered. Progesterone acts through mainly two isoforms of the progesterone receptor (PR), PRA and PRB which have been reported to exhibit different transcriptional activities. Studies examining the expression and function of the PRs in the normal endometrium and myometrium as well as in endometrial cancer and uterine leiomyoma are summarized here. The clinical use of progestins and the transcriptional activity of the PR on genes specific to endometrial cancer and leiomyoma are described. An increased understanding of the differential expression of PRs and response to progesterone in these two diseases is critical in order to develop more efficient and targeted therapies.

I. Introduction The progesterone receptor (PR) has been the focus of extensive analysis over the past few decades given its significance in reproductive tissues. The uterus is one of the most highly responsive organs to progesterone. Based on PR function, certain modalities of treatment for uterine pathologies have involved synthetic progestins or selective progesterone receptor modulators (SPRM). These compounds have proven to be effective in certain cases of endometrial cancer or uterine leiomyoma. Studies investigating the expression of PRs, and action of progesterone through its receptor in endometrial cancer and leiomyoma are summarized here. A brief description of PR expression and progesterone action in the normal endometrium and myometrium followed by a description of the clinical studies using progestins and SPRMs and the transcriptional activity of the PR on genes specific to endometrial cancer and leiomyoma will be presented.

II. The Uterus The uterus is the major female reproductive organ where the fetus develops during pregnancy. During development, the uterus develops from the middle to upper portion of the paramesonephric duct, also known as the Mullerian ducts.1 The uterus further organizes into distinct layers: the outermost layer which consists of smooth muscle is the myometrium and the innermost layer which lines the uterine cavity is the endometrium (Fig. 1A). The endometrium consists of a layer of columnar luminal epithelium supported by cellular stroma containing tubular glands (Fig. 1B). The luminal and glandular cells of the endometrium originate from the paramesonephric duct epithelium while the stroma originates from the mesenchyme surrounding

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A Infundibulum

Uterine tube

Fundus

Uterus

Endometrium Myometrium Fimbriae

Ovary Perimetrium

Vagina

Cervix B Myometrium

Endometrium FIG. 1. (A) The human uterus is comprised of an outer smooth muscle layer termed the myometrium and the innermost layer which lines the uterine cavity termed the endometrium. (B) Cross section of human uterine tissue shows distinct morphology of the myometrium and endometrium. The myometrium consists of smooth muscle cells with supporting stroma and vasculature. The endometrium consist mainly of epithelial glands and stroma.

the urogenital ridge. It is also from this mesenchyme that the myometrium forms. The myometrium consists of an organized network of smooth muscle cells with supporting stromal and vascular tissue (Fig. 1B). During pregnancy, the myometrium stretches by expanding the size and number of the smooth muscle cells and contracts in a coordinated fashion during labor. After pregnancy the uterus returns to its nonpregnant size. Both the endometrium and myometrium are highly responsive to the steroid hormones, estrogen and progesterone, and represent one of the most dynamic sites of hormone action during the menstrual cycle and pregnancy.

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III. Progesterone Action on the Endometrium and Myometrium A. Physiological Response to Progesterone The ovary is the major source of estrogen and progesterone in the human, synthesizing and secreting these hormones in a cyclical fashion.2 Granulosa cells from developing primary follicles biosynthesize and secrete estrogen and after ovulation these granulosa cells mature to form the corpus luteum which actively secretes progesterone and estrogen during the secretory phase of the menstrual cycle. If there is no pregnancy, the corpus luteum regresses resulting in the decline of estrogen and progesterone levels. If there is a pregnancy, the corpus luteum continues to grow and function for several months, after which time, it will regress as the placenta begins to synthesize estrogen and progesterone. The endometrium undergoes extensive remodeling in response to ovarian steroid hormones. Estrogen promotes proliferation and growth of the endometrial lining while progesterone antagonizes estrogen driven growth as well as promotes differentiation in preparation of an impending implantation.3 Specifically, when progesterone levels are high during the luteal phase of the menstrual cycle, the glandular epithelium transforms from relatively inactive cells full of free ribosomes to very active polarized cells, containing giant mitochondrial profiles, intracellular deposits of glycogen/glycoprotein‐rich material, and a complex intranuclear channel system.4 Morphologically, the glands become tortuous and have large lumens due to increased secretory activity. In parallel, the underlying stroma becomes very edematous as a result of increased capillary permeability and the cells begin to appear large and polyhedral, a transformation process termed decidualization. Decidualization begins in the stroma around the spiral arteries when progesterone levels are high during the mid‐ luteal phase, and spreads to the upper two‐thirds of the endometrium.5 If embryo implantation occurs, the reaction is intensified and becomes the decidua of pregnancy. The decidualized cell biochemically expresses new proteins and two of the most abundantly secreted proteins are insulin‐like growth factor binding protein‐1 (IGFBP1) and prolactin (PRL) and as a result are considered markers of decidualization.6 If there is no pregnancy, levels of estrogen and progesterone decline due to the regression of the corpus luteum causing the upper two thirds of the endometrium to be shed. The overall effect of progesterone on the myometrium is to maintain quiescence in both the pregnant and nonpregnant uterus. The nonpregnant uterus contracts throughout the menstrual cycle.7 During the late follicular phase, a progressive increase in uterine contractility parallels the rise in estradiol levels. After ovulation, during the luteal phase, the uterus undergoes a characteristic period of quiescence, strongly implicating progesterone as a

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mediator of this quiescence. At menses, after circulating levels of progesterone decrease, contractility increases and participates in the emptying of the uterine contents. In the pregnant uterus, it is well accepted that progesterone keeps the myometrium quiescent in order to promote and sustain pregnancy. Progesterone promotes myometrial relaxation and thought to actively block the transformation of the myometrium to a contractile phenotype. Although circulating levels of progesterone do not decrease before labor onset,8–10 the withdrawal remains a principal mechanism for the control of human parturition. Synthetic progesterone antagonists, such as RU486 initiate myometrial contractions at all stages of pregnancy.11,12

B. Progesterone Receptors The physiologic actions of progesterone are mediated by interaction with the PR, a member of the nuclear hormone receptor superfamily of ligand‐ activated transcription factors.13,14 There are two predominant PR isoforms, designated PR‐A and PR‐B, transcribed from the same gene by two distinct promoters, with the only difference being that human PR‐B is larger by an additional 164 amino acids at the amino terminus15–17 (Fig. 2). As a result, PR‐ A and PR‐B have distinct transcriptional activities.18–25 Three activation function (AF) domains have been identified in PR as AF1, AF2, and AF3. In many contexts, PRB functions as an activator of progesterone‐responsive genes, while PRA is transcriptionally inactive. In addition, PRA also functions as a strong transdominant repressor of PRB as well as the human estrogen receptor (ER) transcriptional activity.23 The precise mechanism underlying the differential activities of the two PR isoforms is not fully understood. Studies have suggested that PRA and PRB adopt different conformations within the cell which may contribute to its different transactivation functions. Tetel et al.19 demonstrated that the interaction of the amino terminus to the carboxyl terminus in PRB and PRA is different. Giangrande et al.24 demonstrated that PRA is unable to efficiently recruit the coactivators SRC‐1 and GRIP1 upon

AF3 1

PRB

AF1

AF2

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BUS

DBD

LBD

DBD

LBD

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PRA

FIG. 2. Functional domains of PRB and PRA. PRA lacks the 164 amino acids in the N‐terminus and thus the activation function (AF)3 domain. The DNA binding domain (DBD), ligand binding domain (LBD) AF1 and AF2 regions are present in both PRA and PRB.

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agonist binding despite the fact that both PRA and PRB contain sequences within the LBD that binds coactivators. In addition, PRA interacts efficiently with the corepressor SMRT permitting it to function as a transdominant repressor.24 At the promoter level, PR binds to a palindromic consensus sequence called the progesterone response element (PRE). It is thought that multisite PREs promote a more stable complex.23,26 Surprisingly, a survey of proximal promoter regions of endogenous genes regulated by PR reveal a lack of tandem palindromic PREs and an abundance of PRE half‐sites. This suggests that additional mechanisms are in place for PR recruitment to specific sites and activation of genes. Reports have shown that for the glucocorticoid receptor (GR), ‘‘composite’’ response elements which consist of a single hormone response element in tandem with heterologous binding sites leads to synergy between the GR and a variety of factors, including Sp1, NF1, CACCC‐box, and AP1.27–29

C. Coregulators of PR Nuclear receptors recruit the coregulators that perform all of the subsequent reactions needed to induce or repress expression of genes. Coactivators, such as the SRC family (steroid receptor coactivator) and CBP/p300, enhance transcription by liganded receptors while corepressors (such as SMRT and NCOR) repress transcription.30 These coregulators exist and function in large multiprotein complexes which are recruited to the target gene in rapid sequence by nuclear receptors. This complex contains many enzymes that are required for gene expression. Such enzymatic reactions for transcription include chromatin modification and remodeling, initiation of transcription, elongation of RNA chains, RNA splicing, and termination of the transcriptional response. Consequently, it is suggested that genes encoding for coregulators of hormones receptors are the true master genes of eukaryotes.30 It has been shown that upon ligand treatment, PR interacted preferentially with SRC‐1, which recruited CBP and significantly enhanced acetylation at K5 of histone H4.31 In contrast, activated GR preferentially associated with SRC‐2 (TIF‐2/GRIP‐1), which subsequently recruited pCAF and led to specific modification of histone H3, suggesting that specific coactivators are differentially recruited by different steroid hormone receptors which then recruit distinct histone acetyltransferases to modulate the transcription of steroid‐responsive genes. Investigation of the SRC family of coactivators, which consist of SRC‐1, SRC‐2, and SRC‐3 in mice has demonstrated its relevance in steroid responsive tissues. SRC‐1 knockout mice, although viable exhibited decreased responses to steroid hormone treatment.32 Ablation of SRC‐2 resulted in a partial lethal phenotype due to death in the first month of life.33 Those that survived to

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adulthood showed slowed growth and hypofertility due to placental defects. Female SRC‐3 null mice have reduced fertility. However, SRC‐3 is not expressed in the mouse endometrium and the uteri of SRC‐3
/
mice are able to undergo the artificially induced decidual reaction.34 Thus, it was concluded that SRC1 and SRC2 are the critical members of the p160 coactivator family for the regulation of uterine function.34 Recent studies have demonstrated the role of SRC‐2 in the adult uterus using a floxed allele of SRC‐2 crossed to the PR‐Cre mouse which abrogated SRC‐2 function only in cell lineages that express the PR.35 Absence of SRC‐2 in PR‐positive uterine cells was shown to result in infertility due to an early block in embryo implantation. The uterus of these mice was unable to undergo the necessary cellular and molecular changes that precede complete progesterone‐induced decidual progression. The expression of a number of decidualization markers, Bmp2, Cox2, and follistatin was significantly reduced in the partially decidualized PR(Cre/þ) SRC‐2(flox/flox) mice. While Bmp2 induction was negligible, Cox2 and follistatin were partially induced, which was suggested to be due to SRC‐2 being essential for the induction of pathways that lead to Bmp2 expression but that additional coregulators may be required for elaborating the Cox2 and follistatin expression pathways. The incomplete decidual response shown in both the PRCre/þSRC‐2flox/flox mouse and the SRC‐1KO32 suggests that both SRC coregulators may be required together in PR‐mediated signaling cascades that result in a fully decidualized uterus. To support this hypothesis, removal of SRC‐1 in these PR(Cre/þ) SRC‐2(flox/flox) mouse uterus resulted in the complete absence of a decidual response, confirming that both uterine SRC‐2 and ‐1 cooperate in progesterone‐initiated transcriptional programs. It is also widely observed that other transcription factors are able to interact with and modulate function of nuclear receptors. Reports have shown that for GR, PR, and ER‐alpha, other transcription factors, including members of Sp, NF, CACCC‐box, and AP1 families can bind to response elements that occur in tandem to the nuclear receptor response element allowing synergy/antagonism between the steroid receptors and the other transcription factors.27–29 In the absence of canonical PRE sequences, PR can tether to some transcription factors such as Sp1, Ap‐1, Stat5, p65 subunit of NF‐kB,36–38 and FOXO1.39,40 Through mouse models, it has been demonstrated that FKBPs, which are immunophilins interact with PR and influence its localization.41 Specifically, FKBP4 and FKBP5 interact with PR and are expressed in the uterine stroma during implantation. Furthermore, the FKBP4 knockout mice are infertile due to the inability to support implantation or undergo adequate decidualization. In one pioneering study, a genome‐wide scan of chromosomes 21 and 22 was performed in order to identify ER binding regions.42 In doing so, the authors discovered that FOXA1 was necessary for mediating the estrogen response in breast cancer cells. Furthermore, the FOXA1 binding site was

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the most conserved motif proximal to the regions that had an ER element. Another forkhead protein, from a different subfamily, FOXO1 can interact with steroid hormone receptors such as ER‐alpha, retinoic acid receptor, thyroid hormone receptor, and PR and elicits either repressive or activating effects on nuclear receptor mediated gene expression.39,43,44 In endometrial fibroblasts, FOXO1 and PR interacted with each other and bound to tandem DNA sequences in the IGFBP1 promoter.39,40 In addition, FOXO1 modulated PR transactivation of a PRE responsive reporter.39 Gene array studies revealed that many of the genes significantly regulated by FOXO1 during decidualization of endometrial stromal cells are also dependent on PR.40 Given that both transcription factors are involved in important cellular processes in the endometrium associated with growth inhibition and differentiation, the cross talk between these two molecules may be an important mode of endometrial remodeling.

D. Progesterone Receptors in the Endometrium and Myometrium Progesterone is central to the remodeling that occurs in the endometrium for uterine receptivity and acts on both the epithelial and stromal compartments. Studies in mice with selective ablation of PR isoforms revealed that PR‐A is necessary for ovulation and modulates the antiproliferative effects of progesterone in the uterus while PR‐B is required for normal mammary gland development and function.45,46 Recent evidence has confirmed the existence of a functional third isoform, designated PR‐C, which appears to play a critical role in the onset of parturition.47 The presence of multiple PR isoforms potentially increases the specificity and versatility of hormone action in a target tissue. PRs A and B are expressed in cells of the endometrium and its expression is dependent on the hormonal status and cell type. In the glandular epithelium, PR expression is stimulated by estrogen during the proliferative phase but is downregulated by its own ligand in the secretory phase. Prior to ovulation, PR‐A and PR‐B levels are approximately equivalent in glandular epithelium but only PR‐B persists in these cells in the mid‐secretory phase, suggesting that PR‐B is most important for the progesterone driven phenotypic changes in the glands at this time. In the stroma, PR‐A is the dominant isoform throughout the cycle.48–50 Conditions associated with endometrial pathology such as endometrial cancer is due to inadequate progesterone response as subsequently described. Myometrial expression of PR is less dynamically regulated than the endometrium during the menstrual cycle. Studies have demonstrated that both PRA and PRB are expressed and levels are consistent throughout the menstrual cycle.51,52

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IV. Endometrial Cancer Endometrial cancer is the most common cancer of the female reproductive tract, with estimated 39,080 new cases diagnosed in 2007. Despite the frequent detection of early‐stage cancers and the evolving use of adjuvant chemotherapy for advanced disease, the death rate from this malignancy has increased and currently claims 7400 lives among US women per year.53 The incidence of endometrial cancer is rising as life expectancy increases and as key risk factors, including obesity, become more prevalent. A better understanding of the pathophysiology underlying endometrial cancer is the first step to identifying key biomarkers that can improve diagnostic efforts and prevent development of this disease. Endometrial cancer is diagnosed by pathological examination. Approximately 80% of all endometrial carcinomas are endometrioid type, which arise from endometrial glands (Fig. 3A). The malignant phenotype and the varying degrees of differentiation are easily recognizable by microscopy.54 Endometrioid carcinoma is graded histologically from grade 1 to grade 3 depending on the percentage of solid nonsquamous areas and cytological atypia.55 Most endometrioid carcinomas are well to moderately differentiated and present alongside hyperplastic endometrium. These tumors, also known as type 1 endometrial carcinomas, are associated with chronic exposure to estrogen and a lack of opposing progesterone. One source of estrogen is fat tissue, in which peripheral androgens act as aromatase substrates to produce estrone. Estrone is then converted to estradiol by 17‐hydroxysteroid dehydrogenase.56

A

B

Uterine fibroids

Endometrial adenocarcinoma

FIG. 3. Endometrial cancer and uterine leiomyoma. (A) Endometrial adenocarcinoma arises from the endometrial glands exhibiting malignant behavior. (B) Uterine leiomyomas arise from benign overgrowth of smooth muscle cells and can range dramatically in size.

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Through the androgen‐to‐estrone pathway, postmenopausal women produce approximately 100 mg of estrone per day, or more if they are obese.57 Chronic estrogen exposure is also exacerbated by comorbid conditions in obese women, namely hypertension and diabetes. Polycystic ovarian syndrome (PCOS) is also associated with higher levels of estrogen and androgen, as well as low levels of opposing progesterone, leading to a higher risk of endometrial cancer in these patients. Finally, the long‐term use of tamoxifen, with its paradoxical estrogen agonist effect in the uterus, can also lead to type I endometrial cancer.58 Type II endometrial cancer occurs primarily in elderly postmenopausal women, and is neither related to estrogen nor preceded by endometrial hyperplasia. Type II tumors are high grade tumors with serous or clear‐cell morphology and carry a poor prognosis. Other morphological variants of endometrial cancer are placed into this category but occur at much lower rates.58,59 At the molecular level, type 1 tumors are commonly associated with abnormalities of DNA‐ mismatch repair genes, including k‐ras, PTEN, and beta‐catenin. Type 2 tumors are associated with abnormalities of p53 and HER2/neu, although they are not present in all cases.58

V. Progesterone Receptor Action in Endometrial Cancer A. Progestin Therapy in Women Studies have proven the efficacy of progesterone in protecting the endometrium against the hyperplastic effects of estradiol by inducing glandular and stromal differentiation.60 Accordingly, progestins play an essential and effective role in the management of endometrial hyperplasia.61,62 In a study of 52 postmenopausal women diagnosed with atypical hyperplasia or hyperplasia without atypia, 90% of patients had complete remission after treatment with 40 mg megestrol acetate per day for 42 months.63 However, close follow‐up is usually recommended in women who are treated with progestins especially for atypical hyperplasia since there is a significant increased risk of progression to carcinoma.64 Progestins have been used as adjuvant for endometrial cancer in hopes to prevent recurrence. However, several studies have shown that progestin treatment is not beneficial to the overall survival of women postsurgery.65,66 Progestins are also used as primary therapy, especially for premenopausal women as a fertility‐sparing treatment. Approximately 25% of endometrial cancer cases affect premenopausal women, particularly in the setting of obesity, chronic anovulation, and polycystic ovary syndrome.62,67–70 Progestin therapy would only be given when the tumor is well differentiated with positive receptors. There are few studies looking at the efficacy of progestin treatment in these women with the majority of published studies being

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case reports. In a review of articles published between Jan 1966 and Jan 2007 describing patients with endometrial cancer treated with hormonal therapy, 133 patients were identified, who were treated for an average duration of 6 months, and who demonstrated an average response time of 12 weeks.71 Of these 133 patients, 51% demonstrated a lasting complete response, 25% showed a temporary response, and 24% never responded to treatment. It is evident that a larger study is required to demonstrate the true benefit of progestin therapy in endometrial cancer. In regards to sparing fertility, there is no doubt that progestin therapy can be used;72 however, close follow‐up is required because of the substantial rate of recurrence.

B. Progesterone Receptors in Endometrial Cancer Morphological and biochemical evaluations demonstrated that in endometrial cancer, PRA is localized to the nucleus, even in the absence of progesterone.73 In contrast, a large proportion of PRB is cytoplasmic in the absence of ligand, but is rapidly translocated to the nucleus in the presence of progesterone. All endometrial cancer specimens demonstrated cytoplasmic PRB in 50% or more of the cells, and five of the seven tumors that were moderately to poorly differentiated demonstrated no PRB staining in the nuclei. Nuclear PRB was thus significantly associated with increasing tumor differentiation. PRA and PRB exhibit different activating properties and mediate the transcription of different sets of genes in endometrial cancer cells. Smid‐Koppman et al.74 demonstrated that in the presence of progesterone, PRB expressing Ishikawa cells displayed almost complete inhibition of cell growth, while PRA expressing Ishikawa cells only displayed 50% inhibition of cell growth. In an additional study by Hanekamp et al.,75 it was demonstrated that while PRB expressing Ishikawa cells caused more tumor growth in vivo than PRA expressing Ishikawa cells, tumor growth was inhibited after administration of MPA only in the tumors expressing PRB. There is an ongoing debate as to the PR status in endometrial tumors with one study suggesting that PRB is predominant in advanced endometrial tumors,76 another study pointing to the loss of both isoforms in advanced endometrial cancer,77 and a third study that indicates only PRA is expressed in poorly differentiated endometrial carcinoma cell lines.78

C. Genes Regulated by Progestins in Endometrial Cancer Nevertheless, in vitro studies have clearly demonstrated the efficacy of progestins to influence endometrial cancer cell behavior. When endometrial cancer cells are transfected with specifically PR‐A or PR‐B, progesterone can promote cell cycle inhibition, endometrial cancer cell invasion, differentiation

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to a secretory phenotype, induction of replicative senescence, and can downregulate the expression of cellular adhesion molecules.79,80 Regulation of genes such as cyclin D1, matrix metalloproteinase‐1 (MMP‐1), ‐2, ‐7, and ‐9, and Ets‐1 in response to progestins have been implicated to mediate the inhibition of cell growth and invasiveness.81 Primary cells from endometrial tumors also respond to progestins by significantly reducing proMMP‐9, proMMP‐2, and MMP‐2 release.82 Progestins have been shown to induce glycodelin expression in Ishikawa cells83–85 which causes inhibition of G1/S progression and upregulation of CDKIs thereby reducing cell proliferation.86 Progestins can increase FOXO1 protein levels in Ishikawa cells, specifically through PRB87 and promote cell cycle arrest and apoptosis in these cells. Interestingly, levels of FOXO1 protein are dramatically lower in 77%88 or 95.9%87 of endometrial tumor tissues studied compared to normal tissues from cycling endometrium. Shiozawa et al.89 reported that p27 expression in hyperplastic epithelia was negligible before MPA treatment, whereas it was greatly increased after treatment. Watanabe et al. demonstrated that it is through PRB that p21 and p27 expression increases.90 Microarray studies revealed that short term (4 h) and high dose (30 mg/ml)) exposure of Ishikawa cells to progesterone result in 247 differentially expressed genes of which 126 were upregulated and 121 were downregulated. Of these, 135 genes were involved in biological processes like cell cycle, cell proliferation and differentiation, developmental processes, immune responses, intracellular protein traffic, and transport.91 Hanekamp et al.92 reported that MPA inhibits expression of several metastasis‐related genes in a set of endometrial cancer cell lines. Treatment of Hec50co cells transfected with PR with progesterone for 12 h significantly regulated genes associated with cell signaling, DNA remodeling, apoptosis, tumor‐suppressor, and transcription factors. Interestingly, there was a consistent modulation of cytokines consistent to an antiinflammatory environment. Specifically, proinflammatory genes such as TNFalpha, IL‐1beta, and MCP‐1/MCAF‐1 were downregulated and antiinflammatory genes such as TRAP1 and SMAD4 were induced93. Progestins have been shown to modulate proteins in the apoptotic cascade in human endometrial precancers. Women with hyperplasia treated with either system MPA or a levonorgestrel intrauterine device exhibited increased apoptosis in the glandular cells with decreased expression of the antiapoptotic genes, Bcl‐2 and BAX.94 Overexpression of PRA and PRB in endometrial cancer cells resulted in a significant progesterone‐dependent inhibition of expression of a cadre of cellular adhesion molecules, including fibronectin, integrin alpha3, integrin beta1, integrin beta3, and cadherin 6.79,95 Thus, it is apparent that progesterone, through PRA and PRB modulate genes that are involved in processes associated with cell cycle, apoptosis, cell adhesion, differentiation, and inflammation in order to regulate endometrial cancer cell behavior.

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D. Transcriptional Activity of Progesterone Receptors in Endometrial Cancer It has been demonstrated that transcriptional activation by progesterone can involve other transcription factors. For example, liganded PR decreases the transcriptional activity of the activating protein‐1 (AP‐1) transcription factor family, and in particular, c‐Jun. In addition, progesterone strongly inhibited total AP‐1 as well as c‐Jun recruitment to the cyclin D1 promoter, whereas it enhanced AP‐1 occupancy on the p53 and p21 promoters, as shown by chromatin immunoprecipitation assays. This study concluded that in endometrial cancer cells, modulation of AP‐1 activity is a potential pathway of progesterone‐ induced growth inhibition in endometrial cancer cells.96 Another mechanism of progesterone action has been proposed to involve inhibition of NFkappaB transcriptional activity. Specifically, expression of A20 and ABIN‐2 were induced through PRB and these factors bind in a complex and inhibit NFkappaB transcriptional activity.97 EMSAs revealed the complete inhibition of NFkappaB dimer binding to DNA by both PRA and PRB. The inhibition of NFkappaB and its tumorigenic inflammatory and antiapoptotic effects by PR may be one pathway by which progesterone treatment is effective against endometrial hyperplasia and cancer. Glycodelin (GdA) is a progesterone induced gene in normal endometrial epithelial cells and endometrial cancer cells. Studies have shown that ligand‐ activated PR stimulates GdA promoter activity through functional Sp1 sites.98 As on numerous other genes, PR can tether to Sp1 to regulate promoters that do not have PRE sequences. In another study, it was shown that progesterone upregulated COMT protein expression in Ishikawa cells primarily through PRA. COMT converts genotoxic catecholestrogens to anticarcinogenic methoxyestrogens (2‐ME2) in the endometrium. COMT promoter activity was differentially regulated by the three half‐site PREs. Accordingly, high doses of 2‐ME2 inhibited Ishikawa cell proliferation.99 A novel mechanism for PR‐A and PR‐B mediated gene transcription in the uterus has been proposed to involve selected KLF members. Specifically, Kruppel-like factor 9 (also known as BTEB1) interacts with ligand‐activated PRB to increase PRB transactivity. This facilitates the recruitment of the transcriptional integrator CREB‐binding protein within the PR dimer, and is dependent on the structure of the ligand bound by PRB. By contrast, BTEB1 does not influence agonist bound PRA transactivity, but augments PRA inhibition of PRB‐mediated transactivation. Also, BTEB1 potentiates ligand‐independent PRA transcriptional activity in the presence of CREB‐binding protein. Similar observations were made with the BTEB1‐related family members Kru¨ppel‐like family (KLF) 13/ FKLF2/BTEB3 and Sp1 on PRB transactivity.100

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VI. Conclusions and Perspectives of Progesterone Action in Endometrial Cancer Endometrial adenocarcinoma is highly associated with unopposed estrogen action. The significance of progesterone in preventing estrogen‐driven proliferation is underlined by its efficacy in eradicating endometrial hyperplasia and some endometrial cancers. While its role in preventing endometrial cancer may involve independent mechanisms to those that promote tumor cell death and regression, it is obvious that progesterone action is complex and involves numerous pathways and players. Despite the limitations of in vitro systems, which utilize endometrial cancer cell lines that have been propagated over many years, endometrial cancer cell behavior in response to progestins and the specific genes that are regulated have proven to be remarkably similar in these cell lines as those grown in xenograft models as well as in the tumor behavior from women. Thus far, in vivo and in vitro studies have shown that progesterone, through its receptor can regulate genes associated with cell cycle, apoptosis, cell adhesion, differentiation, and inflammation. Progesterone binds to either receptor A or B, which can then bind to specific sequences on promoters, in the presence of numerous other coregulators. Depending on the PR isoform and the predominant coregulators it associates with, progesterone can enhance or repress transcription of genes. The complexity of PR action is demonstrated by its differential action depending on the promoter region, length of progesterone exposure, and cell type. Further investigation is required to elucidate mechanisms of PR action in endometrial cancer. It is also noteworthy that progesterone is key in differentiating the endometrial stroma. While many studies have focused on progesterone action in the stroma in the normal cycling uterus as it pertains to pregnancy and fertility, very little is known on its role in the stroma as it pertains to endometrial cancer. Evidence is strong that progestins are effective in treating endometrial hyperplasia and some endometrial cancers and studies have demonstrated that progestins can regulate endometrial cancer cell behavior. Given the significant role that progesterone plays in the stroma, it would be worth investigating how progesterone acts through the stroma to influence endometrial cancer cells. Thus, progesterone responsiveness may be dictated not only by the hyperplastic or malignant epithelium but also by the stroma.

VII. Uterine Leiomyoma Uterine fibroids, also known as leiomyomas, are benign tumors originating from the myometrium. They are composed of smooth muscle cells and large amounts of extracellular matrix (ECM) (Fig. 3B). These tumors can range from

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a few millimeters to over 20 cm in size. Leiomyomas are common and can occur in up to 77% of women.101 The incidence in African‐American women is 60% at age 35 and over 80% by age 50 whereas Caucasian women have an incidence of 40% by age 35 and almost 70% by age 50.102 Although the tumors are considered benign, they cause significant morbidity, pain and discomfort, and excessive menstrual bleeding. Risk factors for leiomyomas include early menarche, family history, ethnicity, increased body mass index, and tissue injury. Leiomyomas are the primary indication for over 200,000 hysterectomies in the USA.103 Studies have identified possible factors responsible for the development of leiomyomas, including chromosome rearrangements, congenitally elevated ERs, hormonal changes, and injury.104 Once the disease has set in, hormones and growth factors play a prominent role in the growth and expansion of leiomyomas. To date, medical treatments for leiomyomas are limited and this is due to the fact that the mechanisms regulating the development and growth of these tumors remain unclear. There exists only one FDA approved drug for the treatment of uterine leiomyoma and thus there is a desperate need for new treatments for one of the most prevalent chronic public health problems in US women. It is hoped that a better understanding of leiomyomas at the molecular level would lead to a more effective treatment of this disease.

VIII. Progesterone Receptor Action in Leiomyoma A. Relevance of Progesterone in Uterine Leiomyomas Although the initial steps in the pathogenesis of uterine fibroids are most likely due to chromosomal aberrations and/or the effects of specific genes,105 their development is highly dependent on ovarian steroid hormones. Traditionally, estrogen has been considered the major mitogenic factor in the uterus. However, a growing body of evidence from biochemical, histological, clinical, and pharmacological studies indicates that progesterone and PR play a key role in uterine fibroid growth and development.106 Several investigators have shown an increased concentration of both PR‐A and PR‐B in leiomyoma tissue compared with adjacent myometrium.51,107,108 Furthermore, there was an increase in mitotic activity in fibroid tissue relative to the adjacent myometrial tissue during the luteal phase109 and after treatment with medroxyprogesterone acetate.110 Increased expression of the proliferation marker Ki67 in leiomyoma compared with the normal myometrium has also been described, and its upregulation was linked to progesterone.107 Epidermal growth factor (EGF) mRNA was increased in leiomyomata only during the secretory phase of the cycle, suggesting that progesterone, not estrogen, controls the expression of this important growth factor.111 In addition, in vitro studies showed that

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progesterone suppresses apoptosis and stimulates proliferation of leiomyoma cells.112–116 Progesterone markedly increased BCL2 protein expression in primary leiomyoma cell cultures.112–116 Clinical studies with both progestins and RU486 indicate that progesterone may be at least as important as estrogen for regulating fibroid growth. When used as add‐back therapy in combination with GnRH agonists, the synthetic progestins medroxyprogesterone acetate and norethindrone attenuate or reverse the inhibitory effects of GnRH agonists on leiomyoma size.117,118 The effects of pregnancy on leiomyoma size have been studied as a possible model for in vivo exposure to high levels of progesterone.119–122 The greatest increase in volume of uterine leiomyomata occurred before the 10th week of gestation.119 Those investigators who followed the leiomyoma size longitudinally after the first trimester, however, did not observe a further difference between the second and third trimester.120–122 The strongest current evidence for possible in vivo mitogenic effect of progesterone on leiomyoma growth comes from clinical trials indicating that four different antiprogestins, RU486, asoprisnil, proellex, and CDB2914 consistently reduced tumor size.123–131 The original studies of Murphy and coworkers in the 1990s suggested that RU486 might be used in the medical management of uterine leiomyomata. Pilot studies indicated that the size of leiomyomata decreased significantly after treatment with RU486.123–125 Early studies indicated that different doses of RU486 decreased leiomyoma size as well as associated excessive uterine bleeding.126 A similar endometrial histology, characterized by hyperplastic glands and stroma, was observed in patients treated with the antiprogestins RU486 and asoprisnil.127 It was subsequently shown that asoprisnil also acts primarily as a progesterone antagonist in the endometrium.132 A number of investigators have attempted to avoid the side effect of endometrial hyperplasia by decreasing the dose of RU486 to 5 mg/day; this dose has been shown to successfully decrease leiomyoma size and uterine bleeding associated with these tumors.125,130,131 Importantly, treatment with RU486 given at a dose of 5 mg/day did not cause endometrial hyperplasia.131 Despite the number of mechanisms proposed for these effects,133–143 a full understanding of the pathophysiology responsible for progesterone‐ dependent growth and the mechanisms underlying the observed therapeutic effects of antiprogestins remain unclear.

B. Progesterone Action on Genes Associated with Proliferation, Apoptosis, and ECM Deposition Although data focusing on the genes regulated in leiomyoma by progesterone are limited, studies investigating differential gene expression in leiomyomas during the menstrual cycle have provided groundwork for identifying

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those that are influenced by steroid hormones. In one study, the temporal and spatial expression of proliferative and proapoptotic molecules that could participate in leiomyoma pathogenesis was determined.144 For example, levels of Fas ligand (FasL) protein, which is associated with apoptosis, was higher during the secretory phase compared with the proliferative phase in the leiomyoma. Furthermore, higher expression of FasL was found in the leiomyoma compared to myometrium. Levels of proliferating cell nuclear antigen (PCNA), which is associated with cell proliferation, was higher during the proliferative phase in leiomyoma and levels were higher than that of paired myometrium. Lower PTEN expression, which is the phosphatase that is associated with the PI3K/AKT pathway, was detected in the leiomyoma compared to the myometrium. In this study, it was speculated that the higher FasL level in the leiomyoma may correspond to suppression of local immunity by inducing apoptosis of immune cells, while a higher level of PCNA and a lower level of PTEN may be related to increased mitogenesis and decreased apoptosis in leiomyoma. Other studies have demonstrated that leiomyoma tissues have higher PCNA levels than myometrium throughout the menstrual cycle.145 Furthermore, treatment with estradiol or progesterone increases PCNA expression in leiomyoma cells compared to controls.112 Asoprisnil, a SPRM decreased the PCNA positive rate in cultured leiomyoma cells with no difference in myometrial cells.146 The antiapoptotic bcl‐2 gene in leiomyoma has been investigated by several groups. It has been demonstrated that bcl‐2 is more highly expressed in leiomyoma than myometrium.115,147,148 Progesterone and estrogen regulate bcl‐2 expression differently. Progesterone upregulates bcl‐2 mRNA while estrogen downregulates bcl‐2 protein.115 Furthermore, Yin et al.149 found that liganded PR binds to the bcl‐2 promoter and enhances bcl‐2 transcription in primary cultured leiomyoma cells. Overexpression of the dominant negative ER in cultured leiomyoma cells decreased bcl‐2.150 It has been speculated that this reduction in ER activity results in decreased PR expression and hence a decrease in bcl‐2 expression. Asoprisnil decreased antiapoptotic bcl‐2 with a corresponding increase in TUNEL staining, cleaved caspase 3, and cleaved PARP supporting a role for PR in preventing apoptosis in these cells.133,146 ECM components are of high interest in leiomyoma pathology due to large quantities of matrix proteins found in leiomyoma. It has been demonstrated that certain ECM components and proteins are regulated by steroid hormones. Collagen type I and III mRNAs were upregulated in leiomyoma compared to myometrium during the proliferative phase of the menstrual cycle.151 In concert with this is the higher expression of MMPs in leiomyoma compared to myometrium during the secretory phase, while tissue inhibitors of MMPs (TIMPs) are more highly expressed in leiomyoma during the proliferative phase.152 Collagen binding protein fibrododulin (FMOD), which is involved

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in collagen fibril network formation, is more highly expressed in leiomyoma and myometrium during the proliferative phase of the menstrual cycle.153 Levens et al.153 also demonstrated that GnRHa treatments decreased FMOD. Asoprisnil treatment of primary leiomyoma cultures decreased TIMP1, TIMP2, collagen I, and collagen III while increasing MMP‐1, MT1‐MMP, EMMPRIN supporting that progesterone may be in involved in ECM deposition and turnover.154 Recently, investigators have uncovered that miRNA’s may play a role in leiomyoma pathogenesis. MicroRNAs are small noncoding RNA’s that inhibit translation mostly through binding to target mRNA 30 UTR. Marsh et al.155 and Wang et al.156 suggested that certain miRNA’s are differentially expressed in leiomyoma compared to myometrium. Wang et al.156 focused on the let‐7 family of miRNA’s and found that certain let‐7 family miRNA’s may be correlated with tumor size. Pan et al.157 also found that miRNA’s are differentially expressed in leiomyoma compared to myometrium and are regulated by sex steroids. More research is needed to address target genes of differentially regulated miRNA’s.

C. Growth Factor Regulation in Leiomyoma by Progesterone Given the growth properties of leiomyomas, the expression and regulation of growth factors have been studied. Here, the regulation of these growth factors by both progesterone and estrogen is highlighted. Steroid hormones can regulate EGF and its receptor (EGFR) which have been implicated in leiomyoma growth. Both the local growth factor and its receptor are expressed in leiomyoma and myometrial tissues.158 During the secretory phase of the menstrual cycle, EGF mRNA is higher in leiomyoma than myometrium.111 Interestingly, progesterone does not increase EGF‐R while increasing EGF and estrogen increases EGFR but does not increase EGF.145 This supports previous results159 that progesterone treatment but not estradiol increased EGF mRNA. In addition, asoprisnil decreased EGF mRNA.134,146 A dominant negative ER decreased immunoreactive EGF in cultured primary and immortalized leiomyoma cells.150 These data suggest that in the case of EGF and EGFR, estrogen and progesterone alter the response to the same growth factor pathway in different ways. Insulin like growth factors IGF‐I, IGF‐II, and IGF‐II receptor but not receptor type I have been detected in leiomyoma tissues at levels higher than the myometrium.160,161 Studies show that IGF‐I treatment can increase leiomyoma proliferation.162–164 IGF‐I gene expression was most abundant in leiomyomata obtained during the late proliferative phase of the cycle and was undetectable in leiomyomata from hypoestrogenic patients. SPRM, asoprisnil,

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decreased IGF‐I mRNA in leiomyoma while having no effect in myometrial cells.134,146 IGF‐II mRNA expression did not vary with phase of the menstrual cycle.165 Expression levels of IGF‐II receptor were not altered with progesterone and estrogen treatments in cultured leiomyoma cells.166 Both platelet derived growth factor (PDGF) and receptor are expressed in leiomyoma and myometrium and have been implicated in leiomyoma growth.167 PDGF is a potent mitogen for smooth muscle cells and leiomyoma cells.168–170 While it has been shown that leiomyoma tissues express higher levels of PDGF‐A and B chain mRNA levels compared to matched myometrial tissue,161,171 other studies show conflicting observations.144,172–174 Women treated with GnRHa exhibited a reduction in uterine volume which was statistically related to the decrease in PDGF expression.165 While estrogen can upregulate PDGF in cultured leiomyoma cells,164 progesterone has also been implicated in regulating PDGF expression as shown by the increased expression of PDGF‐BB expression during the secretory phase compared to the proliferative phase of the menstrual cycle in leiomyoma tissue.171 The transforming growth factor beta (TGF‐beta) family increases the expression of ECM components and are involved in reproductive tissue development and growth.175,176 Since fibroid tumors are composed mostly of ECM, examining connections between leiomyoma growth and TGF‐beta cytokines and receptors have been of interest. Consistent expression of TGF‐beta receptors type I–II and TGF‐beta 1, 2, 3 have been found in myometrium177,178 although expression in leiomyoma remains discrepant. For example, two studies showed increased expression of TGF‐beta 1 mRNA in leiomyoma compared to myometrium, while another group showed the contrary.168,177,179 In addition, TGF‐beta 2, 3, and their receptors have been found to be more highly expressed in leiomyoma than in myometrium.177 The expression of TGF‐beta 3 has been more consistent demonstrating higher expression in leiomyoma compared to myometrium.180 Furthermore, highest levels of TGF‐beta 3 were found during the secretory phase of the menstrual cycle suggesting progesterone involvement.168,181 Accordingly, the SPRM, asoprisnil, decreased TGF‐beta 3 mRNA in leiomyoma cells.134,146 Treatment with estrogen and progesterone differentially altered TGF‐beta levels in myometrium and leiomyoma.139,180

D. Activation of Signaling Pathways in Leiomyoma by Progesterone and Estrogen The role of nuclear hormone receptors in activating signaling pathways as a mechanism for leiomyoma tumor growth have been proposed in recent years. In concert with the hormonally regulated growth factors described in the above sections EGF, FGF, IGF‐I, HGF, and PDGF receptors were found to be highly

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expressed in leiomyoma tissues compared to myometrium in a receptor tyrosine kinase array.161 Specifically, expression of IGF‐IR beta was more abundant in leiomyoma as well as phosphorylated IGF‐IR and downstream effectors were more highly activated in leiomyoma. The mitogen activated protein kinase (MAPK) pathway regulates a variety of proteins involved in apoptosis and cell growth.182 Estradiol has been shown to rapidly activate the MAPK pathway in primary leiomyoma cells, including the rapid protein tyrosine phosphorylation of a subset of intracellular proteins, such as GAP, PI3K, and PLCgamma.183 Interestingly, activation of this pathway was related to E2‐induced PDGF secretion. In this study, it was proposed that PDGF, alone or in association with other growth factors, is the main growth factor involved in the proliferation response of leiomyoma cells to E2 stimulation. In accordance to this, ER‐ alpha phosphorylation was higher in leiomyoma tissues derived from patients in the proliferative phase of the menstrual cycle and this correlated with an increased phosphorylation of p44/p42 MAPK proteins in leiomyoma.184 In addition, phosphorylated p44/42 colocalized with ER‐alpha phosphorylated on serine 118, suggesting that MAPK can phosphorylate ER‐alpha in leiomyoma. ER‐alpha can also bind to the p85‐alpha regulatory subunit of PI3K, allowing for PI3K activation in MCF‐7 cells.185 There is increasing evidence that progesterone also has rapid, membrane initiated effects independent of gene transcription to alter production of second messenger and cell signal transduction pathway. Some of these rapid nongenomic effects of progesterone have been shown to be mediated through the same nuclear PR that regulates gene transcription.186,187 Pioneering work by Edwards’ group187 demonstrated that PRB can directly bind to the SH3 domain of Src kinase and thereby activate the kinase. Similarly, progesterone‐ mediated regulation of the PI3K/AKT pathway has been demonstrated in breast cancer cells as well as in rat endometrial stromal cells.186–190 Boonyaratanakornkit et al.187 showed that p85 can interact with PR in a GST‐ pull‐down system. Recently, it was shown that progesterone can rapidly phosphorylate AKT in leiomyoma cells.191 AKT phosphorylation was abrogated by PR antagonist RU 486 and PI3K inhibitor LY290004 in primary leiomyoma cells. Furthermore, the downstream targets of AKT, FOXO1, and GSK3 beta were phosphorylated upon progestin treatment indicating activation of downstream signaling components. In leiomyoma, protein levels of AKT as well as the phosphorylated form were higher than myometrium and phospho‐AKT levels dropped in leiomyoma samples taken from menopausal women.147 In concert with this is deactivated PTEN, a negative regulator of PI3K which was also increased in leiomyoma compared to myometrium, although these differences were less dramatic in menopause.192 GnRHa therapy decreased PI3K activity and AKT phosphorylation supporting that AKT activation is hormone

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dependent.193 Given the involvement of the AKT pathway in cell proliferation and survival, its activation by progesterone may be another mechanism by which this hormone promotes leiomyoma growth. Although estrogen and progesterone have distinct functions, the two hormones and their receptors interact with one another. Hodges et al. found that R5020 and MPA inhibited estradiol induced proliferation of ELT3 cells and inhibited ER activated gene transcription. These results suggest that liganded PR transdominantly suppresses ER signaling in leiomyoma. Of note is that estradiol increased expression of both PR isoforms.194 Accordingly, overexpression of dominant‐negative ER decreased PR expression in human leiomyoma cells.150

IX. Conclusions and Perspectives on Progesterone Action in Uterine Leiomyoma The importance of progesterone in promoting leiomyoma growth was initially substantiated by clinical studies as described above. The effectiveness of antiprogestins and SPRMs in reducing leiomyoma size provides strong support of progesterone being mitogenic in leiomyoma. Since the field of progesterone action in leiomyoma is one that has been understudied, it remains unclear as to how progesterone promotes growth of these tumors. Thus far, many studies have demonstrated that expression of genes and proteins are similar in leiomyoma compared to matched myometrium, however, it is the levels of expression and regulation during the menstrual cycle that differ. Whether it is demonstrated by the differential expression of genes during the menstrual phase or in response to progestin or antiprogestin treatment in vivo and in vitro, it is apparent that progesterone promotes expression of genes associated with growth and survival of leiomyoma. Regulation of growth factors and their receptors by progesterone have been proposed as another mode of progesterone action. At the transcriptional level, there are very few studies investigating the role of PR on promoters of genes in leiomyoma. Studying the recruitment of coregulators to various PR binding regions has given insight to how PR functions at the transcriptional level in leiomyomas. Finally, the rapid effects of progesterone on signaling molecules in leiomyoma provide yet another mode of progesterone action on leiomyoma growth. The involvement of classical PR in mediating these rapid effects are physiologically significant and whether progesterone membrane receptors are involved in progesterone mediated activation of kinases are unknown. Given the high incidence of leiomyoma in women, the morbidity that is associated with this disease and the financial burden of over 200,000 hysterectomies per year in the

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US alone, it is imperative that alternate therapies are developed. This can only be done with a better understanding of the molecular mechanisms associated with this disease.

X. Future Directions The significance of progesterone in the uterus is indisputable. While its action remains complex and context‐specific, it is crucial to decipher how PR works in uterine pathologies in order to be able to treat these diseases effectively. The stark contrast in the physiological response to progesterone in endometrial cancer compared to leiomyoma has important clinical implications when using progestins or antiprogestins as a mode of therapy. A clear example is the use of RU486 for decreasing leiomyoma size, which although effective, can promote endometrial hyperplasia. More information is needed on the differential action of PR in endometrial cancer cells and leiomyoma cells. Comparative studies on the mechanisms of action of PR in epithelial cells and the mesenchymal‐derived fibroblasts would provide further insight to the differences in PR action in these two diseases. Alternatively, given the differing response to progesterone in the breast and the endometrium, it would be worthwhile to conduct studies comparing PR action in breast and endometrial cancers. At the molecular level, since PR action is dictated by coregulators, an extensive analysis of proteins that complex with PR on different gene promoters after specific times of progesterone treatment and then combined with gene expression studies would be informational. Global analysis of PR binding regions using chromatin immunoprecipitation techniques, identification of coregulators using mass spectrometry, and analysis of gene expression using microarray combined together with bioinformatics analysis would be one effective approach to decipher differential PR action in uterine cells. Although unraveling the complexity of PR may seem daunting and insurmountable, the information gathered thus far provides solid groundwork to tackle this challenge. Use of innovative and state‐of‐the‐art technology will be key in moving this field forward.

References 1. Kurita T, Nakamura H. Embryology and anatomy of the uterus. In: Aplin J, Fazleabas S, Glasser S, Giudice L, editors. The Endometrium, Molecular, Cellular, and Clinical Perspectives. 2nd ed. Harwood Academic Publishers; 2007. 2. Norman AW, Litwack G. Hormones. Orlando, FL: Academic Press; 1987. 3. Clarke CL, Sutherland RL. Progestin regulation of cellular proliferation. Endocr Rev 1990;11:266–301.

PROGESTERONE IN ENDOMETRIAL CANCER AND LEIOMYOMA

75

4. Dockery P, Li TC, Rogers AW, Cooke ID, Lenton EA. The ultrastructure of the glandular epithelium in the timed endometrial biopsy. Hum Reprod 1988;3:826–34. 5. Wynn RM. Ultrastructural development of the human decidua. Am J Obstet Gynecol 1974;118:652–70. 6. Gellersen B, Brosens J. Cyclic AMP and progesterone receptor cross‐talk in human endometrium: a decidualizing affair. J Endocrinol 2003;178:357–72. 7. de Ziegler D, Bulletti C, Fanchin R, Epiney M, Brioschi PA. Contractility of the nonpregnant uterus: the follicular phase. Ann N Y Acad Sci 2001;943:172–84. 8. Boroditsky RS, Reyes FI, Winter JS, Faiman C. Maternal serum estrogen and progesterone concentrations preceding normal labor. Obstet Gynecol 1978;51:686–91. 9. Tulchinsky D, Hobel CJ, Yeager E, Marshall JR. Plama estradiol, estriol, and progesterone in human pregnancy. II. Clinical applications in Rh‐isoimmunization disease. Am J Obstet Gynecol 1972;113:766–70. 10. Walsh SW, Stanczyk FZ, Novy MJ. Daily hormonal changes in the maternal, fetal, and amniotic fluid compartments before parturition in a primate species. J Clin Endocrinol Metab 1984;58:629–39. 11. Haluska GJ, Cook MJ, Novy MJ. Inhibition and augmentation of progesterone production during pregnancy: effects on parturition in rhesus monkeys. Am J Obstet Gynecol 1997;176:682–91. 12. Selinger M, Mackenzie IZ, Gillmer MD, Phipps SL, Ferguson J. Progesterone inhibition in mid‐trimester termination of pregnancy: physiological and clinical effects. Br J Obstet Gynaecol 1987;94:1218–22. 13. Mangelsdorf DJ, Thummel C, Beato M, Herrlich P, Schutz G, Umesono K, et al. The nuclear receptor superfamily: the second decade. Cell 1995;83:835–9. 14. Robinson‐Rechavi M, Escriva Garcia H, Laudet V. The nuclear receptor superfamily. J Cell Sci 2003;116:585–6. 15. Gronemeyer H, Meyer ME, Bocquel MT, Kastner P, Turcotte B, Chambon P. Progestin receptors: isoforms and antihormone action. J Steroid Biochem Mol Biol 1991;40:271–8. 16. Kastner P, Krust A, Turcotte B, Stropp U, Tora L, Gronemeyer H, et al. Two distinct estrogen‐ regulated promoters generate transcripts encoding the two functionally different human progesterone receptor forms A and B. EMBO J 1990;9:1603–14. 17. Lessey BA, Alexander PS, Horwitz KB. The subunit structure of human breast cancer progesterone receptors: characterization by chromatography and photoaffinity labeling. Endocrinology 1983;112:1267–74. 18. McDonnell DP, Shahbaz MM, Vegeto E, Goldman ME. The human progesterone receptor A‐form functions as a transcriptional modulator of mineralocorticoid receptor transcriptional activity. J Steroid Biochem Mol Biol 1994;48:425–32. 19. Tetel MJ, Giangrande PH, Leonhardt SA, McDonnell DP, Edwards DP. Hormone‐dependent interaction between the amino‐ and carboxyl‐terminal domains of progesterone receptor in vitro and in vivo. Mol Endocrinol 1999;13:910–24. 20. Tung L, Abdel‐Hafiz H, Shen T, Harvell DM, Nitao LK, Richer JK, et al. Progesterone receptors (PR)‐B and ‐A regulate transcription by different mechanisms: AF‐3 exerts regulatory control over coactivator binding to PR‐B. Mol Endocrinol 2006;20:2656–70. 21. Tung L, Mohamed MK, Hoeffler JP, Takimoto GS, Horwitz KB. Antagonist‐occupied human progesterone B‐receptors activate transcription without binding to progesterone response elements and are dominantly inhibited by A‐receptors. Mol Endocrinol 1993;7:1256–65. 22. Vegeto E, Shahbaz MM, Wen DX, Goldman ME, O’Malley BW, McDonnell DP. Human progesterone receptor A form is a cell‐ and promoter‐specific repressor of human progesterone receptor B function. Mol Endocrinol 1993;7:1244–55.

76

KIM ET AL.

23. Chi T, Lieberman P, Ellwood K, Carey M. A general mechanism for transcriptional synergy by eukaryotic activators. Nature 1995;377:254–7. 24. Giangrande PH, Kimbrel EA, Edwards DP, McDonnell DP. The opposing transcriptional activities of the two isoforms of the human progesterone receptor are due to differential cofactor binding. Mol Cell Biol 2000;20:3102–15. 25. Wagner BL, Norris JD, Knotts TA, Weigel NL, McDonnell DP. The nuclear corepressors NCoR and SMRT are key regulators of both ligand‐ and 8‐bromo‐cyclic AMP‐dependent transcriptional activity of the human progesterone receptor. Mol Cell Biol 1998;18:1369–78. 26. Ellwood K, Huang W, Johnson R, Carey M. Multiple layers of cooperativity regulate enhanceosome‐responsive RNA polymerase II transcription complex assembly. Mol Cell Biol 1999;19:2613–23. 27. Miner JN, Yamamoto KR. The basic region of AP‐1 specifies glucocorticoid receptor activity at a composite response element. Genes Dev 1992;6:2491–501. 28. Prefontaine GG, Lemieux ME, Giffin W, Schild‐Poulter C, Pope L, LaCasse E, et al. Recruitment of octamer transcription factors to DNA by glucocorticoid receptor. Mol Cell Biol 1998;18:3416–30. 29. Schule R, Muller M, Kaltschmidt C, Renkawitz R. Many transcription factors interact synergistically with steroid receptors. Science 1988;242:1418–20. 30. O’Malley BW. Coregulators: from whence came these ‘‘master genes’’. Mol Endocrinol 2007;21:1009–13. 31. Li X, Wong J, Tsai SY, Tsai MJ, O’Malley BW. Progesterone and glucocorticoid receptors recruit distinct coactivator complexes and promote distinct patterns of local chromatin modification. Mol Cell Biol 2003;23:3763–73. 32. Xu J, Qiu Y, DeMayo FJ, Tsai SY, Tsai MJ, O’Malley BW. Partial hormone resistance in mice with disruption of the steroid receptor coactivator‐1 (SRC‐1) gene. Science 1998;279:1922–5. 33. Gehin M, Mark M, Dennefeld C, Dierich A, Gronemeyer H, Chambon P. The function of TIF2/GRIP1 in mouse reproduction is distinct from those of SRC‐1 and p/CIP. Mol Cell Biol 2002;22:5923–37. 34. Jeong JW, Lee KY, Han SJ, Aronow BJ, Lydon JP, O’Malley BW, et al. The p160 steroid receptor coactivator 2, SRC‐2, regulates murine endometrial function and regulates progesterone‐ independent and ‐dependent gene expression. Endocrinology 2007;148:4238–50. 35. Mukherjee A, Soyal SM, Fernandez‐Valdivia R, Gehin M, Chambon P, Demayo FJ, et al. Steroid receptor coactivator 2 is critical for progesterone‐dependent uterine function and mammary morphogenesis in the mouse. Mol Cell Biol 2006;26:6571–83. 36. Bamberger AM, Bamberger CM, Gellersen B, Schulte HM. Modulation of AP‐1 activity by the human progesterone receptor in endometrial adenocarcinoma cells. Proc Natl Acad Sci USA 1996;93:6169–74. 37. Gronemeyer H. Transcription activation by estrogen and progesterone receptors. Annu Rev Genet 1991;25:89–123. 38. Kalkhoven E, Wissink S, van der Saag PT, van der Burg B. Negative interaction between the RelA(p65) subunit of NF‐kappaB and the progesterone receptor. J Biol Chem 1996;271:6217–24. 39. Kim JJ, Buzzio OL, Li S, Lu Z. Role of FOXO1A in the regulation of insulin‐like growth factor‐binding protein‐1 in human endometrial cells: interaction with progesterone receptor. Biol Reprod 2005;73:833–9. 40. Takano M, Lu Z, Goto T, Fusi L, Higham J, Francis J, et al. Transcriptional cross talk between the forkhead transcription factor forkhead box O1A and the progesterone receptor coordinates cell cycle regulation and differentiation in human endometrial stromal cells. Mol Endocrinol 2007;21:2334–49.

PROGESTERONE IN ENDOMETRIAL CANCER AND LEIOMYOMA

77

41. Li X, O’Malley BW. Unfolding the action of progesterone receptors. J Biol Chem 2003;278:39261–4. 42. Carroll JS, Liu XS, Brodsky AS, Li W, Meyer CA, Szary AJ, et al. Chromosome‐wide mapping of estrogen receptor binding reveals long‐range regulation requiring the forkhead protein FoxA1. Cell 2005;122:33–43. 43. Schuur ER, Loktev AV, Sharma M, Sun Z, Roth RA, Weigel RJ. Ligand‐dependent interaction of estrogen receptor‐alpha with members of the forkhead transcription factor family. J Biol Chem 2001;276:33554–60. 44. Zhao HH, Herrera RE, Coronado‐Heinsohn E, Yang MC, Ludes‐Meyers JH, Seybold‐ Tilson KJ, et al. Forkhead homologue in rhabdomyosarcoma functions as a bifunctional nuclear receptor‐interacting protein with both coactivator and corepressor functions. J Biol Chem 2001;276:27907–12. 45. Mulac‐Jericevic B, Lydon JP, DeMayo FJ, Conneely OM. Defective mammary gland morphogenesis in mice lacking the progesterone receptor B isoform. Proc Natl Acad Sci USA 2003;100:9744–9. 46. Mulac‐Jericevic B, Mullinax RA, DeMayo FJ, Lydon JP, Conneely OM. Subgroup of reproductive functions of progesterone mediated by progesterone receptor‐B isoform. Science 2000;289:1751–4. 47. Condon JC, Hardy DB, Kovaric K, Mendelson CR. Up‐regulation of the progesterone receptor (PR)‐C isoform in laboring myometrium by activation of nuclear factor‐kappaB may contribute to the onset of labor through inhibition of PR function. Mol Endocrinol 2006;20:764–75. 48. Garcia E, Bouchard P, De Brux J, Berdah J, Frydman R, Schaison G, et al. Use of immunocytochemistry of progesterone and estrogen receptors for endometrial dating. J Clin Endocrinol Metab 1988;67:80–7. 49. Mylonas I, Jeschke U, Shabani N, Kuhn C, Kunze S, Dian D, et al. Steroid receptors ERalpha, ERbeta, PR‐A and PR‐B are differentially expressed in normal and atrophic human endometrium. Histol Histopathol 2007;22:169–76. 50. Snijders MP, de Goeij AF, Debets‐Te Baerts MJ, Rousch MJ, Koudstaal J, Bosman FT. Immunocytochemical analysis of oestrogen receptors and progesterone receptors in the human uterus throughout the menstrual cycle and after the menopause. J Reprod Fertil 1992;94:363–71. 51. Nisolle M, Gillerot S, Casanas‐Roux F, Squifflet J, Berliere M, Donnez J. Immunohistochemical study of the proliferation index, oestrogen receptors and progesterone receptors A and B in leiomyomata and normal myometrium during the menstrual cycle and under gonadotrophin‐releasing hormone agonist therapy. Hum Reprod 1999;14:2844–50. 52. Vienonen A, Miettinen S, Blauer M, Martikainen PM, Tomas E, Heinonen PK, et al. Expression of nuclear receptors and cofactors in human endometrium and myometrium. J Soc Gynecol Investig 2004;11:104–12. 53. Jemal A, Siegel R, Ward E, Hao Y, Xu J, Murray T, et al. Cancer statistics, 2008. CA Cancer J Clin 2008;58:71–96. 54. Clement PB, Young RH. Endometrioid carcinoma of the uterine corpus: a review of its pathology with emphasis on recent advances and problematic aspects. Adv Anat Pathol 2002;9:145–84. 55. Creasman WT, Eddy GL. Recent advances in endometrial cancer. Semin Surg Oncol 1990;6:339–42. 56. Judd HL, Judd GE, Lucas WE, Yen SS. Endocrine function of the postmenopausal ovary: concentration of androgens and estrogens in ovarian and peripheral vein blood. J Clin Endocrinol Metab 1974;39:1020–4.

78

KIM ET AL.

57. Kirschner MA, Schneider G, Ertel NH, Worton E. Obesity, androgens, estrogens, and cancer risk. Cancer Res 1982;42:3281s–5s. 58. Munstedt K, Grant P, Woenckhaus J, Roth G, Tinneberg HR. Cancer of the endometrium: current aspects of diagnostics and treatment. World J Surg Oncol 2004;2:24. 59. Ryan AJ, Susil B, Jobling TW, Oehler MK. Endometrial cancer. Cell Tissue Res 2005;322:53–61. 60. Cohen CJ, Rahaman J. Endometrial cancer. Management of high risk and recurrence including the tamoxifen controversy. Cancer 1995;76:2044–52. 61. Jadoul P, Donnez J. Conservative treatment may be beneficial for young women with atypical endometrial hyperplasia or endometrial adenocarcinoma. Fertil Steril 2003;80:1315–24. 62. Randall TC, Kurman RJ. Progestin treatment of atypical hyperplasia and well‐differentiated carcinoma of the endometrium in women under age 40. Obstet Gynecol 1997;90:434–40. 63. Gal D, Edman CD, Vellios F, Forney JP. Long‐term effect of megestrol acetate in the treatment of endometrial hyperplasia. Am J Obstet Gynecol 1983;146:316–22. 64. Ferenczy A, Gelfand M. The biologic significance of cytologic atypia in progestogen‐treated endometrial hyperplasia. Am J Obstet Gynecol 1989;160:126–31. 65. Lai CH, Huang HJ. The role of hormones for the treatment of endometrial hyperplasia and endometrial cancer. Curr Opin Obstet Gynecol 2006;18:29–34. 66. Martin‐Hirsch PL, Lilford RJ, Jarvis GJ. Adjuvant progestagen therapy for the treatment of endometrial cancer: review and meta‐analyses of published randomised controlled trials. Eur J Obstet Gynecol Reprod Biol 1996;65:201–7. 67. Benshushan A. Endometrial adenocarcinoma in young patients: evaluation and fertility‐ preserving treatment. Eur J Obstet Gynecol Reprod Biol 2004;117:132–7. 68. Crissman JD, Azoury RS, Barnes AE, Schellhas HF. Endometrial carcinoma in women 40 years of age or younger. Obstet Gynecol 1981;57:699–704. 69. Gallup DG, Stock RJ. Adenocarcinoma of the endometrium in women 40 years of age or younger. Obstet Gynecol 1984;64:417–20. 70. Lowe MP, Bender D, Sood AK, Davis W, Syrop CH, Sorosky JI. Two successful pregnancies after conservative treatment of endometrial cancer and assisted reproduction. Fertil Steril 2002;77:188–9. 71. Chiva L, Lapuente F, Gonzalez‐Cortijo L, Carballo N, Garcia JF, Rojo A, et al. Sparing fertility in young patients with endometrial cancer. Gynecol Oncol 2008;111:S101–4. 72. Ushijima K, Yahata H, Yoshikawa H, Konishi I, Yasugi T, Saito T, et al. Multicenter phase II study of fertility‐sparing treatment with medroxyprogesterone acetate for endometrial carcinoma and atypical hyperplasia in young women. J Clin Oncol 2007;25:2798–803. 73. Leslie KK, Stein MP, Kumar NS, Dai D, Stephens J, Wandinger‐Ness A, et al. Progesterone receptor isoform identification and subcellular localization in endometrial cancer. Gynecol Oncol 2005;96:32–41. 74. Smid‐Koopman E, Blok LJ, Kuhne LC, Burger CW, Helmerhorst TJ, Brinkmann AO, et al. Distinct functional differences of human progesterone receptors A and B on gene expression and growth regulation in two endometrial carcinoma cell lines. J Soc Gynecol Investig 2003;10:49–57. 75. Hanekamp EE, Kuhne LM, Grootegoed JA, Burger CW, Blok LJ. Progesterone receptor A and B expression and progestagen treatment in growth and spread of endometrial cancer cells in nude mice. Endocr Relat Cancer 2004;11:831–41. 76. Fujimoto J, Ichigo S, Hori M, Nishigaki M, Tamaya T. Expression of progesterone receptor form A and B mRNAs in gynecologic malignant tumors. Tumour Biol 1995;16:254–60. 77. Arnett‐Mansfield RL, deFazio A, Wain GV, Jaworski RC, Byth K, Mote PA, et al. Relative expression of progesterone receptors A and B in endometrioid cancers of the endometrium. Cancer Res 2001;61:4576–82.

PROGESTERONE IN ENDOMETRIAL CANCER AND LEIOMYOMA

79

78. Kumar NS, Richer J, Owen G, Litman E, Horwitz KB, Leslie KK. Selective down‐regulation of progesterone receptor isoform B in poorly differentiated human endometrial cancer cells: implications for unopposed estrogen action. Cancer Res 1998;58:1860–5. 79. Dai D, Wolf DM, Litman ES, White MJ, Leslie KK. Progesterone inhibits human endometrial cancer cell growth and invasiveness: down‐regulation of cellular adhesion molecules through progesterone B receptors. Cancer Res 2002;62:881–6. 80. Ueda M, Fujii H, Yoshizawa K, Abe F, Ueki M. Effects of sex steroids and growth factors on migration and invasion of endometrial adenocarcinoma SNG‐M cells in vitro. Jpn J Cancer Res 1996;87:524–33. 81. Saito T, Mizumoto H, Tanaka R, Satohisa S, Adachi K, Horie M, et al. Overexpressed progesterone receptor form B inhibit invasive activity suppressing matrix metalloproteinases in endometrial carcinoma cells. Cancer Lett 2004;209:237–43. 82. Di Nezza LA, Jobling T, Salamonsen LA. Progestin suppresses matrix metalloproteinase production in endometrial cancer. Gynecol Oncol 2003;89:325–33. 83. Jaffe RC, Ferguson‐Gottschall SD, Gao W, Beam C, Fazleabas AT. Histone deacetylase inhibition and progesterone act synergistically to stimulate baboon glycodelin gene expression. J Mol Endocrinol 2007;38:401–7. 84. Uchida H, Maruyama T, Nagashima T, Asada H, Yoshimura Y. Histone deacetylase inhibitors induce differentiation of human endometrial adenocarcinoma cells through up‐regulation of glycodelin. Endocrinology 2005;146:5365–73. 85. Uchida H, Maruyama T, Ono M, Ohta K, Kajitani T, Masuda H, et al. Histone deacetylase inhibitors stimulate cell migration in human endometrial adenocarcinoma cells through up‐ regulation of glycodelin. Endocrinology 2007;148:896–902. 86. Ohta K, Maruyama T, Uchida H, Ono M, Nagashima T, Arase T, et al. Glycodelin blocks progression to S phase and inhibits cell growth: a possible progesterone‐induced regulator for endometrial epithelial cell growth. Mol Hum Reprod 2008;14:17–22. 87. Ward EC, Hoekstra AV, Blok LJ, Hanifi‐Moghaddam P, Lurain JR, Singh DK, et al. The regulation and function of the forkhead transcription factor, Forkhead box O1, is dependent on the progesterone receptor in endometrial carcinoma. Endocrinology 2008;149:1942–50. 88. Goto T, Takano M, Albergaria A, Briese J, Pomeranz KM, Cloke B, et al. Mechanism and functional consequences of loss of FOXO1 expression in endometrioid endometrial cancer cells. Oncogene 2008;27:9–19. 89. Shiozawa T, Nikaido T, Nakayama K, Lu X, Fujii S. Involvement of cyclin‐dependent kinase inhibitor p27Kip1 in growth inhibition of endometrium in the secretory phase and of hyperplastic endometrium treated with progesterone. Mol Hum Reprod 1998;4:899–905. 90. Watanabe J, Watanabe K, Jobo T, Kamata Y, Kawaguchi M, Imai M, et al. Significance of p27 as a predicting marker for medroxyprogesterone acetate therapy against endometrial endometrioid adenocarcinoma. Int J Gynecol Cancer 2006;16(Suppl 1):452–7. 91. Paulssen RH, Moe B, Gronaas H, Orbo A. Gene expression in endometrial cancer cells (Ishikawa) after short time high dose exposure to progesterone. Steroids 2008;73:116–28. 92. Hanekamp EE, Kuhne EC, Smid‐Koopman E, de Ruiter PE, Chadha‐Ajwani S, Brinkmann AO, et al. Loss of progesterone receptor may lead to an invasive phenotype in human endometrial cancer. Eur J Cancer 2002;38(Suppl 6):S71–2. 93. Davies S, Dai D, Wolf DM, Leslie KK. Immunomodulatory and transcriptional effects of progesterone through progesterone A and B receptors in Hec50co poorly differentiated endometrial cancer cells. J Soc Gynecol Investig 2004;11:494–9. 94. Vereide AB, Kaino T, Sager G, Orbo A. Bcl‐2, BAX, and apoptosis in endometrial hyperplasia after high dose gestagen therapy: a comparison of responses in patients treated with intrauterine levonorgestrel and systemic medroxyprogesterone. Gynecol Oncol 2005;97:740–50.

80

KIM ET AL.

95. Dai D, Kumar NS, Wolf DM, Leslie KK. Molecular tools to reestablish progestin control of endometrial cancer cell proliferation. Am J Obstet Gynecol 2001;184:790–7. 96. Dai D, Litman ES, Schonteich E, Leslie KK. Progesterone regulation of activating protein‐1 transcriptional activity: a possible mechanism of progesterone inhibition of endometrial cancer cell growth. J Steroid Biochem Mol Biol 2003;87:123–31. 97. Davies S, Dai D, Feldman I, Pickett G, Leslie KK. Identification of a novel mechanism of NF‐ kappaB inactivation by progesterone through progesterone receptors in Hec50co poorly differentiated endometrial cancer cells: induction of A20 and ABIN‐2. Gynecol Oncol 2004;94:463–70. 98. Gao J, Mazella J, Seppala M, Tseng L. Ligand activated hPR modulates the glycodelin promoter activity through the Sp1 sites in human endometrial adenocarcinoma cells. Mol Cell Endocrinol 2001;176:97–102. 99. Salih SM, Salama SA, Jamaluddin M, Fadl AA, Blok LJ, Burger CW, et al. Progesterone‐ mediated regulation of catechol‐O‐methyl transferase expression in endometrial cancer cells. Reprod Sci 2008;15:210–20. 100. Zhang XL, Zhang D, Michel FJ, Blum JL, Simmen FA, Simmen RC. Selective interactions of Kruppel‐like factor 9/basic transcription element‐binding protein with progesterone receptor isoforms A and B determine transcriptional activity of progesterone‐responsive genes in endometrial epithelial cells. J Biol Chem 2003;278:21474–82. 101. Cramer SF, Patel A. The frequency of uterine leiomyomas. Am J Clin Pathol 1990;94:435–8. 102. Day Baird D, Dunson DB, Hill MC, Cousins D, Schectman JM. High cumulative incidence of uterine leiomyoma in black and white women: ultrasound evidence. Am J Obstet Gynecol 2003;188:100–7. 103. Farquhar CM, Steiner CA. Hysterectomy rates in the United States 1990–1997. Obstet Gynecol 2002;99:229–34. 104. Parker WH. Etiology, symptomatology, and diagnosis of uterine myomas. Fertil Steril 2007;87:725–36. 105. Williams AJ, Powell WL, Collins T, Morton CC. HMGI(Y) expression in human uterine leiomyomata. Involvement of another high‐mobility group architectural factor in a benign neoplasm. Am J Pathol 1997;150:911–8. 106. Cermik D, Arici A, Taylor HS. Coordinated regulation of HOX gene expression in myometrium and uterine leiomyoma. Fertil Steril 2002;78:979–84. 107. Brandon DD, Bethea CL, Strawn EY, Novy MJ, Burry KA, Harrington MS, et al. Progesterone receptor messenger ribonucleic acid and protein are overexpressed in human uterine leiomyomas. Am J Obstet Gynecol 1993;169:78–85. 108. Englund K, Blanck A, Gustavsson I, Lundkvist U, Sjoblom P, Norgren A, et al. Sex steroid receptors in human myometrium and fibroids: changes during the menstrual cycle and gonadotropin‐releasing hormone treatment. J Clin Endocrinol Metab 1998;83:4092–6. 109. Kawaguchi K, Fujii S, Konishi I, Nanbu Y, Nonogaki H, Mori T. Mitotic activity in uterine leiomyomas during the menstrual cycle. Am J Obstet Gynecol 1989;160:637–41. 110. Tiltman AJ. The effect of progestins on the mitotic activity of uterine fibromyomas. Int J Gynecol Pathol 1985;4:89–96. 111. Harrison‐Woolrych ML, Charnock‐Jones DS, Smith SK. Quantification of messenger ribonucleic acid for epidermal growth factor in human myometrium and leiomyomata using reverse transcriptase polymerase chain reaction. J Clin Endocrinol Metab 1994;78:1179–84. 112. Maruo T, Matsuo H, Samoto T, Shimomura Y, Kurachi O, Gao Z, et al. Effects of progesterone on uterine leiomyoma growth and apoptosis. Steroids 2000;65:585–92. 113. Kurachi O, Matsuo H, Samoto T, Maruo T. Tumor necrosis factor‐alpha expression in human uterine leiomyoma and its down‐regulation by progesterone. J Clin Endocrinol Metab 2001;86:2275–80.

PROGESTERONE IN ENDOMETRIAL CANCER AND LEIOMYOMA

81

114. Matsuo H, Kurachi O, Shimomura Y, Samoto T, Maruo T. Molecular bases for the actions of ovarian sex steroids in the regulation of proliferation and apoptosis of human uterine leiomyoma. Oncology 1999;57(Suppl 2):49–58. 115. Matsuo H, Maruo T, Samoto T. Increased expression of Bcl‐2 protein in human uterine leiomyoma and its up‐regulation by progesterone. J Clin Endocrinol Metab 1997;82:293–9. 116. Maruo T, Matsuo H, Shimomura Y, Kurachi O, Gao Z, Nakago S, et al. Effects of progesterone on growth factor expression in human uterine leiomyoma. Steroids 2003;68:817–24. 117. Carr BR, Marshburn PB, Weatherall PT, Bradshaw KD, Breslau NA, Byrd W, et al. An evaluation of the effect of gonadotropin‐releasing hormone analogs and medroxyprogesterone acetate on uterine leiomyomata volume by magnetic resonance imaging: a prospective, randomized, double blind, placebo‐controlled, crossover trial. J Clin Endocrinol Metab 1993;76:1217–23. 118. Friedman AJ, Daly M, Juneau‐Norcross M, Rein MS, Fine C, Gleason R, et al. A prospective, randomized trial of gonadotropin‐releasing hormone agonist plus estrogen‐progestin or progestin ‘‘add‐back’’ regimens for women with leiomyomata uteri. J Clin Endocrinol Metab 1993;76:1439–45. 119. Rosati P, Exacoustos C, Mancuso S. Longitudinal evaluation of uterine myoma growth during pregnancy. A sonographic study. J Ultrasound Med 1992;11:511–5. 120. Aharoni A, Reiter A, Golan D, Paltiely Y, Sharf M. Patterns of growth of uterine leiomyomas during pregnancy. A prospective longitudinal study. Br J Obstet Gynaecol 1988;95:510–3. 121. Neiger R, Sonek JD, Croom CS, Ventolini G. Pregnancy‐related changes in the size of uterine leiomyomas. J Reprod Med 2006;51:671–4. 122. Hammoud AO, Asaad R, Berman J, Treadwell MC, Blackwell S, Diamond MP. Volume change of uterine myomas during pregnancy: do myomas really grow?. J Minim Invasive Gynecol 2006;13:386–90. 123. Murphy AA, Kettel LM, Morales AJ, Roberts VJ, Yen SS. Regression of uterine leiomyomata in response to the antiprogesterone RU 486. J Clin Endocrinol Metab 1993;76:513–7. 124. Murphy AA, Morales AJ, Kettel LM, Yen SS. Regression of uterine leiomyomata to the antiprogesterone RU486: dose‐response effect. Fertil Steril 1995;64:187–90. 125. Eisinger SH, Meldrum S, Fiscella K, le Roux HD, Guzick DS. Low‐dose mifepristone for uterine leiomyomata. Obstet Gynecol 2003;101:243–50. 126. Murphy LL, Mahesh VB. Selective release of follicle‐stimulating hormone and luteinizing hormone by 5a`‐dihydroprogesterone and 3a`,5a`‐ tetrahydroprogesterone in pregnant mare’s serum gonadotropin‐ primed immature rats exposed to constant light. Biol Reprod 1985;32:795–803. 127. Williams AR, Critchley HO, Osei J, Ingamells S, Cameron I.T, Han C, et al. The effects of the selective progesterone receptor modulator asoprisnil on the morphology of uterine tissues after 3 months treatment in patients with symptomatic uterine leiomyomata. Hum Reprod 2007. 128. Steinauer J, Pritts EA, Jackson R, Jacoby AF. Systematic review of mifepristone for the treatment of uterine leiomyomata. Obstet Gynecol 2004;103:1331–6. 129. Chwalisz K, Larsen L, Mattia‐Goldberg C, Edmonds A, Elger W, Winkel CA. A randomized, controlled trial of asoprisnil, a novel selective progesterone receptor modulator, in women with uterine leiomyomata. Fertil Steril 2007;87:1399–412. 130. Eisinger SH, Bonfiglio T, Fiscella K, Meldrum S, Guzick DS. Twelve‐month safety and efficacy of low‐dose mifepristone for uterine myomas. J Minim Invasive Gynecol 2005;12:227–33. 131. Fiscella K, Eisinger SH, Meldrum S, Feng C, Fisher SG, Guzick DS. Effect of mifepristone for symptomatic leiomyomata on quality of life and uterine size: a randomized controlled trial. Obstet Gynecol 2006;108:1381–7.

82

KIM ET AL.

132. Madauss KP, Grygielko ET, Deng SJ, Sulpizio AC, Stanley TB, Wu C, et al. A structural and in vitro characterization of asoprisnil: a selective progesterone receptor modulator. Mol Endocrinol 2007;21:1066–81. 133. Chen W, Ohara N, Wang J, Xu Q, Liu J, Morikawa A, et al. A novel selective progesterone receptor modulator asoprisnil (J867) inhibits proliferation and induces apoptosis in cultured human uterine leiomyoma cells in the absence of comparable effects on myometrial cells. J Clin Endocrinol Metab 2006;91:1296–304. 134. Wang J, Ohara N, Wang Z, Chen W, Morikawa A, Sasaki H, et al. A novel selective progesterone receptor modulator asoprisnil (J867) down‐regulates the expression of EGF, IGF‐I, TGFbeta3 and their receptors in cultured uterine leiomyoma cells. Hum Reprod 2006;21:1869–77. 135. Sasaki H, Ohara N, Xu Q, Wang J, DeManno DA, Chwalisz K, et al. A novel selective progesterone receptor modulator asoprisnil activates tumor necrosis factor‐related apoptosis‐inducing ligand (TRAIL)‐mediated signaling pathway in cultured human uterine leiomyoma cells in the absence of comparable effects on myometrial cells. J Clin Endocrinol Metab 2007;92:616–23. 136. Stewart EA, Austin DJ, Jain P, Penglase MD, Nowak RA. RU486 suppresses prolactin production in explant cultures of leiomyoma and myometrium. Fertil Steril 1996;65:1119–24. 137. Hyder SM, Huang JC, Nawaz Z, Boettger‐Tong H, Makela S, Chiappetta C, et al. Regulation of vascular endothelial growth factor expression by estrogens and progestins. Environ Health Perspect 2000;108(Suppl 5):785–90. 138. Wang H, Jin J. [Effects of mifepristone on estrogen and progestin receptors in human uterine leiomyoma]. Zhonghua Fu Chan Ke Za Zhi 2000;35:79–81. 139. Chegini N, Ma C, Tang XM, Williams RS. Effects of GnRH analogues, ‘add‐back’ steroid therapy, antiestrogen and antiprogestins on leiomyoma and myometrial smooth muscle cell growth and transforming growth factor‐beta expression. Mol Hum Reprod 2002;8:1071–8. 140. Jeng YJ, Suarez VR, Izban MG, Wang HQ, Soloff MS. Progesterone‐induced sphingosine kinase‐1 expression in the rat uterus during pregnancy and signaling consequences. Am J Physiol Endocrinol Metab 2007;292:E1110–21. 141. Kettel LM, Murphy AA, Morales AJ, Yen SS. Clinical efficacy of the antiprogesterone RU486 in the treatment of endometriosis and uterine fibroids. Hum Reprod 1994;9(Suppl 1):116–20. 142. Murphy AA, Castellano PZ. RU486: pharmacology and potential use in the treatment of endometriosis and leiomyomata uteri. Curr Opin Obstet Gynecol 1994;6:269–78. 143. Ramachandran S, Song MQ, Lowe E, Dominguez CE, Parthasarathy S, Murphy AA. RU486 inhibits expression of lysophosphatidic acid induced glycodelin. Am J Obstet Gynecol 2005;192:1285–93; discussion 1293–84. 144. Kayisli UA, Berkkanoglu M, Kizilay G, Senturk L, Arici A. Expression of proliferative and preapoptotic molecules in human myometrium and leiomyoma throughout the menstrual cycle. Reprod Sci 2007;14:678–86. 145. Shimomura Y, Matsuo H, Samoto T, Maruo T. Up‐regulation by progesterone of proliferating cell nuclear antigen and epidermal growth factor expression in human uterine leiomyoma. J Clin Endocrinol Metab 1998;83:2192–8. 146. Ohara N, Morikawa A, Chen W, Wang J, DeManno DA, Chwalisz K, et al. Comparative effects of SPRM asoprisnil (J867) on proliferation, apoptosis, and the expression of growth factors in cultured uterine leiomyoma cells and normal myometrial cells. Reprod Sci 2007;14:20–7. 147. Kovacs KA, Lengyel F, Kornyei JL, Vertes Z, Szabo I, Sumegi B, et al. Differential expression of Akt/protein kinase B, Bcl‐2 and Bax proteins in human leiomyoma and myometrium. J Steroid Biochem Mol Biol 2003;87:233–40.

PROGESTERONE IN ENDOMETRIAL CANCER AND LEIOMYOMA

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148. Wu X, Blanck A, Olovsson M, Henriksen R, Lindblom B. Expression of Bcl‐2, Bcl‐x, Mcl‐1, Bax and Bak in human uterine leiomyomas and myometrium during the menstrual cycle and after menopause. J Steroid Biochem Mol Biol 2002;80:77–83. 149. Yin P, Lin Z, Cheng YH, Marsh EE, Utsunomiya H, Ishikawa H, et al. Progesterone receptor regulates Bcl‐2 gene expression through direct binding to its promoter region in uterine leiomyoma cells. J Clin Endocrinol Metab 2007;92:4459–66. 150. Hassan MH, Salama SA, Arafa HM, Hamada FM, Al‐Hendy A. Adenovirus‐mediated delivery of a dominant‐negative estrogen receptor gene in uterine leiomyoma cells abrogates estrogen‐ and progesterone‐regulated gene expression. J Clin Endocrinol Metab 2007;92:3949–57. 151. Stewart EA, Friedman AJ, Peck K, Nowak RA. Relative overexpression of collagen type I and collagen type III messenger ribonucleic acids by uterine leiomyomas during the proliferative phase of the menstrual cycle. J Clin Endocrinol Metab 1994;79:900–6. 152. Dou Q, Tarnuzzer RW, Williams RS, Schultz GS, Chegini N. Differential expression of matrix metalloproteinases and their tissue inhibitors in leiomyomata: a mechanism for gonadotrophin releasing hormone agonist‐induced tumour regression. Mol Hum Reprod 1997;3:1005–14. 153. Levens E, Luo X, Ding L, Williams RS, Chegini N. Fibromodulin is expressed in leiomyoma and myometrium and regulated by gonadotropin‐releasing hormone analogue therapy and TGF‐beta through Smad and MAPK‐mediated signalling. Mol Hum Reprod 2005;11:489–94. 154. Morikawa A, Ohara N, Xu Q, Nakabayashi K, DeManno DA, Chwalisz K, et al. Selective progesterone receptor modulator asoprisnil down‐regulates collagen synthesis in cultured human uterine leiomyoma cells through up‐regulating extracellular matrix metalloproteinase inducer. Hum Reprod 2008;23:944–51. 155. Marsh EE, Lin Z, Yin P, Milad M, Chakravarti D, Bulun SE. Differential expression of microRNA species in human uterine leiomyoma versus normal myometrium. Fertil Steril 2008;89:1771–6. 156. Wang T, Zhang X, Obijuru L, Laser J, Aris V, Lee P, et al. A micro‐RNA signature associated with race, tumor size, and target gene activity in human uterine leiomyomas. Genes Chromosomes Cancer 2007;46:336–47. 157. Pan Q, Luo X, Chegini N. Differential expression of microRNAs in myometrium and leiomyomas and regulation by ovarian steroids. J Cell Mol Med 2008;12:227–40. 158. Rossi MJ, Chegini N, Masterson BJ. Presence of epidermal growth factor, platelet‐derived growth factor, and their receptors in human myometrial tissue and smooth muscle cells: their action in smooth muscle cells in vitro. Endocrinology 1992;130:1716–27. 159. Yeh J, Rein M, Nowak R. Presence of messenger ribonucleic acid for epidermal growth factor (EGF) and EGF receptor demonstrable in monolayer cell cultures of myometria and leiomyomata. Fertil Steril 1991;56:997–1000. 160. Chandrasekhar Y, Heiner J, Osuamkpe C, Nagamani M. Insulin‐like growth factor I and II binding in human myometrium and leiomyomas. Am J Obstet Gynecol 1992;166:64–9. 161. Yu L, Saile K, Swartz CD, He H, Zheng X, Kissling GE, et al. Differential expression of receptor tyrosine kinases (RTKs) and IGF‐I pathway activation in human uterine leiomyomas. Mol Med 2008;14:264–75. 162. Gao Z, Matsuo H, Wang Y, Nakago S, Maruo T. Up‐regulation by IGF‐I of proliferating cell nuclear antigen and Bcl‐2 protein expression in human uterine leiomyoma cells. J Clin Endocrinol Metab 2001;86:5593–9. 163. Strawn EY, Jr., Novy MJ, Burry KA, Bethea CL. Insulin‐like growth factor I promotes leiomyoma cell growth in vitro. Am J Obstet Gynecol 1995;172:1837–43; discussion 1843–4. 164. van der Ven LT, Gloudemans T, Roholl PJ, van Buul‐Offers SC, Bladergroen BA, Welters MJ, et al. Growth advantage of human leiomyoma cells compared to normal smooth‐muscle cells due to enhanced sensitivity toward insulin‐like growth factor I. Int J Cancer 1994;59:427–34.

84

KIM ET AL.

165. Giudice LC, Irwin JC, Dsupin BA, Pannier EM, Jin IH, Vu TH, et al. Insulin‐like growth factor (IGF), IGF binding protein (IGFBP), and IGF receptor gene expression and IGFBP synthesis in human uterine leiomyomata. Hum Reprod 1993;8:1796–806. 166. Yamada T, Nakago S, Kurachi O, Wang J, Takekida S, Matsuo H, et al. Progesterone down‐ regulates insulin‐like growth factor‐I expression in cultured human uterine leiomyoma cells. Hum Reprod 2004;19:815–21. 167. Palman C, Bowen‐Pope DF, Brooks JJ. Platelet‐derived growth factor receptor (beta‐subunit) immunoreactivity in soft tissue tumors. Lab Invest 1992;66:108–15. 168. Arici A, Sozen I. Expression, menstrual cycle‐dependent activation, and bimodal mitogenic effect of transforming growth factor‐beta1 in human myometrium and leiomyoma. Am J Obstet Gynecol 2003;188:76–83. 169. Fayed YM, Tsibris JC, Langenberg PW, Robertson AL, Jr.. Human uterine leiomyoma cells: binding and growth responses to epidermal growth factor, platelet‐derived growth factor, and insulin. Lab Invest 1989;60:30–7. 170. Ross R, Bowen‐Pope DF, Raines EW. Platelet‐derived growth factor: its potential roles in wound healing, atherosclerosis, neoplasia, and growth and development. Ciba Found Symp 1985;116:98–112. 171. Liang M, Wang H, Zhang Y, Lu S, Wang Z. Expression and functional analysis of platelet‐ derived growth factor in uterine leiomyomata. Cancer Biol Ther 2006;5:28–33. 172. Boehm KD, Daimon M, Gorodeski IG, Sheean LA, Utian WH, Ilan J. Expression of the insulin‐like and platelet‐derived growth factor genes in human uterine tissues. Mol Reprod Dev 1990;27:93–101. 173. Mangrulkar RS, Ono M, Ishikawa M, Takashima S, Klagsbrun M, Nowak RA. Isolation and characterization of heparin‐binding growth factors in human leiomyomas and normal myometrium. Biol Reprod 1995;53:636–46. 174. Hwu YM, Li SH, Lee RK, Tsai YH, Yeh TS, Lin SY. Increased expression of platelet‐derived growth factor C messenger ribonucleic acid in uterine leiomyomata. Fertil Steril 2008;89:468–71. 175. Ingman WV, Robertson SA. Defining the actions of transforming growth factor beta in reproduction. Bioessays 2002;24:904–14. 176. Verrecchia F, Mauviel A. Transforming growth factor‐beta and fibrosis. World J Gastroenterol 2007;13:3056–62. 177. Dou Q, Zhao Y, Tarnuzzer RW, Rong H, Williams RS, Schultz GS, et al. Suppression of transforming growth factor‐beta (TGF beta) and TGF beta receptor messenger ribonucleic acid and protein expression in leiomyomata in women receiving gonadotropin‐releasing hormone agonist therapy. J Clin Endocrinol Metab 1996;81:3222–30. 178. Tang XM, Dou Q, Zhao Y, McLean F, Davis J, Chegini N. The expression of transforming growth factor‐beta s and TGF‐beta receptor mRNA and protein and the effect of TGF‐beta s on human myometrial smooth muscle cells in vitro. Mol Hum Reprod 1997;3:233–40. 179. Chegini N, Tang XM, Ma C. Regulation of transforming growth factor‐beta1 expression by granulocyte macrophage‐colony‐stimulating factor in leiomyoma and myometrial smooth muscle cells. J Clin Endocrinol Metab 1999;84:4138–43. 180. De Falco M, Staibano S, D’Armiento FP, Mascolo M, Salvatore G, Busiello A, et al. Preoperative treatment of uterine leiomyomas: clinical findings and expression of transforming growth factor‐beta3 and connective tissue growth factor. J Soc Gynecol Investig 2006;13:297–303. 181. Lee BS, Nowak RA. Human leiomyoma smooth muscle cells show increased expression of transforming growth factor‐beta 3 (TGF beta 3) and altered responses to the antiproliferative effects of TGF beta. J Clin Endocrinol Metab 2001;86:913–20.

PROGESTERONE IN ENDOMETRIAL CANCER AND LEIOMYOMA

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182. McCubrey JA, Steelman LS, Chappell WH, Abrams SL, Wong EW, Chang F, et al. Roles of the Raf/MEK/ERK pathway in cell growth, malignant transformation and drug resistance. Biochim Biophys Acta 2007;1773:1263–84. 183. Barbarisi A, Petillo O, Di Lieto A, Melone MA, Margarucci S, Cannas M, et al. 17‐Beta estradiol elicits an autocrine leiomyoma cell proliferation: evidence for a stimulation of protein kinase‐dependent pathway. J Cell Physiol 2001;186:414–24. 184. Hermon TL, Moore AB, Yu L, Kissling GE, Castora FJ, Dixon D. Estrogen receptor alpha (ERalpha) phospho‐serine‐118 is highly expressed in human uterine leiomyomas compared to matched myometrium. Virchows Arch 2008;453:557–69. 185. Simoncini T, Hafezi‐Moghadam A, Brazil DP, Ley K, Chin WW, Liao JK. Interaction of oestrogen receptor with the regulatory subunit of phosphatidylinositol‐3‐OH kinase. Nature 2000;407:538–41. 186. Carnevale RP, Proietti CJ, Salatino M, Urtreger A, Peluffo G, Edwards DP, et al. Progestin effects on breast cancer cell proliferation, proteases activation, and in vivo development of metastatic phenotype all depend on progesterone receptor capacity to activate cytoplasmic signaling pathways. Mol Endocrinol 2007;21:1335–58. 187. Boonyaratanakornkit V, Scott MP, Ribon V, Sherman L, Anderson SM, Maller JL, et al. Progesterone receptor contains a proline‐rich motif that directly interacts with SH3 domains and activates c‐Src family tyrosine kinases. Mol Cell 2001;8:269–80. 188. Ballare C, Vallejo G, Vicent GP, Saragueta P, Beato M. Progesterone signaling in breast and endometrium. J Steroid Biochem Mol Biol 2006;102:2–10. 189. Lengyel F, Vertes Z, Kovacs KA, Kornyei JL, Sumegi B, Vertes M. Effect of estrogen and inhibition of phosphatidylinositol‐3 kinase on Akt and FOXO1 in rat uterus. Steroids 2007;72:422–8. 190. Vallejo G, Ballare C, Baranao JL, Beato M, Saragueta P. Progestin activation of nongenomic pathways via cross talk of progesterone receptor with estrogen receptor beta induces proliferation of endometrial stromal cells. Mol Endocrinol 2005;19:3023–37. 191. Hoekstra AV, Sefton EC, Berry E, Lu Z, Hardt J, Marsh E, et al. Progestins activate the AKT pathway in leiomyoma cells and promote survival. J Clin Endocrinol Metab 2009 [in review]. 192. Kovacs KA, Lengyel F, Vertes Z, Kornyei JL, Gocze PM, Sumegi B, et al. Phosphorylation of PTEN (phosphatase and tensin homologue deleted on chromosome ten) protein is enhanced in human fibromyomatous uteri. J Steroid Biochem Mol Biol 2007;103:196–9. 193. Bifulco G, Miele C, Pellicano M, Trencia A, Ferraioli M, Paturzo F, et al. Molecular mechanisms involved in GnRH analogue‐related apoptosis for uterine leiomyomas. Mol Hum Reprod 2004;10:43–8. 194. Hodges LC, Houston KD, Hunter DS, Fuchs‐Young R, Zhang Z, Wineker RC, et al. Transdominant suppression of estrogen receptor signaling by progesterone receptor ligands in uterine leiomyoma cells. Mol Cell Endocrinol 2002;196:11–20.

Nuclear Xenobiotic Receptors: Integrating Gene Regulation to Physiological Functions Jinhan He* and Wen Xie*,{ *Center for Pharmacogenetics and Department of Pharmaceutical Sciences, University of Pittsburgh, Pittsburgh, Pennsylvania 15216 {

Department of Pharmacology and Chemical Biology, University of Pittsburgh, Pittsburgh, Pennsylvania 15216

I. Introduction ................................................................................ II. Ligands for Nuclear Receptors......................................................... III. Nuclear Receptor Domain Structures ................................................ A. LBD and the AF‐2 Domain ........................................................ B. DNA‐Binding Domain .............................................................. C. AF‐1 Domain .......................................................................... IV. Xenobiotic Receptor Functions and Their Implications in Physiology and Diseases, A Case Study............................................................. V. Pregnane X Receptor (PXR) ............................................................ A. Cloning and Initial Characterization of PXR ................................... B. PXR in Phase I CYP Enzyme Regulation ....................................... C. PXR in Phase II Enzyme Regulation............................................. D. PXR in Drug Transporter Regulation ............................................ E. Implications of PXR‐Mediated Gene Regulation in Drug Metabolism... F. Endobiotic Functions of PXR...................................................... G. Species Specificity of PXR and the Creation of ‘‘Humanized’’ Mice ...... VI. Constitutive Androstane Receptor (CAR) ........................................... A. Identification of CAR as the Regulator of CYP2B Genes.................... B. CAR in the Regulation of Other DMEs ......................................... C. Mechanism of CAR Activation..................................................... D. Species Differences in the Activation of CAR.................................. VII. Concluding Remarks ..................................................................... References ..................................................................................

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Nuclear receptors (NRs) belong to a superfamily of evolutionarily related DNA‐binding transcription factors that can be activated by steroid and thyroid hormones, and other lipid metabolites. Ligand activated NRs can regulate target gene expression by binding to DNA response elements present in the target gene promoters. Through this regulation, NRs are broadly implicated in

Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87003-8

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physiology and metabolism. In this chapter, we will focus on the more recently identified and characterized xenobiotic receptors and their functions in xeno‐ and endobiotic responses.

I. Introduction Nuclear receptors (NRs) are members of a superfamily of evolutionarily related DNA‐binding transcription factors that regulate programs involved in a broad spectrum of physiological phenomena.1–3 Before the genes encoding these receptors were cloned, the first NR was identified biochemically in the 1960s.3,4 Specifically, Elwood Jensen and his colleagues showed that estradiol was specifically retained in target cells of this hormone, leading to the discovery that the cellular activity of estradiol was mediated by a specific high‐affinity receptor.5 Subsequently, and only 20 years ago, the human glucocorticoid receptor (GR) was one of the first NRs to be cloned by Ronald Evans at the Salk Institute. The estrogen receptor (ER) was cloned shortly after by the laboratories of Pierre Chambon and Geoffrey Greene.6–8 Since then, other NRs were rapidly cloned and their target DNA sequences were identified. Cloning of the steroid and thyroid hormone receptors demonstrated that these receptors share an extensive structural homology, which led to searches for new proteins with similar structure.9 So far, nearly 50 vertebrate NRs have been cloned. Among them, about 40 NRs were cloned before the study of their physiological function, and the ligands for many of them were yet to be identified; and for this reason, they were termed ‘‘orphan receptors.’’ Since their initial cloning, endogenous or synthetic ligands have been identified for many of the orphan receptors, converting these receptors to ‘‘adopted orphans.’’

II. Ligands for Nuclear Receptors NRs are understood primarily as ligand‐regulated transcription factors. The ligands for several initially cloned NRs, including GR, ER, and TR, are high‐affinity endogenous steroids or thyroids hormone, with Kd values in the nanomolar range. In 1988, the first orphan NRs, estrogen receptor‐related receptors (ERRs), were cloned.10 Although they are closely related to ER, ERRs do not bind to, and are not activated by estrogens. Since then, many more orphan NRs have been cloned.

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Since their initial cloning, numerous efforts were made to search for ligands of orphan NRs. Indeed, endogenous or synthetic ligands have been identified for many of the orphan NRs. Interestingly and in contrast to the ligands for GR, ER, and TR, most of the orphan NR ligands bind to the receptors with poor affinity (Kd values in the micromolar range) and have a broad specificity. Peroxisome proliferator‐activated receptors (PPARs), for example, can accommodate many natural and synthetic molecules that fill only a small portion (20%) of a potentially large ligand‐binding pocket. Similarly, the xenobiotic receptors PXR and constitutive androstane receptor (CAR) can be activated by a wide array of xenobiotics with relatively low affinities. Since NRs are broadly implicated in normal physiology and metabolism, activation and deactivation of NRs represent an intriguing means to treat a wide range of human diseases. Pharmaceutical NR agonists or antagonists, such as tamoxifen for ERs (targeted for breast cancer), thiazolidinediones for PPARg (targeted for type II diabetes), or dexamethasone (DEX) for GR (targeted for inflammatory diseases), are among the most commonly prescribed clinical drugs.11

III. Nuclear Receptor Domain Structures The ligand‐binding domain (LBD) and DNA‐binding domain (DBD) are two common structural motifs shared by most, if not all, NRs (Fig. 1A). DBD is connected to LBD by a short amino acid sequence termed hinge region. The complete functional properties of the hinge region are still unclear. It has been reported that the hinge region can be phosphorylated, and phosphorylation is coupled to increased transcriptional activity of the receptor.12–14 Most NRs contain amino acid sequences N‐terminal to the DBD, which contains a transcriptional activation function and was termed AF‐1.

A. LBD and the AF‐2 Domain LBD possesses a number of critical functions. First, as indicated by its name, LBD contains an interior binding pocket specific for its cognate hormone or ligand. The strength and specificity of LBD ligand complexes are based largely on hydrophobic interactions, extensive hydrogen bonding networks, and the steric size and shape of the binding pocket. In the case of steroid receptors, hydrophobic regions within the pocket closely contour the shape of a ligand, and polar groups serve to specifically bind and orient it.12 These structural features ensure the receptors to discriminate among closely related steroidal structures. Within the NR superfamily, the overall size and shape of the binding pockets apparently correlate with receptor function.15 Steroid receptors that maintain high affinity toward only a small number of ligands

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A N-

AF-1

DBD

LBD

AF-2

-C

Ligand- and coactivator-binding

DNA-binding B

NR

RXR

5⬘

3⬘ Target gene AGGTCA

AGGTCA n

DRn

n

IRn

n

ERn

FIG. 1. Structural and functional domains of nuclear receptors. (A) Basic structure of nuclear receptors. AF‐1 and DBD are at the amino (N)‐terminal, and AF‐2 and LBD are at the carboxy (C)‐terminal. (B) Nuclear receptors can recognize and bind to AGGTCA half‐sites spaced by several base pairs as direct repeats (DRn), inverted repeats (IRn), or everted repeats (ERn).

have smaller volumes within their binding pockets; but have extensive polar side chains that can precisely hydrogen bond with the ligand. In contrast, orphan NRs tend to have larger volume binding pockets that can accommodate ligands of different chemical structures.12 LBD also contains a ligand‐regulated transcriptional activation function (AF‐2) necessary for the recruitment of various coregulators. To achieve transcriptional activation of target genes, most NRs require intermediary factors. These proteins, so-called coregulators, often constitute subunits of larger multiprotein complexes that act at several functional levels, such as chromatin remodeling, enzymatic modification of histone tails, or modulation of the preinitiation complex via interactions with RNA polymerase II.16 The structural interface for recruiting coregulators localizes to a hydrophobic groove formed by several helices of the LBD including helix 12 (also called the AF‐2 helix). Coregulators that contain helical LXXLL motifs bind via hydrophobic interactions in the groove. Interaction specificity is determined by charge–clamp electrostatic interactions between LBD and coregulator residues that cap each end of the two-turn LXXLL helix. In the presence of an activating ligand, helix 12 is stabilized against the surface of the LBD, allowing the formation of hydrophobic binding groove and coregulators recruitment.12,17

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B. DNA‐Binding Domain The highly conserved DBD links the receptor to specific promoter regions of its target genes, termed hormone response elements (HREs). As the most conserved NR domain, DBD can recognize DNA response elements that contain one or two consensus core half‐sites related to the hexamer ACAACA (for steroid receptors) or AGGTCA (for estrogen receptors and other NRs).18,19 A HRE contains two NR half‐sites of the consensus hexameric sequences, arranged as inverted, everted or direct repeats with a 3‐ to 6‐bp spacing (Fig. 1B). The DBD of GR was one of the first structures determined for any receptor subunit and still serves as a representative structural model for the NR superfamily.20,21 The GR DBD has two a‐helices; the N‐terminal helix directly interacts with the major groove of each DNA half‐site, making base‐specific contacts, whereas the C‐terminal helix in a perpendicular fashion and contributes to the stabilization of the overall protein structure.12 GR is monomeric in solution but can undergo DNA‐induced dimerization upon binding to a palindromic (inverted repeat) response element. The residues that make up the dimer interface are located within the C‐terminal zinc finger and are defined as the ‘‘D‐box’’; the residues critical to sequence‐specific DNA binding are located within helix 1 and are defined as the ‘‘P‐box.’’ A comparison of the free GR structure with the DNA‐bound GR structure indicates that interactions between the P‐box and the DNA half‐site are coupled to conformational changes in the D‐box necessary for cooperative recruitment of the second monomer.22 Thus, the specific binding sequence acts as an allosteric effector of function.22 Many NRs bind to DNA as heterodimers with RXR, which is a cognate receptor for 9‐cis retinoic acid. The heterodimerization between NRs and RXR works in a similar fashion.

C. AF‐1 Domain AF‐1 domain is the least conserved region of NRs. AF‐1 domains from different receptors vary in both sizes and sequences.16 Consequently, the activation capacity of AF‐1 domains has been shown to vary considerably between NRs. In addition, many receptor isoforms that differ exclusively in their N‐terminal amino acid sequences are generated through alternative splicing and/or alternative promoter usage. Limited proteolysis experiments on the progesterone receptor (PR) N‐terminus suggested a lack of well‐folded structure for isolated AF‐1.23,24 It was suggested, through the analysis of AF‐1 domains of PR and GR, that the AF‐1 domains may be stabilized in the context of the intact receptor.12,23

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IV. Xenobiotic Receptor Functions and Their Implications in Physiology and Diseases, A Case Study Numerous papers have been published on the structures and functions of NRs in divergent systems including endocrine, cardiovascular, nervous, immune, metabolic, and other systems. It is nearly impossible to make a comprehensive review on all that has been published on NRs. In this chapter, we intend to focus on the more recently discovered and characterized xenobiotic receptors PXR and CAR, and to summarize their physiological functions and their implications in physiology and diseases. An important requirement for physiological homeostasis is the metabolism/ detoxification and removal of endogenous chemicals and xenobiotic compounds with biological activities. Much of the detoxification and elimination are carried out by drug metabolizing enzymes (DMEs) that include Phase I cytochrome P450 (CYP) enzymes and Phase II conjugation enzymes, as well as the ‘‘Phase III’’ drug transporters. The products of Phase I metabolism are generally more polar and readily excreted than their parent compounds. The Phase I enzyme‐mediated functionalization also often enables xenobiotics to become better substrates for the Phase II conjugating enzymes. The Phase II metabolism involves conjugation of hydrophilic moieties, such as sugar, glutathione, and sulfate, to increase polarity and water solubility of xenobiotics and therefore promoting their excretion and elimination. In addition to Phase I and Phase II enzymes, equally important is a group of transporter proteins that modulate the absorption, distribution, and excretion of many drugs. It has become evident that NRs play an important role in the transcriptional regulation of DMEs.25–27 The induction profiles of the major DMEs are remarkably linked to the activation of several orphan NRs. For instance, compounds such as rifampicin (RIF) and phenobarbital (PB) are PXR activators and inducers of CYP3A, CYP2B, UGT1A1, CYP2C9, and multidrug resistance protein 1(MDR1/ABCB1); whereas 1,4‐bis[2‐(3,5‐dichloropyridyloxy)]benzene (TCPOBOP) and 6‐(4‐chlorophenyl:imidazo[2,1‐b]thiazole‐5carbaldehyde O‐(3,4‐dichlorobenzyl)oxime (CITCO) induce CYP2B, CYP3A, and UGT1A1 through the activation of CAR.28–31 This association of orphan NRs with particular DMEs not only demonstrates DME specificity but also defines species‐specific differences in the DME induction responses. Here, we focus on recent advances in our knowledge and understanding of the regulation of DMEs and transporters by NRs. We will focus on the xenobiotic receptors PXR and CAR, although it has become increasingly evident that several NRs other than PXR and CAR are also important in regulating the expression of drug disposition genes either directly, or by modulating the activities of PXR and CAR.

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V. Pregnane X Receptor (PXR) A. Cloning and Initial Characterization of PXR In 1998, several groups published the cloning and initial characterization of PXR. Kliewer and colleagues identified mouse PXR on the basis of its sequence homology with other NRs. It was named PXR based on its activation by various natural and synthetic pregnanes.28 The cloning of human PXR (hPXR) was first reported by Blumberg and colleagues, and this group, led by Ronald Evans at the Salk Institute, initially named this receptor ‘‘steroid and xenobiotic receptor (SXR)’’ based on its activation by steroids and xenobiotics.32 hPXR was also subsequently cloned by Bertilsson and colleagues and was alternately referred to as the pregnane‐activated receptor (PAR).33 PXR shares a common NR modular structure with a conserved N‐terminal DBD and a C‐terminal LBD. The DBD mediates interaction with specific DNA sequences known as xenobiotic response elements (XREs). The LBD determines ligand‐binding specificity and heterodimerizes with RXR. Although DBDs of the mammalian PXRs are highly conserved, sharing more than 95% amino acid identity, the LBDs of PXRs are much more divergent across species than those of other NRs. For example, the human and rat PXR share only 76% amino acid identity in their LBD, whereas most human and rodent NR orthologs share more than 90% amino acid identity.34 The LBD sequence divergence of PXR between species is believed to be responsible for the species‐specific response to ligands/drugs. PXR is highly expressed in the liver and small intestine, where most DMEs are also highly expressed and induced. In rodents, lower levels of PXR mRNA have also been detected in kidney, stomach, lung, uterus, ovary, and placenta. PXR can be activated by xenobiotics and endobiotics, which underlie the significance of PXR in both xenobiotic and endobiotic responses. PXR orthologs also exhibit interesting and striking species‐dependent ligand specificity. For example, the antiglucocorticoid pregnenolone‐16a‐carbonitrile (PCN) is an effective activator of mouse and rat PXRs, but has little effect on hPXR. In contrast, the antibiotic RIF can activate the human and rabbit PXRs, but not the mouse and rat PXRs. The XREs bound by PXR–RXR heterodimers are also called PXR response elements (PXREs). The classical PXRE is DR‐3 which is composed of two copies direct repeat of the consensus AG (G/T) TCA NR binding motif and separated by three nucleotides. Other PXREs, including DR‐4, DR‐5, ER‐6 (everted repeat spaced by 6 bp), ER‐8, and IR‐0 (inverted repeat without a spacing nucleotide), have also been reported.35–37

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B. PXR in Phase I CYP Enzyme Regulation The oxidative CYP enzymes catalyze the metabolic conversion of xenobiotics to polar derivatives that are more readily eliminated. Among the 78 mouse and 65 human CYP enzymes itemized within the current build of the Unigene sequence databank, a significant number of these enzymes are known to be induced by various environmental pollutants, pharmaceuticals, or steroid metabolites. Among CYP enzymes, the CYP3A isozymes are of particular medical significance since they are involved in the metabolism of more than 50% of clinical drugs as well as neutraceuticals and herbal medicines.38 In 1998, PXR was isolated as a xenobiotic receptor regulating CYP3A gene expression.28,32,33,39 PXR regulates the expression of both human and rodent CYP3A genes by directly binding to ER‐6 and DR‐3 localized in the promoter regions of the human and rodent CYP3A genes, respectively. Subsequent studies using both the loss‐of‐function gene knockout and gain‐of‐function transgenic mice have provided convincing genetic and pharmacological evidence to support the role of PXR in CYP3A gene regulation. Targeted disruption of the mouse PXR locus abolished the CYP3A xenobiotic response to prototypic inducers such as PCN and DEX.40,41 In contrast, hepatic expression of an activated form of hPXR in transgenic mice resulted in sustained induction of CYP3A enzymes and enhanced protection against xenobiotic toxicants, such as zoxazolamine and tribromoethanol.40 In addition to CYP3A isozymes, PXR has also been shown to regulate CYP2B6,42–45 CYP2B9,46 CYP2C8,47 and CYP2C9.46

C. PXR in Phase II Enzyme Regulation The Phase II conjugating enzymes include broad specificity transferases, such as UDP‐glucuronosyltransferase (UGT), glutathione S‐transferase (GST), and sulfotransferases (SULT).48–51 Phase II conjugation has dual role in drug metabolism. First, conjugation reactions not only terminate the ability of electrophiles to react with DNA and proteins, but they also prevent nucleophiles from interacting with receptor proteins. At the same time, conjugation increases the water solubility of the compounds, which in turn, promotes renal and biliary excretion.52 Therefore, the Phase II reactions play a critical role in xeno‐ and endobiotic detoxification. 1. PXR IN UGT REGULATION Glucuronidation, a major metabolic pathway for endo‐ and xenobiotics, is catalyzed by enzymes belonging to the family of membrane bound UGTs. Using UDP‐glucuronic acid as a sugar donor, UGTs catalyze the transfer of glucuronic acid to a variety of substrates and thus convert small lipophilic molecules to water‐soluble glucuronides. UGT‐mediated glucuronidation functions as the principle means to eliminate steroid, heme metabolites,

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environmental toxin, and drugs.48,53–55 Among UGTs, UGT1A1 is one of the most characterized UGT isoform. UGT1A1 is the critical enzyme responsible for the detoxification of bilirubin. Patients deficient in the expression and/or activity of UGT1A1 can lead to severe accumulation of unconjugated bilirubin, medically termed hyperbilirubinemia, a hallmark of the Crigler–Najjar (CN) syndrome.48 Our study showed that UGT1A1 is under the positive control of PXR. UGT1A1 mRNA and protein expression was upregulated in transgenic mice that express the activated hPXR (VP‐hPXR or VP fusion of hPXR) and in RIF‐treated ‘‘humanized’’ hPXR transgenic mice.56 The microsomal glucuronidation activity toward b‐estradiol (a UGT1A1 substrate), thyroid hormones, corticosterone, and xenobiotics (such as 4‐nitrophenol and 4‐OH‐PhIP) was also increased in the VP‐hPXR transgenic mice. Chen and colleagues showed that PCN, a rodent‐specific PXR agonist, induced UGT expression and increased UGT enzymatic activity in wild‐type mice and this induction was abolished in PXR‐null mice.57 The identification of a DR3‐like PXR responsive element in the human UGT1A1 promoter further established UGT1A1 as a direct target gene of PXR. In addition to UGT1A1, the expression of UGT1A9, but not UGT1A2 and 2B5, was also increased by PXR activation, suggesting that the UGT induction by PXR is isoform‐specific.57 In addition to PXR, the expression of UGT1A1 and several other UGT isoforms have also been reported to be regulated by several other NRs, including CAR56,58,59 and PPARa.60,61 2. PXR IN SULT REGULATION The cytosolic SULTs are another important family of Phase II enzymes that catalyze the conjugation of nucleophilic compounds. SULTs catalyze the transfer of a sulfonyl group from a sulfate donor 30 ‐phosphoadenosine 50 ‐phosphosulfate (PAPS) to hydroxyl or amino groups of acceptor molecules, forming sulfate or sulfamate conjugates. SULTs are specific to sulfonate small lipophilic molecules, such as steroids, bioamines, and therapeutic drugs. It was believed that, although SULTs and UGTs have similar substrate spectrum, SULTs have higher affinities but lower turnover rates than UGTs.62 Thus, UGT activities are likely to play a more significant role when substrates are abundant, whereas SULTs may play a leading role when concentrations of the substrate are lower. SULT‐mediated sulfonation is known to play a significant role in the homeostasis of sex hormones that are present in low concentrations in plasma.63 The expression of rodent hepatic Sult2a9, also called Sult2a1 or dehydroepiandrosterone sulfotransferase (DHEA SULT or STD), is subjected to transcriptional regulation by PXR and an IR‐0 element located in the 50 ‐flanking region of rodent Sult2a gene is required for its activation by PXR.36 The expression of Sult2a9/2a1/STD has also been shown to be regulated by

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FXR,64 CAR,65 and LXR.66 Interestingly, all four receptors share the same IR‐0 response element to regulate the expression of this SULT isoform. The hierarchy and relative contribution of individual NRs in Sult2a regulation remain to be determined. More recent studies showed that the human SULT2A1 is also regulated by PXR.67 3. PXR IN GST REGULATION GSTs are soluble enzymes that use reduced glutathione in conjugation and reduction reactions. GSTs play an important role in the conjugation of electrified chemicals. Since chemical carcinogens are often highly nucleophilic, they also represent potential substrates for GST. For this reason, GSTs are believed to play an important role to protect cells from genotoxic compounds. The cytosolic GST isozymes of rodents and humans can be grouped into several classes, such as alpha, mu, pi, theta‐omega, and zeta, based on their amino acid sequences, immunological properties, and substrate specificities.68,69 The Alpha, Mu, and Pi classes are most abundantly expressed GSTs. GST regulation by PXR was first hinted by several studies of general profiling of gene expression.70,71 It was also reported that GSTA2 expression was induced by PXR and an IR‐6 response element was responsible for this transcriptional regulation.69 A more systemic and comparative analysis of GST regulation by PXR was reported in transgenic mice that bear the expression of activated PXR in the liver and intestine.72 In this study, it was shown that the expression of GST Alpha, Pi, and Mu classes are all under the control of PXR. Interestingly, PXR‐mediated GST regulation exhibited clear isoform‐, tissue‐, and gender‐specificity in that: (1) GST Mu was the only isoform that was upregulated by PXR in both liver and intestine in both sexes; (2) GST Alpha was induced in the small intestine, but not in the liver; and (3) PXR had an opposite effect on hepatic GST Pi expression, inducing this class in females, but suppressing it in males. Paradoxically, although the overall GST expression and activity were increased, activation of PXR sensitized mice to the oxidative xenotoxicant paraquat in vivo and in cultured cancer cells. Moreover, heightened paraquat sensitivity in transgenic mice was female‐specific. Whether the PXR‐mediated, gender‐specific GST regulation accounts for the intact paraquat sensitivity in transgenic males remains to be determined. Nevertheless, the regulation of GSTs by PXR suggests that this regulatory pathway may be relevant to carcinogenesis by sensitizing normal and cancerous tissues to oxidative cellular damage.72

D. PXR in Drug Transporter Regulation In addition to regulating DMEs, PXR also participates in the regulation of drug transporters. Drug transporters are responsible for uptake and efflux of endogenous and exogenous chemicals, including many clinically prescribed drugs.73

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Organic anion‐transporting polypeptides (Oatps) are a family of major uptake drug transporters in the liver. Oatps are localized to the basolateral membrane of hepatocytes and transport compounds, including organic anions, organic cations, and neutral compounds, from blood into hepatocytes. Oatps family includes nine human, 13 rat, 15 mouse members.74 PXR ligand PCN administration induced Oatp2 mRNA expression in the livers of wild‐type mice, but not PXR‐ null mice.75 A DR‐3 type PXRE has been identified in the rat Oatp2 promoter.76 Multidrug resistance associated proteins (MRPs) are efflux transporters for structurally diverse amphipathic chemicals and organic anions. The liver‐ enriched Mrp2/Abcc2 is localized to the canalicular membrane of hepatocytes and is responsible for the hepatobiliary excretion of amphipathic anions.77,78 Natural mutations in MRP2/ABCC2 cause Dubin–Johnson syndrome/hyperbilirubinemia II, a disorder characterized by impaired transfer of anionic conjugates into the bile. Induction of Mrp2/Abcc2 by PXR ligand PCN and DEX was observed in primary hepatocytes isolated from livers of wild‐type mice, but not PXR‐null mice, implicating a role of PXR in the regulation of mouse Oatp2.78 An ER‐8 type of NR response element was identified in the promoter of the rat Mrp2/Abcc2 gene that confers the induction of Mrp2/ Abcc2 by PXR.78 Mrp3/Abcc3 was also reported to be induced by PXR.79 In the intestine, PXR has been shown to stimulate the expression of Mdr1/Abcb1, which encodes an ATP‐dependent efflux pump that transports a wide variety of xenobiotics, including many widely used prescription drugs. In humans, activation of PXR in the intestine may decrease intestinal drug absorption by increasing the expression of MDR1/ABCB1. A DR‐4 type PXRE was identified in MDR1/ABCB1 gene promoter.30

E. Implications of PXR‐Mediated Gene Regulation in Drug Metabolism The implication of PXR‐mediated gene regulation in drug metabolism and drug interaction has been recognized since the cloning of this receptor. It is obvious that the regulation of DMEs by PXR is involved in clinical drug–drug interactions, in which one drug accelerates the metabolism of a second medicine and may change or cause adverse results. Because CYP enzymes can recognize a large spectrum of pharmaceutical substrates, a CYP gene‐inducing drug is potentially capable of affecting the metabolism and clearance of any coconsumed drugs. Indeed, the identification of RIF as a potent hPXR agonist has provided an explanation why this antibiotic drug is prone to drug–drug interactions. In another example, St. John’s Wort (SJW), a popular herbal remedy for mild depression, has been reported to trigger severe adverse interactions with several clinical drugs, such as oral contraceptives, the HIV protease inhibitor indinavir, and the immunosuppressant cyclosporine.

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Such drug–drug interactions are likely results of activation of PXR and consequent induction of CYP3A by SJW and the subsequent increased metabolism and/or decreased bioavailability of cometabolized drugs.80 Several traditional Chinese medicines (TCMs) have been implicated in drug–drug interactions. One clinical concern of herbal product use is the effect of herbal products on the metabolism of coadministered drugs. We showed that two TCM herbs, Wu Wei Zi (Schisandra chinensis Baill) and Gan Cao (Glycyrrhiza uralensis Fisch), can activate PXR and induce the expression of several DMEs and transporters, including CYP3A, CYP2C9, and MRP2/ABCC2 in reporter gene assay and in primary hepatocyte cultures.81 The anticoagulant warfarin is known to be metabolized by CYP2C9 in humans.82 As expected, administration of Wu Wei Zi and Gan Cao extracts in rats resulted in an increased metabolism of coadministered warfarin, reinforcing concerns involving the safe use of herbal medicines and other nutraceuticals to avoid drug–drug interactions.81 Having known the potential of PXR‐activating agents in causing drug–drug interactions, it is important to emphasize that PXR activation alone may not be sufficient to predict the propensity of drug–drug interactions. Sinz and colleagues published the evaluation of 170 xenobiotics in an hPXR transactivation assay and compared these results to known clinical drug–drug interactions. Of the 170 xenobiotics tested, 54% of them demonstrated some level of hPXR transactivation. However, by taking into consideration cell culture conditions (solubility, cytotoxicity, appropriate drug concentration in media), as well as in vivo pharmacokinetics (therapeutic plasma concentration or Cmax, distribution, route of administration, dosing regimen, liver exposure, potential to inhibit CYP3A4), the risk potential of CYP3A4 enzyme induction for most compounds reduced dramatically. By employing this overall interpretation strategy, the final percentage of compounds predicted to significantly induce CYP3A4 reduced to 5%, all of which are known to cause drug–drug interactions in the clinic.83

F. Endobiotic Functions of PXR Even though PXR was initially identified as a ‘‘xenobiotic receptor,’’ emerging evidence has pointed to an equally important role of PXR as an ‘‘endobiotic receptor’’ that responds to a wide array of endogenous chemicals (endobiotics), such as bile acids and their intermediates, as well as certain steroid hormones. Moreover, many endobiotics are substrates of PXR target enzymes and transporters. Activation of PXR by endogenous or xenobiotic ligands has implications in several important physiological and pathological conditions. For this reason, there have been extensive discussions on whether or not PXR can be explored as a therapeutic target.37,84

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1. PXR IN BILE ACID DETOXIFICATION AND CHOLESTASIS One family of endogenous PXR ligands identified shortly after the cloning of PXR is bile acids. Bile acids are catabolic end products of cholesterol metabolism. They are physiologically important in the formation of bile and solublizing biliary lipids and promoting their absorption. However, excessive bile acids are potentially toxic. For example, the secondary bile acid lithocholic acid (LCA) has been shown to cause cholestasis in experimental animals and has long been suspected of doing the same in humans. Therefore, excess bile acid should be efficiently eliminated to avoid the toxic effect. PXR has been demonstrated to act as a LCA sensor and plays an essential role in detoxification of cholestatic bile acids.41,85,86 Studies in different animal models showed that activation of PXR protected mice from severe liver damage induced by LCA. Pretreatment of wild‐type mice, but not the PXR‐null mice, with PCN reduced the toxic effects of LCA. Moreover, genetic activation of PXR by expressing the activated PXR in the liver of transgenic mice was sufficient to confer resistance to the hepatotoxicity of LCA. The cholestatic preventive effect of PXR was initially reasoned to be due to the activation of CYP3A, an important CYP enzyme responsible for bile acid hydroxylation.41,86 Subsequent identification of SULT2A, a bile acid detoxifying hydroxysteroid SULT, as a PXR target gene suggested that additional PXR target genes may have also contributed to the phenotype.36,87 Several follow‐up studies, including those using mice with individual or combined loss of PXR and CAR, have suggested that PXR‐responsive bile acid transporter regulation may also play a role in preventing cholestasis.88–90 Since activation of PXR was sufficient to prevent cholestasis, it has been suggested that PXR agonists may be useful in the treatment of human cholestatic liver disease, a notion that has been supported by several clinical observations. Both RIF and SJW have been empirically used to treat cholestatic liver diseases.35 The relief from cholestasis‐associated pruritis and amelioration of cholestasis by RIF was associated with increased bile acid hydroxylation, which in turn facilitates glucuronidation by UGTs. Both RIF and SJW are potent agonists of hPXR and both CYP3A and UGT may have been PXR target genes, suggesting the anticholestatic effects of RIF and SJW may have been mediated by PXR. 2. PXR IN BILIRUBIN DETOXIFICATION AND CLEARANCE Bilirubin is the catabolic byproduct of heme proteins. Accumulation of bilirubin in the blood is potentially toxic. An insufficiency in expression of UGT1A1, a key enzyme for the conjugation of bilirubin, in the Crigler–Najjar syndrome and Gilbert’s diseases results in severe hyperbilirubinemia. Deficiency of MRP2/ABCC2, a drug transporter responsible for the hepatic excretion of conjugated bilirubin, leads to Dubin–Johnson syndrome, characterized by the

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accumulation of glucuronidated bilirubin. PXR has been shown to induce the expression of multiple key components in the clearance pathway, including UGT1A1, OATP2, GSTA1, and 2 and MRP2/ABCC2. OATP2 facilitates bilirubin uptake from blood into hepatocytes.73 GSTA1 and 2 reduce bilirubin back efflux from hepatocytes into blood. MRP2/ABCC2 promotes the canalicular efflux of conjugated bilirubin. Consistent with the pattern of gene regulation, activation of PXR in transgenic mice has been shown to prevent experimental hyperbilirubinemia.56 3. PXR IN ADRENAL STEROID HOMEOSTASIS AND DRUG–HORMONE INTERACTIONS PXR plays an important endobiotic role in adrenal steroid homeostasis. Our study showed that activation of hPXR in mice markedly increased plasma concentrations of corticosterone and aldosterone, the respective primary glucocorticoid and mineralocorticoid in rodents. The increased levels of corticosterone and aldosterone were associated with activation of adrenal steroidogenic enzymes, including CYP11a1, CYP11b1, CYP11b2, and 3b‐Hsd. The PXR‐activating transgenic mice also exhibited hypertrophy of the adrenal cortex, loss of glucocorticoid circadian rhythm, and lack of glucocorticoid responses to psychogenic stress.91 Interestingly, the VP‐hPXR transgenic mice had normal pituitary secretion of adrenocorticortropic hormone (ACTH) and the corticosterone suppressing effect of DEX was intact, suggesting a functional hypothalamus–pituitary– adrenal (HPA) axis despite a severe disruption of adrenal steroid homeostasis. The ACTH‐independent hypercortisolism in the PXR‐activating transgenic mice is reminiscent of the pseudo‐Cushing’s syndrome in patients, the clinical hallmark of which is the normal DEX suppression despite a high circulating level of glucocorticoid. Pseudo‐Cushing’s syndrome is most seen in alcoholic, depressed, or obese subjects. It is of interest to know whether or not these susceptible patients are associated with increased expression and/or activity of PXR. The glucocorticoid effect appeared to be PXR‐specific, as the activation of CAR in transgenic mice had little effect. We propose that PXR is a potential endocrine disrupting factor that may have broad implications in steroid homeostasis and drug–hormone interactions.91 4. PXR IN LIPID METABOLISM PXR has also recently been shown to play an endobiotic role by impacting lipid homeostasis.92 Expression of an activated PXR in the livers of transgenic mice resulted in an increased hepatic deposit of triglycerides. This PXR‐ mediated lipid accumulation was independent of the activation of the lipogenic transcriptional factor sterol regulatory element‐binding protein 1c (SREBP‐1c) and its primary lipogenic target enzymes, including fatty acid synthase (FAS)

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and acetyl CoA carboxylase 1 (ACC‐1). Instead, the lipid accumulation in transgenic mice was associated with an increased expression of the free fatty acid transporter CD36 and several accessory lipogenic enzymes, such as stearoyl CoA desaturase‐1 (SCD‐1) and long chain free fatty acid elongase (FAE). Studies using transgenic and knockout mice showed that PXR is both necessary and sufficient for CD36 activation. Promoter analyses revealed a DR‐3 type of PXRE in the mouse CD36 gene promoter, establishing CD36 as a direct transcriptional target of PXR. The hepatic lipid accumulation and CD36 induction was also seen in the hPXR ‘‘humanized’’ mice treated with the hPXR agonist RIF. The activation of PXR was also associated with an inhibition of pro‐b‐oxidative genes, such as PPARa and thiolase, and an upregulation of PPARg, a positive regulator of CD36. The cross‐regulation of CD36 by PXR and PPARg suggests that this fatty acid transporter may function as a common target of orphan NRs in their regulation of lipid homeostasis.

G. Species Specificity of PXR and the Creation of ‘‘Humanized’’ Mice 1. CHALLENGES FOR RODENTS AS DRUG METABOLISM MODELS Primary hepatocytes represent a common and important in vitro system to evaluate metabolism, toxicity, and enzyme induction.93–95 However, a significant disadvantage of this system is the lack of routine availability of high quality human liver tissues or cells. Other model systems that may provide more consistent access include immortalized hepatocytes or humanized animal models.40,96 Over the years, it has been perceived that rodent models have limited utility in predicting drug‐related human effects due to significant species differences in DMEs, transporters, and nuclear hormone receptors. For example, PCN is an effective CYP3A inducer in rodents but not in humans, and RIF induces CYP3A in humans but not in rats.97 These findings have been attributed to the species differences in the effect of several drugs on CYP3A expression mediated by PXR.97,98 These differences across species demand the development of humanized animal models to evaluate the potential effect of a chemical in humans using an animal model. 2. SPECIES SPECIFICITY OF THE RODENT AND hPXR Both hPXR and mPXR are highly expressed in the liver and small intestine and share many functional properties, in particular, the regulation of CYP3A genes. However, as discussed earlier, these two orthologs are pharmacologically distinct in that strong activators of one receptor are often poor activators of the other. This species‐specific ligand profile is reflected by the sequence divergence in the LBDs of the mouse and human receptors. The crystal structure of the hPXR LBD has been solved.99 The hydrophobic ligand‐binding cavity of

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hPXR is composed of a large, smooth surface containing only a small number of polar residues, suggesting that it is not necessary for activators to conform to a restricted orientation. Based on site‐directed mutational analysis, the position and nature of these polar residues were found to be crucial for establishing the precise pharmacological activation profile of hPXR.99 Indeed, conversion of four amino acids of mouse PXR that correspond to the hPXR‐specific activator SR12813‐interacting residues in hPXR produces a hybrid mouse–hPXR that was no longer activated by PCN and was only weakly activated by SR12813 in reporter gene assays.99 The structural and pharmacological differences between hPXR and mPXR and that of other species might reflect the difference in the diets of rodents and primates and the evolutionary need to respond to a different set of ingested nutrients and xenobiotics. 3. CREATION AND CHARACTERIZATION OF THE hPXR ‘‘HUMANIZED’’ MICE Based on the hypothesis that the species origin of the receptor is the determining factor for the ligand specificity between species, we have created transgenic mice to determine whether the human receptor is sufficient to establish a human response profile40 (Fig. 2). First, hPXR transgenic mice were generated by expressing the hPXR in the mouse liver. The liver‐specific expression of the transgene was accomplished by using the mouse albumin promoter. Because the resulting mice harbor both mPXR and hPXR in their livers, the transgenic mice exhibited a chimeric or combined CYP3A response to both the rodent‐specific inducer PCN and the human‐specific inducer RIF. These results imply that mice only expressing hPXR could be fully humanized for the xenobiotic response. These animals were created by breeding the hPXR transgene into the mPXR knockout background. In contrast to the null mice that are devoid of CYP3A induction by steroids, replacement of mPXR with transgenic hPXR restores xenobiotic regulation with a humanized response profile. These mice readily responded to human inducers, such as RIF, in the equivalent range of the standard oral dosing regimen in humans and exhibited similar pharmacokinetics of CYP3A regulation.40,100 A ‘‘fully’’ human profile of CYP3A inducibility is obtained in the mPXR null/hPXR transgenic mice. Therefore, these experiments provide compelling evidence that PXR functions as a species‐specific xenosensor mediating the adaptive hepatic response. This is also one of the rare examples in which replacing a single transcriptional regulator enables conversion of species‐specific gene regulation. The original ‘‘humanized’’ mice bear the expression hPXR exclusively in the liver.40 Since both the DMEs and xenobiotic receptors are also highly expressed in the intestines, it is conceivable that mouse models with the humanized receptors expressed in both the liver and intestine would represent a more complete humanized mouse model. The liver and intestine dual

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Cyp3a11 induction RIF PCN

Wild type

rodent inducer

human inducer

+

–

+

+

–

+

hPXR transgene

hPXR transgenic

mPXR knockout

hPXR humanized

FIG. 2. Schematic representation of the hPXR ‘‘humanized’’ mice. The humanization was achieved in the liver only when the liver‐specific albumin promoter was used to direct the transgene expression; or in both the liver and intestine when the fatty acid binding protein promoter was used. PCN, pregnenolone‐16a‐carbonitrile; RIF, rifampicin. ‘‘þ’’ and ‘‘’’ mean induction and lack of induction, respectively.

humanization has been achieved by using the fatty acid binding protein (FABP) gene promoter that targets the expression of hPXR transgene to both the liver and intestine.92 Frank Gonzalez’s laboratory at the National Cancer Institute reported the creation of humanized PXR mouse model by bacterial artificial chromosome (BAC) transgenesis in PXR‐null mice.101 To create the transgenic mice, a BAC clone containing the complete hPXR sequence, as well as 50 ‐ and 30 ‐flanking sequences was linearized and microinjected into fertilized FVB/N mouse eggs. The resulting transgenic mice were bred with PXR‐null mice.41 Quantitative PCR revealed that hPXR was expressed in liver, duodenum, jejunum, and ileum of the humanized mice. Similar to the previously described humanized mice,40 these mice also mimicked the human response to PXR ligands treatment.101 In the same study, it was shown that in RIF‐pretreated

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PXR‐humanized mice, an approximately 60% decrease was observed for both the maximal midazolam serum concentration and the area under the concentration–time curve, as a result of a threefold increase in midazolam 10 ‐hydroxylation. These results illustrate the potential utility of the PXR‐humanized mice in the investigation of drug–drug interactions mediated by CYP3A and suggest that the PXR‐humanized mouse model would be an appropriate in vivo tool for evaluation of the overall pharmacokinetic consequences of hPXR activation by drugs.101 4. SIGNIFICANCE OF HUMANIZED MICE IN DRUG METABOLISM STUDIES AND DRUG DEVELOPMENT The creation of mouse models with humanized xenobiotic response may aid pharmaceutical development by predicting potential drug–drug interactions.35,37,87,102 For decades, rodent models have been standard components in the assessment of potential toxicity for the development of candidate human drugs. However, their reliability as predictors of the human xenobiotic response is limited due to the species‐specificity of the xenobiotic response. To date, there has been no reliable system outside of humans to directly and quantitatively assess the drug–drug interactions. Primary cultures of human hepatocytes are valuable. However, since the hepatocytes are from individual patients, the utility of human hepatocytes is compromised by interindividual variability, limited tissue resources, and high cost. The humanized mice exhibited a ‘‘humanized’’ hepatic xenobiotic response profile, readily responding to the human‐specific inducer RIF in a concentration range equivalent to the standard oral dosing regimen in humans.40 The creation of these mice represents a major step toward generating a humanized rodent toxicological model that is continuously renewable and completely standardized.

VI. Constitutive Androstane Receptor (CAR) A. Identification of CAR as the Regulator of CYP2B Genes The human CAR (NR1I3) was originally isolated by screening a human liver library with degenerate oligonucleotides probes.103 The name CAR was initially defined as ‘‘constitutively activated receptor,’’ because it forms a heterodimer with RXR that binds to retinoic acid response elements (RAREs) and transactivates target genes in the absence of ligands in transfection assays.103,104 Using human CAR cDNA as probe, the mouse counterpart was isolated subsequently.104 In 1998, two androstane metabolites androstanol (5a‐androstan‐3a‐ol) and androstenol (5a‐androstan‐16‐en‐3a‐ol) were found

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to be endogenous antagonists of mouse CAR, giving an alternative name of CAR as ‘‘constitutive androstane receptor.’’ A breakthrough in linking CAR with CYP2B gene expression was pioneered by Masahiko Negishi at the National Institute of Environmental Health Sciences and David Moore at the Baylor College of Medicine.29,43,85,103–105 Xenobiotic induction of CYP2B genes in different species has been realized for a long time. Through a series of elegant studies, a specific sequence termed PB‐responsive element (PBRE) or PB‐responsive unit (PBRU), or later delineated as PB‐responsive enhancer module (PBREM), was identified as the key sequences that are responsible for xenobiotic induction of CYP2B genes.42,106–108 A substantial progress in our understanding of PBREM activation by PB was obtained upon identification of CAR as the predominant regulator of PBREM activation using cell‐based transfection assays.42 Electrophoretic mobility shift assays (EMSAs) confirmed that CAR–RXR heterodimers bound to the NR1 and NR2 motifs of the PBREM.42 The role of CAR in the regulation of PB induction of the CYP2B genes was established definitively through the creation and characterization of CAR‐null mice, in which the activation of CYP2b10 mRNA expression by PB and TCPOBOP was completely abolished.109,110

B. CAR in the Regulation of Other DMEs 1. CAR REGULATION OF CYP2C GENES CYP2C enzymes are responsible for the metabolism of approximately 20% of therapeutic drugs and many endogenous compounds in humans.82 Induction of human CYP2Cs can result in drug tolerance as well as drug–drug interactions. A series of studies by Goldstein and coworkers demonstrated that xenobiotic induction of CYP2Cs is collaboratively mediated by several NRs, including CAR, PXR, GR, and HNF4a.29,57,111 In primary human hepatocytes, CYP2C9 is inducible by xenobiotics including PB, RIF, and DEX.112–114 A role for CAR in CYP2C9 regulation was suggested by evidence that both constitutive and drug inducible CYP2C9 mRNA expression was elevated in HepG2 cell lines stably transfected with mCAR or hCAR, in the absence or presence of TCPOBOP or PB, respectively.29 By analysis of the CYP2C9 promoter region, two CAR‐responsive elements, a DR5 and a DR4, were discovered independently.29,115 CYP2C8 is the most inducible CYP2C isozyme in human hepatocytes. Examination of the CYP2C8 promoter region revealed that a distal PXR/ CAR‐binding site located to 8806 bp of the transcriptional start site confers inducibility of CYP2C8 via the PXR agonist RIF and the CAR agonist CITCO.47 Further analysis of the CYP2C8 basal promoter region revealed several putative binding motifs, including TATA‐box, HNF3, CCAAT enhancer‐binding protein, HNF4a, and GATA‐binding protein, indicating

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many other potential nuclear factors may be involved in controlling the transcription of CYP2C8. Another important member of human CYP2C subfamily, CYP2C19, metabolizes a number of clinically important drugs such as omeprazole, diazepam, propranolol, and S‐mephenytoin. Chen et al. reported that transcriptional regulation of CYP19 gene expression is mediated by CAR/PXR and GR through the consensus binding sites (CAR‐RE; 1891/1876 bp) and (GRE; 1750/1736 bp), respectively.116 In addition, a functional CAR response element has been localized and characterized in the promoter region of mouse cyp2c37 gene.117 Intriguingly, this response element only responded to CAR but not PXR. Overall, these data suggest that CAR has a promiscuous role in the regulation of multiple CYP2Cs. 2. CAR REGULATION OF UGT1A1 GENES PB has long been used in the treatment of hyperbilirubinemia since this compound induces the expression of UGT1A1 enzyme involved in bilirubin glucuronidation.118–120 The molecular mechanism of PB induction of UGT1A1 remained elusive until a 290‐bp distal enhancer module, located from 3483 to 3194 bp of the UGT1A1 promoter, was identified as CAR response element.59 Consisting of three putative NR palindromes, this enhancer module can be activated in transfection assays by human and mouse CAR in HepG2 cells, and primary mouse hepatocytes. Site‐directed mutagenesis of this module abrogated the response. In another study, Huang et al. demonstrated that CAR activation increases hepatic expression of several relevant genes involved in bilirubin‐clearance, including UGT1A1, MRP2/ABCC2, SLC21A6, GSTA1, and GSTA2, in wild type but not CAR‐null mice.58 A similar activation profile was observed also in a line of transgenic mice expressing human CAR but not mouse CAR in the liver. These results suggest that both human and mouse CAR can respond to elevated levels of bilirubin by regulating simultaneously the expression of UGT1A1, MRP2/ABCC3, SLC21A6, GSTA1, and GSTA2. It is of note that within this 290‐bp distal enhancer module, several other XREs have also been recognized subsequently, including a PXRE, two GREs, and an AhR‐specific XRE.56,121,122 3. CAR REGULATION OF DRUG TRANSPORTERS Several efflux transporters, as well as members of the OATP uptake transporter family, have been identified as transcriptional target genes of PXR, CAR, PPAR, and FXR.26,30,76,78,123,124 Multidrug resistance‐associated protein 2 (MRP2/ABCC2) is involved in the transport of organic anions, bile salts, glutathione, and xenobiotics, such as the anticancer drugs cisplatin, anthacyclines, vinca alkaloids, and methotrexate.125 Treatment with the PXR‐selective activator PCN and the mouse CAR‐specific ligand TCPOBOP resulted in MRP2/ABCC2 induction at the mRNA level. By analyzing the 50 flanking

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region of the MRP2/ABCC2 gene, Kast et al. isolated an unusual 26‐bp sequence, located 440 bp upstream of the MRP2/ABCC2 transcription initiation site, that contained an everted repeat of AGTTCA spaced by eight nucleotides (ER8).78 This ER8 motif was shown to bind both CAR and PXR, and to be activated by CAR and PXR ligands in immortalized cells. These findings demonstrated the versatility in the binding abilities of PXR and CAR, and provided the first evidence that CAR and PXR can recognize response elements (ER8) in addition to the previously identified ER6 elements. Interestingly, PB induction of MRP2/ABCC2 was greater in PXR‐null mice compared with wild‐type animals. Given the fact that PB induction of CYP2B and CYP3A expression was unaffected in PXR‐null mice but abolished in CAR‐null mice, it was suggested that mouse PXR may interact negatively with PB and that shared PXR/CAR‐target genes may be induced by PB more efficaciously in mice lacking PXR due to reduced ligand‐binding competition. There are conflicting reports regarding the contribution of CAR in the regulation of MRP3/ABCC3, another ATP‐dependent efflux transporter. Staudinger et al. showed that selective activation of mouse CAR by TCPOBOP mediates the inducible expression of Oatp2 and Mrp3/Abcc3.75 These results were similar to those obtained in PXR‐null mice, where PB induction of both Oatp2 and Mrp3/Abcc3 was enhanced. Another report also demonstrated, by RT‐PCR, that both CAR and PXR play critical roles in regulating the mouse Mrp3/Abcc3 gene expression.71 In contrast, human CAR appears not to be involved in the regulation of human MRP3/ABCC3 gene expression. Both human CAR‐ and mouse CAR‐expressing HepG2 cells have been used to elucidate the role of CAR in regulating induction of human CYP2B6 and UGT1A1 by PB‐like inducers.59,105,126

C. Mechanism of CAR Activation CAR is constitutively activated in all the immortalized cell lines without xenobiotic stimulation, making the investigation of CAR activation more challenging. In contrast, CAR activation in primary cultured hepatocytes and intact liver in vivo is inducer‐dependent, and CAR is localized in the cytoplasm of liver cells and translocated into the nucleus only after exposure to xenobiotic inducers.105,126 The essential feature critical for regulating xenobiotic‐induced CAR activation involves nuclear accumulation. Studies pioneered by Masahiko Negishi’s laboratory showed that PB and TCPOBOP treatments were associated with decreased CAR expression in cytoplasmic fraction compared with nuclear extracts, this finding was substantiated by the increased binding of a CAR/RXR heterodimer to NR1 in liver nuclear extracts.126 These results indicate that nuclear translocation of CAR might be the first activation step in response to PB‐type inducers. As a result, the mechanism by which CAR is retained in the cytoplasm has become a major focus of research interest.

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Inhibition of nuclear translocation by antagonists such as androstanes is one possible mechanism to explain cytoplasmic retention of CAR in primary cells and intact liver. However, physiological serum concentrations of androstane metabolites are several magnitudes lower than concentrations required for repression of CAR activation of the PBREM and induction of CYP2B genes in HepG2 cells.127 Moreover, treatment of mouse primary hepatocytes with high concentrations of androstenol could not inhibit PB‐driven nuclear accumulation of CAR.126 It seems that direct binding is not necessary for a drug to stimulate CAR nuclear translocation. Although PB induces nuclear translocation of both mouse and human CAR, it does not bind to either in various in vitro assays.128,129 Similarly, TCPOBOP treatment resulted in mouse and human CAR accumulation in the nucleus even though human CAR could neither bind nor be activated by TCPOBOP.129 Advances in our understanding of the CAR translocation mechanism include the discovery that okadaic acid (OA), an inhibitor of protein phosphatase 2A, was able to inhibit PB‐ and TCPOBOP‐driven CAR translocation and CYP2b10 induction in mouse primary hepatocytes.126,130,131 Nonetheless, studies have demonstrated that translocation of CAR is necessary but not sufficient to activate this receptor. Calcium/calmodulin‐dependent kinase (CaMK) inhibitors, KN‐93 and KN‐62 could efficiently inhibit PB mediated CYP2B induction without affecting CAR nuclear accumulation.132 Taken together, these observations suggest that tight control of an otherwise constitutively active receptor is achieved by cytosolic sequestration in vivo and primary hepatocytes through complicated yet unknown mechanisms.

D. Species Differences in the Activation of CAR Although significant progress on the molecular mechanisms underlying CAR regulation of DME gene expression has been achieved in rodents, information pertaining to hCAR regulation of human DMEs is more limited. Two apparent issues that remain to be resolved are whether human CAR is regulated in the same fashion as the rodent CAR or CAR is the predominant regulator of CYP2B6 in human liver. The current conclusions on the role of CAR in the regulation of CYP2B or other genes are based almost entirely on rodent CAR.104,133 Although human CAR exhibits some common characteristics with its rodent counterparts, such as undergoing nuclear translocation after PB treatment and binding to the PBREM, there are distinct differences between rodent and human CAR. For example, TCPOBOP, the most potent mCAR ligand identified to date, cannot bind or activate either rat or human CAR. All known mCAR inhibitors, such as androstenol, progesterone, androgens, and CaMK inhibitors, do not inhibit hCAR activation. Overall, current evidence suggests that there are clear species‐specific differences in CYP2B induction and CAR activation, reminiscent of the ligand specificity for PXR.

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Generation of the humanized CAR mice model has made it possible to specifically address the effects of these receptors on DME regulation in vivo. In CAR‐null mice, the induction of CYP2b10 gene expression by PB and TCPOBOP was abolished.109 Similarly, in obese Zucker rats, which express extremely low levels of CAR but have normal levels of PXR expression, PB only poorly induces both CYP2B and CYP3A, whereas CYP3A was significantly induced by the PXR‐specific activator PCN.134 A CAR humanized mouse line has been generated.135 This line specifically expresses the hCAR transgene in the mCAR‐null mouse liver as driven by the albumin promoter. Using this model, opposite responses of mCAR and hCAR to a widely used antiemetic, meclizine have been reported.136

VII. Concluding Remarks In the past decade, our understanding of the physiological functions of NRs has grown significantly. Moreover, the knowledge has led to the successful development of several NR agonists and antagonists to treat human diseases. The understanding of xenobiotic receptors has expanded our ability to predict the potential pharmacological and toxicological properties in the early stage of drug development. The PXR and CAR humanized mice are expected to aid pharmaceutical development by predicting potential drug–drug interactions. Detailed mechanisms of NR signaling in both physiological and pathologic conditions will continue to be the major focus of the field. Acknowledgments The original results from our labs that are described in this chapter were generated with the support of NIH grants ES012479, ES014626, CA107011 (to W.X.). The review on the CAR functions has been greatly assisted by Dr. Hongbing Wang at the University of Maryland, School of Pharmacy.

References 1. Chambon P. The nuclear receptor superfamily: a personal retrospect on the first two decades. Mol Endocrinol 2005;19:1418–28. 2. Evans RM. The nuclear receptor superfamily: a rosetta stone for physiology. Mol Endocrinol 2005;19:1429–38. 3. Germain P, Staels B, Dacquet C, Spedding M, Laudet V. Overview of nomenclature of nuclear receptors. Pharmacol Rev 2006;58:685–704. 4. Jensen EV, Khan SA. A two‐site model for antiestrogen action. Mech Ageing Dev 2004;125:679–82. 5. Jensen EV. On the mechanism of estrogen action. Perspect Biol Med 1962;6:47–59.

110

HE AND XIE

6. Green S, Walter P, Kumar V, Krust A, Bornert JM, Argos P, et al. Human oestrogen receptor cDNA: sequence, expression and homology to v‐erb‐A. Nature 1986;320:134–9. 7. Greene GL, Gilna P, Waterfield M, Baker A, Hort Y, Shine J. Sequence and expression of human estrogen receptor complementary DNA. Science 1986;231:1150–4. 8. Hollenberg SM, Weinberger C, Ong ES, Cerelli G, Oro A, Lebo R, et al. Primary structure and expression of a functional human glucocorticoid receptor cDNA. Nature 1985;318:635–41. 9. Aranda A, Pascual A. Nuclear hormone receptors and gene expression. Physiol Rev 2001;81:1269–304. 10. Giguere V, Yang N, Segui P, Evans RM. Identification of a new class of steroid hormone receptors. Nature 1988;331:91–4. 11. Gronemeyer H, Gustafsson JA, Laudet V. Principles for modulation of the nuclear receptor superfamily. Nat Rev Drug Discov 2004;3:950–64. 12. Bain DL, Heneghan AF, Connaghan‐Jones KD, Miura MT. Nuclear receptor structure: implications for function. Annu Rev Physiol 2007;69:201–20. 13. Knotts TA, Orkiszewski RS, Cook RG, Edwards DP, Weigel NL. Identification of a phosphorylation site in the hinge region of the human progesterone receptor and additional amino‐ terminal phosphorylation sites. J Biol Chem 2001;276:8475–83. 14. Lee YK, Choi YH, Chua S, Park YJ, Moore DD. Phosphorylation of the hinge domain of the nuclear hormone receptor LRH‐1 stimulates transactivation. J Biol Chem 2006;281:7850–5. 15. Li Y, Lambert MH, Xu HE. Activation of nuclear receptors: a perspective from structural genomics. Structure 2003;11:741–6. 16. Warnmark A, Treuter E, Wright AP, Gustafsson JA. Activation functions 1 and 2 of nuclear receptors: molecular strategies for transcriptional activation. Mol Endocrinol 2003;17:1901–9. 17. Shiau AK, Barstad D, Loria PM, Cheng L, Kushner PJ, Agard DA, et al. The structural basis of estrogen receptor/coactivator recognition and the antagonism of this interaction by tamoxifen. Cell 1998;95:927–37. 18. Cairns W, Cairns C, Pongratz I, Poellinger L, Okret S. Assembly of a glucocorticoid receptor complex prior to DNA binding enhances its specific interaction with a glucocorticoid response element. J Biol Chem 1991;266:11221–6. 19. Lee MS, Kliewer SA, Provencal J, Wright PE, Evans RM. Structure of the retinoid X receptor alpha DNA binding domain: a helix required for homodimeric DNA binding. Science 1993;260:1117–21. 20. Hard T, Kellenbach E, Boelens R, Maler BA, Dahlman K, Freedman LP, et al. Solution structure of the glucocorticoid receptor DNA‐binding domain. Science 1990;249:157–60. 21. Luisi BF, Xu WX, Otwinowski Z, Freedman LP, Yamamoto KR, Sigler PB. Crystallographic analysis of the interaction of the glucocorticoid receptor with DNA. Nature 1991;352:497–505. 22. Dahlman‐Wright K, Baumann H, McEwan IJ, Almlof T, Wright AP, Gustafsson JA, et al. Structural characterization of a minimal functional transactivation domain from the human glucocorticoid receptor. Proc Natl Acad Sci USA 1995;92:1699–703. 23. Bain DL, Franden MA, McManaman JL, Takimoto GS, Horwitz KB. The N‐terminal region of the human progesterone A‐receptor. Structural analysis and the influence of the DNA binding domain. J Biol Chem 2000;275:7313–20. 24. Birnbaumer M, Schrader WT, O’Malley BW. Assessment of structural similarities in chick oviduct progesterone receptor subunits by partial proteolysis of photoaffinity‐labeled proteins. J Biol Chem 1983;258:7331–7. 25. Honkakoski P, Negishi M. Regulation of cytochrome P450 (CYP) genes by nuclear receptors. Biochem J 2000;347:321–37.

NUCLEAR RECEPTORS AND THEIR PHYSIOLOGICAL FUNCTIONS

111

26. Synold TW, Dussault I, Forman BM. The orphan nuclear receptor SXR coordinately regulates drug metabolism and efflux. Nat Med 2001;7:584–90. 27. Wang H, LeCluyse EL. Role of orphan nuclear receptors in the regulation of drug‐ metabolising enzymes. Clin Pharmacokinet 2003;42:1331–57. 28. Kliewer SA, Moore JT, Wade L, Staudinger JL, Watson MA, Jones SA, et al. An orphan nuclear receptor activated by pregnanes defines a novel steroid signaling pathway. Cell 1998;92:73–82. 29. Ferguson SS, LeCluyse EL, Negishi M, Goldstein JA. Regulation of human CYP2C9 by the constitutive androstane receptor: discovery of a new distal binding site. Mol Pharmacol 2002;62:737–46. 30. Geick A, Eichelbaum M, Burk O. Nuclear receptor response elements mediate induction of intestinal MDR1 by rifampin. J Biol Chem 2001;276:14581–7. 31. Maglich JM, Parks DJ, Moore LB, Collins JL, Goodwin B, Billin AN, et al. Identification of a novel human constitutive androstane receptor (CAR) agonist and its use in the identification of CAR target genes. J Biol Chem 2003;278:17277–83. 32. Blumberg B, Sabbagh W, Jr., Juguilon H, Bolado J, Jr., van Meter CM, Ong ES, et al. SXR, a novel steroid and xenobiotic‐sensing nuclear receptor. Genes Dev 1998;12:3195–205. 33. Bertilsson G, Heidrich J, Svensson K, Asman M, Jendeberg L, Sydow‐Backman M, et al. Identification of a human nuclear receptor defines a new signaling pathway for CYP3A induction. Proc Natl Acad Sci USA 1998;95:12208–13. 34. Maglich JM, Sluder A, Guan X, Shi Y, McKee DD, Carrick K, et al. Comparison of complete nuclear receptor sets from the human, Caenorhabditis elegans and Drosophila genomes. Genome Biol 2001;2: RESEARCH0029. 35. Kliewer SA, Goodwin B, Willson TM. The nuclear pregnane X receptor: a key regulator of xenobiotic metabolism. Endocr Rev 2002;23:687–702. 36. Sonoda J, Xie W, Rosenfeld JM, Barwick JL, Guzelian PS, Evans RM. Regulation of a xenobiotic sulfonation cascade by nuclear pregnane X receptor (PXR). Proc Natl Acad Sci USA 2002;99:13801–6. 37. Xie W, Uppal H, Saini SP, Mu Y, Little JM, Radominska‐Pandya A, et al. Orphan nuclear receptor‐mediated xenobiotic regulation in drug metabolism. Drug Discov Today 2004;9:442–9. 38. Maurel D, Ixart G, Barbanel G, Mekaouche M, Assenmacher I. Effects of acute tilt from orthostatic to head‐down antiorthostatic restraint and of sustained restraint on the intra‐ cerebroventricular pressure in rats. Brain Res 1996;736:165–73. 39. Lehmann JM, McKee DD, Watson MA, Willson TM, Moore JT, Kliewer SA. The human orphan nuclear receptor PXR is activated by compounds that regulate CYP3A4 gene expression and cause drug interactions. J Clin Invest 1998;102:1016–23. 40. Xie W, Barwick JL, Downes M, Blumberg B, Simon CM, Nelson MC, et al. Humanized xenobiotic response in mice expressing nuclear receptor SXR. Nature 2000;406:435–9. 41. Staudinger JL, Goodwin B, Jones SA, Hawkins‐Brown D, MacKenzie KI, LaTour A, et al. The nuclear receptor PXR is a lithocholic acid sensor that protects against liver toxicity. Proc Natl Acad Sci USA 2001;98:3369–74. 42. Honkakoski P, Zelko I, Sueyoshi T, Negishi M. The nuclear orphan receptor CAR‐retinoid X receptor heterodimer activates the phenobarbital‐responsive enhancer module of the CYP2B gene. Mol Cell Biol 1998;18:5652–8. 43. Wang H, Negishi M. Transcriptional regulation of cytochrome p450 2B genes by nuclear receptors. Curr Drug Metab 2003;4:515–25. 44. Dussault I, Yoo HD, Lin M, Wang E, Fan M, Batta AK, et al. Identification of an endogenous ligand that activates pregnane X receptor‐mediated sterol clearance. Proc Natl Acad Sci USA 2003;100:833–8.

112

HE AND XIE

45. Goodwin B, Moore LB, Stoltz CM, McKee DD, Kliewer SA. Regulation of the human CYP2B6 gene by the nuclear pregnane X receptor. Mol Pharmacol 2001;60:427–31. 46. Dvorak Z, Modriansky M, Pichard‐Garcia L, Balaguer P, Vilarem MJ, Ulrichova J, et al. Colchicine down‐regulates cytochrome P450 2B6, 2C8, 2C9, and 3A4 in human hepatocytes by affecting their glucocorticoid receptor‐mediated regulation. Mol Pharmacol 2003;64:160–9. 47. Ferguson SS, Chen Y, LeCluyse EL, Negishi M, Goldstein JA. Human CYP2C8 is transcriptionally regulated by the nuclear receptors constitutive androstane receptor, pregnane X receptor, glucocorticoid receptor, and hepatic nuclear factor 4alpha. Mol Pharmacol 2005;68:747–57. 48. Tukey RH, Strassburg CP. Human UDP‐glucuronosyltransferases: metabolism, expression, and disease. Annu Rev Pharmacol Toxicol 2000;40:581–616. 49. Hayes JD, Pulford DJ. The glutathione S‐transferase supergene family: regulation of GST and the contribution of the isoenzymes to cancer chemoprotection and drug resistance. Crit Rev Biochem Mol Biol 1995;30:445–600. 50. Salinas AE, Wong MG. Glutathione S‐transferases—a review. Curr Med Chem 1999;6:279–309. 51. Nagata K, Yamazoe Y. Pharmacogenetics of sulfotransferase. Annu Rev Pharmacol Toxicol 2000;40:159–76. 52. Sheweita SA. Drug‐metabolizing enzymes: mechanisms and functions. Curr Drug Metab 2000;1:107–32. 53. Radominska‐Pandya A, Czernik PJ, Little JM, Battaglia E, Mackenzie PI. Structural and functional studies of UDP‐glucuronosyltransferases. Drug Metab Rev 1999;31:817–99. 54. Mackenzie PI, Owens IS, Burchell B, Bock KW, Bairoch A, Belanger A, et al. The UDP glycosyltransferase gene superfamily: recommended nomenclature update based on evolutionary divergence. Pharmacogenetics 1997;7:255–69. 55. Mackenzie PI, Gregory PA, Gardner‐Stephen DA, Lewinsky RH, Jorgensen BR, Nishiyama T, et al. Regulation of UDP glucuronosyltransferase genes. Curr Drug Metab 2003;4:249–57. 56. Xie W, Yeuh MF, Radominska‐Pandya A, Saini SP, Negishi Y, Bottroff BS, et al. Control of steroid, heme, and carcinogen metabolism by nuclear pregnane X receptor and constitutive androstane receptor. Proc Natl Acad Sci USA 2003;100:4150–5. 57. Chen C, Staudinger JL, Klaassen CD. Nuclear receptor, pregname X receptor, is required for induction of UDP‐glucuronosyltranferases in mouse liver by pregnenolone‐16 alpha‐ carbonitrile. Drug Metab Dispos 2003;31:908–15. 58. Huang W, Zhang J, Chua SS, Qatanani M, Han Y, Granata R, et al. Induction of bilirubin clearance by the constitutive androstane receptor (CAR). Proc Natl Acad Sci USA 2003;100:4156–61. 59. Sugatani J, Kojima H, Ueda A, Kakizaki S, Yoshinari K, Gong QH, et al. The phenobarbital response enhancer module in the human bilirubin UDP‐ glucuronosyltransferase UGT1A1 gene and regulation by the nuclear receptor CAR. Hepatology 2001;33:1232–8. 60. Barbier O, Villeneuve L, Bocher V, Fontaine C, Torra IP, Duhem C, et al. The UDP‐glucuronosyltransferase 1A9 enzyme is a peroxisome proliferator‐activated receptor alpha and gamma target gene. J Biol Chem 2003;278:13975–83. 61. Barbier O, Duran-Sandoval D, Pineda-Torra I, Kosykh V, Fruchart JC, Staels B. Peroxisome proliferator-activated receptor alpha induces hepatic expression of the human bile acid glucuronidating UDP-glucuronosyltransferase 2B4 enzyme. J Biol Chem 2003;278:32852–60. 62. Morris ME, Pang KS. Competition between two enzymes for substrate removal in liver: modulating effects due to substrate recruitment of hepatocyte activity. J Pharmacokinet Biopharm 1987;15:473–96. 63. Qian YM, Sun XJ, Tong MH, Li XP, Richa J, Song WC. Targeted disruption of the mouse estrogen sulfotransferase gene reveals a role of estrogen metabolism in intracrine and paracrine estrogen regulation. Endocrinology 2001;142:5342–50.

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64. Song CS, Echchgadda I, Baek BS, Ahn SC, Oh T, Roy AK, et al. Dehydroepiandrosterone sulfotransferase gene induction by bile acid activated farnesoid X receptor. J Biol Chem 2001;276:42549–56. 65. Saini SP, Sonoda J, Xu L, Toma D, Uppal H, Mu Y, et al. A novel constitutive androstane receptor‐mediated and CYP3A‐independent pathway of bile acid detoxification. Mol Pharmacol 2004;65:292–300. 66. Uppal H, Saini SP, Moschetta A, Mu Y, Zhou J, Gong H, et al. Activation of LXRs prevents bile acid toxicity and cholestasis in female mice. Hepatology 2007;45:422–32. 67. Fang HL, Strom SC, Ellis E, Duanmu Z, Fu J, Duniec‐Dmuchowski Z, et al. Positive and negative regulation of human hepatic hydroxysteroid sulfotransferase (SULT2A1) gene transcription by rifampicin: roles of hepatocyte nuclear factor 4alpha and pregnane X receptor. J Pharmacol Exp Ther 2007;323:586–98. 68. Townsend DM, Tew KD. The role of glutathione‐S‐transferase in anti‐cancer drug resistance. Oncogene 2003;22:7369–75. 69. Falkner KC, Pinaire JA, Xiao GH, Geoghegan TE, Prough RA. Regulation of the rat glutathione S‐transferase A2 gene by glucocorticoids: involvement of both the glucocorticoid and pregnane X receptors. Mol Pharmacol 2001;60:611–9. 70. Rosenfeld JM, Vargas R, Jr., Xie W, Evans RM. Genetic profiling defines the xenobiotic gene network controlled by the nuclear receptor pregnane X receptor. Mol Endocrinol 2003;17:1268–82. 71. Maglich JM, Stoltz CM, Goodwin B, Hawkins‐Brown D, Moore JT, Kliewer SA. Nuclear pregnane x receptor and constitutive androstane receptor regulate overlapping but distinct sets of genes involved in xenobiotic detoxification. Mol Pharmacol 2002;62:638–46. 72. Gong H, Singh SV, Singh SP, Mu Y, Lee JH, Saini SP, et al. Orphan nuclear receptor pregnane X receptor sensitizes oxidative stress responses in transgenic mice and cancerous cells. Mol Endocrinol 2006;20:279–90. 73. Klaassen CD, Slitt AL. Regulation of hepatic transporters by xenobiotic receptors. Curr Drug Metab 2005;6:309–28. 74. Hagenbuch B, Meier PJ. The superfamily of organic anion transporting polypeptides. Biochim Biophys Acta 2003;1609:1–18. 75. Staudinger JL, Madan A, Carol KM, Parkinson A. Regulation of drug transporter gene expression by nuclear receptors. Drug Metab Dispos 2003;31:523–7. 76. Guo GL, Staudinger J, Ogura K, Klaassen CD. Induction of rat organic anion transporting polypeptide 2 by pregnenolone‐16alpha‐carbonitrile is via interaction with pregnane X receptor. Mol Pharmacol 2002;61:832–9. 77. Kruh GD, Belinsky MG. The MRP family of drug efflux pumps. Oncogene 2003;22:7537–52. 78. Kast HR, Goodwin B, Tarr PT, Jones SA, Anisfeld AM, Stoltz CM, et al. Regulation of multidrug resistance‐associated protein 2 (ABCC2) by the nuclear receptors pregnane X receptor, farnesoid X‐activated receptor, and constitutive androstane receptor. J Biol Chem 2002;277:2908–15. 79. Teng S, Jekerle V, Piquette‐Miller M. Induction of ABCC3 (MRP3) by pregnane X receptor activators. Drug Metab Dispos 2003;31:1296–9. 80. Moore LB, Goodwin B, Jones SA, Wisely GB, Serabjit‐Singh CJ, Willson TM, et al. St. John’s wort induces hepatic drug metabolism through activation of the pregnane X receptor. Proc Natl Acad Sci USA 2000;97:7500–2. 81. Mu Y, Zhang J, Zhang S, Zhou HH, Toma D, Ren S, et al. Traditional Chinese medicines Wu Wei Zi (Schisandra chinensis Baill) and Gan Cao (Glycyrrhiza uralensis Fisch) activate pregnane X receptor and increase warfarin clearance in rats. J Pharmacol Exp Ther 2006;316:1369–77. 82. Goldstein JA. Clinical relevance of genetic polymorphisms in the human CYP2C subfamily. Br J Clin Pharmacol 2001;52:349–55.

114

HE AND XIE

83. Sinz M, Kim S, Zhu Z, Chen T, Anthony M, Dickinson K, et al. Evaluation of 170 xenobiotics as transactivators of human pregnane X receptor (hPXR) and correlation to known CYP3A4 drug interactions. Curr Drug Metab 2006;7:375–88. 84. Gong H, Sinz MW, Feng Y, Chen T, Venkataramanan R, Xie W. Animal models of xenobiotic receptors in drug metabolism and diseases. Methods Enzymol 2005;400:598–618. 85. Xie W, Evans RM. Orphan nuclear receptors: the exotics of xenobiotics. J Biol Chem 2001;276:37739–42. 86. Xie W, Radominska‐Pandya A, Shi Y, Simon CM, Nelson MC, Ong ES, et al. An essential role for nuclear receptors SXR/PXR in detoxification of cholestatic bile acids. Proc Natl Acad Sci USA 2001;98:3375–80. 87. Sonoda J, Rosenfeld JM, Xu L, Evans RM, Xie W. A nuclear receptor‐mediated xenobiotic response and its implication in drug metabolism and host protection. Curr Drug Metab 2003;4:59–72. 88. Stedman CA, Liddle C, Coulter SA, Sonoda J, Alvarez JG, Moore DD, et al. Nuclear receptors constitutive androstane receptor and pregnane X receptor ameliorate cholestatic liver injury. Proc Natl Acad Sci USA 2005;102:2063–8. 89. Zhang J, Huang W, Qatanani M, Evans RM, Moore DD. The constitutive androstane receptor and pregnane X receptor function coordinately to prevent bile acid‐induced hepatotoxicity. J Biol Chem 2004;279:49517–22. 90. Uppal H, Toma D, Saini SP, Ren S, Jones TJ, Xie W. Combined loss of orphan receptors PXR and CAR heightens sensitivity to toxic bile acids in mice. Hepatology 2005;41:168–76. 91. Zhai Y, Pai HV, Zhou J, Amico JA, Vollmer RR, Xie W. Activation of pregnane X receptor disrupts glucocorticoid and mineralocorticoid homeostasis. Mol Endocrinol 2007;21:138–47. 92. Zhou J, Zhai Y, Mu Y, Gong H, Uppal H, Toma D, et al. A novel pregnane X receptor‐ mediated and sterol regulatory element‐binding protein‐independent lipogenic pathway. J Biol Chem 2006;281:15013–20. 93. Tucker CM, Petersen S, Herman KC, Fennell RS, Bowling B, Pedersen T, et al. Self‐ regulation predictors of medication adherence among ethnically different pediatric patients with renal transplants. J Pediatr Psychol 2001;26:455–64. 94. Weaver RJ. Assessment of drug‐drug interactions: concepts and approaches. Xenobiotica 2001;31:499–538. 95. Weaver SA, Russo MP, Wright KL, Kolios G, Jobin C, Robertson DA, et al. Regulatory role of phosphatidylinositol 3‐kinase on TNF‐alpha‐induced cyclooxygenase 2 expression in colonic epithelial cells. Gastroenterology 2001;120:1117–27. 96. Mills JB, Rose KA, Sadagopan N, Sahi J, de Morais SM. Induction of drug metabolism enzymes and MDR1 using a novel human hepatocyte cell line. J Pharmacol Exp Ther 2004;309:303–9. 97. Kocarek TA, Schuetz EG, Strom SC, Fisher RA, Guzelian PS. Comparative analysis of cytochrome P4503A induction in primary cultures of rat, rabbit, and human hepatocytes. Drug Metab Dispos 1995;23:415–21. 98. Jones SA, Moore LB, Shenk JL, Wisely GB, Hamilton GA, McKee DD, et al. The pregnane X receptor: a promiscuous xenobiotic receptor that has diverged during evolution. Mol Endocrinol 2000;14:27–39. 99. Watkins RE, Wisely GB, Moore LB, Collins JL, Lambert MH, Williams SP, et al. The human nuclear xenobiotic receptor PXR: structural determinants of directed promiscuity. Science 2001;292:2329–33. 100. Kolars JC, Schmiedlin‐Ren P, Schuetz JD, Fang C, Watkins PB. Identification of rifampin‐ inducible P450IIIA4 (CYP3A4) in human small bowel enterocytes. J Clin Invest 1992;90:1871–8.

NUCLEAR RECEPTORS AND THEIR PHYSIOLOGICAL FUNCTIONS

115

101. Ma X, Shah Y, Cheung C, Guo GL, Feigenbaum L, Krausz KW, et al. The PREgnane X receptor gene‐humanized mouse: a model for investigating drug‐drug interactions mediated by cytochromes P450 3A. Drug Metab Dispos 2007;35:194–200. 102. Moore JT, Kliewer SA. Use of the nuclear receptor PXR to predict drug interactions. Toxicology 2000;153:1–10. 103. Baes M, Gulick T, Choi HS, Martinoli MG, Simha D, Moore DD. A new orphan member of the nuclear hormone receptor superfamily that interacts with a subset of retinoic acid response elements. Mol Cell Biol 1994;14:1544–51. 104. Choi HS, Chung M, Tzameli I, Simha D, Lee YK, Seol W, et al. Differential transactivation by two isoforms of the orphan nuclear hormone receptor CAR. J Biol Chem 1997;272:23565–71. 105. Sueyoshi T, Kawamoto T, Zelko I, Honkakoski P, Negishi M. The repressed nuclear receptor CAR responds to phenobarbital in activating the human CYP2B6 gene. J Biol Chem 1999;274:6043–6. 106. Trottier E, Belzil A, Stoltz C, Anderson A. Localization of a phenobarbital‐responsive element (PBRE) in the 50 ‐ flanking region of the rat CYP2B2 gene. Gene 1995;158:263–8. 107. Park Y, Li H, Kemper B. Phenobarbital induction mediated by a distal CYP2B2 sequence in rat liver transiently transfected in situ. J Biol Chem 1996;271:23725–8. 108. Honkakoski P, Negishi M. Characterization of a phenobarbital‐responsive enhancer module in mouse P450 Cyp2b10 gene. J Biol Chem 1997;272:14943–9. 109. Wei P, Zhang J, Egan‐Hafley M, Liang S, Moore DD. The nuclear receptor CAR mediates specific xenobiotic induction of drug metabolism. Nature 2000;407:920–3. 110. Ueda A, Hamadeh HK, Webb HK, Yamamoto Y, Sueyoshi T, Afshari CA, et al. Diverse roles of the nuclear orphan receptor CAR in regulating hepatic genes in response to phenobarbital. Mol Pharmacol 2002;61:1–6. 111. Chen Y, Ferguson SS, Negishi M, Goldstein JA. Induction of human CYP2C9 by rifampicin, hyperforin, and phenobarbital is mediated by the pregnane X receptor. J. Pharmacol Exp Ther 2004;308:495–501. 112. Raucy JL, Mueller L, Duan K, Allen SW, Strom S, Lasker JM. Expression and induction of CYP2C P450 enzymes in primary cultures of human hepatocytes. J Pharmacol Exp Ther 2002;302:475–82. 113. Rae JM, Johnson MD, Lippman ME, Flockhart DA. Rifampin is a selective, pleiotropic inducer of drug metabolism genes in human hepatocytes: studies with cDNA and oligonucleotide expression arrays. J Pharmacol Exp Ther 2001;299:849–57. 114. Gerbal‐Chaloin S, Pascussi JM, Pichard‐Garcia L, Daujat M, Waechter F, Fabre JM, et al. Induction of CYP2C genes in human hepatocytes in primary culture. Drug Metab Dispos 2001;29:242–51. 115. Gerbal‐Chaloin S, Daujat M, Pascussi JM, Pichard‐Garcia L, Vilarem MJ, Maurel P. Transcriptional regulation of CYP2C9 gene. Role of glucocorticoid receptor and constitutive androstane receptor. J Biol Chem 2002;277:209–17. 116. Chen Y, Ferguson SS, Negishi M, Goldstein JA. Identification of constitutive androstane receptor and glucocorticoid receptor binding sites in the CYP2C19 promoter. Mol Pharmacol 2003;64:316–24. 117. Jackson JP, Ferguson SS, Negishi M, Goldstein JA. Phenytoin induction of the cyp2c37 gene is mediated by the constitutive androstane receptor. Drug Metab Dispos 2006;34:2003–2. 118. Schambach K, Menzel K. [Clinical experiences using phenobarbital in the prevention of hyperbilirubinemia in mature newborn infants in a controlled study]. Z Arztl Fortbild (Jena) 1975;69:535–7. 119. Kopecky P, Schwarz I, Schwenzel W. [Antepartal phenobarbital therapy for the improvement of fetal bilirubin conjugation]. Arch Gynakol 1975;219:455–7.

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HE AND XIE

120. Ishii Y, Tsuruda K, Tanaka M, Oguri K. Purification of a phenobarbital‐inducible morphine UDP‐glucuronyltransferase isoform, absent from Gunn rat liver. Arch Biochem Biophys 1994;315:345–51. 121. Yueh MF, Huang YH, Hiller A, Chen S, Nguyen N, Tukey RH. Involvement of the xenobiotic response element (XRE) in Ah receptor‐mediated induction of human UDP‐glucuronosyltransferase 1A1. J Biol Chem 2003;278:15001–6. 122. Kuno T, Togawa H, Mizutani T. Induction of human UGT1A1 by a complex of dexamethasone‐GR dependent on proximal site and independent of PBREM. Mol Biol Rep 2007;35(3), 361–7 [Epub May 26, 2007]. 123. Guo GL, Johnson DR, Klaassen CD. Postnatal expression and induction by pregnenolone‐ 16alpha‐carbonitrile of the organic anion‐transporting polypeptide 2 in rat liver. Drug Metab Dispos 2002;30:283–8. 124. Dussault I, Lin M, Hollister K, Wang EH, Synold TW, Forman BM. Peptide mimetic HIV protease inhibitors are ligands for the orphan receptor SXR. J Biol Chem 2001;276:33309–12. 125. Konig J, Nies AT, Cui Y, Leier I, Keppler D. Conjugate export pumps of the multidrug resistance protein (MRP) family: localization, substrate specificity, and MRP2‐mediated drug resistance. Biochim Biophys Acta 1999;1461:377–94. 126. Kawamoto T, Sueyoshi T, Zelko I, Moore R, Washburn K, Negishi M. Phenobarbital‐responsive nuclear translocation of the receptor CAR in induction of the CYP2B gene. Mol Cell Biol 1999;19:6318–22. 127. Dufort I, Soucy P, Lacoste L, Luu‐The V. Comparative biosynthetic pathway of androstenol and androgens. J Steroid Biochem Mol Biol 2001;77:223–7. 128. Moore LB, Parks DJ, Jones SA, Bledsoe RK, Consler TG, Stimmel JB, et al. Orphan nuclear receptors constitutive androstane receptor and pregnane X receptor share xenobiotic and steroid ligands. J Biol Chem 2000;275:15122–7. 129. Tzameli I, Pissios P, Schuetz EG, Moore DD. The xenobiotic compound 1,4‐bis[2‐(3,5‐ dichloropyridyloxy)]benzene is an agonist ligand for the nuclear receptor CAR. Mol Cell Biol 2000;20:2951–8. 130. Honkakoski P, Negishi M. Protein serine/threonine phosphatase inhibitors suppress phenobarbital‐induced Cyp2b10 gene transcription in mouse primary hepatocytes. Biochem J 1998;330(Pt 2): 889–95. 131. Sidhu JS, Omiecinski CJ. An okadaic acid‐sensitive pathway involved in the phenobarbital‐ mediated induction of CYP2B gene expression in primary rat hepatocyte cultures. J Pharmacol Exp Ther 1997;282:1122–9. 132. Negishi M. Nuclear receptor CAR as a phenobarbital induction signal of CYP2B gene [abstract]. FASEB J 2000;14:1306. 133. Sueyoshi T, Negishi M. Phenobarbital response elements of cytochrome P450 genes and nuclear receptors. Annu Rev Pharmacol Toxicol 2001;41:123–43. 134. Zelko I, Negishi M. Phenobarbital‐elicited activation of nuclear receptor CAR in induction of cytochrome P450 genes. Biochem Biophys Res Commun 2000;277:1–6. 135. Zhang J, Huang W, Chua SS, Wei P, Moore DD. Modulation of acetaminophen‐induced hepatotoxicity by the xenobiotic receptor CAR. Science 2002;298:422–4. 136. Huang W, Zhang J, Wei P, Schrader WT, Moore DD. Meclizine is an agonist ligand for mouse constitutive androstane receptor (CAR) and an inverse agonist for human CAR. Mol Endocrinol 2004;18:2402–8.

Emerging Roles of the Ubiquitin Proteasome System in Nuclear Hormone Receptor Signaling David M. Lonard and Bert W. O’Malley Department of Molecular and Cellular Biology, Baylor College of Medicine, Houston, Texas 77030

I. Introduction: Nuclear Hormone Receptors, Ubiquitin, and the Proteasome....................................................................... II. The Ubiquitin Proteasome System .................................................... III. Nuclear Receptor Interactions with the Ubiquitin Proteasome System....... A. Nuclear Receptor Protein Stability, Targeting by the Ubiquitin Proteasome System and Influences of Ligand on Nuclear Receptor Protein Degradation ................................................................. B. Role of the Ubiquitin Proteasome System in Nuclear Receptor‐ Mediated Transcription ............................................................. IV. Coregulators and the UPS............................................................... V. Coregulators as UPS Targets............................................................ VI. Ubiquitin‐Like Modifications in Nuclear Receptor Signaling ................... VII. Conclusion and Perspective............................................................. References ..................................................................................

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Nuclear receptor (NR)‐mediated transcription is intimately tied to the ubiquitin proteasome system (UPS). The UPS targets numerous NR and coregulator proteins, regulating their stability and altering their transcriptional activities through the posttranslational placement of ubiquitin marks on them. Differences in the manner in which ubiquitin is attached to target proteins or itself have distinct regulatory consequences. Protein monoubiquitination, polyubiquitination, the site of ubiquitin attachment to a target protein, and the type of polyubiquitin chain linkage all lead to different biological outcomes and have an important regulatory function in NR‐mediated transcription. Consistent with its role in protein degradation, the UPS is able to limit the biological actions of both NRs and coregulators by reducing their protein concentrations in the cell. However, in spite of its destructive capabilities, the UPS can play a positive role in facilitating NR‐mediated transcription as well. In addition, ubiquitin‐like modifications such as SUMOylation also modify and regulate

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NRs and coregulators. The UPS forms a key biological system that underlies a sophisticated postranslational regulatory scheme from which complex and dynamic regulation of NR‐mediated transcription can occur.

I. Introduction: Nuclear Hormone Receptors, Ubiquitin, and the Proteasome Nuclear receptors (NRs) accomplish their transcriptional activities through the help of an ever growing list of accessory proteins such as stress proteins and coregulators.1–3 For instance, heat shock proteins have long been known to associate with NRs, maintaining them in a readied state while unliganded, then in the presence of ligand, these proteins are dismissed and coactivator proteins are subsequently recruited.1,4 Since the discovery of coactivators and corepressors, our understanding of NR‐mediated transcription has led to the realization that a diverse array of interacting proteins is needed for NRs to function as transcription factors. A common theme that runs through coactivator and corepressor biology is that these proteins harbor myriad distinct enzymatic activities that contribute either positively or negatively to transcription.1 Corepressor complexes possess enzymatic functions that serve to effect transcriptional repression, such as by contributing histone deacetylase activities that remove activating acetylation marks on histones.5 On the other hand when NRs are bound to agonist ligands, coactivators are recruited that perform enzymatic actions likely to increase gene transcription, such as by acetylating, methylating, or rearranging histones.1–3 It was seen very early on that, in addition to these coregulatory enzymatic activities, ubiquitin proteasome system (UPS)‐related enzyme activity is also involved in modulating NR‐mediated gene expression. Indeed, one of the first identified NR‐interacting proteins, TRIP1, was pulled out of an early search for thyroid hormone (TR)‐interacting proteins and later shown to be a component of the UPS.6,7 Subsequent searches for NR coregulators and work on deciphering the histone code revealed additional UPS‐ related coregulators, substantiating the role of this system in NR‐mediated transcription. The impact that the UPS has on NR‐mediated transcription is complex but important and will be discussed in detail in this chapter.

II. The Ubiquitin Proteasome System A complete description of the UPS is beyond scope of this chapter and is covered in greater detail elsewhere.8–10 It consists of two major functional constituents. The first is a targeting system responsible for directing the covalent attachment of ubiquitin onto target proteins. This includes the sole

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ubiquitin‐activating enzyme, ubiquitin‐conjugating enzymes, and numerous ubiquitin ligases that function collectively to direct the 76 amino acid ubiquitin moiety to internal lysine residues in cellular proteins through a covalent isopeptide linkage. Subsequent ubiquitins then can be attached to the initial ubiquitin linked to the target protein, leading to the formation of a polyubiquitin chain. The second major part of this system consists of the proteasome, a large multisubunit barrel‐shaped protease whose active sites are contained within its central cavity.11 It is misleading to think degradation is the inevitable sole fate for all ubiquitinated proteins. Ubiquitin can function as a signaling molecule independent of protein degradation; for instance, monoubiquitination does not promote protein instability such as in the case of histones H2A and H2B.12 Polyubiquitin chains that are not formed through K48 ubiquitin linkages are also not subjected to degradation. The proteasome consists of a 20S ‘‘core’’ that is responsible for protein hydrolysis. The core proteasome is regulated by different regulatory ‘‘cap’’ complexes. One of these, the 19S particle contains proteins that recognize polyubiquitinated proteins and a hexameric ATPase‐dependent ‘‘unfoldase’’ that facilitates the entry of proteins into the 20S proteolytic core particle. The proteasome can also degrade proteins in an ubiquitin‐ and ATP‐independent manner through its association with alternative regulatory complexes such as REGg.13 Unlike the 19S particle, the REGg regulatory particle lacks unfoldase activity and primarily degrades unstructured proteins and peptides. The UPS is linked with RNA polymerase II‐mediated transcription at a fundamental level. A classic suppressor of galactose‐1 genetic screen identified Sug1/TRIP1 as a protein involved in transcription.14,15 In addition to this, even before ubiquitin was known to be involved in protein degradation, it was a recognized chromatin‐associated protein linked to histones.12 Histones H2A and H2B are extensively monoubiquitinated and their posttranslational modifications play key roles in transcription, telomeric silencing, transcriptional elongation, and transcriptional termination. Ubiquitination of histone H2B is an upstream signaling event leading to subsequent histone H3 lysine‐4 methylation, a histone posttranslational modification strongly associated with actively transcribed genes.16 The placement of ubiquitin on H2B depends upon NR‐interacting proteins such as Sug1/TRIP117 (Fig. 1, #3). Other evidence indicates that the UPS plays a basic role in transcription, histone coding, and chromatin remodeling. Examples reveal the core TFIID complex component, TAF1/TAFII250 to be an ubiquitin‐activating and ‐conjugating enzyme18. Other evidence points to histone ubiquitination as playing a key role in downstream events involved in histone H3 K4 methylation and in response to DNA damage.19,20 All of these findings point to the notion that the UPS is involved in basic mechanisms of transcriptional regulation and leads to the full expectation that it would play a key role in NR‐mediated gene transcription as well.21,22

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2 Ub Ub ligase CoA 3 1

CoA

4 Ub

CoA

Ub NHR NHR

Ub

Ub

RNA pol II

Ub

FIG. 1. The ubiquitin proteasome system interacts with NR‐mediated transcription at multiple levels. (1) Ubiquitin (Ub) ligase coactivators (CoA) can posttranscriptionally modify nuclear hormone receptors (NR), altering their transcriptional activity and promoting their degradation by the proteasome. (2) CoA that are part of a coactivator protein complex are targeted by Ub ligase CoA members of the complex. (3) Ub ligase CoA target histones H2A and H2B, altering the chromatin histone code. (4) RNA polymerase (pol) II is also targeted by the proteasome.

III. Nuclear Receptor Interactions with the Ubiquitin Proteasome System A. Nuclear Receptor Protein Stability, Targeting by the Ubiquitin Proteasome System and Influences of Ligand on Nuclear Receptor Protein Degradation Even before coregulator biology came into being, it was known that the stability of NR proteins is influenced by their cognate ligands.23–25 The cellular concentrations of estrogen receptor‐alpha (ERa) and progesterone receptor (PR) were observed to be downregulated in the presence of their cognate ligands. Subsequent work determined that the UPS plays a key role in the degradation of the ERa protein.26,27 The thyroid hormone receptor (TR),28 retinoic acid receptors (RARs),29–31 glucocorticoid receptor (GR),32,33 and mineralocorticoid receptor (MR)34,35 are also subject to accelerated protein degradation in the presence of their cognate ligands. In contrast, the vitamin D receptor (VDR)36,37 and androgen receptor (AR)38,39 are stabilized by vitamin D and androgens, respectively; pointing out that ligand‐mediated degradation is not a universal phenomenon among NRs. An interesting observation is that agonist ligands for many NRs promote receptor protein degradation at the same time they stimulate NR‐mediated gene expression. Is NR protein degradation inexorably linked to gene

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expression in these cases? Early evidence seemed to support this idea; inhibition of RNA polymerase II‐mediated transcription with actinomycin D or a‐amanitin resulted in the stabilization of ERa and PR.24,40 This line of thinking was supported by the finding that the selective estrogen receptor modulators (SERMs), 4‐hydroxytamoxifen (the biologically active metabolite of tamoxifen) and raloxifene, both were able to stabilize ERa.41,42 On the other hand, the ‘‘pure’’ antiestrogen ICI 182,780 promotes receptor protein degradation without any stimulation of ERa‐dependent gene expression, thus functioning as a selective estrogen receptor downregulator (SERD).42–44 In the case of ICI 182,780, SERD‐mediated degradation of ERa is dependent upon the ability of this ligand to induce a distinct conformation in the receptor LBD on helices 3 and 5.42 This ligand‐induced conformational surface then is likely able to recruit an endogenous protein(s) that promotes degradation of the receptor.

B. Role of the Ubiquitin Proteasome System in Nuclear Receptor‐Mediated Transcription Even now, our understanding of whether ERa degradation promotes or limits transcription remains unclear. However, it is clear that the ERa protein is ubiquitinated and degraded by the proteasome.23,24 Early studies that focused on the consequences of proteasome inhibition on the transcriptional activity of ERa found that proteasome activity was necessary for its ability to drive expression from a reporter gene while in contrast, GR‐mediated transcription was unimpeded by proteasome inhibition.24 ERa stimulation of PR gene expression was abrogated with proteasome inhibitor treatment; however, in another study it was found that another ERa target gene, pS2, was not inhibited.45 Yet another study showed that proteasome inhibition blocks E2‐ dependent ERa‐mediated transcription, but not ligand‐independent function of the receptor.46 In an effort to better understand these observations, the complex receptor‐ and gene‐specific actions that proteasome inhibition has on both ERa and GR were studied through gene expression profiling. This revealed that the proteasome affects both receptors in complex ways.47 For both ERa and GR receptors, proteasome inhibition blocks the expression of many genes, has no effect on others, and stimulates yet another group of NR‐regulated genes. The UPS plays a fundamental role providing for the dynamic interchange of proteins at the promoter of the estrogen‐regulated pS2 gene promoter.40 Chromatin immunoprecipitation (ChIP) analysis revealed that cycling of ERa, coactivators, general transcription machinery, and RNA pol II at the promoter was impaired in the presence of the proteasome inhibitor MG132. This loss of dynamic protein exchange at the ERa‐regulated pS2 gene is thought to be responsible for abrogated receptor‐dependent gene expression.

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In another investigation of the mechanism that the proteasome plays in PR‐ mediated transcription, ChIP analysis was used to determine the recruitment of receptor, coactivators and RNA pol II to a stably integrated PR‐regulated MMTV‐CAT gene.48 PR, SRC family coactivators, and proteasome subunits were not affected by treatment of cells with MG132. However, the recruitment of RNA pol II was inhibited, indicating that while many of the substeps of PR‐ mediated transcription do not rely upon the proteasome, an unidentified part of the transcriptional process downstream of coactivator recruitment requires the proteasome. This study contrasts with that seen for ERa where proteasome inhibition interferes with sequential cofactor recruitment at an early step during transcriptional initiation.40 Proteasome activity is also required for AR‐dependent transcription in prostate cancer cell lines. Treatment of LNCaP and PC‐3 prostate cancer cell lines with the proteasome inhibitor MG132 resulted in impaired AR translocation to the nucleus and reduced interaction with SRC‐2 and ARA70 coactivators.49,50 The targeting of PR for degradation by the proteasome is defective in BRCA1 mutant mice, leading to increased PR‐mediated transcription and subsequent carcinogenesis.51 The resulting elevation in PR in mouse mammary tissue correlates with increased carcinogenesis and suggests that human carriers of the BRCA1 mutation have tumors that are restricted to ERa/PR expressing tissues due to elevations in PR protein content. Consistent with this model, treatment of BRCA1 mutant mice with a PR antagonist RU486 was able to protect against breast cancer, again implicating excess PR‐mediated gene expression as a key molecular event in BRCA1‐mediated carcinogenesis. Unlike ERa and PR, AR is stabilized in the presence of androgens.52 In spite of this, proteasome function is required for AR to function as a transcription factor, suggesting that the degradation of coactivators or other components of the AR‐transcription complex must occur for AR to function as a transcription factor.53 Consistent with this, stabilization of AR is enhanced by the action of the coactivator NRIP, which itself is upregulated by AR and androgens.54,55 Thus, NRIP is part of a feed‐forward mechanism that protects AR from proteasome‐mediated degradation, enhancing its transcriptional activity. Polymorphisms in the polyglutamine tract of human AR play a role in its stability.56–58 Longer polyglutamine tracts are associated with decreased AR‐ mediated transcriptional capacity and decreased receptor protein stability. Phenotypic manifestations of different polyglutamine tract length variations include prostate cancer susceptibility, male infertility, and hirsutism.59 Polyglutamine expansions of greater than 35–40 residues lead to the X‐linked motor neuronal disorder, spinal and bulbar muscular atrophy, also known as Kennedy’s disease. Interestingly, AR and another pathological polyglutamine tract repeat protein termed Huntington, contain related pentapeptide motifs

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FQKLL and FQNLL, respectively, that promote the aggregation of these polyglutamine‐tract proteins into punctate cytoplasmic bodies and their degradation by the proteasome.57 The identification of CHIP, an ubiquitin ligase involved in the degradation of misfolded proteins, has led to a model for ERa that involves two separate mechanisms for receptor degradation.60,61 In its unliganded state the HSP70‐ associated ubiquitin ligase CHIP degrades ERa, while in the presence of estradiol, other ubiquitin ligases take over to facilitate the degradation of the liganded receptor. Overall, it is clear that multiple ubiquitin ligases converge on ERa and other NRs.62 Phosphorylation pathways influence NR protein stability. A recent report indicates that AKT‐mediated stabilization of ERa results in reduced recruitment of the receptor to chromatin and decreased transcription.63 PR proteasome‐mediated degradation depends upon its MAP kinase‐dependent phosphorylation of serine residue 294.64 PR stability is also dependent upon posttranslational modification of lysine residue 388 that doubles as an ubiquitination and SUMOylation site.65 In this case, proteasome‐mediated degradation of PR is necessary for both progesterone and cAMP/PKA pathways to work together to evoke a transcriptionally synergistic response.66 In another study, the previously uncharacterized protein, CUE domain 2 has been shown to interact with PR and reduce receptor SUMOylation at lysine residue 388, promoting ubiquitination at the same lysine residue and enhancing the receptor’s proteasome‐mediated degradation.67 In this case, CUEDC2 reduces the transcriptional potential of PR‐B and abrogates its ability to function in nongenomic MAPK signaling.

IV. Coregulators and the UPS Since it is clear that many NRs are subject to ligand‐regulated proteasome‐ mediated protein degradation, it is a straightforward prediction that UPS‐ related coactivators could promote NR protein degradation (Fig. 1, #1). As mentioned above, TRIP1/Sug1 was identified as a TR‐interacting protein and been shown to promote the degradation of Sp1,68 ERa,69 and VDR.37 The subsequent characterization of TRIP1/sug1 as a component of the 19S proteasome cap seemed at the time to preclude its role as a transcriptional regulator.70 However, subsequent studies now support its role as both a transcriptional regulator and proteasome cap component. For instance, the proteasome is present in the nucleus and components of the 19S cap including TRIP1/sug1 dynamically associate with chromatin and gene promoters.40,71

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The HECT domain ubiquitin ligase E6‐AP is another UPS protein that interacts with NRs.72 E6‐AP acts as a coactivator for PR, ERa, and other receptors, but its ubiquitin ligase activity is not essential for its coactivator function. However, its ubiquitin ligase activity plays a critical role in the manifestation of the Angelman’s syndrome phenotype, an inherited genetic disorder, characterized by mental retardation and gait ataxia, suggesting that E6‐AP has distinct separable biological functions.73,74 Because of this fact, it is unclear exactly how the ubiquitin ligase activity of this protein contributes to NR‐mediated transcription for these NRs. In the case of ERb though, phosphorylation of serine residues in its AF‐1 lead to its targeting by E6‐AP and degradation by the proteasome, indicating that its ubiquitin ligase activity plays a role in regulating the protein stability of this NR.75 Another HECT domain ubiquitin ligase, RPF1, was shown to stimulate PR‐mediated transcription.76 Its yeast homolog RPS5 functions as an ubiquitin ligase that targets RNA polymerase II for ubiquitination, suggesting that the coactivator activity of RPF1 is related to its ability to target RNA polymerase II77 (Fig. 1, #4). Another UPS protein that interacts with NRs is the ubiquitin interacting motif (UIM) containing protein RAP80 that is able to block ERa ubiquitination and degradation.78 RAP80 also plays a role in protein SUMOylation and in response to DNA damage. Further work is required to understand how these different roles of RAP80 might link ERa to DNA damage. The closely related cointegrators, p300 and CBP, are well‐known histone acetyltransferases and this enzymatic function plays a key role in their ability to function as NR coactivators. Unexpectedly though, p300 has also been found to possess ubiquitin ligase activity, acting as an E4 ubiquitin ligase for p53.79,80 Here, Mdm2 functions as an initial E3 ubiquitin ligase that targets p53 for monoubiquitination. Once monoubiquitinated, p53 becomes primed for subsequent E4 polyubiquitination by p300. At first glance, protein ubiquitination can be thought of as a one‐way process, leading to a protein’s proteasome‐mediated destruction. However, protein ubiquitination can be undone through the action of deubiquitinating enzymes (DUBs).81 From genetic and biochemical studies in yeast, DUBs have been found to play critical roles in histone deubiquitination and are required for transcriptional elongation and response to DNA damage. One of these DUBs, USP10, was identified as an AR‐interacting protein in PC3 prostate cancer cells and its enzymatic activity was determined to be necessary for it to enhance AR‐ mediated transcription.82 In another study, a DUB component of the SAGA transcriptional cofactor complex, USP22, was found to be required for full transcriptional activity of AR.83 USP22 targets ubiquitinated histones H2A and H2B, counteracting chromatin silencing and facilitating AR‐dependent transcription through its influence on the histone code. Another DUB, 2A‐DUB, deubiquitinates histone H2A as part of a p/CAF coactivator complex and is

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required for AR‐mediated gene expression.84 The levels of ubiquitinated H2A are significantly reduced in prostate cancers, suggesting a role for 2A‐DUB in the carcinogenesis of androgen‐dependent prostate cancer.

V. Coregulators as UPS Targets Above, we have seen that UPS proteins can act as coactivators by interacting with ligand bound NRs, and in some cases, facilitating transcription through their UPS‐specific enzymatic activities. However, like NRs, coactivators also represent targets of the UPS and this targeting plays a critical role in influencing NR‐mediated transcription (Fig. 1, #2). Inhibition of the proteasome leads to increases in the steady state levels of SRC‐1, SRC‐2, SRC‐3, CBP, p300 as well as the corepressors SMRT and NCoR.27,85,86 In the case of NCoR, the role that its UPS‐mediated destruction plays is perhaps the most straightforward conceptually. NCoR exists as part of a multisubunit protein complex that contains as one of its subunits, the F‐box protein TBL1 and TBLR1.86 Either of these two proteins can function as specificity factors for the Skp/Cullin/F‐box (SCF) ubiquitin ligase complex. TBL1 and TBLR1 contain WD40 motifs that recognize serine/threonine phosphorylated proteins, targeting them for ubiquitination by the SCF complex. In spite of being part of NCoR and SMRT complexes, TBL1 and TBLR1 actually stimulate NR‐ mediated transcription by promoting the proteasome‐mediated degradation of the corepressor, allowing for it to be replaced by coactivators during gene activation. Due to their ability to recognize phosphorylated substrates, TBL1 and TBLR1 act as sensors that detect and destroy phosphorylated NCoR, allowing for its replacement with coactivator complexes. TBL1 and TBLR1 also function as CtBP corepressor complex sensors responsible for the proteasome‐mediated destruction of this corepressor in response to its PKC‐mediated phosphorylation.87 As mentioned above, coactivators are targets of the proteasome and SRC‐3 is the most investigated. Two distinct modes of SRC‐3 proteasome‐mediated degradation have been identified; one is designed to control SRC‐3 protein levels and its activity and the other is coupled to its role in transcription.88,89 REGg is an alternative proteasome cap protein that forms a regulatory particle distinct from the better known 19S proteasome cap.13 The REGg‐proteasome does not recognize ubiquitinated proteins and the spectrum of proteins that it degrades is less well characterized. Known targets include the hepatitis C coat protein90 and the cellular proteins SRC‐388 and p21.91 In this case, SRC‐3 degradation by the proteasome occurs apart from the coactivator’s ubiquitination and involvement in transcription, thus reducing the transcriptional potential of the coactivator. Because SRC‐3 is a powerful growth regulator

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and oncogene, REGg functions to establish a safe cellular concentration ‘‘set point’’ of coactivator that is low enough to prevent uncontrolled cell proliferation. In contrast to its turnover by the REGg‐proteasome, SRC‐3 is also degraded in an ubiquitin‐ and transcription‐dependent manner. In response to estradiol and growth factor signaling, SRC‐3 is phosphorylated at numerous sites that activate the coactivator’s ability to enhance NR‐mediated transcription.92 One of these phosphorylated residues, serine 505, is phosphorylated by GSK3b, promoting the coactivator’s interaction with the SCF Fbw7a ubiquitin ligase complex and its monoubiquitination.89 Another recent study also reported that a GSK3b/S505‐independent phosphorylation event in a C‐terminal PEST region of SRC‐3 can facilitate its ubiquitin‐dependent degradation.93 Ubiquitination is processive; initially SRC‐3 is monoubiquitinated and this event is essential for it to function as an ERa or AR coactivator. Subsequent ubiquitin moieties are attached to SRC‐3 as growing polyubiquitin chains, leading to the coactivator’s eventual destruction by the proteasome. Fbw7a‐mediated SRC‐3 ubiquitination is coupled to SRC‐3’s engagement in transcription, both enhancing its function as a coactivator and limiting the number of times that SRC‐3 can be reused in subsequent rounds of transcription because ubiquitin molecules are added sequentially to the coactivator to form polyubiquitin chains. In this way, the processive nature of ubiquitination is the basis for an unique transcription‐based ‘‘ubiquitin clock.’’94 In yet another twist in this story, other specific SRC‐3 protein phosphorylations protect the coactivator from proteasome‐mediated degradation.95 The atypical PKC kinases PKCz and PKCi target the C‐terminal region of SRC‐3, leading to phosphorylation at distinct residues from those mentioned above. Here, the ubiquitination event described above is presumed to occur still, but these kinases protect the activated SRC‐3 from degradation that would normally occur after its transcriptional activity has been accomplished. This results in a dangerous situation that promotes the accumulation of transcriptionally active SRC‐3. It is possible that the oncogenic potential of elevated atypical PKC activity may be due, in part, to such increases in transcriptionally active SRC‐3. The AKT kinase also protects SRC‐3 from proteasome‐mediated degradation.93 Since AKT activity is frequently enhanced in cancers, it is possible that this is related to elevated cellular concentrations of SRC‐3 as well. A number of other factors also influence SRC‐3 protein stability. SRC‐3 dephosphorylation also plays a dynamic role in its PTM‐coding, influencing its activity and stability.96 Phosphorylation of SRC‐3 facilitates its interaction with the peptidyl isomerase Pin1.97 Pin1 catalyzes a cis/trans isomerization of the amino acid chain at proline residues, leading to conformation state changes in the coactivator that promote its transcriptional activity and degradation. E6‐AP was also shown to target SRC‐3 for ubiquitination.98 SRC‐3 protein

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stability is reduced in serum starved cells. In the presence of serum, the C‐terminus of SRC‐3 interacted with E6‐AP, promoting its ubiquitination and degradation. SRC‐3 protein stability is linked to a nuclear localization sequence (NLS) near the C‐terminus of the coactivator.99 Disruption of the NLS results in its cytoplasmic localization and dramatically higher protein stability. Both ubiquitin‐dependent and ‐independent degradation of SRC‐3 by the Fbw7a SCF ubiquitin ligase complex and REGg occur within the nucleus, consistent with the conclusion that the majority of SRC‐3 protein degradation occurs within the nucleus. Overall, the fate of SRC‐3 rests upon multiple kinase signaling pathways, phosphatases, and distinct modes of proteasome‐mediated destruction. This is intrinsically linked with SRC‐3’s function as a central ‘‘master’’ coregulator that integrates signals from diverse growth factor and hormone signaling pathways. SRC‐2/GRIP‐1 is a target of the UPS. SRC‐2 associates with microscopic ND10 bodies in the nucleus that contain components of the UPS.100 Deletion of the activation domain of SRC‐2 abolished the colocation of SRC‐2 with these proteasome‐enriched nuclear bodies. SRC‐2 protein degradation is accelerated upon its activation by cAMP/PKA. Treatment with compounds that stimulated the cAMP/PKA pathway both enhanced SRC‐2‐mediated transcriptional effects and resulted in greater proteasome‐mediated protein degradation of the coactivator. Treatment with a proteasome inhibitor led to an increase in SRC‐2 recruitment to the pS2 gene in MCF‐7 cells whichstimulated PKA signaling but reduced SRC‐2 recruitment in estradiol‐treated cells.101,102 Consistent with these data, other studies indicate that the consequences of proteasome inhibition on ERa‐dependent gene expression differ between ligand‐dependent and ‐independent pathways; E2 activation of gene expression is proteasome sensitive while PKA activated and ligand‐ independent activation of ERa appears to be resistant to proteasome inhibitors.46 Protein acetylation is linked to proteasome‐mediated protein degradation. These histone acetyltransferases p300, TAF1, and PCAF have been reported to possess ubiquitin ligase activity (TAF1 has both ubiquitin conjugating and ligase activity)18,79,103 and proteins that are modified by acetylation have altered protein stability themselves.104 Protein acetylation plays a protective role by denying lysine ubiquitination at acetylated residues on target proteins. PGC‐1a, like SRC‐3, is a target of an SCF ubiquitin ligase complex, in this case it is ubiquitinated by the SCFCdc4 which leads to reduction in its steady state level, reducing its transcriptional impact on the cell.105 SCFCdc4 expression is downregulated by oxidative stress in neurons, leading to reduced PGC‐ 1a protein degradation. Another study showed that PGC‐1a is also degraded through the proteasome‐independent calpain protease pathway in response to

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oxidative stress while its basal degradation ensues through a proteasome‐ dependent mechanism, indicating that the protein stability of PGC‐1a is subject to complex regulation.106

VI. Ubiquitin‐Like Modifications in Nuclear Receptor Signaling In addition to ubiquitin, a number of other small ubiquitin‐like proteins exist in the cell and can be attached covalently to proteins through an isopeptide bond linkage in the same way as ubiquitin is linked. These include SUMO‐1, SUMO‐2, SUMO‐3, ISG15, and NEDD.107 Each of these moieties is directed to their targets by activating, conjugating, and ligase enzymes similar to ubiquitin targeting, however none of these ubiquitin‐like modifiers target proteins for degradation by the proteasome. Early on, it was found that AR was coactivated by UBC9108,109 and subsequent work went on to reveal that UBC9 is not a ubiquitin conjugating enzyme as its name implies, but rather, the sole SUMO conjugating enzyme in the human genome.110 SUMOylation has primarily been characterized as a covalent modification that functions in transcriptional repression; however in selected instances it actually has the opposite effect.111. The role of SUMOylation in AR‐mediated transcription has been substantiated by the findings that SUMO/sentrin proteases that remove SUMO from SUMOylated proteins can function as AR coactivators.112 Sentrin protease 1 is commonly overexpressed in prostate cancers and its experimental overexpression in the prostate of transgenic mice leads to prostate cancer.113,114 PR and GR are SUMOylated in their AF‐1 as discussed before.65,66 ERa is SUMOylated in its hinge region in a hormone‐dependent manner.115 A SUMOylation‐defective ERa mutant has reduced transcriptional activity while having no influence on the receptor’s nuclear localization. In the SRC‐1 coactivator, conserved lysine residues adjacent to two of its NR boxes facilitate coactivator interaction with NRs when SUMOylated.65 In SRC‐3, more details about the SUMOylation of these conserved lysines are known. Coactivator phosphorylation results in the dismissal of SUMO from lysines 723 and 786, allowing these same sites free to be ubiquitinated.116 SRC‐3 SUMOylation, phosphorylation, monoubiquitination, and polyubiquitination thus form a sequence of posttranslational events involved in the activation and eventual degradation of SRC‐3. RIP140 has also been shown to be SUMOylated at two conserved lysine residues.117 Mutation of these SUMOylation sites impairs RIP140’s function as an NR transcriptional repressor, consistent with the more common observation that SUMOylation is transcriptionally repressive. As an example for protein NEDDylation in the regulation of NR‐mediated transcription, the NEDD8‐activating enzyme Uba3 was identified as a repressor of ERa‐mediated transcription.118 Further work revealed that disruption of

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the NEDDylation system blocked degradation of the ICI 182,780 antagonist‐ bound receptor and conferred resistance to antiestrogens in MCF‐7 breast cancer cells.119 NEDD8 conjugation to the cullin scaffolding protein acts as a posttranslational switch that activates Skp2‐cullin‐F box protein E3 ubiquitin ligase complexes.120

VII. Conclusion and Perspective A large and growing body of evidence exists linking the UPS to NR‐ mediated transcription. In the past, protein degradation did not seem to be an intuitively interesting means of cellular regulation. Now, we see that NR‐ mediated transcription is in large part dependent on the UPS. The regulatory capabilities of the UPS affect NR‐mediated transcription in diverse ways. The initial placement of ubiquitin can be directed to different lysine residues on NRs and coregulators; it can be limited to monoubiquitination, or distinct polyubiquitin chain species (i.e., through K48 or K63), all of which can be edited or removed by DUBs. Histones are predominately monoubiquitinated, a modification that does not lead to protein degradation by the proteasome. Other transcriptional regulators contain a composite of monoubiquitinations and polyubiquitinations, each designed for distinct purposes. The UPS can target many proteins involved in NR‐mediated transcription including NRs, corepressors, coactivators, general transcription factors, chromatin, and RNA polymerase II. Conversely, many of these same proteins are UPS component proteins themselves, indicating that ubiquitin signals both into and out of the NR transcription apparatus. The requirements for and consequences of perturbation of the UPS have varied effects on NR‐mediated transcription, emphasizing its importance for regulation, but making it impossible to establish a simple model for how it influences transcription. References 1. Lonard DM, O’Malley BW. Nuclear receptor coregulators: judges, juries, and executioners of cellular regulation. Mol Cell 2007;27:691–700. 2. Lonard DM, O’Malley BW. The expanding cosmos of nuclear receptor coactivators. Cell 2006;125:411–4. 3. Lonard DM, Lanz RB, O’Malley BW. Nuclear receptor coregulators and human disease. Endocr Rev 2007;28:575–87. 4. Tsai MJ, O’Malley BW. Molecular mechanisms of action of steroid/thyroid receptor superfamily members. Annu Rev Biochem 1994;63:451–86. 5. Rosenfeld MG, Lunyak VV, Glass CK. Sensors and signals: a coactivator/corepressor/epigenetic code for integrating signal‐dependent programs of transcriptional response. Genes Dev 2006;20:1405–28.

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6. Lee JW, Ryan F, Swaffield JC, Johnston SA, Moore DD. Interaction of thyroid‐hormone receptor with a conserved transcriptional mediator. Nature 1995;374:91–4. 7. vom Baur E, Zechel C, Heery D, Heine MJ, Garnier JM, Vivat V, et al. Differential ligand‐ dependent interactions between the AF‐2 activating domain of nuclear receptors and the putative transcriptional intermediary factors mSUG1 and TIF1. EMBO J 1996;15:110–24. 8. Glickman MH, Ciechanover A. The ubiquitin‐proteasome proteolytic pathway: destruction for the sake of construction. Physiol Rev 2002;82:373–428. 9. Ravid T, Hochstrasser M. Diversity of degradation signals in the ubiquitin‐proteasome system. Nat Rev Mol Cell Biol 2008;9:679–90. 10. Konstantinova IM, Tsimokha AS, Mittenberg AG. Role of proteasomes in cellular regulation. Int Rev Cell Mol Biol 2008;267:59–124. 11. Heinemeyer W, Ramos PC, Dohmen RJ. The ultimate nanoscale mincer: assembly, structure and active sites of the 20S proteasome core. Cell Mol Life Sci 2004;61:1562–78. 12. Weake VM, Workman JL. Histone ubiquitination: triggering gene activity. Mol Cell 2008;29:653–63. 13. Hill CP, Masters EI, Whitby FG. The 11S regulators of 20S proteasome activity. Curr Top Microbiol Immunol 2002;268:73–89. 14. Swaffield JC, Bromberg JF, Johnston SA. Alterations in a yeast protein resembling HIV Tat‐ binding protein relieve requirement for an acidic activation domain in GAL4. Nature 1992;357:698–700. 15. Swaffield JC, Melcher K, Johnston SA. A highly conserved ATPase protein as a mediator between acidic activation domains and the TATA‐binding protein. Nature 1995;374:88–91. 16. Briggs SD, Xiao T, Sun ZW, Caldwell JA, Shabanowitz J, Hunt DF, et al. Gene silencing: trans‐ histone regulatory pathway in chromatin. Nature 2002;418:498. 17. Lee D, Ezhkova E, Li B, Pattenden SG, Tansey WP, Workman JL. The proteasome regulatory particle alters the SAGA coactivator to enhance its interactions with transcriptional activators. Cell 2005;123:423–36. 18. Pham AD, Sauer F. Ubiquitin‐activating/conjugating activity of TAFII250, a mediator of activation of gene expression in Drosophila. Science 2000;289:2357–60. 19. Geng F, Tansey WP. Polyubiquitylation of histone H2B. Mol Biol Cell 2008;19:3616–24. 20. Branzei D, Foiani M. Regulation of DNA repair throughout the cell cycle Nat Rev Mol Cell Biol 2008;9:297–308. 21. Dhananjayan SC, Ismail A, Nawaz Z. Ubiquitin and control of transcription. Essays Biochem 2005;41:69–80. 22. Mittenberg AG, Moiseeva TN, Barlev NA. Role of proteasomes in transcription and their regulation by covalent modifications. Front Biosci 2008;13:7184–92. 23. Jensen EV, Suzuki T, Numata M, Smith S, DeSombre ER. Estrogen‐binding substances of target tissues. Steroids 1969;13:417–27. 24. Horwitz KB, McGuire WL. Actinomycin D prevents nuclear processing of estrogen receptor. J Biol Chem 1978;253:6913–22. 25. Mockus MB, Horwitz KB. Progesterone receptors in human breast cancer. Stoichiometric translocation and nuclear receptor processing. J Biol Chem 1983;258:4778–83. 26. Nawaz Z, Lonard DM, Dennis AP, Smith CL, O’Malley BW. Proteasome‐dependent degradation of the human estrogen receptor. Proc Natl Acad Sci USA 1999;96:1858–62. 27. Lonard DM, Nawaz Z, Smith CL, O’Malley BW. The 26S proteasome is required for estrogen receptor‐alpha and coactivator turnover and for efficient estrogen receptor‐a transactivation. Mol Cell 2000;5:939–48. 28. Dace A, Zhao L, Park KS, Furuno T, Takamura N, Nakanishi M, et al. Hormone binding induces rapid proteasome‐mediated degradation of thyroid hormone receptors. Proc Natl Acad Sci USA 2000;97:8985–90.

EMERGING ROLES OF THE UBIQUITIN

131

29. Yoshida H, Kitamura K, Tanaka K, Omura S, Miyazaki T, Hachiya T, et al. Accelerated degradation of PML‐retinoic acid receptor a (PML‐RARa) oncoprotein by all‐trans‐retinoic acid in acute promyelocytic leukemia: possible role of the proteasome pathway. Cancer Res 1996;56:2945–8. 30. Zhu J, Gianni M, Kopf E, Honore´ N, Chelbi‐Alix M, Koken M, et al. Retinoic acid induces proteasome‐dependent degradation of retinoic acid receptor a (RARa) and oncogenic RARa fusion proteins. Proc Natl Acad Sci USA 1999;96:14807–12. 31. Kopf E, Plassat JL, Vivat V, de The´ H, Chambon P, Rochette‐Egly C. Dimerization with retinoid X receptors and phosphorylation modulate the retinoic acid‐induced degradation of retinoic acid receptors a and g through the ubiquitin‐proteasome pathway. J Biol Chem 2000;275:33280–8. 32. Wallace AD, Cidlowski JA. Proteasome‐mediated glucocorticoid receptor degradation restricts transcriptional signaling by glucocorticoids. J Biol Chem 2001;276:42714–21. 33. Schaaf MJ, Cidlowski JA. Molecular determinants of glucocorticoid receptor mobility in living cells: the importance of ligand affinity. Mol Cell Biol 2003;23:1922–34. 34. Yokota K, Shibata H, Kobayashi S, Suda N, Murai A, Kurihara I, et al. Proteasome‐mediated mineralocorticoid receptor degradation attenuates transcriptional response to aldosterone. Endocr Res 2004;30:611–6. 35. Tirard M, Almeida OF, Hutzler P, Melchior F, Michaelidis TM. Sumoylation and proteasomal activity determine the transactivation properties of the mineralocorticoid receptor. Mol Cell Endocrinol 2007;268:20–9. 36. Wiese RJ, Uhland‐Smith A, Ross TK, Prahl JM, DeLuca HF. Up‐regulation of the vitamin D receptor in response to 1,25‐dihydroxyvitamin D3 results from ligandinduced stabilization. J Biol Chem 1992;267:20082–6. 37. Masuyama H, MacDonald PN. Proteasome‐mediated degradation of the vitamin D receptor (VDR) and a putative role for SUG1 interaction with the AF‐2 domain of VDR. J Cell Biochem 1998;71:429–40. 38. Kemppainen JA, Lane MV, Sar M, Wilson EM. Androgen receptor phosphorylation, turnover, nuclear transport, transcriptional activation. Specificity for steroids and antihormones. J Biol Chem 1992;267:968–74. 39. Zhou Z‐X, Lane MV, Kemppainen JA, French FS, Wilson EM. Specificity of ligand‐ dependent androgen receptor stabilization: receptor domain interactions influence ligand dissociation and receptor stability. Mol Endocrinol 1995;9:208–18. 40. Reid G, Hu¨bner MR, Me´tivier R, Brand H, Denger S, Manu D, et al. Cyclic, proteasome‐ mediated turnover of unliganded and liganded ERa on responsive promoters is an integral feature of estrogen signaling. Mol Cell 2003;11:695–707. 41. Kiang DT, Killander RE, Thomas T, Kennedy BJ. Up‐regulation of estrogen receptors by nonsteroidal antiestrogens in human breast cancer. Cancer Res 1989;49:5312–6. 42. Wittmann BM, Sherk A, McDonnell DP. Definition of functionally important mechanistic differences among selective estrogen receptor down‐regulators. Cancer Res 2007;67:9549–60. 43. Dauvois S, Danielian PS, White R, Parker MG. Antiestrogen ICI 164,384 reduces cellular estrogen receptor content by increasing its turnover. Proc Natl Acad Sci USA 1992;89:4037–41. 44. Wijayaratne AL, Nagel SC, Paige LA, Christensen DJ, Norris JD, Fowlkes DM, McDonnell DP. Comparative analyses of mechanistic differences among antiestrogens. Endocrinology 1999;140:5828–40. 45. Fan M, Nakshatri H, Nephew KP. Inhibiting proteasomal proteolysis sustains estrogen receptor‐alpha activation. Mol Endocrinol 2004;18:2603–15. 46. Tsai HW, Katzenellenbogen JA, Katzenellenbogen BS, Shupnik MA. Protein kinase A activation of estrogen receptor a transcription does not require proteasome activity and protects the receptor from ligand‐mediated degradation. Endocrinology 2004;145:2730–8.

132

LONARD AND O’MALLEY

47. Kinyamu HK, Collins JB, Grissom SF, Hebbar PB, Archer TK. Genome wide transcriptional profiling in breast cancer cells reveals distinct changes in hormone receptor target genes and chromatin modifying enzymes after proteasome inhibition. Mol Carcinog 2008;47:845–85. 48. Dennis AP, Lonard DM, Nawaz Z, O’Malley BW. Inhibition of the 26S proteasome blocks progesterone receptor‐dependent transcription through failed recruitment of RNA polymerase II. J Steroid Biochem Mol Biol 2005;94:337–46. 49. Sheflin L, Keegan B, Zhang W, Spaulding SW. Inhibiting proteasomes in human HepG2 and LNCaP cells increases endogenous androgen receptor levels. Biochem Biophys Res Commun 2000;276:144–50. 50. Lin HK, Altuwaijri S, Lin WJ, Kan PY, Collins LL, Chang C. Proteasome activity is required for androgen receptor transcriptional activity via regulation of androgen receptor nuclear translocation and interaction with coregulators in prostate cancer cells. J Biol Chem 2002;277:36570–6. 51. Poole AJ, Li Y, Kim Y, Lin SC, Lee WH, Lee EY. Prevention of Brca1‐mediated mammary tumorigenesis in mice by a progesterone antagonist. Science 2006;314:1467–70. 52. Chen PH, Tsao YP, Wang CC, Chen SL. Nuclear receptor interaction protein, a coactivator of androgen receptors (AR), is regulated by AR and Sp1 to feed forward and activate its own gene expression through AR protein stability. Nucleic Acids Res 2008;36:51–66. 53. Kang Z, Pirskanen A, Ja¨nne OA, Palvimo JJ. Involvement of proteasome in the dynamic assembly of the androgen receptor transcription complex. J Biol Chem 2002;277:48366–71. 54. Stenoien DL, Cummings CJ, Adams HP, Mancini MG, Patel K, DeMartino GN, et al. Polyglutamine‐expanded androgen receptors form aggregates that sequester heat shock proteins, proteasome components and SRC‐1, and are suppressed by the HDJ‐2 chaperone. Hum Mol Genet 1999;8:731–41. 55. Lin HK, Altuwaijri S, Lin WJ, Kan PY, Collins LL, Chang C. Proteasome activity is required for androgen receptor transcriptional activity via regulation of androgen receptor nuclear translocation and interaction with coregulators in prostate cancer cells. J Biol Chem 2002;277:36570–6. 56. Palazzolo I, Gliozzi A, Rusmini P, Sau D, Crippa V, Simonini F, et al. The role of the polyglutamine tract in androgen receptor. J Steroid Biochem Mol Biol 2008;108:245–53. 57. Chandra S, Shao J, Li JX, Li M, Longo FM, Diamond MI. A common motif targets huntingtin and the androgen receptor to the proteasome. J Biol Chem 2008;283:23950–5. 58. Palazzolo I, Gliozzi A, Rusmini P, Sau D, Crippa V, Simonini F, et al. The role of the polyglutamine tract in androgen receptor. J Steroid Biochem Mol Biol 2008;108:245–53. 59. Chandra S, Shao J, Li JX, Li M, Longo FM, Diamond MI. A common motif targets huntingtin and the androgen receptor to the proteasome. J Biol Chem 2008;283:23950–5. 60. Fan M, Park A, Nephew KP. CHIP (carboxyl terminus of Hsc70‐interacting protein) promotes basal and geldanamycin‐induced degradation of estrogen receptor‐a. Mol Endocrinol 2005;19:2901–14. 61. Tateishi Y, Kawabe Y, Chiba T, Murata S, Ichikawa K, Murayama A, et al. Ligand‐dependent switching of ubquitin‐proteasome pathways for estrogen receptor. EMBO J 2004;23:4813–23. 62. Horie K, Urano T, Ikeda K, Inoue S. Estrogen‐responsive RING finger protein controls breast cancer growth. J Steroid Biochem Mol Biol 2003;85:101–4. 63. Park S, Song J, Joe CO, Shin I. Akt stabilizes estrogen receptor alpha with the concomitant reduction in its transcriptional activity. Cell Signal 2008;20:1368–74. 64. Lange CA, Shen T, Horwitz KB. Phosphorylation of human progesterone receptors at serine‐ 294 by mitogen‐activated protein kinase signals their degradation by the 26S proteasome. Proc Natl Acad Sci USA 2000;97:1032–7.

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65. Chauchereau A, Amazit L, Quesne M, Guiochon‐Mantel A, Milgrom E. Sumoylation of the progesterone receptor and of the steroid receptor coactivator SRC‐1. J Biol Chem 2003;278:12335–43. 66. Daniel AR, Faivre EJ, Lange CA. Phosphorylation‐dependent antagonism of sumoylation derepresses progesterone receptor action in breast cancer cells. Mol Endocrinol 2007;21:2890–906. 67. Zhang PJ, Zhao J, Li HY, Man JH, He K, Zhou T, et al. CUE domain containing 2 regulates degradation of progesterone receptor by ubiquitin‐proteasome. EMBO J 2007;26:1831–42. 68. Su K, Yang X, Roos MD, Paterson AJ, Kudlow JE. Human Sug1/p45 is involved in the proteasome‐dependent degradation of Sp1. Biochem J 2000;348:281–9. 69. Masuyama H, Hiramatsu Y. Involvement of suppressor for Gal 1 in the ubiquitin/proteasome‐ mediated degradation of estrogen receptors. J Biol Chem 2004;279:12020–6. 70. Ghislain M, Udvardy A, Mann C. S. cerevisiae 26S protease mutants arrest cell division in G2/ metaphase. Nature 1993;366:358–62. 71. Wo´jcik C, DeMartino GN. Intracellular localization of proteasomes. Int J Biochem Cell Biol 2003;35:579–89. 72. Nawaz Z, Lonard DM, Smith CL, Lev‐Lehman E, Tsai SY, Tsai MJ, O’Malley BW. The Angelman syndrome‐associated protein, E6‐AP, is a coactivator for the nuclear hormone receptor superfamily. Mol Cell Biol 1999;19:1182–9. 73. Jiang YH, Beaudet AL. Human disorders of ubiquitination and proteasomal degradation. Curr Opin Pediatr 2004;16:419–26. 74. Matentzoglu K, Scheffner M. Ubiquitin ligase E6‐AP and its role in human disease. Biochem Soc Trans 2008;36:797–801. 75. Picard N, Charbonneau C, Sanchez M, Licznar A, Busson M, Lazennec G, et al. Phosphorylation of activation function‐1 regulates proteasome‐dependent nuclear mobility and E6‐ associated protein ubiquitin ligase recruitment to the estrogen receptor b. Mol Endocrinol 2008;22:317–30. 76. Imhof MO, McDonnell DP. Yeast RSP5 and its human homolog hRPF1 potentiate hormone‐ dependent activation of transcription by human progesterone and glucocorticoid receptors. Mol Cell Biol 1996;16:2594–605. 77. Huibregtse JM, Yang JC, Beaudenon SL. The large subunit of RNA polymerase II is a substrate of the Rsp5 ubiquitin‐protein ligase. Proc Natl Acad Sci USA 1997;94:3656–61. 78. Yan J, Kim YS, Yang XP, Albers M, Koegl M, Jetten AM. Ubiquitin‐interaction motifs of RAP80 are critical in its regulation of estrogen receptor a. Nucleic Acids Res 2007;35:1673–86. 79. Grossman SR, Deato ME, Brignone C, Chan HM, Kung AL, Tagami H, et al. Polyubiquitination of p53 by a ubiquitin ligase activity of p300. Science 2003;300:342–4. 80. Hoppe T. Multiubiquitylation by E4 enzymes: ‘‘one size’’ doesn’t fit all. Trends Biochem Sci 2005;30:183–7. 81. Nijman SM, Luna‐Vargas MP, Velds A, Brummelkamp TR, Dirac AM, Sixma TK, et al. A genomic and functional inventory of deubiquitinating enzymes. Cell 2005;123:773–86. 82. Faus H, Meyer HA, Huber M, Bahr I, Haendler B. The ubiquitin‐specific protease USP10 modulates androgen receptor function. Mol Cell Endocrinol 2005;245:138–46. 83. Zhao Y, Lang G, Ito S, Bonnet J, Metzger E, Sawatsubashi S, et al. A TFTC/STAGA module mediates histone H2A and H2B deubiquitination, coactivates nuclear receptors, and counteracts heterochromatin silencing. Mol Cell 2008;29:92–101. 84. Zhu P, Zhou W, Wang J, Puc J, Ohgi KA, Erdjument‐Bromage H, et al. A histone H2A deubiquitinase complex coordinating histone acetylation and H1 dissociation in transcriptional regulation. Mol Cell 2007;27:609–21. 85. Zhang J, Guenther MG, Carthew RW, Lazar MA. Proteasomal regulation of nuclear receptor corepressor‐mediated repression. Genes Dev 1998;12:1775–80.

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86. Perissi V, Aggarwal A, Glass CK, Rose DW, Rosenfeld MG. A corepressor/coactivator exchange complex required for transcriptional activation by nuclear receptors and other regulated transcription factors. Cell 2004;116:511–26. 87. Perissi V, Scafoglio C, Zhang J, Ohgi KA, Rose DW, Glass CK, et al. TBL1 and TBLR1 phosphorylation on regulated gene promoters overcomes dual CtBP and NCoR/SMRT transcriptional repression checkpoints. Mol Cell 2008;29:755–66. 88. Li X, Lonard DM, Jung SY, Malovannaya A, Feng Q, Qin J, et al. The SRC‐3/AIB1 coactivator is degraded in a ubiquitin‐ and ATP‐independent manner by the REGg proteasome. Cell 2006;124:381–92. 89. Wu RC, Feng Q, Lonard DM, O’Malley BW. SRC‐3 coactivator functional lifetime is regulated by a phospho‐dependent ubiquitin time clock. Cell 2007;129:1125–40. 90. Mori Y, Moriishi K, Matsuura Y. Hepatitis C virus core protein: its coordinate roles with PA28gamma in metabolic abnormality and carcinogenicity in the liver. Int J Biochem Cell Biol 2008;40:1437–42. 91. Li X, Amazit L, Long W, Lonard DM, Monaco JJ, O’Malley BW. Ubiquitin‐ and ATP‐ independent proteolytic turnover of p21 by the REGg‐proteasome pathway. Mol Cell 2007;26:831–42. 92. Wu RC, Qin J, Yi P, Wong J, Tsai SY, Tsai MJ, et al. Selective phosphorylations of the SRC‐3/ AIB1 coactivator integrate genomic reponses to multiple cellular signaling pathways. Mol Cell 2004;15:937–49. 93. Ferrero M, Avivar A, Garcı´a‐Macı´as MC, de Mora JF. Phosphoinositide 3‐kinase/AKT signaling can promote AIB1 stability independently of GSK3 phosphorylation. Cancer Res 2008;68:5450–9. 94. Lonard DM, O’Malley BW. SRC‐3 transcription‐coupled activation, degradation, and the ubiquitin clock: is there enough coactivator to go around in cells. Sci Signal 2008;1:pe16. 95. Yi P, Feng Q, Amazit L, Lonard DM, Tsai SY, Tsai MJ, et al. Atypical protein kinase C regulates dual pathways for degradation of the oncogenic coactivator SRC‐3/AIB1. Mol Cell 2008;29:465–76. 96. Li C, Liang YY, Feng XH, Tsai SY, Tsai MJ, O’Malley BW. Essential phosphatases and a phospho‐degron are critical for regulation of SRC‐3/AIB1 coactivator function and turnover. Mol Cell 2008;31:835–49. 97. Yi P, Wu RC, Sandquist J, Wong J, Tsai SY, Tsai MJ, et al. Peptidyl‐prolyl isomerase 1 (Pin1) serves as a coactivator of steroid receptor by regulating the activity of phosphorylated steroid receptor coactivator 3(SRC‐3/AIB1). Mol Cell Biol 2005;25:9687–99. 98. Mani A, Oh AS, Bowden ET, Lahusen T, Lorick KL, Weissman AM, et al. E6AP mediates regulated proteasomal degradation of the nuclear receptor coactivator amplified in breast cancer 1 in immortalized cells. Cancer Res 2006;66:8680–6. 99. Li C, Wu RC, Amazit L, Tsai SY, Tsai MJ, O’Malley BW. Specific amino acid residues in the basic helix‐loop‐helix domain of SRC‐3 are essential for its nuclear localization and proteasome‐dependent turnover. Mol Cell Biol 2007;27:1296–308. 100. Baumann CT, Ma H, Wolford R, Reyes JC, Maruvada P, Lim C, et al. The glucocorticoid receptor interacting protein 1 (GRIP1) localizes in discrete nuclear foci that associate with ND10 bodies and are enriched in components of the 26S proteasome. Mol Endocrinol 2001;15:485–500. 101. Fenne I.S, Hoang T, Hauglid M, Sagen JV, Lien EA, Mellgren G. Recruitment of coactivator glucocorticoid receptor interacting protein 1 to an estrogen receptor transcription complex is regulated by the 30 ,50 ‐cyclic adenosine 50 ‐monophosphate‐dependent protein kinase. Endocrinology 2008;149:4336–45.

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102. Hoang T, Fenne S, Cook C, Børud B, Bakke M, Lien EA, et al. cAMP‐dependent protein kinase regulates ubiquitin‐proteasome‐mediated degradation and subcellular localization of the nuclear receptor coactivator GRIP1. J Biol Chem 2004;279:49120–30. 103. Linares LK, Kiernan R, Triboulet R, Chable‐Bessia C, Latreille D, Cuvier O, et al. Intrinsic ubiquitination activity of PCAF controls the stability of the oncoprotein Hdm2. Nat Cell Biol 2007;9:331–8. 104. Brooks CL, Gu W. Ubiquitination, phosphorylation and acetylation: the molecular basis for p53 regulation. Curr Opin Cell Biol 2003;15:164–71. 105. Olson BL, Hock MB, Ekholm‐Reed S, Wohlschlegel JA, Dev KK, Kralli A, et al. SCFCdc4 acts antagonistically to the PGC‐1a transcriptional coactivator by targeting it for ubiquitin‐ mediated proteolysis. Genes Dev 2008;22:252–64. 106. Rasbach KA, Green PT, Schnellmann RG. Oxidants and Ca2þ induce PGC‐1a degradation through calpain. Arch Biochem Biophys 2008;478:130–5. 107. Kerscher O, Felberbaum R, Hochstrasser M. Modification of proteins by ubiquitin and ubiquitin‐like proteins. Annu Rev Cell Dev Biol 2006;22:159–80. 108. Poukka H, Karvonen U, Janne OA, Palvimo JJ. Covalent modification of the androgen receptor by small ubiquitin‐like modifier (SUMO‐1). Proc Natl Acad Sci USA 2000;97: 14145–50. 109. Poukka H, Aarnisalo P, Karvonen U, Palvimo JJ, Ja¨nne OA. Ubc9 interacts with the androgen receptor and activates receptor‐dependent transcription. J Biol Chem 1999;274:19441–6. 110. Anckar J, Sistonen L. SUMO: getting it on. Biochem Soc Trans 2007;35:1409–13. 111. Liu B, Shuai K. Regulation of the sumoylation system in gene expression. Curr Opin Cell Biol 2008;20:288–93. 112. Kotaja N, Karvonen U, Ja¨nne OA, Palvimo JJ. The nuclear receptor interaction domain of GRIP1 is modulated by covalent attachment of SUMO‐1. J Biol Chem 2002;277:30283–8. 113. Bawa‐Khalfe T, Cheng J, Wang Z, Yeh ET. Induction of the SUMO‐specific protease 1 transcription by the androgen receptor in prostate cancer cells. J Biol Chem 2007;282: 37341–9. 114. Cheng J, Bawa T, Lee P, Gong L, Yeh ET. Role of desumoylation in the development of prostate cancer. Neoplasia 2006;8:667–76. 115. Sentis S, Le Romancer M, Bianchin C, Rostan MC, Corbo L. Sumoylation of the estrogen receptor alpha hinge region regulates its transcriptional activity. Mol Endocrinol 2005;19:2671–84. 116. Wu H, Sun L, Zhang Y, Chen Y, Shi B, Li R, et al. Coordinated regulation of AIB1 transcriptional activity by sumoylation and phosphorylation. J Biol Chem 2006;281:21848–56. 117. Rytinki MM, Palvimo JJ. SUMOylation modulates the transcription repressor function of RIP140. J Biol Chem 2008;283:11586–95. 118. Fan M, Long X, Bailey JA, Reed CA, Osborne E, Gize EA, et al. The activating enzyme of NEDD8 inhibits steroid receptor function. Mol Endocrinol 2002;16:315–30. 119. Fan M, Bigsby RM, Nephew KP. The NEDD8 pathway is required for proteasome‐mediated degradation of human estrogen receptor (ER)‐a and essential for the antiproliferative activity of ICI 182,780 in ERa‐positive breast cancer cells. Mol Endocrinol 2003;17:356–65. 120. Parry G, Estelle M. Regulation of cullin‐based ubiquitin ligases by the Nedd8/RUB ubiquitin‐ like proteins. Semin Cell Dev Biol 2004;15:221–9.

Biochemical Analyses of Nuclear Receptor‐Dependent Transcription with Chromatin Templates Donald D. Ruhl* and W. Lee Kraus*,{ *Department of Molecular Biology and Genetics, Cornell University, Ithaca, New York 14853 {

Department of Pharmacology, Weill Medical College of Cornell University, New York, New York 10021

I. Nuclear Receptors (NRs): Transcription Factors (TFs) with Separable Biochemical Activities................................................................... II. Biochemical Analyses of NR Activities, Interactions, and Functions: An Historical View ....................................................................... A. Elucidation of NR Biochemical Activities...................................... B. NRs and the RNA Polymerase II Transcription Machinery ............... C. NR Coregulators..................................................................... D. Transcriptional Regulation by NRs: Early Studies ........................... III. Role of Chromatin in NR‐Dependent Transcription ............................. A. Chromatin: The Physiological Template for NR‐Dependent Transcription ......................................................................... B. Linking NR Function to Chromatin............................................. IV. Biochemical Methods for the Analysis of NR‐Dependent Transcription .... A. Components of Activator‐Dependent In Vitro Transcription Systems .. B. DNA Templates ...................................................................... C. Transcription Machinery........................................................... D. Detection of Transcripts ........................................................... E. Purification of NR and Coregulator Proteins.................................. V. Biochemical Methods for the Assembly and Analysis of Chromatin.......... A. DNA Templates ...................................................................... B. Histones ............................................................................... C. Methods for Assembling Chromatin In Vitro ................................. D. Methods for Analyzing Chromatin Assembled In Vitro..................... E. Putting It All Together: In Vitro Transcription with Chromatin Templates............................................................... VI. What have We Learned About NR‐Dependent Transcription from In Vitro Chromatin Assembly and Transcription Studies? ................................. A. Role of Ligands ...................................................................... B. Role of Basal TFs.................................................................... C. Role of Coactivators................................................................. D. Role of Corepressors................................................................

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E. Role of Nucleosome‐Binding Proteins.......................................... F. Order and Dynamics ............................................................... VII. Future Directions ........................................................................ A. Repression of Transcription by NRs............................................. B. Single Molecule Studies ........................................................... C. NR Biochemistry in the Postgenomic Era ..................................... VIII. Summary ................................................................................... References.................................................................................

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Chromatin, the physiological template for transcription, plays important roles in gene regulation by nuclear receptors (NRs). It can (1) restrict the binding of NRs or the transcriptional machinery to their genomic targets, (2) serve as a target of regulatory posttranslational modifications by NR coregulator proteins with histone‐directed enzymatic activities, and (3) function as a binding scaffold for a variety of transcription‐related proteins. The advent of in vitro or ‘‘cell‐free’’ systems that accurately recapitulate ligand‐dependent transcription by NRs with chromatin templates has allowed detailed analyses of these processes. Biochemical studies have advanced our understanding of the mechanisms of gene regulation, including the role of ligands, coregulators, and nucleosome remodeling. In addition, they have provided new insights about the dynamics of NR‐mediated transcription. This chapter reviews the current methodologies for assembling, transcribing, and analyzing chromatin in vitro, as well as the new information that has been gained from these studies.

I. Nuclear Receptors (NRs): Transcription Factors (TFs) with Separable Biochemical Activities NRs comprise a large, evolutionarily conserved family of DNA‐binding TFs that play key roles in a wide variety of physiological and disease processes.1,2 NRs share a conserved structural and functional organization, including a DNA‐binding domain (DBD) located centrally or at the amino‐terminus and a ligand‐binding domain (LBD) located at the carboxyl‐terminus (Fig. 1). These domains confer a number of distinct biochemical activities on NRs,

N

AF-1

DBD

A/B

C

LBD/AF-2 D

E

F C

FIG. 1. General structural and functional organization of NRs. Schematic representation of the general structural and functional organization of NRs, including an amino‐terminal activation function‐1 (AF‐1), a centrally located DNA-binding domain (DBD), a carboxyl‐terminal ligand‐ binding domain (LBD), and a carboxyl‐terminal AF‐2.

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including (1) ligand‐binding, (2) DNA‐binding, (3) homo or heterodimerization, (4) protein–protein interactions, and (5) transcriptional activation, with the latter generally dependent on the first four. The modular nature of NRs has made it possible to study these biochemical activities in isolation. A complete understanding of NR function, however, requires an in‐depth understanding of how these biochemical activities function together to regulate transcriptional outcomes.

II. Biochemical Analyses of NR Activities, Interactions, and Functions: An Historical View A. Elucidation of NR Biochemical Activities The concept of a steroid hormone receptor was first suggested in the 1960s by Jensen and colleagues, a scientific breakthrough made possible by the development of high specific activity radiolabeled ligands for the estrogen receptor (ER).3 This discovery led to the purification of steroid hormone receptors and an initial characterization of their biochemical activities over the next two decades. These early studies focused on (1) the ligand‐binding properties of the receptors, (2) their subcellular localization, and (3) their ligand‐regulated association with protein chaperones, including heat shock proteins.3,4 The cloning of the glucocorticoid receptor (GR) cDNA, the first cDNA encoding an NR, in 19855,6 led to a new wave of analysis of NR function, including identification and characterization of (1) the conserved domain structure of NRs,7 (2) the DNA‐binding properties of NRs,8–11 (3) the transactivation functions (AFs) of NRs,12 (4) the dimerization properties of NRs,13,14 and (5) the NR superfamily and its evolutionary relationships.15 The subsequent purification and crystallization of isolated NR domains16,17 and, more recently, full length NRs18 have provided structural insights to support our understanding of the biochemical functions of NRs.

B. NRs and the RNA Polymerase II Transcription Machinery The identification and characterization of NR‐interacting proteins has been a key focus area over the past two decades. These proteins include components of the RNA polymerase II (Pol II) transcription machinery, as well as a large, diverse group of proteins called ‘‘coregulators’’ (Fig. 2). The initiation of transcription (i.e., mRNA synthesis) by Pol II involves the direct or indirect binding of core promoter DNA elements by a collection of ‘‘basal’’ TFs (including TFIIA, IIB, IID, IIE, IIF, IIH, and IIS).19,20 Some of the first transcriptionally relevant interactions characterized for NRs were with basal TFs.21–24

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Histone modifying enzymes

Mediator

p300/CBP

TRAP/DRIP Med1

SRC + AcCoA Ac

RNA Pol ll Basal TFs

NR NR HRE

Basal machinery Ac NCoR

Brg1

HDAC

Swi/Snf

Histone demodifying enzymes

Chromatin remodeling complexes

+ ATP

FIG. 2. A diverse array of coregulators interact physically and functionally with NRs to control hormone‐dependent gene regulation. Four classes of coregulators (histone‐modifying enzymes, HMEs; histone‐demodifying enzymes, HDEs; Mediator, and chromatin remodeling complexes, CRCs), as well as the basal transcription machinery, are shown. Key proteins within the complexes, such as SRC, NCoR, Brg1, Med1, and the basal transcription factors (TFs; e.g., TBP and TFIIB), make direct contacts with the NRs. Ac, acetylation; AcCoA, acetyl‐CoA; HRE, hormone response element.

TFIID, a complex of proteins containing the TATA‐binding protein (TBP) and a collection of 10–12 polypeptides called TBP‐associated factors (TAFs),25 is one basal factor that plays a key role in directing the binding of Pol II to many promoters. The TAFs in the TFIID complex are required for transcriptional activation by a number of different DNA‐binding activators, including NRs.23,25 Several TAFs in TFIID, as well as TBP itself, make direct contacts with NRs during the ligand‐regulated transcription process.23,26 Pol II and the basal TFs may be recruited to promoters after a stimulus (e.g., hormone treatment) or they may ‘‘preload’’ at the promoter in a poised or paused state in anticipation of a stimulus.27–29 In either case, the binding of ligand‐activated NRs to DNA response elements (‘‘enhancers’’) in the promoter or regulatory regions of a hormone‐responsive gene provides the trigger that promotes transcription initiation and/or elongation.

C. NR Coregulators Knowledge of the biochemical properties NRs and their roles in gene regulation, as well as an emerging understanding of the components and activities of the Pol II transcription machinery, led to the hypothesis in the

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early 1990s that transcriptional regulation by NR would require a set of receptor‐interacting ‘‘transcriptional intermediary factors (TIFs)’’ (now more commonly called ‘‘coregulators’’) distinct from the basal transcription machinery.30 In vitro ‘‘squelching’’ and complementation experiments with activation function‐1 (AF‐1) of progesterone receptor (PR),31 in vitro binding assays with the LBD of ERa,32 and yeast‐based genetic and biochemical experiments33 provided the first evidence for receptor‐interacting coregulators. Numerous subsequent studies, which continue today, led to the identification and characterization of many NR coregulators, some that enhance NR‐dependent transcription (‘‘coactivators,’’ such as the p160/steroid receptor coactivator (SRC) family of proteins) and some that inhibit NR‐dependent transcription (‘‘corepressors,’’ such as the NR corepressor (NCoR)/silencing mediator for retinoid and thyroid hormone receptors (SMRT) proteins).26,34,35 The main classes of coregulators are (1) bridging factors, (2) histone‐ modifying (and factor modifying) enzymes (HMEs), (3) histone‐demodifying (and factor demodifying) enzymes (HDEs), (4) chromatin remodeling complexes (CRCs), and (5) Mediator complexes35,36 (Fig. 2). Bridging factors, such as SRCs and NCoR/SMRT, interact directly with DNA‐bound NRs and recruit HMEs, such as p300 and HDEs, such as histone deacetylases (HDACs), to target promoters, respectively.37,38 The covalent modification of nucleosomal histones and TFs by HMEs (e.g., acetylation by p300) alters the chromatin environment and TF activity, respectively, to modulate the transcription process.35 CRCs, such as Swi/Snf, are ATPase‐containing protein complexes that use the energy stored in ATP to mobilize nucleosomes. Like HMEs, CRCs also alter the chromatin environment to allow the recruitment of the basal transcription machinery, other coregulators, and Pol II.39 Mediator complexes interact with NRs and Pol II, helping to promote the receptor‐dependent assembly of transcription initiation complexes at target promoters.40,41 Some general conclusions about NR coregulators can be drawn from the existing wealth of information in the literature. These include the following: (1) NR coregulators interact directly or indirectly with NRs, and these interactions are often regulated by ligands. (2) Coregulators may interact directly or indirectly with components of the Pol II transcription machinery. (3) Coregulators generally do not possess DNA-binding activity and have little or no transcriptional activity in the absence of a DNA‐bound NR. (4) Coregulators may possess enzymatic activities, including kinase, acetyltransferase, methyltransferase, ATPase, and ubiquitin ligase activities. (5) Multiple coregulators with distinct activities may be found together in high molecular weight complexes.

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(6) Coregulators may be shared by other classes of DNA-binding transcriptional activators. The sum of the actions of the NR coregulators at a given target promoter plays a key role in determining the receptor‐dependent transcriptional outcome.

D. Transcriptional Regulation by NRs: Early Studies Once the biochemical activities of NRs were elucidated and many of the key molecular interactions involving NRs were defined, the attention of the field turned toward the specific contributions of NRs to the transcription process. These studies, many of which were performed during the late‐1980s to mid‐1990s, focused on the requirement for different NR biochemical activities (e.g., ligand‐binding, dimerization, DNA‐binding, interactions with the basal transcription machinery) at specific steps during the transcription process.1,23,24 The methodologies used for these studies included in vitro (or ‘‘cell‐ free’’) transcription systems, as well as cell‐based transient transfection reporter gene assays. Although they provided a wealth of information about the molecular biology of transcriptional regulation by NRs, these approaches were limited in a number of important respects, particularly with respect to the role of chromatin in NR‐dependent transcription (discussed in more detail in the following section). In this regard, the early in vitro studies of NR‐ dependent transcription were generally carried out with ‘‘nonchromatin’’ or ‘‘naked’’ DNA templates. Furthermore, although nonreplicating transiently transfected DNA templates assemble to some degree into nucleosome‐like structures, key functional differences exist between the chromatin structure of these DNA templates and native genes.42 A closer look at the early in vitro studies of NR‐dependent transcription reveals some of the limitations that were encountered. Although the amount of transcriptional activation in these naked DNA experiments was found to correlate with the binding of the NRs to the DNA templates, the role of the ligand or receptor was not always clear or consistent. For example, in some studies, the NR bound constitutively to DNA and ligand‐independent transcriptional activation was observed.31,43–48 In other studies, ligand‐dependent transcriptional activation was observed upon the ligand‐dependent binding of the receptor to DNA.49–51 These latter studies provided more information about the DNA‐binding activities of the receptors than they did about the role of the receptors in transcriptional regulation. In one early study of ERa‐dependent transcription that used reconstituted chromatin templates, the role of ligand was not examined.52 With the advent of biochemical systems for the convenient assembly of physiologically spaced and dynamic nucleosomal arrays,53–55 reconstitution of a robust and fully ligand‐dependent in vitro

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transcription system for NRs was finally possible.56 The following sections describe how in vitro chromatin assembly and transcription systems have been a useful tool for studying the role of chromatin in NR‐dependent transcription.

III. Role of Chromatin in NR‐Dependent Transcription A. Chromatin: The Physiological Template for NR‐Dependent Transcription Chromatin is the physiological template for nuclear processes involving genomic DNA, including transcription, replication, and repair.57 The repeating unit of chromatin, the nucleosome, is a protein‐DNA structure comprising two copies each of four core histones (H2A, H2B, H3, and H4) forming a histone octamer around which 1.7 turns of genomic DNA are wrapped58 (Fig. 3). The electrostatic attraction between the highly positively charged histone core and the highly negatively charged DNA ensures a stable, yet accessible, structure. Chromatin is a dynamic polymer whose biochemical and biophysical properties play important and specific roles in determining the structure and function of A

Histone N-terminal tails

Dyad axis

5.6 nm Linker DNA

11 nm B 2x

+

+ H2A/H2B

(H3/H4)2

146 bp of DNA

FIG. 3. Anatomy of a nucleosome. (A) The DNA double helix wraps 1.7 times around an octamer of core histones. The eight amino‐terminal tails of the nucleosomal histones are unstructured, but contain the bulk of the known targets for histone modifications. The dimensions of the nucleosome and the dyad axis are indicated. (B) The nucleosome is composed of two heterodimers of H2A/H2B, a heterotetramer of H3 and H4 [(H3/H4)2] and 146 bp of DNA.

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chromatin in vivo.59,60 The dynamic properties of chromatin include (1) structural alterations within a nucleosome (e.g., removal of H2A/H2B dimers), (2) mobilization of nucleosomes along a length of DNA (e.g., translational repositioning), and (3) internucleosome interactions leading to the compaction of nucleosomes into higher order structures (e.g., chromatin condensation)59,60 (Fig. 4). The biochemical and biophysical properties of chromatin are determined by the protein composition of the nucleosomal protein core (e.g., canonical vs. variant histones) and association with nucleosome‐binding proteins (e.g., linker histones).61,62

B. Linking NR Function to Chromatin A number of cell‐based studies performed in the mid‐1980s to the mid‐1990s established a clear connection between NR function and chromatin. The first studies linking NR function to specific changes in chromatin structure showed that steroid hormones, such as glucocorticoids and progestins, can induce alterations in DNA cleavage by nucleases (e.g., DNaseI or MNase) or chemical agents (e.g., methidiumpropyl‐EDTA‐Fe(II)).63–65 Together, these studies showed that movement or specific positioning of nucleosomes at hormone‐responsive promoters and enhancers occurs upon hormone‐dependent binding of the receptor. The mouse mammary tumor virus (MMTV) long terminal repeat (LTR), a DNA sequence containing a glucocorticoid‐ and progestin‐inducible promoter with binding sites for the cognate NRs, GR and PR respectively, was a particularly useful model for this type of study.66,67 Subsequent ‘‘template comparison’’ assays, in which GR‐ and PR‐dependent transcription from genomically integrated versus transiently transfected MMTV LTR was monitored, highlighted the importance of chromatin context for proper ligand‐regulated transcriptional outcomes.68,69 Similar results were observed using a Xenopus oocyte

(1) (3)

(2)

FIG. 4. Structural alterations of chromatin. DNA is a dynamic molecule that can be structurally altered through (1) the removal of H2A/H2B dimers, (2) mobilization of nucleosomes along a length of DNA, and (3) internucleosome interactions leading to the compaction of nucleosomes into higher order structures.

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system in which thyroid hormone receptor beta (TRb)‐dependent transcription from replicated (i.e., chromatin‐assembled) versus nonreplicated (i.e., unassembled) DNA was compared.70,71 Additional evidence connecting NR function to chromatin included the identification of components of the Swi/Snf CRC as NR‐interacting proteins and coregulators. The involvement of Swi/Snf complexes in NR‐dependent transcription was originally suggested by studies in yeast and mammalian cells which showed a stimulatory effect of Swi/Snf on NR‐dependent activity.33,72–74 Since then, additional biochemical and cell‐based approaches have supported these results, including experiments showing a requirement for Swi/Snf–receptor interactions in NR‐dependent gene regulation75–81 and chromatin immunoprecipitation (ChIP) experiments showing the ligand‐induced recruitment of Swi/Snf subunits to NR‐regulated promoters.76,82,83 These results, coupled with the general failure of in vitro transcription assays with naked DNA templates to accurately recapitulate the robustness and ligand regulation of NR‐dependent transcription, highlight the important functional connections between NRs and chromatin.

IV. Biochemical Methods for the Analysis of NR‐Dependent Transcription A. Components of Activator‐Dependent In Vitro Transcription Systems In vitro transcription systems have been an invaluable tool for studying the mechanisms of activator (or repressor)‐dependent transcriptional regulation by Pol II.84,85 All of the systems described to date share a number of common components. These include the following: (1) A DNA template containing activator (or repressor) binding sites, a promoter, and a reporter gene. The DNA template may be naked or, as described in Section V, assembled into chromatin. (2) A source of the Pol II transcription machinery, which may be a crude extract, purified native factors, or purified recombinant factors. (3) An assay to detect transcription of the reporter gene, such as primer extension or G‐less cassette. (4) A source of purified DNA‐binding activator (or repressor). In the case of NRs, this may be purified native receptor, but more typically is recombinant receptor expressed in bacteria or insect cells. (5) A source of purified coregulators, small molecule drugs, chromatin‐ binding proteins, etc. that might influence the activity of NRs.

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The combination of these components in a specific order (e.g., order‐of‐ addition experiments) and under specific conditions (e.g., single round transcription assays, template competition assays) allows for a detailed analysis of activator‐dependent transcription.84,85

B. DNA Templates For studies of transcriptional regulation by DNA‐binding activators, such as NRs, several elements are incorporated into the DNA template to be used, which is typically in the form of a highly purified closed circular supercoiled plasmid. These elements include (1) an NR-binding sequence or ‘‘response element,’’ which may be multimerized to give more robust transcriptional responses, (2) a TATA box and initiator sequence (or other basal promoter elements) to nucleate preinitiation complex (PIC) formation (e.g., those from the adenovirus E4 or major late promoters), and (3) a reporter gene to be transcribed (e.g., G‐less cassette) (Fig. 5). If primer extension or nuclease protection will be used as the detection method, no particular reporter gene sequences are required. In some cases, commonly used NR‐responsive luciferase or chloramphenicol acetyltransferase reporter plasmids used in transient transfection assays can also be used as templates for in vitro transcription. +1

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Another option is to use gene fragments containing native arrangements of response elements and a promoter that have been cloned into a plasmid vector. This alternative is useful when studying the native regulation of a selected gene of interest.

C. Transcription Machinery In vitro transcription assays require a source of the Pol II transcription machinery (i.e., Pol II and the basal factors), which may be a crude extract, purified native factors, or purified recombinant factors. Most transcription extracts used today are made using variations of the protocol described by Dignam and colleagues in 1983.86 The purification and characterization of Pol II in the mid‐1970s by Roeder and colleagues87–89 was followed by numerous studies in the 1980s and early‐1990s leading to the purification of all of the basal TFs and the cloning of their cDNAs.90 Most of these factors are now available in recombinant forms.90 The details of these systems are beyond the scope of this review and are described elsewhere.85 The choice of transcription system depends on the goal of the experiment. Minimal purified in vitro transcription systems support basal transcription, but require additional factors that allow activator‐dependent transcription, especially with chromatin templates. Such systems allow for complementation assays that can be used to identify coregulator activities. Extract‐based systems are useful when required coregulators have not yet been identified, but are present in an extract that supports transcription by a particular DNA‐binding activator protein.

D. Detection of Transcripts In vitro transcription systems require a method for the detection and quantification of transcripts, and some of these approaches are noted here. The details of these systems are beyond the scope of this chapter, but they can be found elsewhere.56,84,91,92 Common assays include (1) G‐less cassette, (2) primer extension, and (3) nuclease‐based detection (Fig. 6). Quantitative real‐time PCR may also be a useful approach that could be adapted for detection of transcripts. For the G‐less cassette assay, a DNA template containing a coding strand lacking G-residues for a stretch of 50–400 bp is used.91 In a reaction containing radiolabeled UTP or CTP, but lacking GTP, transcription proceeds until it terminates at the first G‐residue encountered. In this way, a transcript of a defined length is produced (Fig. 6A). For the primer extension assay, the RNA product is reverse transcribed using reverse transcriptase with an end‐labeled transcript‐specific primer, again yielding a product of defined length56,92 (Fig. 6B). In the nuclease‐based detection assays, such as the RNase protection or S1 nuclease protection, the RNA products are hybridized to radiolabeled complementary RNA or DNA probes. Digestion with

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FIG. 6. Methods for the detection of RNA transcripts in vitro. Shown are schematic representations of three methods used to detect RNA generated from in vitro transcription assays. (A) The G‐less cassette method directly labels RNA transcripts through the incorporation of a radiolabeled nucleotide triphosphate (NTP). The length of the transcript is dictated by the length of the G‐less cassette used. Because of the direct labeling of RNA, incomplete transcripts may also be detected on a gel. (B) The primer extension method uses an indirect measurement of the RNA transcripts through reverse transcription (RT) with a radiolabeled primer. The RT products are of uniform size due the specificity and end‐labeling of the primer. (C) Nuclease‐based detection methods use hybridization of a radiolabeled complementary RNA or DNA probe to the RNA transcripts. After annealing, digestion of single-stranded nucleic acids with specific nucleases (e.g., S1 nuclease) leaves a labeled product of uniform size. See the text for more details about these methods.

single‐strand‐specific RNase or S1 nuclease preserves the annealed hybrids, but digests the unannealed probe and transcript, leaving a product of defined length (Fig. 6C). The resulting products from all of these assays can be analyzed by standard acrylamide gel electrophoresis and quantified by phosphorimaging analysis.92

E. Purification of NR and Coregulator Proteins In vitro transcription assays with NRs and coregulators requires a source of purified proteins. Like the basal factors, these can be purified from native sources, but recombinant sources have generally proven superior with respect to ease of purification, yield, and activity. The use of recombinant proteins also

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allows the examination of mutant versions of the proteins. A variety of recombinant expression systems have been used for NRs and their coregulators, including bacteria and insect cells.56 Many NRs are fairly labile in their purified forms, free of chaperones, and they may lose ligand‐binding or DNA‐binding activity. In most cases, rapid single‐step affinity purification using epitope‐ or nickel‐binding tags is the best way to achieve purification of active proteins. Several laboratories have had good success purifying FLAG epitope‐tagged human ERa expressed in insect cells using a baculovius system. The purified protein is >90% pure and is highly active for ligand‐binding, DNA‐binding, and transactivation.93 Similar approaches can be used to purify coregulators.56,94 Prior to use in transcription assays, the biochemical activities of the purified proteins should be confirmed in appropriate ligand‐binding assays, DNA-binding assays (e.g., electrophoretic mobility shift assays), and enzyme assays (e.g., histone modification assays).

V. Biochemical Methods for the Assembly and Analysis of Chromatin A. DNA Templates DNA templates designed for in vitro transcription assays, such as those described in Section IV.B, can be used in chromatin assembly reactions in circular or linear forms without modifications (Fig. 7). In some cases, however, the addition of one or more nucleosome positioning elements (NPE) may be useful to achieve efficient assembly of physiologically spaced nucleosomes (Fig. 5). This is certainly the case with ‘‘passive’’ chromatin assembly systems, such as salt dialysis, that do not actively space the nucleosomes in a physiological manner. Although the overall contribution of DNA sequence to the positioning of nucleosomes throughout the genome is still debated, sequences have been identified that can function as strong NPEs and can position nucleosomes at specific locations along the DNA template. These include native sequences (e.g., Xenopus or sea urchin 5S rDNA sequence)95,96 or sequences isolated through in vitro selection protocols, such as SELEX (e.g., the 601 element from Widom and colleagues).97 When used in a transcription template, multiple NPEs are typically placed upstream and downstream of the promoter.

B. Histones A core histone octamer is made up of two copies each of four core histones, H2A, H2B, H3, and H4, which are required for the assembly of canonical nucleosomes (Fig. 3). Core histones can either be purified in their native form from eukaryotic cells (commonly mammalian, chicken, Drosophila, or yeast

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FIG. 7. In vitro assembly of chromatin. Short or long, linear or circular DNA templates, such as the plasmid template shown here, can be assembled into mononucleosomes or polynucleosomal arrays using a variety of chromatin assembly methodologies (e.g., salt dilution/dialysis, extract‐ based, or recombinant; see text for details). The assembled chromatin templates can be used in several biochemical assays, including in vitro transcription.

cells),98–100 or expressed in a recombinant form in bacteria.101,102 When purified carefully from either source, the core histones in the peak fractions will be present in stoichiometrically equal amounts. The equimolar ratio is important because a slight variation in the molar amount of the core histones can cause inefficient nucleosome assembly. The experimental question to be addressed will dictate the source of histones used. Native core histones purified from animal cells will contain a wide spectrum of posttranslational modifications (e.g., acetylation, methylation, phosphorylation), dictated by the physiology of the source cells, and will contain both canonical and variant histones. Therefore, native histones may

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not be ideal for studying specific posttranslational modifications, or perhaps even histone variants, since the histone preparations may be contaminated with them. Recombinant histones expressed in bacteria are free of either and may be a more suitable source. If the intent is to study a particular posttranslational modification, the histones can be covalently modified prior to use enzymatically (e.g., with a specific recombinant histone‐modifying enzyme103), chemically (e. g., with acetic anhydride104), or through peptide ligation, where a specifically modified histone tail is ligated to an acceptor histone globular domain.105 The use of recombinant histones with site‐specific mutations is also an effective means to study the function of histone modifications, as well as the histone‐ modifying enzymes that catalyze their deposition.106 Recombinant expression systems are also an effective means of generating histone octamers containing histone variants (e.g., H2ABbd107).

C. Methods for Assembling Chromatin In Vitro The two goals for assembling chromatin in vitro are (1) proper association of core histone proteins with DNA to form a nucleosome and (2) proper spacing of individual nucleoproteins along the length of DNA sequence to recapitulate physiological nucleosome spacing (Fig. 7). A variety of methods have been used to assemble chromatin in vitro and they vary in their ability to accomplish these two goals. Chromatin assembly systems fall into three general categories: (1) salt dilution/dialysis, (2) extract‐based, and (3) purified recombinant, which are described in the following sections. 1. SALT DILUTION/DIALYSIS CHROMATIN ASSEMBLY SYSTEMS In 1967, the selective dissociation of calf thymus histones from nucleosomes as a function of increasing salt concentration was observed.108 Subsequently, this process was found to be reversible, allowing for an in vitro method of assembling histone octamers on DNA.109,110 Briefly, histones and DNA are incubated in the presence of a high concentration (e.g., 2 M) of NaCl and are then subjected to stepwise dilution or dialysis to slowly lower the NaCl concentration to about 0.2 M. The slow dilution/dialysis allows for the spontaneous assembly of nucleosomes from the histones and DNA. A protocol detailing this procedure is nicely described by Dyer et al.111 In comparison to the other chromatin assembly methods described in the following sections, salt dilution/dialysis is a relatively easy method to efficiently assemble chromatin. Circular or linear DNA from any source over a wide range of lengths (>146 bp) and with nearly any sequence can be used to assemble individual nucleosomes or polynucleosomal arrays. The drawback of using this method is that physiological nucleosome spacing will not be achieved with polynucleosomal arrays. The nucleosomes do not position with regular spacing

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along the linear path of the DNA as seen in vivo; they may be closely packed with no linker DNA between them or have gaps between them considerably larger than the typical nucleosome repeat length. To circumvent this problem, DNA‐containing NPEs can be used in conjunction with the salt dilution/dialysis method, as noted above. 2. EXTRACT‐BASED CHROMATIN ASSEMBLY SYSTEMS A number of cell extracts competent for chromatin assembly have been described. Several different sources of cells have been used to prepare these extracts, including Xenopus eggs112 and oocytes,113,114 Drosophila embryos,54,115 and mammalian cells.116,117 Key components of these extracts are the histone chaperones that facilitate delivery of the histones to the site of nucleosome assembly. Although more difficult and involved, extract‐based chromatin assembly has distinct advantages over salt dialysis/dilution. For example, it uses physiological conditions for the assembly of nucleosomes and it promotes the assembly of chromatin with physiological nucleosome spacing, thus the inclusion of NPEs into the template DNA is not necessary. Since these systems support DNA‐replication independent chromatin assembly, establishing conditions for replication of the DNA template in the chromatin assembly reactions is not an obligatory step in the protocol. The first extract reported to successfully assemble chromatin in vitro was made from Xenopus eggs, as reported by Laskey et al. in 1977.112 This extract promoted the assembly of a regularly spaced nucleosome array on simian virus 40 DNA, as analyzed by micrococcal nuclease (MNase) digestion. Studies using this extract were the first to demonstrate that reconstituted chromatin could resemble native chromatin in terms of nucleosomal spacing. Modifications over time led to the use of Xenopus oocytes, which produce extracts that give a more reproducible assembly of chromatin.113,114,118 Exogenous pools of histones are not necessary with these extracts because of the large pools of soluble histones in Xenopus oocytes.112,119 The major H2A form present in these extracts, however, is not the same as in somatic cells, nor are the posttranslational modifications.120 Thus, the chromatin assembled will not resemble somatic chromatin with respect to histone composition and modification status. The first chromatin assembly extract prepared from Drosophila embryos was described in 1979,121 although it was not until over a decade later that the Wu and Kadonaga labs devised protocols for producing extracts that give reproducible and robust chromatin assembly.54,115 The Wu lab protocol uses preblastoderm embryos (0–2 h postlaying) to produce the ‘‘S‐150’’ extract, while the Kadonaga lab protocol uses primarily postblastoderm embryos (0–6 h postlaying) to produce the ‘‘S‐190’’ extract. The S‐150 extract, like its

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Xenopus oocyte extract counterpart, contains sufficient endogenous histones for chromatin assembly. In contrast, the S‐190 extract is deficient in histones and must be supplemented with exogenous histones to allow efficient chromatin assembly to proceed. This creates an opportunity to program the extract with histones from a wide variety of sources, including mammalian and recombinant. The chromatin assembled with the S‐150 extract or the S‐190 extract resembles native chromatin in terms of nucleosomal spacing and transcriptional regulation. Chromatin assembly extracts have also been successfully prepared from mammalian cells, including CV‐1 cells (monkey kidney), HEK 293 cells (human embryonic kidney), and HeLa cells (human cervix).116,117 Although these extracts can promote the assembly of chromatin with native nucleosome spacing, the efficiency of assembly is not typically as robust as the Xenopus oocyte and Drosophila embryo extract systems. As such, mammalian cell extracts are not widely used for the assembly of chromatin in vitro. Some of these extracts, however, are competent for DNA replication‐coupled chromatin assembly, making them useful for studying aspects of DNA replication.

3. DEFINED CHROMATIN ASSEMBLY SYSTEMS Biochemical fractionation of extracts competent for chromatin assembly revealed a number of factors required for chromatin assembly. These included histone chaperones such as N1/N2,122 nucleoplasmin,123 NAP‐1 (nucleosome assembly protein‐1),124 and CAF‐1 (chromatin assembly factor‐1),125–127 which can promote nucleosome assembly in vitro, although the assembled nucleosomes have non-native spacing. In 1997, Ito et al. purified an ATP‐utilizing chromatin assembly and remodeling factor (ACF) from Drosophila embryo extracts that functions with NAP‐1 and CAF‐1 to mediate nucleosome deposition and generate regularly spaced nucleosome arrays.124 ACF contains a SNF2‐like ATPase called imitation switch (ISWI), which is found in other chromatin modulating complexes, such as nucleosome remodeling factor (NuRF).128 A complex similar to ACF, called RSF (remodeling and spacing factor), has been purified from human cells.129 The cloning and expression of recombinant ACF and RSF has yielded completely defined and efficient systems for the assembly of physiologically spaced nucleosomal arrays in which every component is known (DNA, histones, ACF or RSF, histone chaperones, and ATP).124,129 This purity is a key advantage of the defined systems over the extract‐based systems, which produce chromatin contaminated with proteins that remain associated with the chromatin template even after subsequent purification.

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D. Methods for Analyzing Chromatin Assembled In Vitro A wide variety of methods are available for analyzing reconstituted chromatin, many of which are similar to methods used to analyze native chromatin. These methods fall into three main categories (1) nuclease digestion, (2) analysis of physical properties, and (3) imaging. Although a comprehensive description of these methods is beyond the scope of this chapter, we highlight a few of the most useful methods for analyzing chromatin assembled in vitro in the following sections.

1. NUCLEASE DIGESTION OF CHROMATIN ASSEMBLED IN VITRO A variety of nucleases can be used to analyze different aspects of chromatin assembled in vitro. For example, restriction endonucleases (REs) can be used to determine the relative accessibility of specific DNA sequences assembled into nucleosomes. Deoxyribonuclease I (DNase I) can be used to map contacts of the histone octamer with the DNA in a nucleosome and map the rotational positioning of the DNA relative to the histone octamer.130–132 MNase can be used to determine the spacing between nucleosomes in a polynucleosomal array.133,134 Digestion of chromatin assembled in vitro with REs, DNase I, and MNase can be used under experimental conditions to address specific mechanistic questions, for example, the location and effects of TF binding, the effects of CRCs on nucleosome positioning and nucleosomal DNA accessibility, and the effects of histone modifications on nucleosome accessibility. The same enzymes can be used to analyze the quality and properties of chromatin assembled in vitro. Of the three enzymes noted above, MNase is the most useful for examining the outcome of in vitro chromatin assembly reactions. MNase is much more efficient at cleaving ‘‘naked’’ DNA, such as that found in the linker region between nucleosomes, than nucleosomal DNA. If an array of nucleosomes has regular spacing, then digestion with a limiting amount of MNase, which is insufficient to cleave all of the linker DNA regions in the sample, will yield a ‘‘ladder’’ with the ‘‘rungs’’ repeating in increments of the fundamental nucleosomal DNA size (150 bp) upon agarose gel electrophoresis54,115 (Fig. 8A). This assay can be coupled with a Southern blotting protocol, using a probe to a specific region of the template, to examine the spacing of nucleosomes in a specific region of the DNA template.135 In addition, variations of the MNase assay can be used in an indirect labeling protocol to map nucleosome positions in a specific region of the DNA template.135

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FIG. 8. Analysis of chromatin assembly. (A) Chromatin assembled on a 3.2‐kb plasmid using Drosophila S-190 extract and core histones was subjected to a limiting digest with MNase and run on 1.3% agarose gel. The DNA was visualized using ethidium bromide staining. M, marker (a 123‐bp repeating ladder). (B) Atomic force microscopy (AFM) imaging of a 10.5‐kb plasmid assembled into chromatin using S-190 extract. Top panel: Scan probe oscillation amplitude image. The length scale is indicated by the white bar. Bottom panel: Topographical image. The height scale is shown along the bottom of the image.

2. ANALYSIS OF THE PHYSICAL PROPERTIES OF CHROMATIN ASSEMBLED IN VITRO Analysis of the physical properties of chromatin, such as the extent of supercoiling of the constituent DNA, can be informative. In the presence of topoisomerase activity, nucleosome assembly on closed circular DNA molecules promote the introduction of one negative supercoil per nucleosome, after the histone proteins are removed from the DNA. Molecules with differing superhelical densities can be separated by agarose gel electrophoresis and a determination of the number of nucleosomes present per DNA molecule can be made.136 The extent of negative supercoiling indicates the number of nucleosomes originally assembled onto the DNA template.136 Although limited digestion with MNase can give an indication of the quality of chromatin assembled in vitro, as indicated by the regular spacing of nucleosomes, the extent of supercoiling can give an indication of the efficiency of chromatin assembly (i.e., the number of nucleosomes deposited). A combination of MNase digestion assays and supercoiling assays will give the best assessment of chromatin assembled in vitro.

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The sedimentation properties of chromatin in glycerol or sucrose gradients can also be informative. For example, more highly compacted chromatin, as well as chromatin with a higher molecular weight due to the binding of chromatin‐interacting proteins, will sediment near the bottom of a glycerol or sucrose gradient run under the appropriate conditions.137 This property of chromatin makes sedimentation assays useful for addressing specific mechanistic questions, such as the conditions that promote or reverse chromatin compaction. 3. IMAGING OF CHROMATIN ASSEMBLED IN VITRO Analysis of chromatin using nuclease digestion, supercoiling, or sedimentation is informative, but each of these approaches is indirect. With the advent of methods for imaging chromatin, visual observations of chromatin can be made. Information about chromatin that can be obtained from images includes core particle size, linker length, number of nucleosomes per DNA molecule, and the extent of higher order structures. In 1975, the first clear images of single strands of chromatin were generated by using electron microscopy, leading to the descriptive term ‘‘beads on a string.’’109 Atomic force microscopy (AFM), developed in 1986, is a useful alternative method for imaging chromatin.138 AFM gives a true three‐ dimensional topographic image of the chromatin sample (Fig. 8B). One of the most recent developments in AFM is recognition imaging, in which the sampling nanotip of the AFM instrument is ‘‘functionalized’’ with an antibody or aptamer directed against an epitope in the chromatin sample.139–142 As the sample is scanned, a signal is generated upon interaction of the functionalized nanotip with the epitope in the sample, yielding a map of the location of the epitope. Bash et al. have used this approach to explore the features of Swi/Snf‐ remodeled nucleosomes under aqueous conditions, including the location of the H2A/H2B dimers using a tip functionalized with an antibody to H2A.143 Electron microscopy or AFM imaging of chromatin provides an excellent complement to enzymatic or physical assays of the properties of chromatin.

E. Putting It All Together: In Vitro Transcription with Chromatin Templates The in vitro transcription systems described in Section IV are compatible with the in vitro chromatin assembly systems described in this section. The combination of these approaches allows one to study the mechanisms of NR‐ dependent transcription with chromatin templates (Fig. 7). The details of how these two systems can be brought together are beyond the scope of this chapter, but can be found elsewhere.56,144–146

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VI. What have We Learned About NR‐Dependent Transcription from In Vitro Chromatin Assembly and Transcription Studies? In vitro chromatin assembly and transcription studies have provided a wealth of information about ligand‐dependent transcriptional regulation by NRs (Fig. 2). We have summarized the key findings from these studies in the following sections. In addition, in some places, we have also summarized key findings from related studies using other DNA-binding transcriptional activators for comparison.

A. Role of Ligands The effects of ligands on NR‐mediated transcription had been studied for over two decades prior to the first use of in vitro chromatin assembly and transcription systems in the mid‐ to late 1990s. This approach, however, has revealed new molecular details about the biochemistry and molecular biology of ligand‐dependent transcriptional regulation by NRs. Three important findings using this approach highlight the role of ligands in NR‐mediated transcription: (1) the repressive effects of chromatin can be overcome through a hormone‐dependent mechanism, even though transcription with naked DNA can occur through a hormone‐independent mechanism, (2) the actions of antihormones are, at least in part, mediated through chromatin‐dependent mechanisms, and (3) coregulators, such as the SRC proteins and p300/ CREB-binding protein (CBP), facilitate the derepression required for transcriptional activation by promoting the covalent modification of histones. This section details the studies leading to these findings. 1. AGONIST LIGANDS ARE REQUIRED FOR NR‐DEPENDENT TRANSCRIPTION WITH CHROMATIN TEMPLATES As noted in Section II.D, the early biochemical studies of transcriptional activation by NRs were performed using naked DNA templates. Although these studies provided information about the role of DNA binding by NRs in the transcription process, they fell short with respect to the role of ligand. One possible explanation for the ligand‐independent activation observed for steroid receptors, such as ERa, in the early in vitro transcription assays with naked DNA is the lack of chaperone proteins in the receptor preparations. In vivo, unliganded steroid receptors are bound by chaperones, such as the heat shock proteins (e.g., hsp70 and hsp90), which sequesters the receptors and prevents them from binding to DNA.4 Ligand binding promotes the release of the chaperones from the receptor, allowing the receptor to bind to its cognate hormone response element. The purified native or recombinant receptors used in biochemical assays are typically free of heat shock proteins and, thus, are

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able to bind DNA in the absence of ligand. In this regard, assays using cell extracts as a source of receptor for in vitro transcription assays yielded ligand‐ dependent transcriptional activation, presumably because the receptors in the extract were complexed with chaperones.147,148 The absence of chaperones may account for some, but certainly not all, of the hormone‐independent transcription observed with naked DNA templates. With the advent of in vitro chromatin assembly systems, a physiological template became available to assess the effects of ligands in NR‐dependent transcription. Assays with these systems produced three outcomes that were generally not observed with naked DNA: (1) low basal transcription, (2) low NR‐dependent transcriptional activity in the absence of ligand, even though the receptors were bound to the DNA, and (3) bona fide ligand‐dependent activation. These outcomes were nicely demonstrated in early studies with ERa, which bound to DNA in the chromatin template, but exhibited low transcriptional activity in the absence of agonist ligand.93 Upon addition of the agonist 17b‐estradiol (E2), a 25‐fold activation of transcription was observed. The ligand‐dependent activation by E2 was blocked by the addition of antiestrogens (i.e., either trans‐hydroxytamoxifen or ICI 164,384) without inhibiting DNA-binding activity.93 The effects of antiestrogens required a chromatin template. These initial observations of ligand‐dependent transcription by ERa with chromatin templates have been extended to include other NRs, such as the PR, retinoid acid receptor (RAR), vitamin D receptor (VDR), and thyroid hormone receptor (TR).149–152 Collectively, these studies show that agonist ligands are required for NR‐ dependent transcription with chromatin templates. Agonist ligands promote the activation of transcription by NRs above the low basal levels that are observed with a restrictive chromatin environment. In the absence of chromatin, basal levels of transcription are elevated considerably, in many cases masking the effect of ligand. As discussed in the following section, the primary action of agonist ligands is to promote the association of coregulators that function with NRs to counteract the restrictive nature of the chromatin environment. 2. AGONIST LIGANDS PROMOTE PRODUCTIVE INTERACTIONS WITH NR COREGULATORS In vitro chromatin assembly and transcription systems have been useful for exploring the role of ligands in promoting productive interactions between NRs and their coregulators. For example, ERa‐dependent transcription was shown to be activated approximately sixfold in the presence of exogenous recombinant p300, a histone acetyltransferase,93 but only in the presence of E2. Moreover, E2 was shown to be required for the SRC‐dependent recruitment of p300 to DNA‐bound ERa, interactions that led to the acetylation of nucleosomal

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histones by p300.153,154 Additionally, transcription by RARa/RXRa heterodimers and TRb/RXRa heterodimers was enhanced by p300 in a manner dependent on agonist ligand (all‐trans retinoic acid or thyroid hormone, respectively).151,152 Likewise, transcription by PR was potentiated by SRC‐1 in the presence of a progestin ligand.150 These studies, in conjunction with a variety of in vitro binding and cell‐based transactivation studies performed contemporarily,26,34 highlight the role of NR ligands in promoting transcriptionally productive interactions with coregulator proteins, which in many cases leads to specific covalent modifications of the chromatin template.

B. Role of Basal TFs As noted in Section II.B, some of the first transcriptionally relevant interactions characterized for NRs were with components of the basal transcription machinery.21–24 In vitro transcription systems have been useful for exploring the functional relevance of these interactions. Several TAFs in TFIID, as well as TBP itself, make direct contacts with NRs during the ligand‐regulated transcription process23,26 (Fig. 2). For example, TAF10 (a.k.a. hTAFII30) binds to ERa, an interaction critical for ERa‐dependent transcription.155 Such interactions can help to recruit or stabilize the binding of TFIID at the promoter, a process that is enhanced by the binding of some TAFs to the core promoter elements.25 Although some activators do not require TAFs,156–158 many activators require them for transcriptional activation, including several NRs. For example, TFIID, but not TBP alone, supported ERa‐dependent transcription in an in vitro chromatin assembly and transcription system.159 This effect was shown to be independent of the carboxyl‐terminal activation function (AF‐2) of ERa, as a point mutation that destroys the AF‐2 domain failed to reduce TFIID‐dependent transcriptional activation. A similar requirement for TAFs was also shown for VDR.81 Furthermore, TFIID, but not TBP alone, can act synergistically with coregulator complexes, such as Mediator and Swi/Snf, to potentiate transcription by NRs.25,40,81 In vitro transcription systems have also been used to explore the role of TFIIB, a basal TF that stabilizes the binding of TBP/TFIID to DNA,160 with NRs. TFIIB was shown to interact with and enhance transcription by NRs, including PR and the orphan NRs COUP‐TF and HNF4.21,161 A similar effect of TFIIB with VDR was demonstrated using cell‐based assays.162,163 TFIIB also interacts with TRb, but this interaction leads to a repression of transcription,164 suggesting NR‐specific or context‐specific effects for NR–TFIIB interactions. TFIIB, as well as TBP, also interact with the NR coactivator SRC‐1,165,166 highlighting the complex set of interactions that occur between NRs, the basal transcription machinery, and coregulator proteins that are required for transcription with chromatin templates.

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C. Role of Coactivators As noted in Section II.C, the coregulator hypothesis led to the eventual (and continued) identification of a wide variety of NR coregulators, proteins whose interactions with NRs (either binding or release) are regulated by ligands. The development of in vitro chromatin assembly and transcription systems for studying NRs went hand‐in‐hand with the identification and characterization of the coregulators (Fig. 2). In the following sections, we discuss how in vitro chromatin assembly and transcription systems have been used to characterize the biochemical activities of a variety of NR coregulators.

1. P160/SRC PROTEINS Members of the p160/SRC family of NR coactivators (referred to herein as the SRC family) were some of the first mammalian NR coregulators, after Swi/Snf as noted in Section III.B, to be cloned and characterized, starting with the founding family member, SRC‐1, in 1994.32 The p160/SRC family has three members, which are referred to by a wide variety of names, but can simply be called SRC‐1, SRC‐2, and SRC‐3.37 All three interact directly with AF‐2 of ligand‐bound NRs via motifs resembling Leu‐X‐X‐Leu‐Leu (‘‘LXXLL’’ motifs or NR boxes) and stimulate NR transcriptional activity. Some studies,167,168 but not others,169,170 have found the SRC family members to possess intrinsic histone acetyltransferase activity. If present, it appears to be weak at best, and is dispensable for NR‐mediated transcription with chromatin templates. This latter observation was made for PR, ERa, and TRb using in vitro chromatin assembly and transcription systems with purified SRC proteins lacking the putative acetyltransferase domain, but containing the NR interaction domain.151,153,171 The ability of the SRC proteins to promote NR‐dependent transcription with chromatin templates seems to be more related to their ability to recruit HMEs to liganded NRs, rather than an intrinsic HME activity. In this regard, SRC‐1 has been shown to have little coactivator activity with naked DNA templates.171 Interactions between the SRC proteins and p300 or CBP (two related, but functionally distinct histone acetyltransferases; see Section VI.C.2.) play a major role in this activity. Functional interplay between SRCs and p300/ CBP was first demonstrated in cell‐based assays, but was functionally dissected in a series of in vitro transcription assays with chromatin templates. For example, deletion of the p300/CBP interacting domain from SRC‐1 blocked its ability to potentiate PR‐mediated transcription on chromatin templates.171 Likewise, inhibitory peptides that prevent SRC‐p300/CBP interactions inhibited ERa‐ and TRb‐dependent transcription.151,153 In sum, SRCs act as

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bridging factors that bind directly and simultaneously to NRs and key HMEs to bring histone‐modifying activities to target promoters to aid in the relief of chromatin‐mediated repression. 2. P300 AND CBP ACETYLTRANSFERASES HMEs facilitate NR‐mediated transcription by establishing a pattern of covalent histone modifications (a ‘‘histone code’’ of acetylation, methylation, phosphorylation, ubiquitylation, etc.) that (1) modifies the physical properties of chromatin changing the electrostatic interactions of the DNA and histones (e.g., through acetylation‐mediated charge neutralization) and (2) creates a set of modification‐dependent binding sites for chromatin and transcription regulatory proteins. The end result is a change in nucleosome structure and the complement of nucleosome‐bound proteins, which dictates the NR‐dependent transcriptional outcome. p300 and CBP are highly related acetyltransferases (commonly referred to collectively as p300/CBP) that can modify a number of different lysine residues in the amino‐terminal tails of core histones, ultimately destabilizing chromatin structure to create a more permissive state for transcriptional activation (Fig. 2). p300 and CBP were originally shown to interact directly with NRs in a ligand‐dependent manner to activate transcription,172,173 although as noted above the primary mode of interaction is likely through SRC proteins. p300/ CBP is required by NRs for maximal transcriptional activation with chromatin templates. This was demonstrated clearly using a variety of approaches coupled with in vitro chromatin assembly and transcription, including the addition of (1) exogenous recombinant p300,93,151–154,171 (2) competition assays using inhibitory polypeptides that disrupt the interaction of p300/CBP with SRCs,151,153 and (3) a p300 deletion mutant lacking the SRC interaction domain, as well as a deletion mutant of SRC‐1 lacking the p300 interacting domain.151,154,171 The coactivator effects of p300/CBP in these in vitro assays required p300/CBP acetyltransferase activity151,153,154 and, not unexpectedly, acetyl‐coenzyme A (acetyl‐CoA, the acetyl donor used by p300).174 In this regard, point mutations within the acetyltransferase domain dramatically reduced p300 transcriptional activity with liganded ERa and TRb.151,153,154 The requirement for p300/CBP acetyltransferase activity in the transcription assays with chromatin templates was partially circumvented by the addition of the histone deacetylase inhibitor, trichostatin A (TSA), which blocks the activity of endogenous HDACs in the transcription extracts.151 Collectively, these biochemical studies highlight one of the primary roles of p300/CBP in NR‐ dependent transcription with chromatin templates, namely the acetylation of nucleosomal histones, although recent studies have demonstrated that NRs themselves may also be targets of p300/CBP‐mediated acetylation.175–179

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Some of the transcriptional activity of p300/CBP may also be mediated through interactions with components of the basal transcription machinery, such as TFIIB, TBP, and Pol II.26,34 Interactions of p300/CBP with Pol II, which occur through the CH3 domain of p300/CBP, facilitate NR‐dependent transcription with chromatin templates. In this regard, deletion of the CH3 domain of p300 blocked its coactivator activity with ERa and TRb, without affecting histone acetylation.151,154 In vitro transcription assays, however, showed no enhancement of NR‐dependent transcription by p300 with nonchromatin or naked DNA templates,93,152 suggesting that the chromatindependent actions of p300 are obligatory and act upstream of interactions with the Pol II machinery.

3. OTHER HMES In addition to acetyltransferases, other proteins with histone‐modifying activities have been implicated in NR‐dependent transcription, including methyltransferases and kinases. As with p300/CBP, in vitro chromatin assembly and transcription systems have been useful experimental approaches for examining the activity of these enzymes. The activity of the protein arginine methyltransferases PRMT1 and PRMT4 (a.k.a. CARM1), HMEs that preferentially methylate arginine residues in the amino‐terminal tails of histones H4 and H3, respectively, enhance NR‐dependent transcription. Both PRMTs were shown to synergistically activate transcription with p300/CBP and SRCs.180,181 Moreover, methylation of HNF4 by PRMT1 enhanced HNF4 DNA-binding activity,181 while methylation of CBP by PRMT4 promoted a switch in the signaling specificity of CBP from one type of activator to another (i.e., from CREB to NRs).180 PRMT4 was also shown to stimulate the ATP‐dependent chromatin remodeling activity of Swi/Snf.182 A key role for PRMT1 and PRMT4 catalytic activity in modulating these effects was shown using a catalytically inactive point mutant180 or recombinant histones with point mutations in the target arginine residues in the histone tails.181 The latter approach illustrates the power of the in vitro chromatin assembly system as a tool to explore mechanistic issues that cannot be readily addressed in intact mammalian cells.106 Phosphorylation of histone tails has also been shown to regulate NR‐ dependent transcription. PR‐dependent transcription with a MMTV promoter construct assembled into chromatin in vitro was shown to be activated by mitogen and stress activated protein kinase 1 (Msk1).183 Msk1 phosphorylated serine 10 on histone H3 (H3S10), increasing the association of Swi/Snf with the histone octamer. Chemical inhibition of Msk1 kinase activity, as well as mutation of H3S10, reduced PR‐dependent transcription.183 Collectively, the

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studies highlighted in this section illustrate the utility of in vitro chromatin assembly and transcription systems for studying the actions of HMEs, as well as the mechanistic conclusions that can be made from such studies. 4. ATP‐DEPENDENT CRCS ATP‐dependent CRCs mobilize or structurally alter nucleosomes, increasing the accessibility of the underlying DNA to TFs and the Pol II transcription machinery184–186 (Fig. 2). The identification and functional characterization of CRCs went hand‐in‐hand with the development of methods for assembling and analyzing nucleosomes and chromatin in vitro. As noted in Section III.B, the Swi/Snf CRC was the first NR coregulator identified, and in vitro chromatin assembly and transcription systems have been useful for studying Swi/Snf activity with NRs. For example, Swi/Snf was shown to be required for full ligand‐dependent transcriptional activation by RARa in a manner dependent on its ATPase activity and functional synergism with p300/SRC.75 Swi/Snf has two highly related forms in humans, PBAF and BAF, which differ in their complement of Brg1‐associated factors (BAFs).185,186 Although these two complexes can be separated biochemically, a functional distinction between them was not apparent until in vitro studies with VDR and peroxisome proliferator activated receptor gamma (PPARg).81 PBAF, but not BAF, was shown to be a potent activator of VDR‐ and PPARg‐dependent transcription with chromatin templates.81 These results illustrate the subunit specificity of Swi/Snf function with NRs (BAF180 vs. BAF250 for PBAF and BAF, respectively). Functional specificity of the Swi/Snf ATPase subunits, Brg1 and Brm, has also been demonstrated using in vitro chromatin assembly and transcription systems.78 Brg1 was shown to interact with and regulate transcription mediated through zinc finger proteins, while Brm showed a similar specificity and activity for ankyrin repeat proteins.78 These results also highlight the utility of using in vitro chromatin assembly and transcription systems for studying the activity of coregulator complexes. The function of the chromatin remodeling factor ISWI, an ATPase, with NRs has also been explored using in vitro chromatin assembly and transcription systems. ISWI was shown to be required for full PR‐ and RARa‐dependent transcription.75,187 Unlike RARa, however, PR does not require Swi/Snf.187 ISWI enhanced the binding of RARa/RXRa heterodimers to DNA and altered the chromatin structure of the promoter region in an ATP‐dependent manner.75 Collectively, these results indicate that ATP‐dependent CRCs, such as Swi/Snf and ISWI, play important and specific roles in overcoming the repressive effects of chromatin in NR‐dependent transcription. Part of the activity of CRCs with NRs requires functional interplay with other coregulators, such as HMEs.75,182,183

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5. MEDIATOR Interactions between DNA‐bound NRs and the Pol II machinery help to promote the formation of stable transcription complexes at the promoter, an event that is especially important in the restrictive chromatin environment of the nucleus (Fig. 2). Mediator (a.k.a. TRAP and DRIP) is a group of highly related coactivator complexes that can interact simultaneously with NRs and Pol II.40,41 Mediator was originally identified and purified from yeast, and subsequently from mammalian cells.188 The TRAP and DRIP complexes were purified using an affinity chromatography strategy with the LBDs of TRb and VDR, respectively.189,190 Other related complexes with variations in subunit composition and activities (e.g., SMCC, PC2, CRSP, and ARC) were subsequently purified and characterized.188 At least two individual subunits of Mediator interact directly with NRs, as shown by a variety of biochemical assays.40,41 Med1 (a.k.a. TRAP220, DRIP205, Med220) binds in a ligand‐dependent manner to the LBDs of NRs through a receptor interaction domain containing two ‘‘LXXLL’’ motifs, while Med14 (a.k.a. TRAP170, DRIP150, Med150) binds to the amino‐ terminal region of some steroid hormone receptors without a requirement for ligand. Med1 promotes the association of the entire Mediator complex with a variety of NRs in vitro and likely plays a key role in promoting the recruitment of Mediator to the promoters of NR‐regulated genes in response to ligand in vivo. The critical role of Med1‐NR interactions for Mediator coactivator activity has been demonstrated in biochemical assays using Med1 mutants,191 as well as inhibitor polypeptides that selectively block Med1‐NR interactions.192 Moreover, Mediator purified from Med1 / mouse embryonic fibroblasts was shown to have impaired coactivator activity with TRb, but not the synthetic activator Gal4‐VP16, which does not interact with Med1.191 Mediator has been shown to enhance transcriptional activation with both naked and chromatin templates for several NRs, including TRb,189,191 VDR,81,149,190,193 ERa,192,194,195 PPARg,196 and HNF4.197 In spite of the ligand‐dependent interactions between Mediator and NRs, ligand has little or no effect on the ability of Mediator to enhance NR‐dependent transcription with naked DNA templates,81,149,190 indicating at least part of Mediator’s coactivator affect with NRs is through a ligand‐independent mechanism. In contrast, ligand is required for Mediator’s coactivator affect with chromatin templates,81,149 again illustrating the need for ligand to stabilize critical transcriptionally productive interactions in a chromatin environment. A recent biochemical study has demonstrated a role for the chromatin and transcription regulatory factor poly(ADP‐ribose) polymerase‐1 (PARP‐1) in regulating the function of Mediator.198 PARP‐1 is an abundant nuclear enzyme with an intrinsic enzymatic activity that catalyzes the polymerization of

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ADP‐ribose units from donor NADþ molecules on target proteins199 PARP‐1 localizes to the vast majority of actively transcribed promoters200 and regulates transcription through a number of distinct mechanisms, including alterations in chromatin structure, as well as classical coregulator functions.201,202 The coregulator functions of PARP‐1 have been shown to play an important role in NR‐dependent transcription.198,203–206 In a purified in vitro chromatin assembly and transcription system with RARa, PARP‐1 was shown to function as an ‘‘exchange factor,’’ promoting a switch from an inactive Mediator complex (i.e., Cdk8‐containing) to an active Mediator complex (i.e., without Cdk8).198 In the absence of PARP‐1, this switch did not occur and transcription was abrogated, highlighting the importance of the composition of Mediator in determining its function.

D. Role of Corepressors The regulation of NR‐dependent transcription requires a carefully orchestrated interplay between coactivators, which enhance transcriptional responses, and corepressors, which attenuate them. Two well‐studied corepressors of NR‐mediated transcription are NCoR and SMRT. NCoR and SMRT are found in complexes with HDACs and CRCs.207–216 These corepressor complexes are recruited to unliganded NRs through direct interactions between the receptors and NCoR or SMRT217,218 (Fig. 2). NCoR‐ and SMRT‐ containing corepressor complexes act to repress transcription by deacetylating nucleosomal histones through the actions of HDACs and establishing repressive chromatin structures through the actions of CRCs.35 Ligand binding to NRs acts as a switch to promote the exchange of corepressors for coactivators.34 The cellular levels of coactivators and corepressors can dictate the nature of a ligand‐dependent transcriptional response, especially with respect to the actions of selective receptor modulator (SRM) ligands, which exhibit contextdependent agonist or antagonist activity.219 This was nicely illustrated in cell‐based assays with the selective ER modulator tamoxifen.220 Although transcriptional repression has been difficult to study with conventional in vitro chromatin assembly and transcription systems due to the very low basal level of transcription in these assays, some studies have found ways to use these systems to explore the function of corepressors. For example, Liu et al. examined the role of coactivator/corepressor ratios on PR‐dependent transcription in the presence of the mixed agonist, RU486.221 RU486 repressed PR‐mediated transcription in nuclear extracts with a low SRC‐1/p300 to NCoR/SMRT ratio, but enhanced PR‐mediated transcription in nuclear extracts with a high ratio. Interestingly, addition of purified SRC‐1 to extracts with a low SRC‐1/p300 to NCoR/SMRT ratio relieved the repression. Likewise, addition of purified SMRT to extracts with a high SRC‐1/p300 to NCoR/SMRT

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ratio promoted repression. In contrast, the pure agonist, progesterone, and the pure antagonist, ZK98299 were unaffected by the coregulator ratios.221 Another corepressor, receptor‐interacting protein 140 (RIP140), which interacts with NRs in the presence of agonist ligands, was shown to repress retinoic acid‐induced transcription by RARa with chromatin templates. RIP140 repressed transcription by inhibiting the recruitment of the histone acetyltransferase, p/CAF, and promoting the recruitment of histone deacetylases.222 Finding additional ways to exploit in vitro chromatin assembly and transcription systems to explore the mechanisms of corepressor‐mediated transcriptional repression by NRs is an important future direction.

E. Role of Nucleosome‐Binding Proteins Proteins that bind to nucleosomes can modulate chromatin structure and, hence, regulate transcription. Linker histones, such as H1, bind at or near the dyad axis of the nucleosome and promote the formation of higher order chromatin structures that are generally repressive to transcription223,224 (Fig. 9A). In contrast, HMGN1/N2 (formerly HMG14/17), charged proteins that bind with high affinity to two distinct sites on nucleosomes225,226 and stabilize core particles,227–229 enhance transcription with chromatin templates.230 The ability to assemble chromatin with or without specific

A Dyad axis

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FIG. 9. Nucleosome‐binding proteins can promote the compaction of chromatin into higher order structures. (A) PARP‐1 and the linker histone H1 bind to overlapping sites on the nucleosome at or near the dyad axis (indicated by the dotted circle). (B) AFM scan probe oscillation amplitude image of a 10.5‐kb plasmid assembled into chromatin using S-190 extract in the absence (left) or presence (right) of PARP‐1. The length scale is indicated by the white bars.

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nucleosome‐binding proteins has made in vitro systems particularly useful for examining the role of these proteins in the regulation of chromatin structure and transcription. The role of histone H1 (H1) in the regulation of NR‐dependent transcription with chromatin templates has been examined using in vitro assays. Studies with three different NRs have yielded three different results. With RARa, the incorporation of H1 into chromatin had no effect on ligand‐ and receptor‐ dependent transcription, whether it was basal or activated, or in the presence or absence of p300 or Swi/Snf.75 In contrast, the incorporation of H1 into chromatin repressed ligand‐ and receptor‐dependent transcription with ERa.231 This occurred without a reduction in ERa binding to the chromatin template or histone acetylation levels, but the promoter had a less accessible chromatin structure and a reduced stability of the Pol II PIC in the presence of H1. With PR, the incorporation of H1 into chromatin enhanced the binding of the receptor to the chromatin template and increased transcription initiation.232 This effect was due to increased homogeneity of nucleosome positioning at the promoter which was more favorable for the binding of PR. Although possible, the different results observed in these three studies are unlikely to be due to some fundamental differences between the three receptors. Rather, they are likely due to differences in the promoter architecture, protein cofactors present in the chromatin assembly and transcription reactions, and perhaps the covalent modification state of the histones used. Together, these studies highlight the complex nature of H1 in transcriptional regulation. Studies using in vitro chromatin assembly and transcription assays have also led to the discovery of an H1‐like nucleosome-binding property of PARP‐ 1. Kim et al. showed that PARP‐1 and H1 bind to an overlapping site and compete for binding to nucleosomes137 (Fig. 9A). The incorporation of PARP‐1 into chromatin in the absence of NADþ (a condition not observed in vivo) promoted the formation of compact nucleosome structures that are repressive to ERa‐dependent transcription137,233 (Fig. 9B). This repression was relieved upon the addition of NADþ. In vivo, the PARP‐1 content at promoters, nuclear NADþ levels, and other signal‐dependent regulatory inputs are likely to dictate the functional outcome of PARP‐1 binding to promoter nucleosomes.201 Another recent study used in vitro reconstitutions with chromatin templates to examine the functional interplay between PARP‐1, the histone chaperone SET (a.k.a. TAF‐I and INHAT), and the chromatin‐associated factor DEK in VDR‐dependent transcription.234 PARP‐1 and DEK repressed transcription in the absence of SET and NADþ by restricting access of the Pol II machinery to chromatin. In contrast, SET potently enhanced transcription in the absence of NADþ by promoting the release of PARP‐1 and DEK. Upon the addition of NADþ, PARP‐1 poly(ADP‐ribosyl)ated and evicted itself and DEK from chromatin permitting the loading of the Pol II machinery in a SET‐

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independent manner.234 SET has also been shown to function as an inhibitor of p300/CBP‐ and PCAF‐mediated histone acetylation by binding to histones and masking them from targeting as acetyltransferase substrates.235 Collectively, the studies highlighted in this section illustrate the utility of using in vitro chromatin assembly and transcription systems to explore the role of nucleosome‐binding proteins in regulating NR‐dependent transcription.

F. Order and Dynamics 1. TEMPORAL ASPECTS OF NR‐DEPENDENT TRANSCRIPTION Understanding the details of NR‐dependent transcription, from ligand binding through transcription initiation and elongation by Pol II, can provide insights into the overall mechanisms of gene control by NRs. As outlined above, many coregulators play key roles in gene regulation. Coordination of the recruitment and activation of these factors is essential for proper gene expression. Although the pathway from NR activation to transcription by Pol II may have some promoter‐ and cell type‐specific nuances, general principles have been identified that are applicable to many NRs and their target genes. A simplified outline of the steps leading to NR‐dependent transcription via the recruitment of Pol II is listed below. One major consideration is whether the ‘‘apo’’ or unliganded NR is complexed with chaperones or is constitutively bound to DNA: (1a) For NRs complexed with chaperones (e.g., steroid hormone receptors): Binding of ligand by the receptor, release of chaperones from the receptor, and binding of the receptor to specific sites in the genome. The latter can occur by direct binding to conserved hormone response elements or, in some cases, indirect binding through other DNAbinding TFs. (1b) For NRs constitutively bound to DNA (e.g., RXR‐containing heterodimers): Binding of ligand by the receptor and release of corepressor proteins. (2) Recruitment of coactivators by the liganded receptor, including HMEs and CRCs, leading to a more open chromatin architecture in the promoter region. This may occur at distal receptor binding sites (i.e., enhancers), with ‘‘delivery’’ of the DNA‐bound receptor/coregulator complexes to the promoter region through a looping mechanism. (3) Recruitment of the Pol II transcriptional machinery, whose binding may be stabilized by the actions of Mediator. (4) Transcription initiation, with subsequent elongation through the nucleosome‐containing body of the gene. The latter may be facilitated by elongation factors with histone chaperoning activities.

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One caveat for this sequence of events is that many genes have a preloaded or promoter proximally paused Pol II prior to signal‐dependent activation28,29 (Fig. 10). In fact, estrogen‐regulated genes are enriched for genes with paused Pol II.27 In these cases, the promoter chromatin will have already been modified (as in 2 above) and Pol II will have already initiated transcription prior to the binding of the liganded receptor to DNA. In either case, this outline provides a useful framework for considering the steps leading to NR‐dependent transcription.

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FIG. 10. NR‐dependent transcription occurs by the regulation of RNA Pol II binding or activity. (A) Control of hormone‐dependent gene expression by regulating Pol II binding. Agonist ligands acting through NRs, which may bind in the proximal promoter region or at distal enhancers (not shown), promote the recruitment of Pol II. The transition from transcription initiation to transcription elongation is associated with phosphorylation of specific residues within the heptapeptide repeat of the Pol II Rpb1 subunit carboxyl‐terminal domain (commonly referred to as the ‘‘Pol II CTD’’). For example, phosphorylation at serine 5 (5P) of the Pol II CTD generally occurs early in the transcription cycle. In contrast, phosphorylation at serine 2 (2P), which is catalyzed primarily by the cyclin‐dependent kinase‐9 (Cdk9) of the positive transcription elongation factor‐b (P‐TEFb), generally occurs upon productive elongation. (B) Control of hormone‐dependent gene expression by regulating the activity of preloaded Pol II. Preloaded Pol II remains in the proximal promoter region prior to hormone treatment in association with negative elongation factor (NELF), which prevents elongation. Agonist ligands acting through NRs (not shown) promote productive elongation by Pol II, possibly involving the inactivation, but not release, of NELF. The movement of Pol II into the body of the gene occurs concomitantly with hormone‐dependent Pol II CTD phosphorylation, primarily at serine 2 (2P), as well as Cdk9 recruitment. Phosphorylation of serine 5 (5P) also increases to some extent as well. This model is based on studies with estrogen signaling and ERa,27 but should be applicable to many NRs.

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In vitro transcription assays provide a unique opportunity to address questions concerning the order of protein recruitment and activity during the process of NR‐dependent transcription. As opposed to cell‐based assays using knockdown and overexpression strategies or transient transfection of plasmid‐based reporters, in vitro transcription assays allows for a direct analysis of the temporal order of factor requirement by controlling the addition of factors to certain time points in the analysis. Furthermore, in vitro transcription assays are well suited to the use of functionally inactive or dominant negative mutant proteins, as well as chemical inhibitors and activators, to study the requirement for specific activities and interactions at discrete steps in the transcription process. Biochemical assays have enhanced our understanding of the steps leading to NR‐dependent transcription. For example, the spatial and temporal actions of coactivators have been explored by bypassing the ligand and receptor components of the system. In this regard, tethering the p300/CBP‐binding domain of SRC to DNA through a heterologous DBD (e.g., from the yeast GAL4 TF) supports potent p300/CBP‐ and histone acetylation‐dependent transcription with chromatin templates in the absence of an NR153 (Fig. 11A and B). The need for SRC can be bypassed as well, by tethering the acetyltransferase domain of p300/CBP to the GAL4 DBD236 (Fig. 11C). These experiments have been complemented by order‐of‐addition experiments. The addition of SRC before, and p300 during, the formation of a stable PIC leads to efficient transcription.171 Reversing the timing of the addition of SRC and p300, however, results in reduced levels of transcription. Other coregulators may act downstream of p300/CBP. For example, the methyltransferase PRMT4 acts synergistically with p300 in activating transcription, possibly due to higher enzymatic activity in the presence of acetylated histones.180 Thus, the primary actions of PRMT may occur after histone acetylation. Together, these results demonstrate that coactivators act downstream of the receptor and upstream of chromatin modulation. In addition, they suggest an order for the actions of coactivators, such as SRC and p300/CBP. A key role for p300/CBP at NR‐regulated promoters is for the acetylation of the amino‐terminal tails of nucleosomal histones. Without acetylation of histones, PIC formation is impaired, indicating that acetylation of histone tails precedes PIC formation.75 Supporting this, histone acetylation by p300 does not require interactions between p300 and the Pol II machinery, as demonstrated using an inhibitory polypeptide that blocks p300–Pol II interactions and transcription, but not histone acetylation.153 Several studies have examined the temporal requirement for histone acetylation by using acetyl‐CoA, the acetyl donor used by p300/CBP, and TSA, and inhibitor of class I and II HDACs. In vitro, acetyl‐CoA is required for activator‐dependent transcription with chromatin templates.75,174,196 Addition of acetyl‐CoA at different times during the transcription process

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A p300/CBP SRC Ac

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SRC(PID) Gal4(DBD) GAL

C p300(HAT) Ac Gal4(DBD) GAL FIG. 11. Experimental systems for analyzing the spatial and temporal order of SRC and p300/ CBP actions. (A) Agonist‐bound NRs can recruit SRC, which in turn recruits p300/CBP, resulting in the acetylation of histones and transcriptional activation. (B) The requirement for NRs can be bypassed by fusing the p300/CBP‐binding domain (PID) of SRC to a heterologous DNA‐binding domain (e.g., GAL4 DBD). This fusion is sufficient to recruit p300/CBP to promoters, resulting in the acetylation of histones and transcriptional activation. (C) The requirement for NRs and SRC can be bypassed by fusing the histone acetyltransferase (HAT) domain to a heterologous DNA‐binding domain (e.g., GAL4 DBD). This fusion is sufficient to promote the acetylation of histones, as well as transcriptional activation.

(i.e., before PIC assembly, during PIC assembly, at transcription initiation) indicates that it is required at a step prior to PIC formation.75,174 These data fit well with experiments showing that the requirement for p300/CBP can be circumvented by the addition of TSA.192 CRCs also play a role at specific points during NR‐dependent transcription with chromatin templates. For example, ISWI acts early in the process to facilitate the receptor binding to HREs in chromatin.75,187 In contrast, Swi/Snf facilitates the assembly of basal TFs onto the promoter.75 The actions of HMEs and CRCs are interdependent and coordinated, allowing multiple opportunities for regulation.75 In sum, the concerted actions of HMEs and CRCs, which alter the patterns of histone modifications and nucleosome positioning, create a chromatin landscape that allows PIC formation.237

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Many of the chromatin‐directed actions outlined above occur independently of the Pol II machinery (e.g., Pol II, TAFs, GTFs, Mediator),238–240 but they are not sufficient for NR‐dependent transcription with chromatin templates. Components of the Pol II machinery are required as well. For example, NRs require both Mediator and TAFs for transcriptional activation.81,149,159,189–197 A well‐documented interaction between the LBD of NRs and the Med1 subunit of Mediator plays an important role in transcriptional activation, and inhibiting this interaction reduces the level of transcriptional activation.191,192,241 Although the recruitment of Mediator and Pol II occurs after histone acetylation,238–240,242,243 the mode of recruitment is not clear. NRs play a role, but may not be absolutely required to bring Mediator to the promoter. For example, Mediator and Pol II have been shown to bind at promoters in the absence of an interaction with an NR (e.g., HNF4), although both the receptor and Mediator were required for transcription to occur.197 Interestingly, there may be selectivity in the recruitment of Mediator and SRC to NRs, even though they bind to overlapping sites on the receptor.192 In fact, a failure to bind and recruit SRCs impairs subsequent recruitment and actions of Mediator.192 Together, these studies support the conclusion that various coregulators act in a concerted manner to promote histone modifications and Pol II recruitment/stabilization, both of which are required at specific points in the process of NR‐dependent transcription with chromatin templates. 2. INITIATION AND REINITIATION Although most studies of NR‐dependent transcription have focused on transcription initiation, other steps in the transcription process, such as elongation and reinitiation are also subject to regulation (Fig. 12). Transcription initiation requires the recruitment of a complete set of all the proteins comprising the Pol II basal transcription machinery. Some of these factors (e.g., TFIID and TFIIA) remain associated with the promoter after transcription initiation.244–246 Transcription reinitiation makes use of these remaining factors (the ‘‘reinitiation scaffold’’) to reassemble a complete and transcriptionally competent complex for subsequent rounds of transcription.245,247 Although similar to initiation, the process of reinitiation requires a distinct set of factors from those required for initiation.245,248 In vitro transcription assays have been a valuable tool in assessing the role of specific proteins for their involvement in initiation and reinitiation. Common experimental approaches include (1) determining the factors that remain bound to the template after transcription initiation and (2) comparing the relative levels of RNA produced from a single round of transcription versus multiple rounds of transcription. The latter is achieved by the addition of the ionic detergent Sarkosyl immediately after the addition of nucleoside

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< reinitiation scaffold > (4) FIG. 12. Steps in the transcription cycle. A schematic representation of four steps in the transcription cycle, from transcription initiation to transcription reinitiation: (1) initiation: the initiation of Pol II transcription by NRs requires multiple proteins, including basal TFs and coregulators (e.g., Mediator); (2) promoter escape; (3) elongation; (4) reinitiation. Only a subset of the proteins required for initiation are required for reinitiation, including NRs, Pol II, Mediator, and some basal TFs. Factors that remain associated with the promoter after initiation (e.g., TFIIA and TFIID) form part of a ‘‘reinitiation scaffold.’’

triphosphates to initiate transcription.249–251 Sarkosyl blocks the initiation of transcription, but allows transcriptionally engaged Pol II to continue elongating, thus limiting transcription to a single round. By using this strategy, NRs and coregulators have been assessed for their contribution to initiation and reinitiation. HNF4 and VDR were shown to be required for transcription initiation with naked DNA templates.161,252 ERa was shown to have a dual role in the transcription process with chromatin templates, participating in both initiation and reinitiation.93 Different types of coregulators have different effects on initiation and reinitiation. For example, SRC proteins were shown to play a role in initiation, but have only a modest or no role in reinitiation during NR‐ dependent transcription.75,153,171 Similarly, p300 and Swi/Snf were shown to play a role in initiation, but not reinitiation.75,93,153,171 In contrast, the Mediator complex is required for both initiation and reinitiation.195 In this regard, Mediator might function as part of the reinitiation platform, remaining at the promoter for subsequent rounds of transcription. These results suggest that once histone modification and chromatin remodeling are complete, the chromatin template is competent for multiple rounds of transcription. Hence, HMEs and CRCs are only required for the first initiation event. In contrast, the recruitment and stabilization of Pol II is needed for each round of transcription, pointing to a role for NRs and Mediator in both initiation and reinitiation.

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3. KINETICS OF NR BINDING TO CHROMATIN Typical models of NR‐dependent transcription, especially those from biochemical assays, depict the NR as stably bound to its cognate DNA element in the presence of ligand. Studies in single living cells, however, have illuminated a more dynamic nature of NR association with DNA, with DNA residence times on the order of seconds. For example, GR was shown to rapidly exchange on integrated tandem arrays of the MMTV promoter in single cell experiments using fluorescence recovery after photobleaching (FRAP) analysis.253 These dynamic interactions with the DNA are not limited to GR, as ERa, PR, and androgen receptor (AR) have also been shown to have short residence times on DNA in cells.254–256 Not surprisingly, NR‐recruited coregulators such as SRC‐1, SRC‐2 (a.k.a. GRIP1), CBP, Brg1, and Brm also interact transiently with DNA in living cells, indicating that dynamic association is not limited to the DNA-binding component.256–259 Interestingly, the rapid exchange of GR that occurs in living cells has also been demonstrated in vitro with chromatin templates.260,261 The residence time of NRs on DNA directly correlates with transcriptional output and, thus, may be a mechanism by which transcriptional activity is regulated.254,262,263 The residence is regulated, in part, by the specific ligand. For example, the binding of the agonist R1881 resulted in a decrease in AR mobility relative to unliganded or antagonist (i.e., hydroxyflutamide)‐bound AR.264,265 The molecular chaperone p23 may also play a role in NR dynamics at HREs, as it was found to disassemble TRb/RXR complexes and reduce transcriptional activation with chromatin templates in vitro.266 Another molecular chaperone, hsp90, as well as Swi/Snf and subcomplexes of the proteasome, have also been shown to play a role in the dynamic association of NRs with DNA.255,256,263 These data indicate that ligand‐ and receptor‐associated proteins can regulate the residence time of NRs on DNA, which ultimately regulates transcriptional activity.

VII. Future Directions Biochemical analyses, including those using in vitro chromatin assembly and transcription assays, have been extremely useful for dissecting the molecular mechanisms of NR‐dependent transcription. These methods will continue to be useful for testing new hypotheses regarding molecular mechanisms that arise from cell, animal, and genomic studies. Future studies using these methods will not only increase our depth of knowledge, but will also expand our understanding with new fundamental concepts regarding transcriptional activation. In the following sections, we highlight some of the future directions using in vitro chromatin assembly and transcription assays.

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A. Repression of Transcription by NRs Ligand‐dependent transcriptional regulation by NRs can result in decreased, as well as increased, transcription. This has been illustrated quite dramatically in microarray expression experiments focusing on agonist ligands for several NRs in which a large fraction of the regulated genes were repressed by the ligand treatment in both cell‐based and whole animal studies.267–274 The use of in vitro chromatin assembly and transcription assays to study the agonist‐ dependent repression of transcription by NRs has been limited. This is, in part, because the low basal transcription observed with in vitro‐assembled chromatin templates is difficult to repress further. Nonetheless, aspects of the repression of transcription by NRs should be amenable to these types of biochemical analyses, and future studies will undoubtedly make use of biochemical assays for this purpose. The use of in vitro chromatin assembly and transcription assays to study NR‐dependent transcriptional repression may be best applied to corepressors that interact with agonist‐bound NRs (as opposed to those that are recruited in the presence of antagonists under conditions where transcription is strongly repressed). These include RIP140 and ligand‐dependent corepressor (LCoR), which exert their repressive effects through the recruitment of HDACs.275 Key questions include: (1) How do NRs interpret the promoter landscape and signal for activation or repression of transcription? and (2) Is the ‘‘choice’’ of repression dictated by the underlying DNA sequence, or potentially through the expression levels of various coregulators? The development of an in vitro transcription system that models NR‐dependent repression with chromatin templates would provide a useful approach for the study of these questions.

B. Single Molecule Studies Single molecule biophysical analyses have illuminated structural and functional details of nucleosome and chromatin structure, as well as the effect of enzymes that modulate this structure (i.e., Swi/Snf, PARP‐1) that were not appreciated in biochemical or cell‐based assays.143,233,276–278 For instance, AFM has been used to observe the dose‐dependent compaction of chromatin by PARP‐1.233 AFM coupled with molecular identification (i.e., the use of a scanning tip functionalized with a specific antibody, e.g., against H2A) was used to monitor the release of H2A (and presumably H2B, as well) from nucleosomes upon remodeling by Swi/Snf.143 Likewise, optical traps, which allow one to apply forces to a single molecule of DNA to stretch or unzip the DNA, have also yielded new insights. For example, optical traps have been used to map histone–DNA contacts to near base pair resolution and explore the effects of histone acetylation on the mechanical stability of the nucleosome.103,279 In addition, optical traps have been used to determine changes in nucleosome

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position (i.e., translocation) and compare histone–DNA interactions within a nucleosome before and after remodeling by Swi/Snf.277 A definitive advantage of single molecule biophysical techniques is that they allow for the study of a process as a single event, rather than an average of events across a population. The extension of these techniques to the study of questions about NR‐dependent transcription will help to provide a more complete understanding of these processes.

C. NR Biochemistry in the Postgenomic Era Genomic studies have revealed several interesting characteristics of NR molecular biology, including (1) a large percentage of genomic NR-binding sites are located distally (>1 kb from the transcription start site), (2) indirect association of NRs with DNA in a ‘‘tethering pathway,’’ and (3) an enrichment of other TF binding sites adjacent to NR binding sites (for a review see Ref. 280) (Fig. 13). However, much of the analysis has been correlative, with causative effects largely lacking. Biochemical methods, such as in vitro chromatin assembly and transcription, will be an effective approach to address the underlying mechanistic details of the observations from genomic analyses. For example, the importance of NR binding at distal sites is unclear. This mode of binding, however, suggests a need for long‐range interactions, perhaps through a chromatin looping mechanism, as suggested by studies with AR and PR281,282 (Fig. 13A). Establishing a long‐range in vitro transcription system will aid in the identification of the required DNA elements and factors for loop formation. These approaches can also be coupled with imaging techniques, such as AFM, for the visualization of NR‐dependent chromatin looping. Genomic analyses suggest that ‘‘tethering’’ of NRs to DNA through other DNA‐bound TFs is a widespread mechanism, especially with regard to ERa283,284 (Fig. 13B). Activator protein‐1 (AP‐1) and Sp1 have been shown to tether ERa to DNA, allowing for the regulation of transcription in an estrogen‐dependent manner.285,286 In vitro chromatin assembly and transcription analyses, which accurately recapitulate ERa‐dependent transcription in the tethering pathway, have demonstrated key differences between ERE‐ mediated and tethering‐mediated transcriptional regulation by ERa with respect to ligand responses and coregulator usage.287 These biochemical approaches will continue to be useful to dissect the molecular mechanisms of tethering‐mediated transcriptional regulation by NRs. Genomic analyses have identified DNA motifs enriched adjacent to particular NR‐binding sites (Fig. 13C). The factors that bind to these sites may act in conjunction with the adjacent NRs in a common transcriptional regulatory pathway. For instance, FOXA1‐binding elements are enriched near

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>1 kb HRE NR NR TF

TF

Prox. site

B NR TF

TF

Teth. site

C NR NR

Collab. factor

HRE

Adj. site

FIG. 13. Using in vitro chromatin assembly and transcription assays to study hypotheses generated from genomic analyses. (A) NR binding at distal (>1 kb) enhancers may regulate transcription by interacting with TFs bound at the proximal promoter through a looping mechanism. NR, nuclear receptor; HRE, hormone response element; TF, transcription factor; Prox. Site, TF binding site in the proximal promoter. (B) NRs may bind indirectly to DNA through interactions with other DNA‐bound TFs. For example, ERs can be tethered through members of the AP‐1 family of TFs. Teth. Site, tethering site. (C) Binding sites for other TFs are enriched adjacent to NR-binding sites, suggesting functional collaboration. For example, FOXA1 motifs are enriched near ERa-binding sites and C/EBP motifs are enriched near PPARg-binding sites. Adj. site, adjacent site; Collab. factor, collaborating factor.

ERa‐binding sites and CCAAT/enhancer‐binding protein (C/EBP) motifs are enriched near PPARg‐binding sites.288–290 Using alterations in the NR‐binding sequences, as well as differential addition of NRs, TFs, and ligands, in an in vitro transcription assay with chromatin templates, one could address the importance of motifs enriched adjacent to NR‐binding sites for NR‐dependent gene expression. Such analyses would go well beyond the bioinformatic correlations from genomic datasets. Genomic analyses have provided new insights into NR molecular biology through the identification of NR genomic binding patterns. Biochemical analyses, including in vitro chromatin assembly and transcription, will reveal the functional significance of the NR‐binding patterns identified in the genomic assays.

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VIII. Summary In vitro or ‘‘cell‐free’’ systems that accurately recapitulate transcriptional regulation with chromatin templates have allowed detailed analyses of ligand‐ dependent transcription by NRs. The wealth of information gained from these studies, as described herein, has advanced our understanding of the mechanisms of NR‐dependent gene regulation, including the role of ligands, coregulators, and nucleosome remodeling. These biochemical approaches will be very useful for addressing the underlying mechanistic details of observations made with animal models and genomic approaches.

References 1. Evans RM. The nuclear receptor superfamily: a rosetta stone for physiology. Mol Endocrinol 2005;19:1429–38. 2. Mangelsdorf DJ, Thummel C, Beato M, Herrlich P, Schutz G, Umesono K, et al. The nuclear receptor superfamily: the second decade. Cell 1995;83:835–9. 3. Jensen EV, Jordan VC. The estrogen receptor: a model for molecular medicine. Clin Cancer Res 2003;9:1980–9. 4. Pratt WB, Welsh MJ. Chaperone functions of the heat shock proteins associated with steroid receptors. Semin Cell Biol 1994;5:83–93. 5. Hollenberg SM, Weinberger C, Ong ES, Cerelli G, Oro A, Lebo R, et al. Primary structure and expression of a functional human glucocorticoid receptor cDNA. Nature 1985;318:635–41. 6. Weinberger C, Hollenberg SM, Ong ES, Harmon JM, Brower ST, Cidlowski J, et al. Identification of human glucocorticoid receptor complementary DNA clones by epitope selection. Science 1985;228:740–2. 7. Weinberger C, Hollenberg SM, Rosenfeld MG, Evans RM. Domain structure of human glucocorticoid receptor and its relationship to the v‐erb‐A oncogene product. Nature 1985;318:670–2. 8. Green S, Kumar V, Theulaz I, Wahli W, Chambon P. The N‐terminal DNA‐binding ‘‘zinc finger’’ of the oestrogen and glucocorticoid receptors determines target gene specificity. EMBO J 1988;7:3037–44. 9. Umesono K, Giguere V, Glass CK, Rosenfeld MG, Evans RM. Retinoic acid and thyroid hormone induce gene expression through a common responsive element. Nature 1988;336:262–5. 10. Naar AM, Boutin JM, Lipkin SM, Yu C, Holloway JM, Glass CK, et al. The orientation and spacing of core DNA‐binding motifs dictate selective transcriptional responses to three nuclear receptors. Cell 1991;65:1267–79. 11. Umesono K, Murakami KK, Thompson CC, Evans RM. Direct repeats as selective response elements for the thyroid hormone, retinoic acid, and vitamin D3 receptors. Cell 1991;65:1255–66. 12. Webster NJ, Green S, Jin JR, Chambon P. The hormone‐binding domains of the estrogen and glucocorticoid receptors contain an inducible transcription activation function. Cell 1988;54:199–207.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

179

13. Kumar V, Chambon P. The estrogen receptor binds tightly to its responsive element as a ligand‐induced homodimer. Cell 1988;55:145–56. 14. Leid M, Kastner P, Lyons R, Nakshatri H, Saunders M, Zacharewski T, et al. Purification, cloning, and RXR identity of the HeLa cell factor with which RAR or TR heterodimerizes to bind target sequences efficiently. Cell 1992;68:377–95. 15. Evans RM. The steroid and thyroid hormone receptor superfamily. Science 1988;240:889–95. 16. Chasse SA, Rastinejad F. Physical structure of nuclear receptor‐DNA complexes. Methods Mol Biol 2001;176:91–103. 17. Moras D, Gronemeyer H. The nuclear receptor ligand‐binding domain: structure and function. Curr Opin Cell Biol 1998;10:384–91. 18. Chandra V, Huang P, Hamuro Y, Raghuram S, Wang Y, Burris TP, et al. Structure of the intact PPAR‐gamma‐RXR‐alpha nuclear receptor complex on DNA. Nature 2008;456:350–6. 19. Orphanides G, Lagrange T, Reinberg D. The general transcription factors of RNA polymerase II. Genes Dev 1996;10:2657–83. 20. Reese JC. Basal transcription factors. Curr Opin Genet Dev 2003;13:114–8. 21. Ing NH, Beekman JM, Tsai SY, Tsai MJ, O’Malley BW. Members of the steroid hormone receptor superfamily interact with TFIIB (S300‐II). J Biol Chem 1992;267:17617–23. 22. Brou C, Wu J, Ali S, Scheer E, Lang C, Davidson I, et al. Different TBP‐associated factors are required for mediating the stimulation of transcription in vitro by the acidic transactivator GAL‐VP16 and the two nonacidic activation functions of the estrogen receptor. Nucleic Acids Res 1993;21:5–12. 23. Klein‐Hitpass L, Schwerk C, Kahmann S, Vassen L. Targets of activated steroid hormone receptors: basal transcription factors and receptor interacting proteins. J Mol Med 1998;76:490–6. 24. Beato M, Sanchez‐Pacheco A. Interaction of steroid hormone receptors with the transcription initiation complex. Endocr Rev 1996;17:587–609. 25. Albright SR, Tjian R. TAFs revisited: more data reveal new twists and confirm old ideas. Gene 2000;242:1–13. 26. McKenna NJ, Xu J, Nawaz Z, Tsai SY, Tsai MJ, O’Malley BW. Nuclear receptor coactivators: multiple enzymes, multiple complexes, multiple functions. J Steroid Biochem Mol Biol 1999;69:3–12. 27. Kininis M, Isaacs GD, Core LJ, Hah N, Kraus WL. Postrecruitment regulation of RNA polymerase II directs rapid signaling responses at the promoters of estrogen target genes. Mol Cell Biol 2009;29:1123–33. 28. Price DH. Poised polymerases: on your mark. . . get set. . . go! Mol Cell 2008;30:7–10. 29. Core LJ, Lis JT. Transcription regulation through promoter‐proximal pausing of RNA polymerase II. Science 2008;319:1791–2. 30. Le Douarin B, vom Baur E, Zechel C, Heery D, Heine M, Vivat V, et al. Ligand‐dependent interaction of nuclear receptors with potential transcriptional intermediary factors (mediators). Philos Trans R Soc Lond B Biol Sci 1996;351:569–78. 31. Shemshedini L, Ji JW, Brou C, Chambon P, Gronemeyer H. In vitro activity of the transcription activation functions of the progesterone receptor. Evidence for intermediary factors. J Biol Chem 1992;267:1834–9. 32. Halachmi S, Marden E, Martin G, MacKay H, Abbondanza C, Brown M. Estrogen receptor‐ associated proteins: possible mediators of hormone‐induced transcription. Science 1994;264:1455–8. 33. Yoshinaga SK, Peterson CL, Herskowitz I, Yamamoto KR. Roles of SWI1, SWI2, and SWI3 proteins for transcriptional enhancement by steroid receptors. Science 1992;258:1598–604. 34. Glass CK, Rosenfeld MG. The coregulator exchange in transcriptional functions of nuclear receptors. Genes Dev 2000;14:121–41.

180

RUHL AND KRAUS

35. Kraus WL, Wong J. Nuclear receptor‐dependent transcription with chromatin. Is it all about enzymes. Eur J Biochem 2002;269:2275–83. 36. Acevedo ML, Kraus WL. Transcriptional activation by nuclear receptors. Essays Biochem 2004;40:73–88. 37. Leo C, Chen JD. The SRC family of nuclear receptor coactivators. Gene 2000;245:1–11. 38. Lazar MA. Nuclear receptor corepressors. Nucl Recept Signal 2003;1:e001. 39. Aoyagi S, Trotter KW, Archer TK. ATP‐dependent chromatin remodeling complexes and their role in nuclear receptor‐dependent transcription in vivo. Vitam Horm 2005;70:281–307. 40. Malik S, Roeder RG. Transcriptional regulation through Mediator‐like coactivators in yeast and metazoan cells. Trends Biochem Sci 2000;25:277–83. 41. Rachez C, Freedman LP. Mediator complexes and transcription. Curr Opin Cell Biol 2001;13:274–80. 42. Smith CL, Hager GL. Transcriptional regulation of mammalian genes in vivo. a tale of two templates. J Biol Chem 1997;272:27493–6. 43. Bagchi MK, Tsai SY, Weigel NL, Tsai MJ, O’Malley BW. Regulation of in vitro transcription by progesterone receptor. Characterization and kinetic studies. J Biol Chem 1990;265:5129–34. 44. De Vos P, Schmitt J, Verhoeven G, Stunnenberg HG. Human androgen receptor expressed in HeLa cells activates transcription in vitro. Nucleic Acids Res 1994;22:1161–6. 45. Elliston JF, Fawell SE, Klein‐Hitpass L, Tsai SY, Tsai MJ, Parker MG, et al. Mechanism of estrogen receptor‐dependent transcription in a cell‐free system. Mol Cell Biol 1990; 10:6607–12. 46. Klein‐Hitpass L, Tsai SY, Weigel NL, Allan GF, Riley D, Rodriguez R, et al. The progesterone receptor stimulates cell‐free transcription by enhancing the formation of a stable preinitiation complex. Cell 1990;60:247–57. 47. Tsai SY, Srinivasan G, Allan GF, Thompson EB, O’Malley BW, Tsai MJ. Recombinant human glucocorticoid receptor induces transcription of hormone response genes in vitro. J Biol Chem 1990;265:17055–61. 48. Schmitt J, Stunnenberg HG. The glucocorticoid receptor hormone binding domain mediates transcriptional activation in vitro in the absence of ligand. Nucleic Acids Res 1993;21:2673–81. 49. Bagchi MK, Tsai SY, Tsai MJ, O’Malley BW. Identification of a functional intermediate in receptor activation in progesterone‐dependent cell‐free transcription. Nature 1990;345: 547–50. 50. Beekman JM, Allan GF, Tsai SY, Tsai MJ, O’Malley BW. Transcriptional activation by the estrogen receptor requires a conformational change in the ligand binding domain. Mol Endocrinol 1993;7:1266–74. 51. Elliston JF, Beekman JM, Tsai SY, O’Malley BW, Tsai MJ. Hormone activation of baculovirus expressed progesterone receptors. J Biol Chem 1992;267:5193–8. 52. Schild C, Claret FX, Wahli W, Wolffe AP. A nucleosome‐dependent static loop potentiates estrogen‐regulated transcription from the Xenopus vitellogenin B1 promoter in vitro. EMBO J 1993;12:423–33. 53. Bulger M, Kadonaga JT. Biochemical reconstitution of chromatin with physiological nucleosome spacing. Methods Mol Genet 1994;5:241–62. 54. Kamakaka RT, Bulger M, Kadonaga JT. Potentiation of RNA polymerase II transcription by Gal4‐VP16 during but not after DNA replication and chromatin assembly. Genes Dev 1993;7:1779–95. 55. Kamakaka RT, Kadonaga JT. Biochemical analysis of the role of chromatin structure in the regulation of transcription by RNA polymerase II. Cold Spring Harb Symp Quant Biol 1993;58:205–12. 56. Kraus WL, Kadonaga JT. Ligand‐ and cofactor‐regulated transcription with chromatin templates. In: Picard D, editor. Steroid/nuclear receptor superfamily: a practical approach. Oxford/New York: Oxford University Press; 1999. p. 167–89.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

181

57. Morales V, Giamarchi C, Chailleux C, Moro F, Marsaud V, Le Ricousse S, et al. Chromatin structure and dynamics: functional implications. Biochimie 2001;83:1029–39. 58. Luger K, Mader AW, Richmond RK, Sargent DF, Richmond TJ. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 1997;389:251–60. 59. Horn PJ, Peterson CL. Molecular biology. Chromatin higher order folding–wrapping up transcription. Science 2002;297:1824–7. 60. Quina AS, Buschbeck M, Di Croce L. Chromatin structure and epigenetics. Biochem Pharmacol 2006;72:1563–9. 61. McBryant SJ, Adams VH, Hansen JC. Chromatin architectural proteins. Chromosome Res 2006;14:39–51. 62. Robinson PJ, Rhodes D. Structure of the ‘‘30 nm’’ chromatin fibre: a key role for the linker histone. Curr Opin Struct Biol 2006;16:336–43. 63. Richard‐Foy H, Hager GL. Sequence‐specific positioning of nucleosomes over the steroid‐ inducible MMTV promoter. EMBO J 1987;6:2321–8. 64. Zaret KS, Yamamoto KR. Reversible and persistent changes in chromatin structure accompany activation of a glucocorticoid‐dependent enhancer element. Cell 1984;38:29–38. 65. Jantzen K, Fritton HP, Igo‐Kemenes T, Espel E, Janich S, Cato AC, et al. Partial overlapping of binding sequences for steroid hormone receptors and DNaseI hypersensitive sites in the rabbit uteroglobin gene region. Nucleic Acids Res 1987;15:4535–52. 66. Archer TK, Fryer CJ, Lee HL, Zaniewski E, Liang T, Mymryk JS. Steroid hormone receptor status defines the MMTV promoter chromatin structure in vivo. J Steroid Biochem Mol Biol 1995;53:421–9. 67. Hager GL. MMTV as a model for gene expression in mammary tissue. Cancer Treat Res 1988;40:267–81. 68. Archer TK, Lee HL, Cordingley MG, Mymryk JS, Fragoso G, Berard DS, et al. Differential steroid hormone induction of transcription from the mouse mammary tumor virus promoter. Mol Endocrinol 1994;8:568–76. 69. Smith CL, Htun H, Wolford RG, Hager GL. Differential activity of progesterone and glucocorticoid receptors on mouse mammary tumor virus templates differing in chromatin structure. J Biol Chem 1997;272:14227–35. 70. Wong J, Shi YB, Wolffe AP. A role for nucleosome assembly in both silencing and activation of the Xenopus TR beta A gene by the thyroid hormone receptor. Genes Dev 1995;9:2696–711. 71. Wong J, Shi YB, Wolffe AP. Determinants of chromatin disruption and transcriptional regulation instigated by the thyroid hormone receptor: hormone‐regulated chromatin disruption is not sufficient for transcriptional activation. EMBO J 1997;16:3158–71. 72. Cairns BR, Levinson RS, Yamamoto KR, Kornberg RD. Essential role of Swp73p in the function of yeast Swi/Snf complex. Genes Dev 1996;10:2131–44. 73. Chiba H, Muramatsu M, Nomoto A, Kato H. Two human homologues of Saccharomyces cerevisiae SWI2/SNF2 and Drosophila brahma are transcriptional coactivators cooperating with the estrogen receptor and the retinoic acid receptor. Nucleic Acids Res 1994;22:1815–20. 74. Muchardt C, Yaniv M. A human homologue of Saccharomyces cerevisiae SNF2/SWI2 and Drosophila brm genes potentiates transcriptional activation by the glucocorticoid receptor. EMBO J 1993;12:4279–90. 75. Dilworth FJ, Fromental‐Ramain C, Yamamoto K, Chambon P. ATP‐driven chromatin remodeling activity and histone acetyltransferases act sequentially during transactivation by RAR/ RXR in vitro. Mol Cell 2000;6:1049–58. 76. DiRenzo J, Shang Y, Phelan M, Sif S, Myers M, Kingston R, Brown M. BRG‐1 is recruited to estrogen‐responsive promoters and cooperates with factors involved in histone acetylation. Mol Cell Biol 2000;20:7541–9.

182

RUHL AND KRAUS

77. Fryer CJ, Archer TK. Chromatin remodelling by the glucocorticoid receptor requires the BRG1 complex. Nature 1998;393:88–91. 78. Kadam S, Emerson BM. Transcriptional specificity of human SWI/SNF BRG1 and BRM chromatin remodeling complexes. Mol Cell 2003;11:377–89. 79. Nagaich AK, Walker DA, Wolford R, Hager GL. Rapid periodic binding and displacement of the glucocorticoid receptor during chromatin remodeling. Mol Cell 2004;14:163–74. 80. Hsiao PW, Fryer CJ, Trotter KW, Wang W, Archer TK. BAF60a mediates critical interactions between nuclear receptors and the BRG1 chromatin‐remodeling complex for transactivation. Mol Cell Biol 2003;23:6210–20. 81. Lemon B, Inouye C, King DS, Tjian R. Selectivity of chromatin‐remodelling cofactors for ligand‐activated transcription. Nature 2001;414:924–8. 82. Belandia B, Orford RL, Hurst HC, Parker MG. Targeting of SWI/SNF chromatin remodelling complexes to estrogen‐responsive genes. EMBO J 2002;21:4094–103. 83. Metivier R, Penot G, Carmouche RP, Hubner MR, Reid G, Denger S, et al. Transcriptional complexes engaged by apo‐estrogen receptor‐alpha isoforms have divergent outcomes. EMBO J 2004;23:3653–66. 84. Reines D, Dvir A, Conaway JW, Conaway RC. Assays for investigating transcription by RNA polymerase II in vitro. Methods 1997;12:192–202. 85. Ge H, Martinez E, Chiang CM, Roeder RG. Activator‐dependent transcription by mammalian RNA polymerase II: in vitro reconstitution with general transcription factors and cofactors. Methods Enzymol 1996;274:57–71. 86. Dignam JD, Lebovitz RM, Roeder RG. Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res 1983;11:1475–89. 87. Roeder RG. Multiple forms of deoxyribonucleic acid‐dependent ribonucleic acid polymerase in Xenopus laevis. Isolation and partial characterization. J Biol Chem 1974;249:241–8. 88. Schwartz LB, Sklar VE, Jaehning JA, Weinmann R, Roeder RG. Isolation and partial characterization of the multiple forms of deoxyribonucleic acid‐dependent ribonucleic acid polymerase in the mouse myeloma, MOPC 315. J Biol Chem 1974;249:5889–97. 89. Sklar VE, Schwartz LB, Roeder RG. Distinct molecular structures of nuclear class I, II, and III DNA‐dependent RNA polymerases. Proc Natl Acad Sci USA 1975;72:348–52. 90. Maldonado E, Drapkin R, Reinberg D. Purification of human RNA polymerase II and general transcription factors. Methods Enzymol 1996;274:72–100. 91. Sawadogo M, Roeder RG. Factors involved in specific transcription by human RNA polymerase II: analysis by a rapid and quantitative in vitro assay. Proc Natl Acad Sci USA 1985;82:4394–8. 92. Kamakaka RT, Kraus WL. In vitro transcription. Curr Protoc Cell Biol 2001;11:16. 93. Kraus WL, Kadonaga JT. p300 and estrogen receptor cooperatively activate transcription via differential enhancement of initiation and reinitiation. Genes Dev 1998;12:331–42. 94. Thackray VG, Nordeen SK. High‐yield purification of functional, full‐length steroid receptor coactivator 1 expressed in insect cells. Biotechniques 2002;32(260):262–3. 95. Simpson RT, Stafford DW. Structural features of a phased nucleosome core particle. Proc Natl Acad Sci USA 1983;80:51–5. 96. Simpson RT, Thoma F, Brubaker JM. Chromatin reconstituted from tandemly repeated cloned DNA fragments and core histones: a model system for study of higher order structure. Cell 1985;42:799–808. 97. Lowary PT, Widom J. New DNA sequence rules for high affinity binding to histone octamer and sequence‐directed nucleosome positioning. J Mol Biol 1998;276:19–42. 98. Brandt WF, Patterson K, von Holt C. The histones of yeast. The isolation and partial structure of the core histones. Eur J Biochem 1980;110:67–76.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

183

99. Prevelige PE, Jr., Fasman GD. Structural studies of acetylated and control inner core histones. Biochemistry 1987;26:2944–55. 100. Yager TD, McMurray CT, van Holde KE. Salt‐induced release of DNA from nucleosome core particles. Biochemistry 1989;28:2271–81. 101. Levenstein ME, Kadonaga JT. Biochemical analysis of chromatin containing recombinant Drosophila core histones. J Biol Chem 2002;277:8749–54. 102. Luger K, Rechsteiner TJ, Flaus AJ, Waye MM, Richmond TJ. Characterization of nucleosome core particles containing histone proteins made in bacteria. J Mol Biol 1997;272:301–11. 103. Brower‐Toland B, Wacker DA, Fulbright RM, Lis JT, Kraus WL, Wang MD. Specific contributions of histone tails and their acetylation to the mechanical stability of nucleosomes. J Mol Biol 2005;346:135–46. 104. Parsons XH, Garcia SN, Pillus L, Kadonaga JT. Histone deacetylation by Sir2 generates a transcriptionally repressed nucleoprotein complex. Proc Natl Acad Sci USA 2003;100:1609–14. 105. Shogren‐Knaak MA, Fry CJ, Peterson CL. A native peptide ligation strategy for deciphering nucleosomal histone modifications. J Biol Chem 2003;278:15744–8. 106. An W, Kim J, Roeder RG. Ordered cooperative functions of PRMT1, p300, and CARM1 in transcriptional activation by p53. Cell 2004;117:735–48. 107. Angelov D, Verdel A, An W, Bondarenko V, Hans F, Doyen CM, et al. SWI/SNF remodeling and p300‐dependent transcription of histone variant H2ABbd nucleosomal arrays. EMBO J 2004;23:3815–24. 108. Ohlenbusch HH, Olivera BM, Tuan D, Davidson N. Selective dissociation of histones from calf thymus nucleoprotein. J Mol Biol 1967;25:299–315. 109. Oudet P, Gross‐Bellard M, Chambon P. Electron microscopic and biochemical evidence that chromatin structure is a repeating unit. Cell 1975;4:281–300. 110. Richards BM, Pardon JF. The molecular structure of nucleohistone (DNH). Exp Cell Res 1970;62:184–96. 111. Dyer PN, Edayathumangalam RS, White CL, Bao Y, Chakravarthy S, Muthurajan UM, et al. Reconstitution of nucleosome core particles from recombinant histones and DNA. Methods Enzymol 2004;375:23–44. 112. Laskey RA, Mills AD, Morris NR. Assembly of SV40 chromatin in a cell‐free system from Xenopus eggs. Cell 1977;10:237–43. 113. Rodriguez‐Campos A, Shimamura A, Worcel A. Assembly and properties of chromatin containing histone H1. J Mol Biol 1989;209:135–50. 114. Shimamura A, Tremethick D, Worcel A. Characterization of the repressed 5S DNA minichromosomes assembled in vitro with a high‐speed supernatant of Xenopus laevis oocytes. Mol Cell Biol 1988;8:4257–69. 115. Becker PB, Wu C. Cell‐free system for assembly of transcriptionally repressed chromatin from Drosophila embryos. Mol Cell Biol 1992;12:2241–9. 116. Banerjee S, Cantor CR. Nucleosome assembly of simian virus 40 DNA in a mammalian cell extract. Mol Cell Biol 1990;10:2863–73. 117. Gruss C, Gutierrez C, Burhans WC, DePamphilis ML, Koller T, Sogo JM. Nucleosome assembly in mammalian cell extracts before and after DNA replication. EMBO J 1990;9:2911–22. 118. Glikin GC, Ruberti I, Worcel A. Chromatin assembly in Xenopus oocytes: in vitro studies. Cell 1984;37:33–41. 119. Adamson ED, Woodland HR. Histone synthesis in early amphibian development: histone and DNA syntheses are not co‐ordinated. J Mol Biol 1974;88:263–85. 120. Shechter D, Nicklay JJ, Chitta RK, Shabanowitz J, Hunt DF, Allis CD. Analysis of Histones in Xenopus laevis: I. a distinct index of enriched variants and modifications exists in each cell type and is remodeled during developmental transitions. J Biol Chem 2009;284:1064–74.

184

RUHL AND KRAUS

121. Nelson T, Hsieh TS, Brutlag D. Extracts of Drosophila embryos mediate chromatin assembly in vitro. Proc Natl Acad Sci USA 1979;76:5510–4. 122. Kleinschmidt JA, Franke WW. Soluble acidic complexes containing histones H3 and H4 in nuclei of Xenopus laevis oocytes. Cell 1982;29:799–809. 123. Laskey RA, Honda BM, Mills AD, Finch JT. Nucleosomes are assembled by an acidic protein which binds histones and transfers them to DNA. Nature 1978;275:416–20. 124. Ito T, Bulger M, Pazin MJ, Kobayashi R, Kadonaga JT. ACF, an ISWI‐containing and ATP‐utilizing chromatin assembly and remodeling factor. Cell 1997;90:145–55. 125. Bulger M, Ito T, Kamakaka RT, Kadonaga JT. Assembly of regularly spaced nucleosome arrays by Drosophila chromatin assembly factor 1 and a 56‐kDa histone‐binding protein. Proc Natl Acad Sci USA 1995;92:11726–30. 126. Kamakaka RT, Bulger M, Kaufman PD, Stillman B, Kadonaga JT. Postreplicative chromatin assembly by Drosophila and human chromatin assembly factor 1. Mol Cell Biol 1996;16:810–7. 127. Smith S, Stillman B. Purification and characterization of CAF‐I, a human cell factor required for chromatin assembly during DNA replication in vitro. Cell 1989;58:15–25. 128. Dirscherl SS, Krebs JE. Functional diversity of ISWI complexes. Biochem Cell Biol 2004;82:482–9. 129. Loyola A, LeRoy G, Wang YH, Reinberg D. Reconstitution of recombinant chromatin establishes a requirement for histone‐tail modifications during chromatin assembly and transcription. Genes Dev 2001;15:2837–51. 130. Lutter LC. Precise location of DNase I cutting sites in the nucleosome core determined by high resolution gel electrophoresis. Nucleic Acids Res 1979;6:41–56. 131. Noll M. Internal structure of the chromatin subunit. Nucleic Acids Res 1974;1:1573–8. 132. Prunell A, Kornberg RD, Lutter L, Klug A, Levitt M, Crick FH. Periodicity of deoxyribonuclease I digestion of chromatin. Science 1979;204:855–8. 133. Noll M. Subunit structure of chromatin. Nature 1974;251:249–51. 134. Hewish DR, Burgoyne LA. Chromatin sub‐structure. The digestion of chromatin DNA at regularly spaced sites by a nuclear deoxyribonuclease. Biochem Biophys Res Commun 1973;52:504–10. 135. Pazin MJ, Kamakaka RT, Kadonaga JT. ATP‐dependent nucleosome reconfiguration and transcriptional activation from preassembled chromatin templates. Science 1994;266:2007–11. 136. Germond JE, Hirt B, Oudet P, Gross‐Bellark M, Chambon P. Folding of the DNA double helix in chromatin‐like structures from simian virus 40. Proc Natl Acad Sci USA 1975;72: 1843–7. 137. Kim MY, Mauro S, Gevry N, Lis JT, Kraus WL. NADþ‐dependent modulation of chromatin structure and transcription by nucleosome binding properties of PARP‐1. Cell 2004;119: 803–14. 138. Binnig G, Quate CF, Gerber C. Atomic force microscope. Phys Rev Lett 1986;56:930–3. 139. Lin L, Hom D, Lindsay SM, Chaput JC. In vitro selection of histone H4 aptamers for recognition imaging microscopy. J Am Chem Soc 2007;129:14568–9. 140. Lohr D, Wang H, Bash R, Lindsay SM. Recognition imaging of chromatin and chromatin‐ remodeling complexes in the atomic force microscope. Methods Mol Biol 2009;464:123–38. 141. Stroh C, Wang H, Bash R, Ashcroft B, Nelson J, Gruber H, et al. Single‐molecule recognition imaging microscopy. Proc Natl Acad Sci USA 2004;101:12503–7. 142. Raab A, Han W, Badt D, Smith‐Gill SJ, Lindsay SM, Schindler H, et al. Antibody recognition imaging by force microscopy. Nat Biotechnol 1999;17:901–5. 143. Bash R, Wang H, Anderson C, Yodh J, Hager G, Lindsay SM, et al. AFM imaging of protein movements: histone H2A‐H2B release during nucleosome remodeling. FEBS Lett 2006;580:4757–61.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

185

144. An W, Roeder RG. Reconstitution and transcriptional analysis of chromatin in vitro. Methods Enzymol 2004;377:460–74. 145. Loyola A, He S, Oh S, McCafferty DG, Reinberg D. Techniques used to study transcription on chromatin templates. Methods Enzymol 2004;377:474–99. 146. Pazin MJ, Kadonaga JT. Transcriptional and structural analysis of chromatin assembled in vitro. In: Gould H, editor. Chromatin: a practical approach. Oxford/New York: Oxford University Press; 1998. p. 173–94. 147. Suen CS, Chin WW. Ligand‐dependent, Pit‐1/growth hormone factor‐1 (GHF‐1)‐independent transcriptional stimulation of rat growth hormone gene expression by thyroid hormone receptors in vitro. Mol Cell Biol 1993;13:1719–27. 148. Corthesy B, Hipskind R, Theulaz I, Wahli W. Estrogen‐dependent in vitro transcription from the vitellogenin promoter in liver nuclear extracts. Science 1988;239:1137–9. 149. Rachez C, Lemon BD, Suldan Z, Bromleigh V, Gamble M, Naar AM, et al. Ligand‐dependent transcription activation by nuclear receptors requires the DRIP complex. Nature 1999;398:824–8. 150. Liu Z, Wong J, Tsai SY, Tsai MJ, O’Malley BW. Steroid receptor coactivator‐1 (SRC‐1) enhances ligand‐dependent and receptor‐dependent cell‐free transcription of chromatin. Proc Natl Acad Sci USA 1999;96:9485–90. 151. Lee KC, Li J, Cole PA, Wong J, Kraus WL. Transcriptional activation by thyroid hormone receptor‐beta involves chromatin remodeling, histone acetylation, and synergistic stimulation by p300 and steroid receptor coactivators. Mol Endocrinol 2003;17:908–22. 152. Dilworth FJ, Fromental‐Ramain C, Remboutsika E, Benecke A, Chambon P. Ligand‐dependent activation of transcription in vitro by retinoic acid receptor alpha/retinoid X receptor alpha heterodimers that mimics transactivation by retinoids in vivo. Proc Natl Acad Sci USA 1999;96:1995–2000. 153. Kim MY, Hsiao SJ, Kraus WL. A role for coactivators and histone acetylation in estrogen receptor alpha‐mediated transcription initiation. EMBO J 2001;20:6084–94. 154. Kraus WL, Manning ET, Kadonaga JT. Biochemical analysis of distinct activation functions in p300 that enhance transcription initiation with chromatin templates. Mol Cell Biol 1999;19:8123–35. 155. Jacq X, Brou C, Lutz Y, Davidson I, Chambon P, Tora L. Human TAFII30 is present in a distinct TFIID complex and is required for transcriptional activation by the estrogen receptor. Cell 1994;79:107–17. 156. Moqtaderi Z, Bai Y, Poon D, Weil PA, Struhl K. TBP‐associated factors are not generally required for transcriptional activation in yeast. Nature 1996;383:188–91. 157. Wu SY, Kershnar E, Chiang CM. TAFII‐independent activation mediated by human TBP in the presence of the positive cofactor PC4. EMBO J 1998;17:4478–90. 158. Fondell JD, Guermah M, Malik S, Roeder RG. Thyroid hormone receptor‐associated proteins and general positive cofactors mediate thyroid hormone receptor function in the absence of the TATA box‐binding protein‐associated factors of TFIID. Proc Natl Acad Sci USA 1999;96:1959–64. 159. Wu SY, Thomas MC, Hou SY, Likhite V, Chiang CM. Isolation of mouse TFIID and functional characterization of TBP and TFIID in mediating estrogen receptor and chromatin transcription. J Biol Chem 1999;274:23480–90. 160. Deng W, Roberts SG. TFIIB and the regulation of transcription by RNA polymerase II. Chromosoma 2007;116:417–29. 161. Malik S, Karathanasis SK. TFIIB‐directed transcriptional activation by the orphan nuclear receptor hepatocyte nuclear factor 4. Mol Cell Biol 1996;16:1824–31. 162. Blanco JC, Wang IM, Tsai SY, Tsai MJ, O’Malley BW, Jurutka PW, et al. Transcription factor TFIIB and the vitamin D receptor cooperatively activate ligand‐dependent transcription. Proc Natl Acad Sci USA 1995;92:1535–9.

186

RUHL AND KRAUS

163. Masuyama H, Jefcoat SC, Jr., MacDonald PN. The N‐terminal domain of transcription factor IIB is required for direct interaction with the vitamin D receptor and participates in vitamin D‐mediated transcription. Mol Endocrinol 1997;11:218–28. 164. Baniahmad A, Ha I, Reinberg D, Tsai S, Tsai MJ, O’Malley BW. Interaction of human thyroid hormone receptor beta with transcription factor TFIIB may mediate target gene derepression and activation by thyroid hormone. Proc Natl Acad Sci USA 1993;90:8832–6. 165. Ikeda M, Kawaguchi A, Takeshita A, Chin WW, Endo T, Onaya T. CBP‐dependent and independent enhancing activity of steroid receptor coactivator‐1 in thyroid hormone receptor‐mediated transactivation. Mol Cell Endocrinol 1999;147:103–12. 166. Takeshita A, Yen PM, Misiti S, Cardona GR, Liu Y, Chin WW. Molecular cloning and properties of a full‐length putative thyroid hormone receptor coactivator. Endocrinology 1996;137:3594–7. 167. Chen H, Lin RJ, Schiltz RL, Chakravarti D, Nash A, Nagy L, et al. Nuclear receptor coactivator ACTR is a novel histone acetyltransferase and forms a multimeric activation complex with P/CAF and CBP/p300. Cell 1997;90:569–80. 168. Spencer TE, Jenster G, Burcin MM, Allis CD, Zhou J, Mizzen CA, et al. Steroid receptor coactivator‐1 is a histone acetyltransferase. Nature 1997;389:194–8. 169. Sheppard HM, Harries JC, Hussain S, Bevan C, Heery DM. Analysis of the steroid receptor coactivator 1 (SRC1)‐CREB binding protein interaction interface and its importance for the function of SRC1. Mol Cell Biol 2001;21:39–50. 170. Voegel JJ, Heine MJ, Tini M, Vivat V, Chambon P, Gronemeyer H. The coactivator TIF2 contains three nuclear receptor‐binding motifs and mediates transactivation through CBP binding‐dependent and ‐independent pathways. EMBO J 1998;17:507–19. 171. Liu Z, Wong J, Tsai SY, Tsai MJ, O’Malley BW. Sequential recruitment of steroid receptor coactivator‐1 (SRC‐1) and p300 enhances progesterone receptor‐dependent initiation and reinitiation of transcription from chromatin. Proc Natl Acad Sci USA 2001;98:12426–31. 172. Kamei Y, Xu L, Heinzel T, Torchia J, Kurokawa R, Gloss B, et al. A CBP integrator complex mediates transcriptional activation and AP‐1 inhibition by nuclear receptors. Cell 1996;85:403–14. 173. Chakravarti D, LaMorte VJ, Nelson MC, Nakajima T, Schulman G, Juguilon H, et al. Role of CBP/P300 in nuclear receptor signalling. Nature 1996;383:99–103. 174. Jiang W, Nordeen SK, Kadonaga JT. Transcriptional analysis of chromatin assembled with purified ACF and dNAP1 reveals that acetyl‐CoA is required for preinitiation complex assembly. J Biol Chem 2000;275:39819–22. 175. Fu M, Rao M, Wang C, Sakamaki T, Wang J, Di Vizio D, et al. Acetylation of androgen receptor enhances coactivator binding and promotes prostate cancer cell growth. Mol Cell Biol 2003;23:8563–75. 176. Fu M, Wang C, Wang J, Zhang X, Sakamaki T, Yeung YG, et al. Androgen receptor acetylation governs trans activation and MEKK1‐induced apoptosis without affecting in vitro sumoylation and trans‐repression function. Mol Cell Biol 2002;22:3373–88. 177. Kim MY, Woo EM, Chong YT, Homenko DR, Kraus WL. Acetylation of estrogen receptor alpha by p300 at lysines 266 and 268 enhances the deoxyribonucleic acid binding and transactivation activities of the receptor. Mol Endocrinol 2006;20:1479–93. 178. Lin HY, Hopkins R, Cao HJ, Tang HY, Alexander C, Davis FB, et al. Acetylation of nuclear hormone receptor superfamily members: thyroid hormone causes acetylation of its own receptor by a mitogen‐activated protein kinase‐dependent mechanism. Steroids 2005;70:444–9. 179. Wang C, Fu M, Angeletti RH, Siconolfi‐Baez L, Reutens AT, Albanese C, et al. Direct acetylation of the estrogen receptor alpha hinge region by p300 regulates transactivation and hormone sensitivity. J Biol Chem 2001;276:18375–83.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

187

180. Xu W, Chen H, Du K, Asahara H, Tini M, Emerson BM, et al. A transcriptional switch mediated by cofactor methylation. Science 2001;294:2507–11. 181. Barrero MJ, Malik S. Two functional modes of a nuclear receptor‐recruited arginine methyltransferase in transcriptional activation. Mol Cell 2006;24:233–43. 182. Xu W, Cho H, Kadam S, Banayo EM, Anderson S, Yates JR, III, et al. A methylation‐mediator complex in hormone signaling. Genes Dev 2004;18:144–56. 183. Vicent GP, Ballare C, Nacht AS, Clausell J, Subtil‐Rodriguez A, Quiles I, et al. Induction of progesterone target genes requires activation of Erk and Msk kinases and phosphorylation of histone H3. Mol Cell 2006;24:367–81. 184. Lusser A, Kadonaga JT. Chromatin remodeling by ATP‐dependent molecular machines. Bioessays 2003;25:1192–200. 185. Schnitzler GR. Control of nucleosome positions by DNA sequence and remodeling machines. Cell Biochem Biophys 2008;51:67–80. 186. Sif S. ATP‐dependent nucleosome remodeling complexes: enzymes tailored to deal with chromatin. J Cell Biochem 2004;91:1087–98. 187. Di Croce L, Koop R, Venditti P, Westphal HM, Nightingale KP, Corona DF, et al. Two‐step synergism between the progesterone receptor and the DNA‐binding domain of nuclear factor 1 on MMTV minichromosomes. Mol Cell 1999;4:45–54. 188. Lewis BA, Reinberg D. The mediator coactivator complex: functional and physical roles in transcriptional regulation. J Cell Sci 2003;116:3667–75. 189. Fondell JD, Ge H, Roeder RG. Ligand induction of a transcriptionally active thyroid hormone receptor coactivator complex. Proc Natl Acad Sci USA 1996;93:8329–33. 190. Rachez C, Suldan Z, Ward J, Chang CP, Burakov D, Erdjument‐Bromage H, et al. A novel protein complex that interacts with the vitamin D3 receptor in a ligand‐dependent manner and enhances VDR transactivation in a cell‐free system. Genes Dev 1998;12:1787–800. 191. Malik S, Guermah M, Yuan CX, Wu W, Yamamura S, Roeder RG. Structural and functional organization of TRAP220, the TRAP/mediator subunit that is targeted by nuclear receptors. Mol Cell Biol 2004;24:8244–54. 192. Acevedo ML, Lee KC, Stender JD, Katzenellenbogen BS, Kraus WL. Selective recognition of distinct classes of coactivators by a ligand‐inducible activation domain. Mol Cell 2004;13: 725–38. 193. Taatjes DJ, Schneider‐Poetsch T, Tjian R. Distinct conformational states of nuclear receptor‐ bound CRSP‐Med complexes. Nat Struct Mol Biol 2004;11:664–71. 194. Kang YK, Guermah M, Yuan CX, Roeder RG. The TRAP/Mediator coactivator complex interacts directly with estrogen receptors alpha and beta through the TRAP220 subunit and directly enhances estrogen receptor function in vitro. Proc Natl Acad Sci USA 2002;99: 2642–7. 195. Acevedo ML, Kraus WL. Mediator and p300/CBP‐steroid receptor coactivator complexes have distinct roles, but function synergistically, during estrogen receptor alpha‐dependent transcription with chromatin templates. Mol Cell Biol 2003;23:335–48. 196. Wallberg AE, Yamamura S, Malik S, Spiegelman BM, Roeder RG. Coordination of p300‐ mediated chromatin remodeling and TRAP/mediator function through coactivator PGC‐ 1alpha. Mol Cell 2003;12:1137–49. 197. Malik S, Wallberg AE, Kang YK, Roeder RG. TRAP/SMCC/mediator‐dependent transcriptional activation from DNA and chromatin templates by orphan nuclear receptor hepatocyte nuclear factor 4. Mol Cell Biol 2002;22:5626–37. 198. Pavri R, Lewis B, Kim TK, Dilworth FJ, Erdjument‐Bromage H, Tempst P, et al. PARP‐1 determines specificity in a retinoid signaling pathway via direct modulation of mediator. Mol Cell 2005;18:83–96.

188

RUHL AND KRAUS

199. Kim MY, Zhang T, Kraus WL. Poly(ADP‐ribosyl)ation by PARP‐1: ‘‘PAR‐laying’’ NADþ into a nuclear signal. Genes Dev 2005;19:1951–67. 200. Krishnakumar R, Gamble MJ, Frizzell KM, Berrocal JG, Kininis M, Kraus WL. Reciprocal binding of PARP‐1 and histone H1 at promoters specifies transcriptional outcomes. Science 2008;319:819–21. 201. Kraus WL. Transcriptional control by PARP‐1: chromatin modulation, enhancer‐binding, coregulation, and insulation. Curr Opin Cell Biol 2008;20:294–302. 202. Kraus WL, Lis JT. PARP goes transcription. Cell 2003;113:677–83. 203. Ju BG, Lunyak VV, Perissi V, Garcia‐Bassets I, Rose DW, Glass CK, et al. A topoisomerase IIbeta‐ mediated dsDNA break required for regulated transcription. Science 2006;312:1798–802. 204. Ohkura N, Nagamura Y, Tsukada T. Differential transactivation by orphan nuclear receptor NOR1 and its fusion gene product EWS/NOR1: possible involvement of poly(ADP‐ribose) polymerase I, PARP‐1. J Cell Biochem 2008;105:785–800. 205. Huang D, Yang C, Wang Y, Liao Y, Huang K. PARP‐1 suppresses adiponectin expression through poly(ADP‐ribosyl)ation of PPAR gamma in cardiac fibroblasts. Cardiovasc Res 2009;81:98–107. 206. Miyamoto T, Kakizawa T, Hashizume K. Inhibition of nuclear receptor signalling by poly (ADP‐ribose) polymerase. Mol Cell Biol 1999;19:2644–9. 207. Heinzel T, Lavinsky RM, Mullen TM, Soderstrom M, Laherty CD, Torchia J, et al. A complex containing N‐CoR, mSin3 and histone deacetylase mediates transcriptional repression. Nature 1997;387:43–8. 208. Huang EY, Zhang J, Miska EA, Guenther MG, Kouzarides T, Lazar MA. Nuclear receptor corepressors partner with class II histone deacetylases in a Sin3‐independent repression pathway. Genes Dev 2000;14:45–54. 209. Kao HY, Downes M, Ordentlich P, Evans RM. Isolation of a novel histone deacetylase reveals that class I and class II deacetylases promote SMRT‐mediated repression. Genes Dev 2000;14:55–66. 210. Li J, Wang J, Wang J, Nawaz Z, Liu JM, Qin J, et al. Both corepressor proteins SMRT and N‐CoR exist in large protein complexes containing HDAC3. EMBO J 2000;19:4342–50. 211. Nagy L, Kao HY, Chakravarti D, Lin RJ, Hassig CA, Ayer DE, et al. Nuclear receptor repression mediated by a complex containing SMRT, mSin3A, and histone deacetylase. Cell 1997;89:373–80. 212. Tong JK, Hassig CA, Schnitzler GR, Kingston RE, Schreiber SL. Chromatin deacetylation by an ATP‐dependent nucleosome remodelling complex. Nature 1998;395:917–21. 213. Wade PA, Gegonne A, Jones PL, Ballestar E, Aubry F, Wolffe AP. Mi‐2 complex couples DNA methylation to chromatin remodelling and histone deacetylation. Nat Genet 1999;23:62–6. 214. Xue Y, Wong J, Moreno GT, Young MK, Cote J, Wang W. NURD, a novel complex with both ATP‐dependent chromatin‐remodeling and histone deacetylase activities. Mol Cell 1998;2: 851–61. 215. Zhang Y, LeRoy G, Seelig HP, Lane WS, Reinberg D. The dermatomyositis‐specific autoantigen Mi2 is a component of a complex containing histone deacetylase and nucleosome remodeling activities. Cell 1998;95:279–89. 216. Underhill C, Qutob MS, Yee SP, Torchia J. A novel nuclear receptor corepressor complex, N‐CoR, contains components of the mammalian SWI/SNF complex and the corepressor KAP‐1. J Biol Chem 2000;275:40463–70. 217. McKenna NJ, Lanz RB, O’Malley BW. Nuclear receptor coregulators: cellular and molecular biology. Endocr Rev 1999;20:321–44. 218. Rosenfeld MG, Lunyak VV, Glass CK. Sensors and signals: a coactivator/corepressor/epigenetic code for integrating signal‐dependent programs of transcriptional response. Genes Dev 2006;20:1405–28.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

189

219. Smith CL, O’Malley BW. Coregulator function: a key to understanding tissue specificity of selective receptor modulators. Endocr Rev 2004;25:45–71. 220. Shang Y, Brown M. Molecular determinants for the tissue specificity of SERMs. Science 2002;295:2465–8. 221. Liu Z, Auboeuf D, Wong J, Chen JD, Tsai SY, Tsai MJ, et al. Coactivator/corepressor ratios modulate PR‐mediated transcription by the selective receptor modulator RU486. Proc Natl Acad Sci USA 2002;99:7940–4. 222. Hu X, Chen Y, Farooqui M, Thomas MC, Chiang CM, Wei LN. Suppressive effect of receptor‐interacting protein 140 on coregulator binding to retinoic acid receptor complexes, histone‐modifying enzyme activity, and gene activation. J Biol Chem 2004;279:319–25. 223. Brown DT. Histone H1 and the dynamic regulation of chromatin function. Biochem Cell Biol 2003;81:221–7. 224. Woodcock CL, Skoultchi AI, Fan Y. Role of linker histone in chromatin structure and function: H1 stoichiometry and nucleosome repeat length. Chromosome Res 2006;14:17–25. 225. Catez F, Lim JH, Hock R, Postnikov YV, Bustin M. HMGN dynamics and chromatin function. Biochem Cell Biol 2003;81:113–22. 226. Mardian JK, Paton AE, Bunick GJ, Olins DE. Nucleosome cores have two specific binding sites for nonhistone chromosomal proteins HMG 14 and HMG 17. Science 1980;209:1534–6. 227. Crippa MP, Alfonso PJ, Bustin M. Nucleosome core binding region of chromosomal protein HMG‐17 acts as an independent functional domain. J Mol Biol 1992;228:442–9. 228. Paton AE, Wilkinson‐Singley E, Olins DE. Nonhistone nuclear high mobility group proteins 14 and 17 stabilize nucleosome core particles. J Biol Chem 1983;258:13221–9. 229. Yau P, Imai BS, Thorne AW, Goodwin GH, Bradbury EM. Effect of HMG protein 17 on the thermal stability of control and acetylated HeLa oligonucleosomes. Nucleic Acids Res 1983;11:2651–64. 230. Paranjape SM, Krumm A, Kadonaga JT. HMG17 is a chromatin‐specific transcriptional coactivator that increases the efficiency of transcription initiation. Genes Dev 1995;9:1978–91. 231. Cheung E, Zarifyan AS, Kraus WL. Histone H1 represses estrogen receptor alpha transcriptional activity by selectively inhibiting receptor‐mediated transcription initiation. Mol Cell Biol 2002;22:2463–71. 232. Koop R, Di Croce L, Beato M. Histone H1 enhances synergistic activation of the MMTV promoter in chromatin. EMBO J 2003;22:588–99. 233. Wacker DA, Ruhl DD, Balagamwala EH, Hope KM, Zhang T, Kraus WL. The DNA binding and catalytic domains of poly(ADP‐ribose) polymerase 1 cooperate in the regulation of chromatin structure and transcription. Mol Cell Biol 2007;27:7475–85. 234. Gamble MJ, Erdjument‐Bromage H, Tempst P, Freedman LP, Fisher RP. The histone chaperone TAF‐I/SET/INHAT is required for transcription in vitro of chromatin templates. Mol Cell Biol 2005;25:797–807. 235. Seo SB, McNamara P, Heo S, Turner A, Lane WS, Chakravarti D. Regulation of histone acetylation and transcription by INHAT, a human cellular complex containing the set oncoprotein. Cell 2001;104:119–30. 236. Martinez‐Balbas MA, Bannister AJ, Martin K, Haus‐Seuffert P, Meisterernst M, Kouzarides T. The acetyltransferase activity of CBP stimulates transcription. EMBO J 1998;17:2886–93. 237. Morse RH. Transcription factor access to promoter elements. J Cell Biochem 2007;102: 560–70. 238. Shang Y, Hu X, DiRenzo J, Lazar MA, Brown M. Cofactor dynamics and sufficiency in estrogen receptor‐regulated transcription. Cell 2000;103:843–52. 239. Sharma D, Fondell JD. Temporal formation of distinct thyroid hormone receptor coactivator complexes in HeLa cells. Mol Endocrinol 2000;14:2001–9.

190

RUHL AND KRAUS

240. Sharma D, Fondell JD. Ordered recruitment of histone acetyltransferases and the TRAP/ Mediator complex to thyroid hormone‐responsive promoters in vivo. Proc Natl Acad Sci USA 2002;99:7934–9. 241. Ren Y, Behre E, Ren Z, Zhang J, Wang Q, Fondell JD. Specific structural motifs determine TRAP220 interactions with nuclear hormone receptors. Mol Cell Biol 2000;20:5433–46. 242. Agalioti T, Lomvardas S, Parekh B, Yie J, Maniatis T, Thanos D. Ordered recruitment of chromatin modifying and general transcription factors to the IFN‐beta promoter. Cell 2000;103:667–78. 243. Cosma MP, Tanaka T, Nasmyth K. Ordered recruitment of transcription and chromatin remodeling factors to a cell cycle‐ and developmentally regulated promoter. Cell 1999;97: 299–311. 244. Sandaltzopoulos R, Becker PB. Heat shock factor increases the reinitiation rate from potentiated chromatin templates. Mol Cell Biol 1998;18:361–7. 245. Yudkovsky N, Ranish JA, Hahn S. A transcription reinitiation intermediate that is stabilized by activator. Nature 2000;408:225–9. 246. Zawel L, Kumar KP, Reinberg D. Recycling of the general transcription factors during RNA polymerase II transcription. Genes Dev 1995;9:1479–90. 247. Rani PG, Ranish JA, Hahn S. RNA polymerase II (Pol II)‐TFIIF and Pol II‐mediator complexes: the major stable Pol II complexes and their activity in transcription initiation and reinitiation. Mol Cell Biol 2004;24:1709–20. 248. Dieci G, Sentenac A. Detours and shortcuts to transcription reinitiation. Trends Biochem Sci 2003;28:202–9. 249. Gariglio P, Buss J, Green MH. Sarkosyl activation of RNA polymerase activity in mitotic mouse cells. FEBS Lett 1974;44:330–3. 250. Hawley DK, Roeder RG. Separation and partial characterization of three functional steps in transcription initiation by human RNA polymerase II. J Biol Chem 1985;260:8163–72. 251. Hawley DK, Roeder RG. Functional steps in transcription initiation and reinitiation from the major late promoter in a HeLa nuclear extract. J Biol Chem 1987;262:3452–61. 252. Lemon BD, Fondell JD, Freedman LP. Retinoid X receptor:vitamin D3 receptor heterodimers promote stable preinitiation complex formation and direct 1,25‐dihydroxyvitamin D3‐ dependent cell‐free transcription. Mol Cell Biol 1997;17:1923–37. 253. McNally JG, Muller WG, Walker D, Wolford R, Hager GL. The glucocorticoid receptor: rapid exchange with regulatory sites in living cells. Science 2000;287:1262–5. 254. Klokk TI, Kurys P, Elbi C, Nagaich AK, Hendarwanto A, Slagsvold T, et al. Ligand‐specific dynamics of the androgen receptor at its response element in living cells. Mol Cell Biol 2007;27:1823–43. 255. Rayasam GV, Elbi C, Walker DA, Wolford R, Fletcher TM, Edwards DP, et al. Ligand‐specific dynamics of the progesterone receptor in living cells and during chromatin remodeling in vitro. Mol Cell Biol 2005;25:2406–18. 256. Stenoien DL, Patel K, Mancini MG, Dutertre M, Smith CL, O’Malley BW, et al. FRAP reveals that mobility of oestrogen receptor‐alpha is ligand‐ and proteasome‐dependent. Nat Cell Biol 2001;3:15–23. 257. Becker M, Baumann C, John S, Walker DA, Vigneron M, McNally JG, et al. Dynamic behavior of transcription factors on a natural promoter in living cells. EMBO Rep 2002;3:1188–94. 258. Johnson TA, Elbi C, Parekh BS, Hager GL, John S. Chromatin remodeling complexes interact dynamically with a glucocorticoid receptor‐regulated promoter. Mol Biol Cell 2008;19:3308–22. 259. Stenoien DL, Nye AC, Mancini MG, Patel K, Dutertre M, O’Malley BW, et al. Ligand‐ mediated assembly and real‐time cellular dynamics of estrogen receptor alpha‐coactivator complexes in living cells. Mol Cell Biol 2001;21:4404–12.

ANALYSES OF

NR‐DEPENDENT TRANSCRIPTION IN VITRO

191

260. Fletcher TM, Ryu BW, Baumann CT, Warren BS, Fragoso G, John S, et al. Structure and dynamic properties of a glucocorticoid receptor‐induced chromatin transition. Mol Cell Biol 2000;20:6466–75. 261. Fletcher TM, Xiao N, Mautino G, Baumann CT, Wolford R, Warren BS, et al. ATP‐dependent mobilization of the glucocorticoid receptor during chromatin remodeling. Mol Cell Biol 2002;22:3255–63. 262. Karpova TS, Kim MJ, Spriet C, Nalley K, Stasevich TJ, Kherrouche Z, et al. Concurrent fast and slow cycling of a transcriptional activator at an endogenous promoter. Science 2008;319:466–9. 263. Stavreva DA, Muller WG, Hager GL, Smith CL, McNally JG. Rapid glucocorticoid receptor exchange at a promoter is coupled to transcription and regulated by chaperones and proteasomes. Mol Cell Biol 2004;24:2682–97. 264. Farla P, Hersmus R, Trapman J, Houtsmuller AB. Antiandrogens prevent stable DNA‐binding of the androgen receptor. J Cell Sci 2005;118:4187–98. 265. Marcelli M, Stenoien DL, Szafran AT, Simeoni S, Agoulnik U, Weigel NL, et al. Quantifying effects of ligands on androgen receptor nuclear translocation, intranuclear dynamics, and solubility. J Cell Biochem 2006;98:770–88. 266. Freeman BC, Yamamoto KR. Disassembly of transcriptional regulatory complexes by molecular chaperones. Science 2002;296:2232–5. 267. Frasor J, Danes JM, Komm B, Chang KC, Lyttle CR, Katzenellenbogen BS. Profiling of estrogen up‐ and down‐regulated gene expression in human breast cancer cells: insights into gene networks and pathways underlying estrogenic control of proliferation and cell phenotype. Endocrinology 2003;144:4562–74. 268. Barish GD, Atkins AR, Downes M, Olson P, Chong LW, Nelson M, et al. PPARdelta regulates multiple proinflammatory pathways to suppress atherosclerosis. Proc Natl Acad Sci USA 2008;105:4271–6. 269. Lee CH, Olson P, Evans RM. Minireview: lipid metabolism, metabolic diseases, and peroxisome proliferator‐activated receptors. Endocrinology 2003;144:2201–7. 270. Lee CH, Olson P, Hevener A, Mehl I, Chong LW, Olefsky JM, et al. PPARdelta regulates glucose metabolism and insulin sensitivity. Proc Natl Acad Sci USA 2006;103:3444–9. 271. Lin R, Nagai Y, Sladek R, Bastien Y, Ho J, Petrecca K, et al. Expression profiling in squamous carcinoma cells reveals pleiotropic effects of vitamin D3 analog EB1089 signaling on cell proliferation, differentiation, and immune system regulation. Mol Endocrinol 2002;16: 1243–56. 272. Rogatsky I, Wang JC, Derynck MK, Nonaka DF, Khodabakhsh DB, Haqq CM, et al. Target‐ specific utilization of transcriptional regulatory surfaces by the glucocorticoid receptor. Proc Natl Acad Sci USA 2003;100:13845–50. 273. Wang TT, Tavera‐Mendoza LE, Laperriere D, Libby E, MacLeod NB, Nagai Y, et al. Large‐ scale in silico and microarray‐based identification of direct 1,25‐dihydroxyvitamin D3 target genes. Mol Endocrinol 2005;19:2685–95. 274. Way JM, Harrington WW, Brown KK, Gottschalk WK, Sundseth SS, Mansfield TA, et al. Comprehensive messenger ribonucleic acid profiling reveals that peroxisome proliferator‐ activated receptor gamma activation has coordinate effects on gene expression in multiple insulin‐sensitive tissues. Endocrinology 2001;142:1269–77. 275. Gurevich I, Flores AM, Aneskievich BJ. Corepressors of agonist‐bound nuclear receptors. Toxicol Appl Pharmacol 2007;223:288–98. 276. Bai L, Santangelo TJ, Wang MD. Single‐molecule analysis of RNA polymerase transcription. Annu Rev Biophys Biomol Struct 2006;35:343–60. 277. Shundrovsky A, Smith CL, Lis JT, Peterson CL, Wang MD. Probing SWI/SNF remodeling of the nucleosome by unzipping single DNA molecules. Nat Struct Mol Biol 2006;13:549–54.

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278. Zhang Y, Smith CL, Saha A, Grill SW, Mihardja S, Smith SB, et al. DNA translocation and loop formation mechanism of chromatin remodeling by SWI/SNF and RSC. Mol Cell 2006;24:559–68. 279. Hall MA, Shundrovsky A, Bai L, Fulbright RM, Lis JT, Wang MD. High‐resolution dynamic mapping of histone–DNA interactions in a nucleosome. Nat Struct Mol Biol 2009;16:124–9. 280. Kininis M, Kraus WL. A global view of transcriptional regulation by nuclear receptors: gene expression, factor localization, and DNA sequence analysis. Nucl Recept Signal 2008;6:e005. 281. Theveny B, Bailly A, Rauch C, Rauch M, Delain E, Milgrom E. Association of DNA‐bound progesterone receptors. Nature 1987;329:79–81. 282. Wang Q, Carroll JS, Brown M. Spatial and temporal recruitment of androgen receptor and its coactivators involves chromosomal looping and polymerase tracking. Mol Cell 2005;19:631–42. 283. Carroll JS, Meyer CA, Song J, Li W, Geistlinger TR, Eeckhoute J, et al. Genome‐wide analysis of estrogen receptor binding sites. Nat Genet 2006;38:1289–97. 284. Kininis M, Chen BS, Diehl AG, Isaacs GD, Zhang T, Siepel AC, et al. Genomic analyses of transcription factor binding, histone acetylation, and gene expression reveal mechanistically distinct classes of estrogen‐regulated promoters. Mol Cell Biol 2007;27:5090–104. 285. Kushner PJ, Agard DA, Greene GL, Scanlan TS, Shiau AK, Uht RM, et al. Estrogen receptor pathways to AP‐1. J Steroid Biochem Mol Biol 2000;74:311–7. 286. Porter W, Saville B, Hoivik D, Safe S. Functional synergy between the transcription factor Sp1 and the estrogen receptor. Mol Endocrinol 1997;11:1569–80. 287. Cheung E, Acevedo ML, Cole PA, Kraus WL. Altered pharmacology and distinct coactivator usage for estrogen receptor‐dependent transcription through activating protein‐1. Proc Natl Acad Sci USA 2005;102:559–64. 288. Carroll JS, Liu XS, Brodsky AS, Li W, Meyer CA, Szary AJ, et al. Chromosome‐wide mapping of estrogen receptor binding reveals long‐range regulation requiring the forkhead protein FoxA1. Cell 2005;122:33–43. 289. Lefterova MI, Zhang Y, Steger DJ, Schupp M, Schug J, Cristancho A, et al. PPARgamma and C/EBP factors orchestrate adipocyte biology via adjacent binding on a genome‐wide scale. Genes Dev 2008;22:2941–52. 290. Nielsen R, Pedersen TA, Hagenbeek D, Moulos P, Siersbaek R, Megens E, et al. Genome‐ wide profiling of PPARgamma:RXR and RNA polymerase II occupancy reveals temporal activation of distinct metabolic pathways and changes in RXR dimer composition during adipogenesis. Genes Dev 2008;22:2953–67.

Chromatin Remodeling and Nuclear Receptor Signaling Manop Buranapramest and Debabrata Chakravarti Division of Reproductive Biology Research, Department of Obstetrics and Gynecology, Robert H. Lurie Comprehensive Cancer Center, Feinberg School of Medicine, Northwestern University, Chicago, Illinois 60611

I. Introduction ................................................................................. II. NR Classification and Structure ........................................................ A. NR Classification ...................................................................... B. NR Structure ........................................................................... III. NR Coregulators ........................................................................... A. Coactivators............................................................................. B. Corepressors............................................................................ IV. Chromatin as an NR Coregulator Substrate ......................................... A. Nucleosomes: The Basic Unit of Chromatin .................................... B. Post‐translational Histone Modifications and the Histone Code............ C. Role of INHATs in NR Signaling and Beyond .................................. D. Nucleosomes in Gene Regulation ................................................. V. ATP‐Dependent Chromatin Remodelers in NR Gene Regulation.............. A. Overview ................................................................................ B. SWI/SNF Complexes and NR Regulation ....................................... C. Acf1 and ISWI Complexes and NR Regulation ................................ VI. Future Directions.......................................................................... References...................................................................................

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Nuclear receptors (NRs) constitute a large family of ligand‐dependent transcription factors that play key roles in development, differentiation, metabolism, and homeostasis. They participate in these processes by coordinating and regulating the expression of their target genes. The eukaryotic genome is packaged as chromatin and is generally inhibitory to the process of transcription. NRs overcome this barrier by recruiting two classes of chromatin remodelers, histone modifying enzymes and ATP‐dependent chromatin remodelers. These remodelers alter chromatin structure at target gene promoters by posttranslational modification of histone tails and by disrupting DNA‐histone interactions, respectively. In the presence of ligand, NRs promote transcription by recruiting remodeling enzymes that increase promoter accessibility to the basal transcription machinery. In the absence of ligand a subset of NRs recruit remodelers that establish and maintain a closed chromatin environment, to Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87006-3

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ensure efficient gene silencing. This chapter reviews the chromatin remodeling enzymes associated with NR gene control, with an emphasis on the mechanisms of NR‐mediated repression.

I. Introduction A wide variety of small lipophilic molecules, including hormones and vitamins, function as chemical signals to regulate development, differentiation, metabolism, and homeostasis.1 Nuclear receptors (NRs) capable of binding these ligands act as transcription factors to regulate the expression of target genes and are, therefore, crucial intermediates in communicating extra‐ and intracellular information to the genome. Ligands that bind to and activate NRs include steroid hormones, such as estrogen and androgen, nonsteroid hormones, such as thyroid hormone, retinoic acid, and Vitamin D, and a variety of metabolic products among others.2 In the presence of ligand, NRs undergo a dramatic conformational change, which simultaneously allows: (1) receptor binding to cognate hormone response elements (HREs), (2) release of corepressors, and (3) recruitment of coactivators to DNA. These coregulators influence NR‐mediated transcription in two major ways: by modulating chromatin structure and by stabilizing the basal transcription machinery. The expression of numerous genes is regulated by NRs. The ligands that activate these receptors, therefore, have profound effects on the physiology of organisms. Many of these genes are associated with disease, explaining why NRs are molecular targets for approximately 13% of FDA approved drugs.3 These pharmaceutical agents treat disease by modulating NR activity. Excellent examples include tamoxifen for the treatment of breast cancer, thiazoidinedione for the treatment of diabetes, and glucocorticoids for the treatment of a variety of inflammatory conditions.4 A deeper understanding of how NRs and their coregulators modulate chromatin structure is essential to a complete understanding of NR‐mediated transcription, but more importantly, may lead to the development of novel therapeutic drugs.

II. NR Classification and Structure NRs were predicted to exist based on studies carried out by Jensen and colleagues in the 1960s using radio‐labeled estrogen ligands.5 Early studies using purified estrogen receptor (ER) allowed researchers to biochemically characterize their ligand‐binding properties and cellular localization.5

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The cloning of the first NR cDNA encoding human glucocorticoid receptor (GR) in 1985 lead to an explosion in the field of NR biology.6 In the two decades following, researchers characterized the DNA‐binding properties, functional domains, dimerization properties, and sequence conservation of members of the NR superfamily (discussed below). Information gleaned from these studies reveals a striking diversity, not only in the number of ligands NRs are capable of responding to, but also in the number of genes they can regulate. Surprisingly, evolutionary conservation of sequence as well as structure between family members, suggests common mechanisms of action by each of the members.

A. NR Classification The NR superfamily is commonly divided into four classes based on DNA‐ binding and dimerization properties.1 Class I NRs include the steroid hormone receptors ER, GR, mineralocorticoid receptor (MR), progesterone receptor (PR), and androgen receptor (AR). These NRs are generally found in the cytoplasm associated with chaperone proteins in the absence of ligand. In the presence of ligand, they form homodimers, translocate to the nucleus, and recognize and bind inverted HREs. Some Class I receptors, such as the ER, are found primarily in the nucleus and are capable of binding DNA, even in the absence of hormone.7 Class II receptors are found constitutively in the nucleus and bind DNA in a ligand‐independent manner. These NRs include the nonsteroid binding NRs such as Vitamin D receptor (VDR), retinoic acid receptor (RAR), thyroid hormone receptor (TR), and peroxisome proliferator‐activated receptor (PPAR).1 Class II NRs form heterodimers with retinoid X receptor (RXR) and bind to direct repeat (DR) HREs containing the sequence AGGTCA. The binding specificity for receptors in this class depends on the number of nucleotides between repeats, known as DRs.8 For example, PPAR binds to DRs with a single nucleotide spacer (DR1), TR binds to DRs with four nucleotide spacers (DR4), and RAR optimally binds to DRs with five nucleotide spacers (DR5).9 In the absence of ligand, NRs in this class bind to HREs and actively repress transcription by recruiting corepressor proteins.10 Ligand–receptor interaction further stabilizes DNA binding, and facilitates the release of corepressors and the recruitment of coactivators to DNA.11 The constitutive binding of Class II NRs to DNA allows them to quickly alter rates of transcription. Class III NRs bind DR HREs as homodimers, whereas Class IV NRs bind half‐sites as a monomer or as dimers.1 The receptors in these two classes often do not have a known ligand, and are referred to as orphan receptors. Orphan receptors include the NRs, nerve growth factor IB‐like receptor (NGFI‐B), and RAR‐related orphan receptor (ROR), and were originally identified based on homology to other NRs.12 Once a ligand is identified, the receptors are

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referred to as ‘‘adopted orphans.’’ Identifying novel ligands for orphan receptors and understanding their mechanisms of action present a challenge, but several have been identified. Examples include Rev‐erb, which has recently been shown to function as a heme sensor.13–15 The action of some orphan receptors may not require the binding of ligand, and may therefore function as ligand‐independent transcription factors.16

B. NR Structure NRs are a large family of evolutionary conserved transcription factors. NRs are modular in nature and share a similar structural and functional organization generally consisting of six conserved domains (A–F).17 Each domain possesses a distinct biochemical function. The A and B domains are located in the N‐terminal region and contain the activation function (AF‐1) domain, which is constitutively active in the absence of ligand.18,19 Domain C contains two highly conserved type II zinc fingers, which together constitute the NR DNA‐ binding domain (DBD).20 The P box, located in the first zinc finger, makes direct contacts with the major groove nucleotides and mediates sequence‐ specific recognition and binding of the NR to HREs.9 Region D contains a hinge region, which confers conformational flexibility, and contains a nuclear localization signal.8 Region E, also referred to as the ligand‐binding domain (LBD), is the largest domain and contains the activation function 2 (AF‐2) domain. Unlike AF‐1, activation by AF‐2 is ligand‐dependent. This region is responsible for ligand recognition and binding, receptor dimerization, and ligand‐dependent transcriptional regulation.17 The hydrophobic pocket of the LBD binds ligand causing a dramatic structural alteration, resulting from the movement of a‐helix 12, located within the C‐terminus of the LBD.21–23 This ligand induced conformational change inhibits the association of corepressors to the LBD, leading to their release, and simultaneously allows the association of coactivators. NRs therefore function as ligand‐dependent molecular switches. The relevance and function of the F region at the C‐terminus is not yet clear. In 2008, Rastinejad and colleagues revealed the crystal structure of the full‐ length PPARg–RXR complex bound to ligand and DNA.24 Results from this study not only support previous biochemical and structural studies, but also provide additional insight into the structure and function of a DNA bound NR complex. The binding of the PPARg–RXR complex involves direct contact between the DBDs and DR1 elements, as well as contact between the PPARg hinge peptide and a 50 AAAT extension sequence. This latter interaction plays a role in correctly positioning the DBD of the heterodimer with respect to the recognition site. The crystal structure also provides information regarding contacts made between the two heterodimeric partners. The NR heterodimer forms a nonsymmetric complex with three major contacts, some of which are

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dependent on DNA binding. Interestingly, the LBD of PPARg lies in a central position and makes contacts with the DBD of RXR. The LBD remains fully accessible to coactivator binding in this position. Disruption of this contact by a single phenylalanine mutation prevents DNA binding by the DBD as well as subsequent activation by the receptor. Surprisingly, the crystal structures of the PPARg–RXR complex are not substantially different when bound to a variety of ligands (rosiglitazone, 9‐cis‐retinoic acid, BVT.13, GW9662).

III. NR Coregulators The activation of transcription by ligand‐bound NRs involves not only binding to HREs, but also the recruitment of a variety of cofactors to DNA. As discussed in the previous section, ligand binding induces a conformational change in the receptor and allows the receptor to interact with a variety of cofactors. Transcription initiation of protein coding genes involves the binding of basal transcription factors and RNA Polymerase II (Pol II) to the core promoter. NRs stabilize the binding of these factors by directly interacting with components of the preinitiation complex (PIC) as well as by recruiting chromatin remodeling complexes that increase DNA accessibility. In fact, some of the first NR interactors identified were basal transcription factors.25,26 Several TAFs (TBP‐associated factors) in TFIID as well as TBP (TATA‐binding protein) make direct contacts with NRs in the presence of ligand.27,28 NRs can also directly interact with Pol II.27,29 These interactions stabilize the PIC and are required for full transcriptional activation.27,30 In addition to direct interactions, NRs also recruit the mediators, TR‐associated protein/VDR‐interacting protein (TRAP/DRIP), which act to further stabilize recruitment of Pol II to DNA.30 NRs recruit chromatin remodeling enzymes to modulate chromatin accessibility. In the absence of ligand, NRs recruit corepressors that function to actively repress transcription. In the presence of ligand, NRs undergo a conformational change that leads to the release of corepressor proteins that associate with histone deacetylases (HDACs) and the recruitment of coactivators that associate with histone acetyltransferases (HATs). Extensive evidence suggests that histone acetylation strongly correlates with gene activation. Therefore, the association of these coregulators suggests the importance of histone acetylation status in NR mediated gene regulation. In addition to histone acetylation, studies also demonstrate a role for histone methylation and ATP‐depending chromatin remodeling of nucleosomes in NR mediated activation and repression.

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A. Coactivators A number of coregulators are required for ligand‐dependent transcriptional activation by NRs. In the presence of ligand, the p160/steroid receptor coactivator (SRC) family of coactivators, including SRC‐1, ‐2, and ‐3, bind directly to the LBD of NRs.31 These coactivators possess LXXLL motifs that directly interact with AF‐2 and stimulate transcription. Alone, these coactivators contain only a small amount of intrinsic HAT activity. Therefore, activation of gene transcription involves the recruitment of more potent HATs, CBP/p300, which directly associate with the coactivators mentioned above.32,33 Recruitment of CBP/p300 to NR target genes results in histone acetylation, a post‐translational modification (PTM) often associated with gene activation.34–36 Other histone modifying enzymes associate with the SRC family of coactivators, including the histone methyltransferase (HMT), CARM‐1, which methylates a specific arginine residue on histone H3 and the HMT, PRMT‐1, which methylates a specific residue on histone H4.34,37 Therefore, in addition to HATs, HMTs can also be recruited to chromatin by NRs and function as ligand‐dependent NR coactivators. These PTMs influence the subsequent recruitment and activity of additional cofactors as discussed in Section IV.B.38,39 In addition to histone modifying enzymes, ATP‐dependent chromatin remodeling enzymes have also been shown to be involved in NR‐mediated gene transcription.40,41 In the presence of ligand, SWI/SNF ATPases function as coactivators for a variety of NRs, including GR, AR, TR, VDR, and RAR.42–46 These enzymes facilitate transcription by restructuring chromatin (nucleosomal dissociation, sliding, or relocation).47

B. Corepressors NR corepressors, as their name suggests, function in opposition to coactivators to repress transcription. Before NR corepressor proteins had been identified, transcriptional silencing by some NRs, including TR and RAR, were shown to be dependent on the C‐terminal region of these NRs.48 The repressive functions of DNA bound NRs were found to be inhibited by the LBDs of these NRs, suggesting the existence and association of corepressor proteins with unliganded NR LBD.49–51 In rapid succession, several laboratories identified the first corepressors involved in NR-mediated repression, 52,53 with nuclear receptor corepressor (NCoR) and silencing mediator of retinoid and thyroid receptors (SMRT) being the most extensively studied. Additional NR corepressors have since been discovered including RIP‐140, Hairless, Alien, SHARP, SUNCoR, PSF, LCoR, and SLIRP.48 These proteins mediate NR repression by distinct mechanisms.

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1. NCOR/SMRT STRUCTURE A critical role for NCoR/SMRT in NR‐mediated repression has been extensively demonstrated. More recently, both NCoR and SMRT have been implicated as corepressors for other nonreceptor transcription factors, broadening the role of these proteins in other physiological processes.54 NCoR and SMRT are large 270 kDa proteins that share several highly conserved domains (Fig. 1A). NCoR/SMRT bind directly to the LBD of NRs via the NR interaction domains (NIDs).55 The NIDs contain motifs consisting of an L/IXXI/VI sequence, referred to as the CoRNR box, which are essential for NR interaction.56–58 The CoRNR box, in a similar fashion to the LXXLL recognition motif of NR coactivators, is predicted to form an extended a‐helical domain. This domain docks in a complementary groove in the NR LBD and is available for corepressor binding in the absence of hormone. Hormone‐ induced release of NCoR/SMRT is a result of structural changes in the LBD. This role is evident by the observation that v‐ErbA, which is unable to bind hormone, is constitutively bound to NCoR/SMRT.52 Although NCoR and SMRT are highly homologous, differing amino acid sequences in their LBDs determine preferred association with specific NRs or transcription factors.59–61 The domains responsible for transcriptional repression are primarily located at the N‐terminus of NCoR and SMRT. Three distinct repression domains (RD1, RD2, RD3) were originally characterized by their ability to autonomously repress transcription as Gal4 fusions.53,62 These RDs, which mediate the interaction with NCoR/SMRT associated proteins, are required for corepressor complex formation as discussed below. Two distinct SANT (SWI3, ADA2, NCoR, TFIIB) motifs lie between RD1 and RD2 and have been shown to be critical for NCoR/SMRT function. A deacetylase activation domain (DAD) encompassing the first SANT motif is required for the association and subsequent activation of HDAC3.63–65 The DAD when fused to DNA‐binding proteins can recruit HDAC3 activity to DNA templates, deacetylate histones, and repress transcription.61 The second SANT motif contains the histone A

NCoR/SMRT

RD1

RD2

RID1/2

RD3

B

Acf1

WAC

DDT

BAZ1

BAZ2

WAKZ PHD

BRD

FIG. 1. Structures of NCoR/SMRT and Acf1. (A) Domain structure of NCoR/SMRT indicating known repressor (RD) and receptor interaction (RID) domains. (B) Domain structure of Acf1 containing the WAC/NID (NCoR interaction domain).

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interaction domain (HID) and interacts directly with deacetylated histone H4 N‐terminus tails. Because the HDAC3 containing NCoR/SMRT complex deacetylates histones and NCoR/SMRT interacts with the final product of this reaction (deacetylated histones) a feed‐forward model of NCoR/SMRT‐ chromatin binding has been proposed.66,67

2. NCOR/SMRT COMPLEXES Numerous proteins, in addition to HDAC3, have been identified as components of either NCoR or SMRT complexes. Several biochemical purifications of the corepressors have identified a core complex containing NCoR or SMRT, HDAC3, transducin (beta)‐like protein (TBL1) or TBL1‐related protein (TBLR1), and G‐protein pathway suppressor 2 (GPS2).63,68–70 Both TBL1 and GPS2 associate directly with RD1 of NCoR/SMRT.70 TBL1 associates with histones H2B and H4 via a WD40‐repeat domain and mediates repression by stabilizing the recruitment of NCoR/SMRT to chromatin.69 TBL1 and TBLR1 have also been suggested to play a role in degrading NCoR and SMRT in a proteasome‐dependent manner upon ligand binding, thereby facilitating the transition from a repressed state to an activated state.71 GPS2 may function to stabilize NCoR/SMRT complexes via its association with TBL1, but otherwise little is known about this component.70 NCoR and SMRT have also been shown in vitro to associate with a mammalian switch‐independent 3 (mSin3)/HDAC1/HDAC2 complex.72,73 Several biochemical purification schemes fail to purify these components as part of an NCoR/SMRT complex.68–70,74 Furthermore, neither HDAC1 nor HDAC2 have been found at NRs target genes, making the physiologic relevance for this association controversial. 61,75 Nevertheless, biochemical fractionation of complexes containing NCoR from Xenopus oocyte extracts demonstrates the existence of Sin3‐dependent complexes suggesting that NCoR may associate in many distinct complexes.76 Other HDACs, including, HDAC4, HDAC5, HDAC7 have also been shown, at least in vitro, to interact directly with NCoR and SMRT, and may also contribute to repression.77,78 The orphan NR small heterodimer partner (SHP) was recently shown to recruit a complex containing Sin3A, NCoR, and HDAC4 to mediate repression.79 Surprisingly, the involvement of RD2 in NCoR‐mediated repression has not been well characterized (see Section V.C) In the absence of ligand, Class II NRs interact strongly with NCoR and SMRT.52,53,80 These corepressors mediate repression by recruiting histone modifying enzymes, such as HDACs, to chromatin. The targeting of the NCoR/HDAC3 complex to NR HREs leads to the deacetylation of local histones and consequently repression of TR mediated transcription.61,63,64 In addition to associating with HDACs, NCoR/SMRT has also been shown to

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directly interact with the HMT SUV39H1 (Amy Ewing and D. Chakravarti, unpublished work). This histone modifying enzyme places a methylation mark at H3K9 residues and is involved in the repression of TR target genes.81 A biochemical purification of endogenous NCoR identified several SWI/ SNF components, including the ATPase subunit, Brahma‐Related Gene 1 (BRG1) and several BRG1/brm‐associated factors (BAFs).82 SWI/SNF complexes are involved in activation as well as repression of genes in yeast.83 It is possible that SWI/SNF complexes function at some NCoR targets to repress transcription. The SWI/SNF ATPase may confer chromatin remodeling activity to the NCoR complex however, a role for SWI/SNF in NCoR‐mediated repression has not been documented. The Mi2/NuRD complex, which possesses both HDAC and ATPase activity, is required for the repression of a TR target gene.75 However, it does not directly interact with NCoR or SMRT. The complex is constitutively recruited to chromatin in a nontargeted fashion and results in the global deacetylation of histones. Therefore, repression by the Mi2/NuRD complex at TR target genes is thought to occur in a TR‐ and NCoR‐independent fashion. Most recently the remodeling activity of Snf2h, a human ISWI homolog, was shown to be required for the repression of TR target genes.84 In addition, Acf1, a Snf2h‐binding partner, was identified in our laboratory by a yeast‐two hybrid screen as an NCoR interactor and was shown to be required for repression of several endogenous VDR and TR target genes.85 Biochemical purifications of NCoR/SMRT complexes suggest that multiple corepressor complexes may exist. The formation of specific complexes may be dictated by cell type, cellular signals, corepressor levels, histone modifications, or NR type.54 For example, TR preferentially recruits NCoR complexes, whereas RAR preferentially recruits SMRT complexes.59 Repression by TR is lost upon NCoR knockdown, whereas both NCoR and SMRT contribute to the repression mediated by the related Rev‐erb.61 In addition, NCoR knockout mice are embryonic lethal, suggesting that SMRT cannot fully compensate for NCoR loss.86 Although a high sequence homology between NCoR and SMRT suggests similar mechanisms of action, it is obvious from these studies that the physiological roles do not appear to be redundant. Additional studies investigating the functional differences between these two corepressors are needed to shed light on their distinct roles in vivo.

IV. Chromatin as an NR Coregulator Substrate Gene transcription by NRs in vivo occurs on a chromatin template. Many of the coactivators and corepressors recruited by NRs, function in distinct ways to modulate and alter chromatin. A basic understanding of the nature of chromatin is therefore essential to appreciate the mechanisms by which these NR coregulators function and are discussed in the following section.

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A. Nucleosomes: The Basic Unit of Chromatin Eukaryotic cells package their DNA into a structure called chromatin. The basic unit of chromatin is the nucleosome, which is composed of 147 bp of negatively charged DNA tightly wound around a positively charged histone octamer. The histone octamer is composed of two of each of the highly conserved canonical histones, H2A, H2B, H3, and H4.87 Crystal structures reveal that each core particle contains two functional domains, a structured histone‐ fold domain, and unstructured N‐ and C‐terminal domains (or tails). The histone tails extend out from the nucleosome core.88 Amino acids in this domain are subject to a variety of PTMs, including acetylation, phosphorylation, methylation, ubiquitylation, sumoylation, and ADP‐ribosylation. These modifications influence the process of transcription by recruiting cofactors to DNA. In addition, PTMs have also been shown to play a role in higher‐order chromatin structure as removal of the tails blocks chromatin condensation.89 The packaging of the genome into chromatin allows 2 m of DNA to fit into a nucleus only 5 mm in diameter. The basic histones function to neutralize the negative DNA backbone and allow compaction of the genome by a factor of up to 10,000. DNA wrapped around histones is significantly less accessible to factors involved in transcription, replication, and DNA‐damage repair, and has therefore previously been viewed as an obstacle for these DNA‐ based processes. Recent studies now suggest that chromatin formation may offer advantages beyond just compacting and packaging the genome.90 In general, only a small percentage of genes are activated in a eukaryotic cell, at any given time. Therefore, chromatin structure may exist to maintain the repression of the vast majority of genes where transcription is not required. This packaging strategy may also be used advantageously to direct DNA‐ binding regulators including NRs and the transcription machinery to appropriate sites on DNA, by limiting regions of binding. Chromatin structure correlates well with transcription status; euchromatic regions containing loosely associated DNA are generally active, whereas heterochromatin regions containing highly packaged DNA are silent. It is not entirely clear whether these relaxed euchromatic regions are generated as a result of active transcription, or whether euchromatin is a prerequisite for active transcription. Structural alterations to chromatin, including the disruption of DNA‐histone contacts and nucleosome eviction/deposition/repositioning/destabilization, confer a certain level of fluidity to the genome. These alterations are carried out primarily in two ways: by PTMs of the histone tails and by ATP‐ dependent chromatin remodeling. These two classes of chromatin remodelers allow regulated access to DNA. Nucleosomes therefore should not be viewed as obstacles to DNA‐based processes, but rather facilitators of DNA‐based processes, such as replication, repair, and transcription.

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B. Post‐translational Histone Modifications and the Histone Code The PTMs of histones by histone modifying enzymes is currently a highly active area of research. The role these modifications play in regulating numerous DNA‐based processes, including transcriptional regulation by NRs, is slowly being elucidated. A number of histone modifications have been described and include: acetylation, methylation, and ubiquitylation of lysine residues, phosphorylation of serines and threonines, and methylation of arginines.91 The primary function of these modifications is to recruit or release effector proteins to chromatin by serving as docking modules for factors containing modification‐recognition motifs. As previously mentioned, PTMs may also play a direct role in regulating chromatin structure. The presence of histone‐modifying enzymes that place as well as remove specific modifications, suggests that the PTM of histones can be highly dynamic. This particular feature allows precise regulation of transcription, as chromatin can be quickly converted between transcriptionally active euchromatin and transcriptionally silent heterochromatin. PTMs were originally thought to alter chromatin structure by altering the charge of amino acids. In this model, histone modifications regulate the interaction between the negatively charged DNA backbone and the positively charged histone tails, to directly interfere with histone–histone or histone–DNA interactions. Histone tails, for example, contain positively charged lysines that can be neutralized upon addition of an acetyl residue by HATs. These interactions are hypothesized to be critical for chromatin compaction. A recent investigation challenged this theory by using recombinant histones to determine the role of the tails in chromatin condensation. This study found that the deletion of all four histone tails (except residues at the base of histone H4) did not alter the formation of compact chromatin structures.92 Another proposed mechanism by which the PTM of histone proteins alters the chromatin structure is by creating bindings sites for protein domains that recognize specific histone modifications. Proteins containing bromodomains, for example, specifically recognize acetylated lysines and chromodomain‐ containing proteins associate with methylated lysines.93,94 Distinct combinations of PTMs were found to correlate with specific transcriptional states; leading to the proposal of the histone code hypothesis.38,39 This hypothesis suggests that the patterns of modifications are read as a code to recruit and release the cellular machinery to and from chromatin. In the process of transcription, these recruited proteins are the effectors of downstream processes, leading to activation or repression of genes. The histone code is believed to function in two major ways: as a transient signaling mediator for gene activation or repression or as a stable marking system to define specific chromatin states.

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As the same set of histone modifications does not always dictate the same transcriptional output, the term ‘‘histone code’’ cannot be interpreted too strictly. The transcriptional readout of a particular set of modifications seems to be dependent on the neighboring modifications, on the particular gene, on the position within the gene, and on the local chromatin environment. While a universal ‘‘histone code’’ might not exist, it is possible that ‘‘gene‐specific’’ histone codes are used between different or related species. 1. HISTONE ACETYLATION The most extensively studied PTM is acetylation. HATs catalyze the transfer of an acetyl group from acetyl‐CoA to the e‐amino groups of lysines on histones. This mark has long been associated with gene activation. Multiple transcriptional coactivators (including PCAF and CBP/p300), which interact with a variety of transcription factors (including nuclear hormone receptors) as well as TAFII250, the largest subunit of TFIID, possess intrinsic HAT activity.95–98 In yeast, acetylation of histone H3K9 and H3K14, as well as the general acetylation of histone H4 are localized predominately at promoters that are transcriptionally active.99 This modification can be rapidly reversed by HDACs, which catalyze the removal of acetyl groups.100 Transcriptional corepressor proteins such as NCoR and mSin3, identified as corepressors for many transcription factors including nuclear hormone receptors, associate with HDACs and require HDAC activity for transcriptional repression.72,101 As mentioned above, acetylation of lysine can also function as a docking site for a protein module known as a bromodomain. These domains are found in many transcription and chromatin regulators102 and are critical for the acetylation‐dependent association of transcriptional activators such as Gcn5 and SWI/SNF ATPase with chromatin.103 Members of the basal transcription machinery such as TAFII250 also contain bromodomains capable of recognizing acetylated lysines and may stabilize their interaction with the promoter.104 Factors containing bromodomains and HAT activity, such as Gcn5 and the NR coactivator PCAF may function in a feed‐forward mechanism to recruit and stabilize transcription activation. Removal of the acetyl groups by HDACs inhibits transcriptional activity by tightening DNA‐histone association, releasing bromodomain containing factors, and serving as a binding site for corepressor proteins. For example, deacetylated histones are a preferred template for core components of the NCoR/SMRT complex and likely enable stable recruitment of the complex to deacetylated chromatin.66,67,69 Hypoacetylated histones are recognized by the corepressor INHAT (inhibitor of acetyltransferases) subunits pp32 and Set/TAFIb. INHATs integrate chromatin hypoacetylation and transcriptional

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repression by (1) binding to hypoacetylated histones and (2) recruiting HDACs, thereby propagating a hypoacetylated state.105 A role for these proteins in NR signaling is discussed in Section IV.C. 2. HISTONE METHYLATION Methylation of lysines and arginines on the tails of histones H3 and H4 tails has been extensively characterized. Unlike acetylation, which is generally associated with transcriptional activation, histone methylation, depending on the extent and site of modification, can signal either activation or repression. HMTs catalyze the transfer of methyl groups from S‐adenosyl‐methionine to create mono‐, di‐, or trimethylated histones. Methylation of lysine and arginine residues creates novel binding sites for proteins that specifically recognize these modifications. Recent investigations of histone methylation have identified at least three motifs that are capable of specific interaction with methylated lysines residues: the chromodomain, the Tudor domain, and the WD‐40 repeat domain.106 These methyl‐binding proteins contain modules that recognize methylated histones based on the specific residue and the extent of its methylation. A major breakthrough in understanding the role of methylation in transcription came from the finding that a well known transcriptional repressor, Drosophila, Su(var)3–9, has HMT activity.107,108 The human homologue, SUV39H1, was also shown to play a role in heterochromatin formation and associate with M31, the human homolog of HP1 (heterochromatin‐binding protein 1).109 Subsequent studies have shown that SUV39H1 methylates histone H3K9, which creates a binding site for the chromodomain of HP1/ M31.110–112 In addition to establishing heterochromatin regions, methylation of histone H3K9 as well as H3K27 (by the HMTs G9a and Polycomb protein, respectively) is found at transcriptionally silent genes located in euchromatic regions.113,114

C. Role of INHATs in NR Signaling and Beyond Nuclear hormone receptor signaling can be attenuated by the adenoviral protein E1A. Since E1A can directly bind to NR coactivators CBP/p300, it was proposed that such binding squelches/inactivates the coactivators from participating in NR mediated gene activation. Subsequent to the discovery that CBP/p300 are also HATs, it was shown that blockade of this enzymatic activity by E1A also contributes to attenuation of NR mediated gene activation.115–118 Following this discovery of a viral mechanism of inactivation of the HAT activity of coactivators, the search was on to identify cellular HAT regulatory activities. Using a biochemical purification scheme in our laboratory, Seo et al. reported the first discovery of cellular proteins termed INHAT or inhibitors of

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acetyltransferase which blocked nuclear hormone receptor signaling at least in part by inhibiting the HAT enzymatic function of coactivators such as CBP/ p300 and PCAF.119,120 The oncoprotein Set/template activating factor‐Ib, TAF‐ Ia, and the phosphoprotein pp32 which are acidic proteins were shown to have such activity in the initial report. Overexpression of these proteins blocked reporter gene activation by Gal4 fusions of coactivators and RARs. Surprisingly, however, in contrast to E1A which functions by binding to the enzyme itself, the INHAT proteins bound the substrate histones and prevented their acetylation. Since acetylation of histones plays a critical role in gene activation, this novel mechanism termed histone masking provided additional mechanisms for regulation of hormone signaling by blockade of HAT functions of coactivators. While this first observation established that the INHAT proteins associate with chromatin, no direct proof was provided as to whether they are indeed involved in transcriptional regulation. Several reports now confirm the original discovery by Seo et al. and Loven et al. demonstrated that Set/TAF‐Ib and pp32 directly associate with ER in vitro and in intact cells. Moreover, overexpression of pp32 and Set/TAFIb decreased ligand activated estrogen responsive reporter activity. These studies also showed that these INHAT proteins in addition to blocking histone acetylation also blocked acetylation of ER which is important for its transcriptional activation function (AF). Not surprisingly, the INHAT proteins also associated with thyroid receptor, progesterone hormone receptors, and PPARs.121,122 Hong and Chakravarti123 noted that the human proliferating cell nuclear antigen (PCNA) also associates with p300, regulates its HATactivity, and blocks RAR mediated reporter gene activation. The involvement of INHAT proteins in nuclear hormone receptor signaling was further biochemically demonstrated by the Nardulli laboratory. Using agarose gel electrophoresis coupled with protein sequencing the authors also identified INHAT proteins pp32, TAF‐Ia, and TAF‐Ib, and PCNA as ER interacting proteins among others.121,122,124 These independent studies therefore place INHAT proteins as additional regulatory components of the nuclear hormone receptor signaling pathways. Further mechanistic studies from our laboratory and by Schneider et al. subsequently showed that acetylation of histones prevents INHAT binding to chromatin.105,125 Kutney et al. further demonstrated that the INHAT proteins are recruited to the hypoacetylated/repressed and are released from the hyperacetylated/activated EB 1 gene upon hormone treatment, consistent with their roles in transcriptional repression by nuclear hormone receptors. Additionally, the INHAT proteins associate with HDACs.105 These interactions and functions were therefore proposed to explain INHAT mediated NR regulation. Using a chromatin integrated GR responsive reporter gene, Ichijo et al. came to a similar conclusion that upon ligand activation, INHAT proteins are released from GR target genes thereby allowing transcription activation to occur.126 Consequently, siRNA mediated knockdown of Set/TAF‐Ib led to

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enhanced expression of a well known glucocorticoid responsive TAT gene providing the first confirmation of an endogenous hormone responsive gene that is negatively regulated by INHAT. Consistent with this result, chromatin immunoprecipitation data showed that endogenous Set/TAF‐Ib and pp32 were bound to TAT GREs in the absence of dexamethasone and were released from the endogenous promoters upon dexamethasone treatment of HTC cells. In this context, it is important to note that a Set‐Can fusion has been found in leukemia; however how this fusion protein contributes to leukemia is unknown.127 Ichijo et al. showed that in contrast to ligand mediated release of Set‐TAF‐Ib from GRE, the Set‐Can fusion protein remained bound to the promoter even under hormone treatment and thus blocked gene activation. These results therefore provide a plausible mechanism contributing to glucocorticoid resistance in acute undifferentiated leukemia cells.126 In the context of this discussion, it was recently observed that the leucine rich acidic nuclear protein LNAP (pp32) also directly regulates expression of neurofilament light (NF‐L) chain, by binding to the promoter and modulating histone acetylation levels and neuronal differentiation. Consequently, in LNAP knockout mice, increased histone acetylation is observed at NF‐L promoter accompanied by increased levels of NF-L mRNA.128 The histone acetylation regulatory activity of the INHAT protein pp32 was clearly established by the studies of Fan et al.129 In Jurkat T cells silencing of pp32 led to increased mRNA expression of IL2 gene accompanied by hyperacetylation of IL2 promoter as determined by ChIP assays. The authors demonstrated that pp32 is specifically targeted to the IL2 gene promoter and overexpression of pp32 led to a significant decrease in histone H3 acetylation of promoter region of IL2. In contrast to the above results Adegbola and Pasternack reported that pp32 associates with AR and functions as a coactivator in transfected reporter assays130 implying that INHAT proteins might have bi‐functional coregulator function in NR regulation. In agreement with this view, Gamble et al. used an in vitro chromatin template to determine the role of Set/TAF‐Ib in Vitamin D3 mediated transcription.131 The authors found that recombinant Set/TAF‐Ib as well as a HeLa nuclear fraction containing Set/TAF‐Ib promoted activation of VDR/Vitamin D3 mediated transcription from in vitro assembled chromatin. Additionally, the authors found that higher concentration Set/TAF‐Ib blocked Vitamin D3 mediated gene activation. Finally, Set/TAF‐Ib appeared to function at a step postchromatin assembly but prior to transcription elongation. While the mechanism for this dual function is not clear, it appears that the INHAT‐target gene architecture, cellular concentration of INHAT proteins, and the cellular context might play important roles in their biologic activity. It is also not clear whether the histone chaperone and chromatin assembly activity of INHAT proteins are critical for their role in regulating nuclear hormone

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receptor function. Nonetheless, the INHAT proteins probably use multiple mechanisms for targeting to responsive promoters and to regulate gene expression by nuclear hormone receptors and probably by other transcription factors. It is important to note that INHAT proteins are ‘‘multitasking’’ proteins: For example, TAF‐I proteins have been shown to play important roles in chromosome assembly and viral DNA replication, and transcription.132 The crystal structure of Set/TAF‐Ib lacking the acidic INHAT domain has been solved providing insight into the molecular function of the protein.133 A dimeric headphone structure in which each subunit provides an earmuff domain is observed. The bottom surface of the earmuff domain is important for histone chaperone as well as double stranded DNA‐binding activity. These histone chaperone proteins have also been shown to play critical roles in apoptosis, cell proliferation, differentiation, and mRNA trafficking.134–136 While chromosomal translocation of Set/TAF‐Ib has been noted in acute undifferentiated leukemia,127 a recent study showed a significant decrease of nuclear TAF‐I proteins and pp32 in lymphoblastoid cells treated with bleomycin implying their potential role in DNA double strand break response.137 These observations raise an important question as to whether all the activities of INHAT proteins are independently or interdependently regulated.123 Is the INHAT activity restricted to Set/TAF‐Ib, TAF‐Ia, and pp32 proteins? The prediction would be no. Consistent with that prediction, Hong et al. demonstrated that the Ets family oncoprotein PU.1 not only blocks CBP/ p300‐dependent histone acetylation and transcription, but also efficiently blocks acetylation of critical hematopoietic transcription factors such as GATA 1, NF‐E2, and EKLF.138 In addition PU.1 inhibits the differentiation associated increase in histone acetylation of the b‐globin gene locus suggesting that INHAT mediated gene regulation might be a general rather than nuclear hormone specific mechanism. Additionally, Ko et al. recently described the proto-oncoprotein Dek, as also having INHAT activity although no specific gene targets were identified.139 Finally, an RNA binding protein, TLS (translocated in liposarcoma) has recently been shown to bind and inhibit CBP/p300 HAT activity and to repress cyclin D1 gene transcription. Recruitment of TLS to cyclin D1 promoter is directed by non-coding RNA (ncRNA). These results provide a novel mechanism integrating histone hypoacetylation and transcriptional repression via recruitment of a ncRNA-RNA binding protein complex to target genes.203 JDP2 (Jun dimerization protein 2) represses transactivation of Jun family proteins by a mechanism that involves dimerization. Like the INHAT proteins discussed earlier, JDP2 also represses transcription by recruiting HDACs to its target gene and blocks retinoic acid mediated differentiation of F9 cells.140 Interestingly, JDP2 also functions as a coactivator of PR.141 To determine additional mechanisms by which JDP2 might function, Jin et al. found that JDP2 blocks histone acetylation by p300 in vitro and in intact cells.

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Overexpression of JDP2 decreased retinoic acid‐dependent histone H4 hyperacetylation of the c‐Jun promoter, a target of JDP2. Like the INHAT proteins Set/TAF‐Ib and TAF‐Ia, JDP2 also has histone chaperone and nucleosome assembly activity and the histone‐binding domain is necessary but not sufficient for the HAT inhibitory activity. Further mutational analysis indicated that the basic DBD of JDP2 is also required for INHAT activity. This is in contrast to the function of the INHAT proteins where the C‐terminal acidic domain was necessary for the INHAT activity. More importantly using site directed mutants the authors show that both the histone binding and INHAT activity of JDP2 are critical for retinoic acid induced differentiation of F9 cells. The Yokoyama group further extended the studies on JDP2 by creating JDP2 knockout mice.142 The authors showed that JDP2 is directly targeted to the C/EBP gene which is a critical transcription factor necessary for C/EBPa and nuclear hormone receptor PPARg mediated adipocyte differentiation. Such a binding repressed C/EBP mRNA expression by blocking histone acetylation strongly suggesting JDP2 as a negative regulator of adipocyte differentiation. Together these results established that JDP2 has INHAT activity which is critical for regulation of its target genes. The emerging concept that INHAT activity operates beyond regulating nuclear hormone receptor target genes come from the identification of NIR as a novel INHAT repressor of p53 transcriptional activity.143 NIR function is not blocked by HDAC inhibitors, suggesting that NIR does not function by attracting HDACs to its target promoters. Instead, NIR directly binds to nucleosomes and core histones and blocks their acetylation by HATs. A clue as to how NIR might regulate specific gene expression came from tandem affinity purification that identified p53 as a NIR interacting protein. The authors showed that NIR is indeed recruited to the p53 target genes such as p21, PIG3 in a p53‐dependent manner. Consistent with that observation NIR did not interact with nuclear hormone receptors suggesting that these histone chaperone proteins might have transcription factor specific INHAT activity. Consistent with that, siRNA mediated knockdown of NIR upregulated p21 expression only in p53 positive but not p53 negative HCT116 cells accompanied by significant increase in histone acetylation of the target promoter. Why would these histone chaperone proteins such as Set, TAF‐I, JDP2, and NIR be necessary for transcriptional regulation? We suggest that they play important roles by functioning as a gate keeper of transcription. In that scenario, recruitment of the histone chaperone proteins to specific sites on DNA via their interactions with transcription factors and/or subsequent recognition and association with histones and DNA stabilizes transcription complexes. Consistent with that view, the above discussion clearly indicate that these histone chaperone and assembly proteins play critical roles in regulating NR signaling and other transcriptional events. It would be important in the future to determine whether these proteins communicate or interact with each

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other to form a supra INHAT complex and whether the INHAT proteins interact with Acf1 which is a chromatin assembly and remodeling protein to mediate transcriptional repression by NRs (see below).

D. Nucleosomes in Gene Regulation A long held assumption for the activation of transcription is that the loss of promoter nucleosomes allows the basal transcriptional machinery access to underlying DNA. Initial evidence for this premise came from the discovery that nucleosome free regions commonly occurred at known transcriptional regulatory regions such as enhancers and promoters.144,145 Early evidence that these regions may be free of histones came from the report that a 115‐bp fragment was released upon restriction enzyme digestion at the active chicken adult‐b‐globin locus promoter.146 Investigations into the location and occupancy of nucleosomes in promoters by ChIP assay, showed that in the Saccharomyces cerevisiae genome, while most nucleosomes are well‐positioned, RNA Pol II transcribed genes contain a nucleosome free gap of 200 bp over the promoter, flanked by two well positioned nucleosomes.147 The loss of nucleosomes in actively transcribed regions was also observed in lower resolution genome‐wide microarrays.148,149 Taken together, these experiments show that there is a depletion of nucleosomes at active regulatory elements and an inverse correlation between nucleosome occupancy at upstream regulatory elements and rates of transcription of the corresponding ORF (open reading frame). Loss of specific histones and of entire nucleosomes from gene promoters upon induction was also observed at genes regulated by NRs and other transcription factors. For example, there is a depletion of histone H3 and H4 from interleukin‐2 gene during T‐cell activation.150 In S. cerevisiae, heat shock treatment causes a loss of histones over entire heat‐shock genes. Progesterone activation of the mouse mammary tumor virus promoter leads to a loss of H2A and H2B from the nucleosome bound by the PR151 and Vitamin D activation of RANKL genes is accompanied by loss of histones H3 and H4 from promoters.85 The mechanism for histone depletion is not well understood. Many theories, currently under investigation may help us understand this phenomenon. Early models proposed that transcription factors compete for DNA during DNA replication to form stable complexes that exclude nucleosomes from the promoters of active genes during chromatin assembly.152 More recent evidence shows that the underlying DNA sequence itself may be responsible for excluding nucleosomes from certain regions of chromatin. Nucleosome‐free gaps are enriched in sequences of poly dAdT, for example.147 These regions can act as upstream activating elements not by binding transcription factors, but rather by excluding nucleosomes. Alternatively certain DNA sequences facilitate nucleosome binding. It has therefore been recently proposed that genomes encode an intrinsic nucleosome organization that can explain about 50% of the in vivo nucleosome positions.153

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While these features may explain why there are nucleosome‐free areas in certain parts of the genome, it does not account for the active displacement of histones during gene activation. Upon gene induction, various transcription factors can destabilize the nucleosome making it more susceptible to displacement. At the PHO5 gene, for example, the HAT GCN5 acetylates histones, which causes the nucleosome to become unfolded and free of the surrounding DNA.154,155 Another class of transcription factors, the ATP‐dependent nucleosome remodeling complexes, can function together with histone modifying enzymes to facilitate histone displacement. The SWI/SNF complex, for instance displaced acetylated nucleosomes from a nucleosome array. This results in the generation of nucleosome‐free DNA surrounding activator‐binding sites.156 Recent studies have shed light on the role nucleosome positions play in regulating transcription. The bulk of these nucleosome mapping studies have been performed in yeast, but several recent studies have also been done in human cells.150,157,158 Using high‐throughput sequencing technologies, Schones et al. have generated a genome‐wide map of nucleosome positions for resting and activated T cells.158 T cell activation induces extensive nucleosome repositioning at promoters and enhancers to allow transcriptional activation or repression. At repressed genes, the first nucleosome downstream of the start site, called the þ1 nucleosome generally adopts one of two nucleosome positions. At nonstalled promoters, the þ1 nucleosome appears over the start site, whereas at stalled promoters, in the presence of Pol II binding, the þ1 is predominately located just downstream of the start site. At actively expressed genes, the downstream position predominates.159 This study demonstrates a strong conservation of nucleosome positioning at the promoters across species (yeast to human) and demonstrate the importance of nucleosome dynamics in transcriptional regulation. However, it is still not clear how nucleosome positioning functions to dictate transcriptional outcomes. NRs have been demonstrated to utilize both nucleosome eviction and nucleosome positioning strategies to regulate transcription of target genes and are discussed in Section V. Nucleosome modulation by NRs occurs primarily by posttranslational histone modifications and ATP‐dependent chromatin remodeling, although additional mechanisms do exist. One good example is the NR corepressor Alien, which has been shown to repress transcription of TR and VDR targets.160 Alien interacts directly with TR‐a and ‐b and VDR in the absence of hormone. Like NCoR/SMRT, ligand binding leads to a release of the corepressor (along with associated HDACs). Alien mediates repressive activity by modulating NAP1 nucleosome assembly.161 Alien binds histone H3 and H4, thereby inhibiting accessibility of NAP1 to histones, and preventing nucleosome disassembly. Additional research is clearly necessary to determine the general role of hormone induced histone eviction, replacement, and nucleosome positioning as mechanisms in NR function.

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V. ATP‐Dependent Chromatin Remodelers in NR Gene Regulation A. Overview Nucleosome positioning plays a major role in determining transcriptional outcomes and to a limited extent is dictated by the underlying DNA sequence. This strategy, however, is unlikely to regulate the position of the majority of nucleosomes in the eukaryotic nucleus. In addition to histone modifying enzymes, ATP‐dependent chromatin remodeling enzymes are one of the major contributors to the dynamic nature of chromatin. ATP‐dependent chromatin remodelers use the energy stored in ATP to restructure, mobilize, and position nucleosomes.162–164 These actions serve to modulate transcription by exposing or occluding DNA sites required for binding transcription factors, the basal transcriptional machinery, and Pol II. These complexes play an important role, not only in the activation and repression of transcription, but also for chromatin assembly, DNA replication, and DNA repair.165 While the basic enzymatic mechanism is similar among chromatin remodeling ATPases, these enzymes have evolved into multiple families which have specialized roles in vivo because they affect structure of nucleosomes and arrays in distinct ways. At least five families of nucleosome remodelers have been characterized in eukaryotes and they include: SWI/SNF, ISWI, NuRD/Mi‐2/CHD, INO80, and SWR1.163 The ATPase component is shared between members within a family and serves to function as the DNA translocation ‘‘motor.’’ Accessory proteins associate with the ATPase ‘‘motor’’ component to alter the qualitative and quantitative aspects of nucleosome mobilization.165 A growing body of evidence suggests that these ATP‐dependent chromatin remodelers are targeted to chromatin by site‐specific DNA‐binding proteins including NRs.166 How the nonenzymatic subunits of nucleosome impart unique functions however remains largely unknown. Eukaryotic cells harbor hundreds of different combinations of these remodeling complexes. And it appears likely that each complex possesses a distinct function in vivo. The huge variety of motor associated factors, suggest a wealth of distinctly different outputs generated by these complexes. Complexes in the SWI/SNF and ISWI families have been studied extensively in vitro and in vivo, and demonstrate involvement in both transcriptional activation and as well as repression by NRs as discussed below.165

B. SWI/SNF Complexes and NR Regulation SWI/SNF (mating type SWItching/sucrose nonfermentors) was the first nucleosome remodeling enzyme discovered. Genes coding for SWI and SNF were identified in genetic screens in S. cerevisiae as positive regulators of HO

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and SUC2 transcription, respectively.167,168 Subsequent analysis revealed SWI2 and SNF2 to be the same gene and code for a protein that functions in complexes that possess ATP‐dependent remodeling activity.169,170 Unlike ISWI complexes which are primarily involved in chromatin assembly and nucleosome phasing, the SWI/SNF ATPase‐containing complexes function instead by disordering and reorganizing the nucleosome array to allow for transcription factor binding and activation.171 In vivo, SWI/SNF associates with hyperacetylated chromatin through a bromodomain and is found to colocalize with Pol II.172 This strongly suggests a role for SWI/SNF in transcriptional activation. Additional evidence reveals a role for SWI/SNF in transcriptional repression, suggesting that the context of the remodeling event is a key determinant in transcriptional regulation.166 1. COACTIVATION OF CLASS I NRS BY SWI/SNF A role for SWI/SNF in NR‐mediated gene regulation has been extensively reviewed elsewhere,40,173–175 so this process will only be touched upon here. SWI/SNF was the first reported ATP‐dependent chromatin remodeling complex involved in regulating transcription by NRs, and was shown to be critical for GR activation.43,46,176 Follow‐up studies demonstrate that SWI/SNF influences ligand‐induced activation of a host of NRs, including ER, AR, TR, VDR, RAR, and others.177 The steroid hormone‐responsive MMTV promoter is the most established model for studying NR activation by chromatin remodelers.43 The MMTV promoter has a highly organized nucleosomal structure of phased nucleosomes when integrated into chromatin. Binding sites for nuclear factor‐1 (NF‐1), octamer transcription factor (OTF), and TATA binding protein (TBP) are found wrapped around a specific nucleosome, termed nucleosome B.178 These sites allow strong promoter activation by glucocorticoids through mechanisms involving chromatin remodeling. The incorporation of DNA into nucleosomes negatively influences the ability of transcription factors, including NRs, to bind cognate sites.179 Rotational positioning of the nucleosome allows access to DNA facing away from the nucleosome, and is therefore a critical determinant in allowing GR to bind its cognate HRE on nucleosome B. The utilization of SWI/SNF chromatin remodeling complexes by steroid receptors was initially determined based on the reported interactions of GR and ER with SWI/SNF subunits.43,180 Experimental evidence supports this view, implicating the BRG1 complex in GR‐mediated chromatin remodeling at the MMTV promoter. ISWI or Mi‐2 based complexes cannot rescue GR‐ mediated activation in the absence of SWI/SNF.181 The targeting of SWI/ SNF to specific gene promoters is thought to take place through the binding of transcription factors, coactivators, or members of the general transcriptional machinery. BAF subunits with bromodomains are known to target acetylated histone tails.182 The ER has also been shown to require BRG1 activity for

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ER‐mediated transcriptional activity.176,183 In BRG1‐null cells, ER‐mediated transcriptional initiation of estrogen‐responsive genes was greatly diminished. Reintroduction of BRG1 restores ER transcription activity.181 The MMTV promoter has also been used to demonstrate a role for SWI/SNF in PR and AR‐mediated activation.43,44 In 2004, Beato and colleagues identified a novel role for histone eviction in hormone‐induced activation.151 Utilizing breast cancer cells containing a chromosomally integrated MMTV promoter, they demonstrated that progesterone induction recruits not only PR, but also a BRG1‐containing complex to the MMTV sequence. Concurrently, histone H2A and H2B are depleted from nucleosome B, but not from adjacent upstream and downstream nucleosomes. They further demonstrate in vitro that SWI/SNF catalyzes the sliding of a mononucleosome (consisting of the MMTV sequence and recombinant histones) and subsequent displacement of H2A/H2B dimers. Depletion of H2A/ H2B dimers by SWI/SNF appears to be dependent on DNA sequence, as SWI/ SNF‐mediated displacement was not seen on mononucleosomes containing the rDNA sequence. This study strongly suggests a role for nucleosome destabilization in transcriptional activation by NRs. 2. THE VDR–WINAC CONNECTION A novel SWI/SNF containing complex, designated the WSTF including nucleosome remodeling complex (WINAC), was isolated by Kato and coworkers in 2003.184 The purified complex contains, in addition to WSTF, SWI/ SNF ATPase subunits (BRG1/hBRM), proteins associated with DNA replication (TopoIIb, CAF‐1p150), a transcription elongation factor (FACTp140), as well as several additional BRG1/hBRM associated factors (BAFs). Surprisingly, Snf2h, which is associated with WSTF in the WICH complex, was not found in the WINAC complex. In the presence of ATP, WINAC promotes the assembly of nucleosome arrays. In addition, the complex appears to function much like the histone chaperone complexes, CAF‐1 and NAP‐1, to reconstitute chromatin upon newly replicated DNA. Knockdown of WSTF or BRG1/hBRM ATPases prevents normal S‐phase progression and results in alterations in cell cycle. The WINAC complex associates with VDR in a ligand‐independent manner and mediates recruitment of the receptor to both positive (human 1a,25‐dihydroxyvitamin D3 24‐hydroxylase [24(OH)ase]) and negative (human 25‐hydroxyvitamin D3 1a‐hydroxylase [1a(OH)ase]) VDREs.184,185 The mechanisms of WINAC‐mediated VDR recruitment differs between positive and negative VDREs and is discussed below. At classic or positive VDREs [24(OH)ase], ChIP experiments demonstrate that WINAC and VDR are recruited to the promoter in a ligand‐independent manner.184 As expected, the recruitment of coactivators (TRAP220 and TIF2)

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and subsequent activation of gene expression by VDR requires ligand binding. Knockdown of WSTF prevents the recruitment of VDR to the promoter. As a consequence, ligand‐induced activation is severely attenuated, even in the presence of ectopically expressed TRAP220/TIF2. Conversely, overexpression of WSTF and either ATPase subunit (BRG1 or hBRM) enhances ligand‐ mediated activation. In functional assays, WINAC is able to disrupt nucleosome arrays in the region of VDR binding. Thus, WINAC, in a similar fashion to other SWI/SNF complexes, enables targeting of VDR to promoters by disrupting chromatin structure in and around the VDRE. In addition to regulating positive VDREs, WINAC is also required for VDR‐mediated transrepression of a negative VDRE present in the 1a(OH) ase promoter.185 The 1a(OH)ase promoter lacks a classical or positive VDRE. Therefore, repression is not through direct binding of VDR. Instead, a bHLH‐ type activator, VDR‐interacting repressor (VDIR) directly binds to the negative VDRE present in the 1a(OH)ase promoter to mediate activation or transrepression. In the absence of ligand, VDIR associates strongly with coactivators (p300) and only weakly with VDR. Recruitment of p300 leads to activation of 1a(OH)ase gene expression. In the presence of ligand, VDR strongly associates with VDIR and NCoR. Ligand‐induced recruitment of VDR and NCoR to the 1a(OH)ase promoter results in HDAC‐mediated transrepression. Knockdown of VDIR, VDR, or WSTF relieves ligand‐induced transrepression at 1a(OH) ase gene, suggesting a critical role for all three complexes. In the absence of ligand, the VDR/WINAC complex is present at the 1a (OH)ase promoter.185 Ablation of VDR prevents WINAC from associating with the 1a(OH)ase promoter, suggesting that the weak VDR–VDIR interaction is necessary for directing the VDR/WINAC complex to the 1a(OH)ase promoter in the absence of ligand. Conversely, knockdown of WSTF prevents VDR recruitment, suggesting that WINAC facilitates retention of VDR on the promoter in the absence of ligand. WSTF cannot directly interact with VDIR, but does harbor two well characterized chromatin‐binding domains: a bromodomain and a PHD finger. The WSTF bromodomain is sufficient to bind acetylated histones in vitro and preferentially binds to acetylated H3K14 in vivo. Deletion of the WSTF bromodomain prevents recruitment of the VDR/WINAC complex to the 1a(OH)ase promoter and inhibits ligand‐ mediated transrepression of gene expression. The WSTF gene is deleted in Williams syndrome patients. Infants with the disease display abnormal Vitamin D metabolism. WSTF serves as a platform protein for WINAC complex assembly and directly interacts with VDR.184 WINAC regulates the expression of two genes, whose products are involved in regulating Vitamin D metabolism [24(OH)ase and 1a(OH)ase].184,185 Therefore, WSTF is proposed to play a critical role in VDR function. Overexpression of WSTF in fibroblasts obtained from a Williams syndrome patient rescues

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VDR recruitment to 24(OH)ase and 1a(OH)ase promoters as well as responsiveness to Vitamin D. These results strongly suggest that disrupted Vitamin D metabolism in these patients is likely due to WINAC dysfunction.184,185 WINAC does not associate with ER in the presence or absence of estrogen ligand.184 In addition, it does not appear to be required for transactivation of other members of the NR family, including PPARg. These studies, therefore, involve the WINAC complex exclusively in VDR function. It is not clear what the determinant of this exclusive interaction is, and whether other nuclear hormone receptors that were not included in the above studies interact with WSTF. Although WINAC is clearly required for VDR regulation of 24(OH)ase and 1a(OH)ase, its involvement in regulating other VDR target genes has not been characterized.184,185 At a positive VDRE [24(OH)ase promoter], WINAC restructures chromatin to allow binding of VDR and/or VDR coregulators.184 It is not known if all VDR targets also require a similar WINAC‐mediated restructuring. It is entirely possible that the chromatin structure at a subset of VDREs is amenable to VDR binding in the absence of WINAC remodeling. Two nonmutually exclusive models could be envisioned to explain the function of WINAC in VDR action: (1) WINAC is required for the stable ligand‐independent recruitment of VDR to a subset of VDREs and (2) WINAC is required at steps following VDR recruitment. In the former scenario, regulation of VDR targets genes would primarily be mediated by the action of known coregulators of VDR action (i.e., NCoR/SMRT or SRC‐1/p300/CBP). In the latter scenario, WINAC could restructure chromatin at the promoter, possibly in a ligand‐dependent manner, to prevent or allow recruitment of additional coregulators. WINAC‐mediated recruitment of VDR to a negative VDRE [1a(OH)ase] requires the bromodomain of WSTF.185 This domain is responsible for anchoring the VDR/WINAC complex to acetylated histones. Whether this domain is required for the recruitment of the VDR/WINAC complex to positive VDREs is not known. The bromodomain of WSTF could be postulated to further stabilize recruitment of the NR in the presence of ligand (highly acetylated histones).

C. Acf1 and ISWI Complexes and NR Regulation Chromatin remodeling complexes containing the ISWI ATPase were first identified in in vitro nucleosome‐remodeling assays of Drosophila embryo extracts.186 Homologous proteins have since been discovered in yeast and mammals. The ISWI family members, known as SNF2h in mammals, include ACF (ATP‐utilizing chromatin assembly and remodeling complex), CHRAC (chromatin accessibility complex, composed of ACF plus two additional subunits), WSTF (Williams syndrome transcription factor), NURF, WICH, NoRC, and RSF (remodeling and spacing factor).186,187 In vivo and in vitro studies

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suggest that ISWI family members function quite differently from SWI/SNF family members. ISWI members are capable of assembling chromatin with evenly spaced nucleosomes in vitro. In vivo they have been shown to play major roles in DNA replication and transcriptional repression.188 In Drosophila, ISWI is associated with hundreds of euchromatic sites in salivary gland polytene chromosomes in a pattern that is nonoverlapping with Pol II.189 This pattern contrasts dramatically with SWI/SNF complexes, suggesting that ISWI complexes function primarily to transcriptionally repress genes. Genetic studies in yeast also suggest a role for ISWI in repression, as yeast ISWI (Isw1/Isw2) mutants exhibit derepression of meiotic genes. Isw2 can establish nuclease‐inaccessible chromatin structures at specific sites in vivo and repress early meiotic genes in parallel with Sin3‐Rpd3 (a yeast HDAC).190 These mutants exhibit defects during early stages of sporulation. In vitro, Isw2 can counteract transcriptional activation by sliding nucleosomes at specific positions in vitro and displacing bound activators from their chromatin target sites.191–193 ISWI complexes are crucial for mammalian development as well, as SNF2h null mice die during the peri‐implantation stage and SNF2h knockdown inhibits cytokine induced differentiation of human hematopoietic progenitor cells into erythrocytes.194 The ACF complex, consisting of Acf1 and SNF2h assembles phased nucleosome arrays in vitro, signifying a chromatin assembly function.186 In Drosophila, loss of Acf1 results in decreased periodicity of nucleosome arrays.189 Periodic nucleosomes are thought to be repressive to DNA‐based processes (i.e., replication, transcription) suggesting a role for Acf1 in the formation of repressive chromatin. Null animals display additional phenotypic defects, including suppression of PEV (position effect variegation) and increased expression of Polycomb repressed genes, further implicating Acf1 in gene repression.189 In addition, Acf1 and ISWI are required for the basal repression of Wingless targets.195 A role for Acf1 and Snf2h in NR‐mediated repression has also recently been described.84,85 While the ACF complex is predominantly associated with transcriptional repression, the NURF complex primarily functions to activate transcription. 1. ISWI COMPLEXES AND NR ACTIVATION Transactivation by the PR is enhanced by Drosophila NURF.196 Cell‐free studies suggest a two step synergy by which liganded PR first binds to the HRE, thereby provoking the NURF ATP‐dependent nuclear remodeling event. The altered chromatin structure renders additional internal receptor promoter‐ binding sites available thus facilitating transactivation or repression by PR. There is in vivo evidence to support a role for NURF in transcriptional activation by NRs. Disruption of the Drosophila NURF complex by null

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mutation interferes with transactivation of several genes targeted by the steroid hormone, ecdysone.197 The ecdysone receptor (EcR) immunoprecipitates with NURF suggesting that EcR targets may require NURF for activation. 2. ISWI COMPLEXES AND NR REPRESSION Transcription can occur on naked DNA in the absence of ligand, whereas ligand is required for transcription on chromatin templates. This suggests that chromatin is inherently inhibitory to transcriptional processes and that the recruitment of chromatin remodeling activities by NRs overcomes this barrier (likely by disrupting chromatin structure). In the absence of ligand, Class II NRs are capable of binding DNA and recruiting cofactors to actively repress transcription. The roles of histone modifying enzymes (HDACs and HMTs) in NR‐mediated transcriptional repression have been extensively studied. And while, ATP‐dependent chromatin remodelers have been linked to gene activation by NRs, their role in NR‐mediated repression is not clear. Using purified chromatin templates, Dilworth et al. showed that two ATP‐ dependent chromatin remodelers are required at distinct stages of transcription by the RAR/RXR heterodimer.42 Snf2h‐containing complexes, isolated from HeLa cell extracts, are required for strong binding of the RAR/RXR heterodimer to the chromatin template. In the absence of ATP, binding was substantially weakened, suggesting a direct role for ATP‐dependent chromatin remodeling at this stage. The addition of p300/TIF2 and/or SWI/SNF complexes were unable to compensate for the Snf2h‐containing complexes, but were involved in activating transcription in a steps subsequent to NR binding. It is not clear from these studies, which Snf2h complex or complexes are involved in the remodeling event and whether they are specifically recruited to HREs by the NR complex. Furthermore, this study did not address whether the chromatin architecture established by the Snf2h‐containing complex was required to mediate repression. Alenghat et al. recently identified the ATPase Snf2h as a component that contributes to repression by unliganded TR.84 Knockdown of Snf2h leads to a concomitant increase in the expression of a genomically integrated TR‐regulated reporter gene as well as the endogenous TR target, dio1. Knockdown of NCoR prevents recruitment of Snf2h to the genomically integrated promoter. As NCoR and Snf2h do not interact by immunoprecipitation, the recruitment of Snf2h to chromatin by NCoR is mostly likely indirect. Snf2h has previously been shown to associate with deacetylated H4 tails (more specifically deacetylated H4K16). Therefore, NCoR/HDAC3 may indirectly recruit Snf2h by deacetylating histones. Two pieces of evidence support this hypothesis: (1) HDAC3 knockdown increased acetylation at the genomically integrated promoter leading to decreased recruitment of Snf2h. (2) HDAC inhibitor treatment also increased promoter acetylation and decreased recruitment of Snf2h at the endogenous dio1 promoter. Interestingly, although HDAC inhibition

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decreased recruitment of Snf2h, it did not alter binding of TR and NCoR. Although not directly tested, this suggests that derepression of TR targets by Snf2h disruption is not due to perturbation of TR/NCoR function. Snf2h or HDAC3 knockdown similarly altered MNase accessibility at the genomically integrated promoter: decreased protection at the transcriptional start site and increased protection at promoter. Together, these results suggest that Snf2h is required to repress TR target genes and that it does so by altering chromatin structure. The mechanism of repression mediated by Snf2h and whether it functions alone is not entirely clear. In vitro and in vivo, Snf2h‐interacting partners (Acf1, WSTF, NURF, TIP5) qualitatively and quantitatively modulate Snf2h ATPase activity. For example, the ACF complex remodels nucleosomes an order of magnitude more efficiently than Snf2h alone.198,199 Although Snf2h possesses nucleosome remodeling activities in vitro, it has never been purified from nuclear extracts on its own. Does Snf2h act alone to repress TR target genes, or does it function as part of a complex? Evidence suggests that chromatin remodeling enzymes are recruited to chromatin by site‐specific DNA factors. It has been proposed that the nonenzymatic subunits of these ATP‐dependent chromatin remodeling complexes may guide the complexes to specific sites. Is Snf2h recruited globally to all deacetylated H4K16 sites or do Snf2h‐interacting partners provide recruitment specificity to TR target genes? At the genomically integrated promoter, Snf2h appears to maintain a specific chromatin architecture. How Snf2h contributes to this architecture, how this architecture mediates repression, and whether a similar architecture is required to repress endogenous TR (or additional NR) target genes is not clear. In the absence of ligand, NRs recruit NCoR/SMRT complexes to establish repression at NR target genes. NCoR/SMRT functions as a scaffold to assemble corepressors, including chromatin remodeling enzymes. However, evidence of a direct involvement of ATP‐dependent chromatin remodeling activity in NCoR/SMRT‐mediated repression has been limited. In an attempt to identify novel NCoR interaction partners in our laboratory, Ewing et al. utilized a yeast‐two hybrid screen, using an NCoR repression domain (RD2) as a bait.85 This strategy identified a novel interaction with Acf1, a subunit of the ACF complex (Fig. 1B). Subsequent studies demonstrate a strong association between the N‐terminus of Acf1 and both RD1 and RD2 of NCoR. The N‐terminal amino acids (1–313) of Acf1 are sufficient for interaction with both NCoR and SMRT. This domain was thus named the NCoR‐interaction domain (NID). The NID is necessary for NCoR‐Acf1 interaction, as deletion of this domain perturbs interaction with RD1. The NID encompasses a WAC domain, which has previously been demonstrated to bind DNA and heterochromatin. Endogenous Acf1 coimmunoprecipitates NCoR suggesting an association in vivo.

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Overexpression of Acf1 enhances repression at the VDR‐regulated genes IGFBP3 and RANKL and a TR‐regulated gene IGF‐I.85 Overexpression of the NID domain of Acf1 or siRNA‐mediated knockdown of endogenous Acf1, leads to derepression of the same set of genes, but not the VDR‐regulated gene 24(OH)lase, or the RAR‐regulated gene IGFBP6. The expression of RANKL, IGFBP3, and IGF‐I mRNA by Acf1 knockdown closely mimics that is seen upon hormone induced activation. These results suggest that: (1) Acf1 is required for the repression of a subset of Class II NR targets, (2) release of Acf1 may be sufficient to activate some NR targets, and (3) additional Acf1 domains beyond the NID are required to mediate the repressive effects of Acf1. In the absence of hormone, NCoR and Acf1 are recruited to the promoters of IGFBP3 and RANKL (assayed by ChIP).85 Hormone treatment leads to release of NCoR and Acf1 from both promoters, further supporting an interaction between these two proteins. Interestingly, Acf1 knockdown also releases NCoR from both promoters, suggesting a role for Acf1 in stabilizing NCoR binding to the promoter. Different mechanisms appear to regulate Acf1‐ mediated repression of RANKL and IGFBP3. At the IGFBP3 promoter, hormone treatment or Acf1 knockdown leads to a decrease in trimethylation at H3K9. This modification is present at heterochromatic regions as well as in the promoters of transcriptionally repressed genes. At the RANKL promoter however, hormone treatment or Acf1 knockdown decreases occupancy of H3/ H4 and increases MNase accessibility at the promoter, suggesting hormone induced nucleosome remodeling. Many studies have established a role for the ACF complex in chromatin assembly and nucleosome phasing.186,189 Several studies have suggested that Acf1 may also be involved in transcriptional repression because it plays a role in heterochromatin replication.200 However, the role of Acf1 in the repression of nonheterochromatic genes had not been investigated. This study clearly demonstrates a role for Acf1 in transcriptional repression of mammalian euchromatic gene targets.85 More specifically, Acf1 is recruited to VDR targets as part of an NCoR complex, stabilizes the binding of NCoR to chromatin, and modifies chromatin to establish a repressive chromatin structure. As Acf1 knockdown leads to gene expression equivalent to hormone‐induced activation, derepression may be a major mechanism of gene activation for some NR targets. Acf1 knockdown does not alter the expression of 24(OH)ase or IGFBP6, suggesting that Acf1 is not absolutely required for the repression of all NR target genes. Alternatively, Acf1 is required for repression, but knockdown of Acf1 alone is not sufficient for derepression. Consistent with this suggestion is the observation that in yeast, the Isw2 complex mediates repression of Ume6 targets in parallel with the HDAC, Rpd3.190 Full derepression of some Ume6 targets requires disruption of both Isw2/Rpd3. Future studies could determine whether Acf1 is recruited to the 24(OH)ase promoter and whether Acf1 and NCoR/HDAC3 cooperate to mediate repression. Additional

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transcription factors, including the nuclear orphan receptor COUP‐TF, vErbA, and most recently AP‐1 were shown to recruit NCoR/SMRT to mediate repression. A requirement for Acf1 in the repression of these and other Class II NR targets warrants further investigation. WSTF associates in a ligand‐independent manner with VDR as part of the WINAC complex184 The association is mediated by the WSTFm1 domain (VDR interaction domain) located C‐terminal to the WAC domain of WSTF. Acf1 and WSTF are paralogs and share similar structural features, including the domains, WAC, DDT, WAKZ, PHD, and a bromodomain. When associated with Snf2h, they form ACF and WICH complexes, which display distinct functions in vivo. It is not clear whether Acf1 and WSTF display some redundancy in regards to NR gene regulation or also play specific roles.85 Several interesting questions remain to be investigated. WSTF shares a highly homologous WAC domain with Acf1. Can NCoR/SMRT recruit the WINAC and/or WICH complex to NR gene targets? Conversely, can Acf1 interact directly with VDR through its own VDR interaction domain? This last scenario is unlikely as Acf1 was not identified as a VDR interactor in the biochemical purification scheme used to identify the WINAC complex. The general role Acf1 has in NR signaling also remains to be determined. Acf1 and Snf2h are the mammalian homologs of the yeast Itc1p and Isw2 subunits.190 The yeast Isw2 complex is recruited to DNA by the transcriptional repressor Ume6 through interaction with the Itc1p subunit and mediates repression by sliding/positioning nucleosomes.193 Acf1 is recruited to DNA in a similar fashion via interaction with corepressors NCoR/SMRT and is required for the repression of a subset of VDR and TR target genes. Whether Snf2h is recruited to chromatin by Acf1 and whether a functional ACF complex is required to establish repression at these targets is not clear. Snf2h may be recruited to deacetylated histones in a nonspecific manner. This supports the observation that Acf1 knockdown does not appear to alter recruitment of Snf2h.85 Snf2h alone has limited functionality in vitro and requires the association of additional subunits (Acf1, WSTF) for full activity. As deletion of these associated subunits perturbs chromatin based processes, it is unlikely that Snf2h alone is sufficient to mobilize nucleosomes in vivo. These additional subunits, recruited to specific sites on DNA, may function to dictate Snf2h activity, and ultimately nucleosome positioning. Acf1 knockdown as well as hormone treatment significantly reduced H3/H4 at the transcription start sit of RANKL, suggesting a loss or sliding of nucleosomes from the region. These results strongly suggest that a novel role for hormone is to promote histone eviction to prepare a gene for hormonal activation. Whether this histone eviction is accompanied by histone exchange at target promoters is not clear and should be investigated. Finally, determining whether nucleosome positioning by ACF plays a role in NR‐mediated repression would also be critical. We propose that Acf1 together with Snf2h function to stabilize a repressive nuclear

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architecture. Hormone treatment releases the Acf1‐containing NCoR/SMRT complex allowing nucleosome mobilization and subsequent recruitment of coactivators, the basal transcription machinery, and Pol II (Fig. 2). In addition to its hypothesized role in ATP‐dependent remodeling of nucleosomes as a component of the ACF complex, Acf1 alone may also possess unique repression functions. Acf1 contains PHD and bromodomains that are dispensable for chromatin assembly by ACF. Interestingly, these domains are not present in the yeast homolog of Acf1, Itc1p. The function of these domains for Acf1 is not known, but has been characterized for other family members. For example, the PHD and bromodomain of TIP5, a component of the NoRC complex, is required for the recruitment of NoRC to acetylated rDNA.201 In addition, both domains are required for recruiting HDACs and DNMTs to rDNA promoters.202 Therefore, TIP5 functions to (1) position nucleosomes by directing Snf2h ATPase activity and (2) recruit repressive histone modifying enzymes to the promoter. The bromodomain of WSTF is required for recruitment of WINAC to acetylated negative VDREs.185 Immediately following removal of hormone, NR target promoters remain hyperacetylated. The bromodomain of Acf1 may then be required for efficient recruitment of the NCoR/SMRT complex to hyperacetylated chromatin templates to reestablish repression. In addition, the dual module PHD/bromodomain may function to recruit additional corepressor molecules to promoters. A requirement for PHD and/or bromodomain‐mediated recruitment of NCoR/SMRT to HREs could potentially explain why Acf1 knockdown destabilizes recruitment of NCoR at these sites.85 Alternatively, a nucleosome positioning function of ACF may be required to organize chromatin structure, allowing NRs and coregulators to bind chromatin. Acf1 knockdown did not derepress all NR targets genes tested.85 We, therefore, hypothesize that Acf1 is required to maintain repression at only a subset of NR targets. Derepression of RANKL and IGFBP3 by Acf1 knockdown results in an equivalent level of gene expression as compared to ligand treatment, suggesting that derepression may be sufficient to activate some genes. Acf1 knockdown did not affect 24(OH)ase gene expression, suggesting either an Acf1‐independent mechanism of gene repression, or a coactivator‐ dependent mechanisms of gene activation (Fig. 3).

VI. Future Directions Despite the wealth of knowledge regarding the coregulators that are involved in NR action, it is not entirely clear how the various functions of these coregulators are coordinated and integrated in vivo. The PTMs on histones appear to specify particular transcriptional states. How this ‘‘histone code’’ functions to coordinate and recruit ATP‐dependent chromatin remodelers to DNA, and exactly how these enzymes function to direct

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FIG. 2. Chromatin remodelers play a role in NR‐mediated activation and repression. In the absence of ligand, NRs recruit corepressor complexes such as NCoR/SMRT. NCoR/SMRT functions as a scaffold to bring chromatin remodeling factors to chromatin. HDACs, such as HDAC3, deacetylate histones, while ATP‐dependent remodelers, such as the ACF complex, potentially alter nucleosome position and chromatin structure to repress transcription. In the presence of ligand, NRs undergo a conformational change, which leads to the release of corepressor complexes and subsequent recruitment of coactivator complexes. Acetylation of histones by HATs and nucleosome remodeling by SWI/SNF increase accessibility of the DNA to general transcription factors and activate transcription.

transcription are only now being elucidated. Recent genome‐wide nucleosome mapping studies suggest that nucleosome positions somehow direct transcriptional machinery to appropriate sites. How nucleosome positions are affected by NR transcription has not been extensively studied. Therefore, understanding the mechanisms by which the underlying DNA template, as well as histone modifying enzymes function to dictate these nucleosome positions may shed light on the role coregulators play in the transcriptional process. NRs modulate chromatin structure to regulate transcriptional processes. Numerous studies have investigated the role of NR coregulators, including histone modifying enzymes and ATP‐dependent chromatin remodelers, in

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FIG. 3. Differential regulation of NR target genes. (A) Gene activation at a subset of NR target genes may only require release of NCoR/SMRT‐mediated repression, whereas other targets may require the additional recruitment of coactivators (B).

transcriptional activation. The role these coregulators play in NR‐mediated repression are not entirely clear. Class II NRs recruit corepressors that actively function to repress gene transcription. A role for HDAC activity in repression has been well documented. However, the role ATP‐dependent chromatin remodelers play in NR‐mediated repression has not been well characterized. At Class II NR promoters, do ATP‐dependent chromatin remodelers specify ‘‘repressive’’ nucleosome positions? The role of Isw2 in maintaining repression of meiotic target genes is well established. Could the ACF complex function in a similar fashion to repress NR targets? How do non class II NRs repress target gene transcription? Does the underlying DNA sequence at these promoters dictate a default ‘‘off’’ state? Future experiments utilizing promoter specific and genome‐wide nucleosome positioning assays in an NR model will uncover the answer to some of these questions. Acknowledgments The original work performed in our laboratory and described in this review was supported by an NIH grant R01DK65148 (to D.C.). We thank Amy Ewing for contributing the illustrations in Fig. 3.

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References 1. Mangelsdorf DJ, Thummel C, Beato M, Herrlich P, Schutz G, Umesono K, et al. The nuclear receptor superfamily: the second decade. Cell 1995;83:835–9. 2. Francis GA, Fayard E, Picard F, Auwerx J. Nuclear receptors and the control of metabolism. Annu Rev Physiol 2003;65:261–311. 3. Overington JP, Al‐Lazikani B, Hopkins AL. How many drug targets are there? Nat Rev Drug Discov 2006;5:993–6. 4. Gronemeyer H, Gustafsson JA, Laudet V. Principles for modulation of the nuclear receptor superfamily. Nat Rev Drug Discov 2004;3:950–64. 5. Jensen EV, Jordan VC. The estrogen receptor: a model for molecular medicine. Clin Cancer Res 2003;9:1980–9. 6. Hollenberg SM, Weinberger C, Ong ES, Cerelli G, Oro A, Lebo R, et al. Primary structure and expression of a functional human glucocorticoid receptor cDNA. Nature 1985;318:635–41. 7. Htun H, Holth LT, Walker D, Davie JR, Hager GL. Direct visualization of the human estrogen receptor alpha reveals a role for ligand in the nuclear distribution of the receptor. Mol Biol Cell 1999;10:471–86. 8. Tsai MJ, O’Malley BW. Molecular mechanisms of action of steroid/thyroid receptor superfamily members. Annu Rev Biochem 1994;63:451–86. 9. Umesono K, Evans RM. Determinants of target gene specificity for steroid/thyroid hormone receptors. Cell 1989;57:1139–46. 10. Glass CK, Rosenfeld MG. The coregulator exchange in transcriptional functions of nuclear receptors. Genes Dev 2000;14:121–41. 11. Ribeiro RC, Kushner PJ, Apriletti JW, West BL, Baxter JD. Thyroid hormone alters in vitro DNA binding of monomers and dimers of thyroid hormone receptors. Mol Endocrinol 1992;6:1142–52. 12. Laudet V, Adelmant G. Nuclear receptors. Lonesome orphans. Curr Biol 1995;5:124–7. 13. Yin L, Wu N, Curtin JC, Qatanani M, Szwergold NR, Reid RA, et al. Rev‐erbalpha, a heme sensor that coordinates metabolic and circadian pathways. Science 2007;318:1786–9. 14. Raghuram S, Stayrook KR, Huang P, Rogers PM, Nosie AK, McClure DB, et al. Identification of heme as the ligand for the orphan nuclear receptors REV‐ERBalpha and REV‐ERBbeta. Nat Struct Mol Biol 2007;14:1207–13. 15. Burris TP. Nuclear hormone receptors for heme: REV‐ERBalpha and REV‐ERBbeta are ligand‐regulated components of the mammalian clock. Mol Endocrinol 2008;22:1509–20. 16. Benoit G, Malewicz M, Perlmann T. Digging deep into the pockets of orphan nuclear receptors: insights from structural studies. Trends Cell Biol 2004;14:369–76. 17. Bourguet W, Germain P, Gronemeyer H. Nuclear receptor ligand‐binding domains: three‐ dimensional structures, molecular interactions and pharmacological implications. Trends Pharmacol Sci 2000;21:381–8. 18. Giguere V, Hollenberg SM, Rosenfeld MG, Evans RM. Functional domains of the human glucocorticoid receptor. Cell 1986;46:645–52. 19. Kumar V, Green S, Stack G, Berry M, Jin JR, Chambon P. Functional domains of the human estrogen receptor. Cell 1987;51:941–51. 20. Luisi BF, Xu WX, Otwinowski Z, Freedman LP, Yamamoto KR, Sigler PB. Crystallographic analysis of the interaction of the glucocorticoid receptor with DNA. Nature 1991;352:497–505. 21. Barettino D, Vivanco Ruiz MM, Stunnenberg HG. Characterization of the ligand‐dependent transactivation domain of thyroid hormone receptor. EMBO J 1994;13:3039–49.

226

BURANAPRAMEST AND CHAKRAVARTI

22. Danielian PS, White R, Lees JA, Parker MG. Identification of a conserved region required for hormone dependent transcriptional activation by steroid hormone receptors. EMBO J 1992;11:1025–33. 23. Durand B, Saunders M, Gaudon C, Roy B, Losson R, Chambon P. Activation function 2 (AF‐2) of retinoic acid receptor and 9‐cis retinoic acid receptor: presence of a conserved autonomous constitutive activating domain and influence of the nature of the response element on AF‐2 activity. EMBO J 1994;13:5370–82. 24. Chandra V, Huang P, Hamuro Y, Raghuram S, Wang Y, Burris TP, et al. Structure of the intact PPAR‐gamma‐RXR‐alpha nuclear receptor complex on DNA. Nature 350–356. 25. Ing NH, Beekman JM, Tsai SY, Tsai MJ, O’Malley BW. Members of the steroid hormone receptor superfamily interact with TFIIB (S300‐II). J Biol Chem 1992;267:17617–23. 26. Brou C, Wu J, Ali S, Scheer E, Lang C, Davidson I, et al. Different TBP‐associated factors are required for mediating the stimulation of transcription in vitro by the acidic transactivator GAL‐VP16 and the two nonacidic activation functions of the estrogen receptor. Nucleic Acids Res 1993;21:5–12. 27. Klein‐Hitpass L, Schwerk C, Kahmann S, Vassen L. Targets of activated steroid hormone receptors: basal transcription factors and receptor interacting proteins. J Mol Med 1998;76:490–6. 28. McKenna NJ, Xu J, Nawaz Z, Tsai SY, Tsai MJ, O’Malley BW. Nuclear receptor coactivators: multiple enzymes, multiple complexes, multiple functions. J Steroid Biochem Mol Biol 1999;69:3–12. 29. Kininis M, Isaacs GD, Core LJ, Hah N, Kraus WL. Postrecruitment regulation of RNA polymerase II directs rapid signaling responses at the promoters of estrogen target genes. Mol Cell Biol 2009;29:1123–33. 30. Belakavadi M, Fondell JD. Role of the mediator complex in nuclear hormone receptor signaling. Rev Physiol Biochem Pharmacol 2006;156:23–43. 31. Onate SA, Tsai SY, Tsai MJ, O’Malley BW. Sequence and characterization of a coactivator for the steroid hormone receptor superfamily. Science 1995;270:1354–7. 32. Kamei Y, Xu L, Heinzel T, Torchia J, Kurokawa R, Gloss B, et al. A CBP integrator complex mediates transcriptional activation and AP‐1 inhibition by nuclear receptors. Cell 1996;85:403–14. 33. Chakravarti D, LaMorte VJ, Nelson MC, Nakajima T, Schulman IG, Juguilon H, et al. Role of CBP/P300 in nuclear receptor signalling. Nature 1996;383:99–103. 34. Chen H, Lin RJ, Xie W, Wilpitz D, Evans RM. Regulation of hormone‐induced histone hyperacetylation and gene activation via acetylation of an acetylase. Cell 1999;98:675–86. 35. Shang Y, Hu X, DiRenzo J, Lazar MA, Brown M. Cofactor dynamics and sufficiency in estrogen receptor‐regulated transcription. Cell 2000;103:843–52. 36. Sharma D, Fondell JD. Ordered recruitment of histone acetyltransferases and the TRAP/ Mediator complex to thyroid hormone‐responsive promoters in vivo. Proc Natl Acad Sci USA 2002;99:7934–9. 37. Koh SS, Chen D, Lee YH, Stallcup MR. Synergistic enhancement of nuclear receptor function by p160 coactivators and two coactivators with protein methyltransferase activities. J Biol Chem 2001;276:1089–98. 38. Jenuwein T, Allis CD. Translating the histone code. Science 2001;293:1074–80. 39. Strahl BD, Allis CD. The language of covalent histone modifications. Nature 2000;403:41–5. 40. Chen J, Kinyamu HK, Archer TK. Changes in attitude, changes in latitude: nuclear receptors remodeling chromatin to regulate transcription. Mol Endocrinol 2006;20:1–13. 41. Urnov FD, Wolffe AP. A necessary good: nuclear hormone receptors and their chromatin templates. Mol Endocrinol 2001;15:1–16.

CHROMATIN REMODELING AND NUCLEAR RECEPTOR SIGNALING

227

42. Dilworth FJ, Fromental‐Ramain C, Yamamoto K, Chambon P. ATP‐driven chromatin remodeling activity and histone acetyltransferases act sequentially during transactivation by RAR/ RXR In vitro. Mol Cell 2000;6:1049–58. 43. Fryer CJ, Archer TK. Chromatin remodelling by the glucocorticoid receptor requires the BRG1 complex. Nature 1998;393:88–91. 44. Huang ZQ, Li J, Sachs LM, Cole PA, Wong J. A role for cofactor‐cofactor and cofactor‐histone interactions in targeting p300, SWI/SNF and Mediator for transcription. EMBO J 2003;22:2146–55. 45. Muchardt C, Yaniv M. ATP‐dependent chromatin remodelling: SWI/SNF and Co. are on the job. J Mol Biol 1999;293:187–98. 46. Yoshinaga SK, Peterson CL, Herskowitz I, Yamamoto KR. Roles of SWI1, SWI2, and SWI3 proteins for transcriptional enhancement by steroid receptors. Science 1992;258:1598–604. 47. Johnson CN, Adkins NL, Georgel P. Chromatin remodeling complexes: ATP‐dependent machines in action. Biochem Cell Biol 2005;83:405–17. 48. Baniahmad A. Nuclear hormone receptor co‐repressors. J Steroid Biochem Mol Biol 2005;93:89–97. 49. Baniahmad A, Leng X, Burris TP, Tsai SY, Tsai MJ, O’Malley BW. The tau 4 activation domain of the thyroid hormone receptor is required for release of a putative corepressor(s) necessary for transcriptional silencing. Mol Cell Biol 1995;15:76–86. 50. Casanova J, Helmer E, Selmi‐Ruby S, Qi JS, Au‐Fliegner M, Desai‐Yajnik V, et al. Functional evidence for ligand‐dependent dissociation of thyroid hormone and retinoic acid receptors from an inhibitory cellular factor. Mol Cell Biol 1994;14:5756–65. 51. Tong GX, Jeyakumar M, Tanen MR, Bagchi MK. Transcriptional silencing by unliganded thyroid hormone receptor beta requires a soluble corepressor that interacts with the ligand‐ binding domain of the receptor. Mol Cell Biol 1996;16:1909–20. 52. Chen JD, Evans RM. A transcriptional co‐repressor that interacts with nuclear hormone receptors. Nature 1995;377:454–7. 53. Horlein AJ, Naar AM, Heinzel T, Torchia J, Gloss B, Kurokawa R, et al. Ligand‐independent repression by the thyroid hormone receptor mediated by a nuclear receptor co‐repressor. Nature 1995;377:397–404. 54. Jepsen K, Rosenfeld MG. Biological roles and mechanistic actions of co‐repressor complexes. J Cell Sci 2002;115:689–98. 55. Privalsky ML. The role of corepressors in transcriptional regulation by nuclear hormone receptors. Annu Rev Physiol 2004;66:315–60. 56. Hu X, Lazar MA. The CoRNR motif controls the recruitment of corepressors by nuclear hormone receptors. Nature 1999;402:93–6. 57. Nagy L, Kao HY, Love JD, Li C, Banayo E, Gooch JT, et al. Mechanism of corepressor binding and release from nuclear hormone receptors. Genes Dev 1999;13:3209–16. 58. Perissi V, Staszewski LM, McInerney EM, Kurokawa R, Krones A, Rose DW, et al. Molecular determinants of nuclear receptor‐corepressor interaction. Genes Dev 1999;13:3198–208. 59. Cohen RN, Brzostek S, Kim B, Chorev M, Wondisford FE, Hollenberg AN. The specificity of interactions between nuclear hormone receptors and corepressors is mediated by distinct amino acid sequences within the interacting domains. Mol Endocrinol 2001;15:1049–61. 60. Hu X, Li Y, Lazar MA. Determinants of CoRNR‐dependent repression complex assembly on nuclear hormone receptors. Mol Cell Biol 2001;21:1747–58. 61. Ishizuka T, Lazar MA. The N‐CoR/histone deacetylase 3 complex is required for repression by thyroid hormone receptor. Mol Cell Biol 2003;23:5122–31. 62. Ordentlich P, Downes M, Xie W, Genin A, Spinner NB, Evans RM. Unique forms of human and mouse nuclear receptor corepressor SMRT. Proc Natl Acad Sci USA 1999;96:2639–44.

228

BURANAPRAMEST AND CHAKRAVARTI

63. Guenther MG, Barak O, Lazar MA. The SMRT and N‐CoR corepressors are activating cofactors for histone deacetylase 3. Mol Cell Biol 2001;21:6091–101. 64. Ishizuka T, Lazar MA. The nuclear receptor corepressor deacetylase activating domain is essential for repression by thyroid hormone receptor. Mol Endocrinol 2005;19:1443–51. 65. Webb P, Anderson CM, Valentine C, Nguyen P, Marimuthu A, West BL, et al. The nuclear receptor corepressor (N‐CoR) contains three isoleucine motifs (I/LXXII) that serve as receptor interaction domains (IDs). Mol Endocrinol 2000;14:1976–85. 66. Hartman HB, Yu J, Alenghat T, Ishizuka T, Lazar MA. The histone‐binding code of nuclear receptor co‐repressors matches the substrate specificity of histone deacetylase 3. EMBO Rep 2005;6:445–51. 67. Yu J, Li Y, Ishizuka T, Guenther MG, Lazar MA. A SANT motif in the SMRT corepressor interprets the histone code and promotes histone deacetylation. EMBO J 2003;22:3403–10. 68. Li J, Wang J, Nawaz Z, Liu JM, Qin J, Wong J. Both corepressor proteins SMRT and N‐CoR exist in large protein complexes containing HDAC3. EMBO J 2000;19:4342–50. 69. Yoon HG, Chan DW, Huang ZQ, Li J, Fondell JD, Qin J, et al. Purification and functional characterization of the human N‐CoR complex: the roles of HDAC3, TBL1 and TBLR1. EMBO J 2003;22:1336–46. 70. Zhang J, Kalkum M, Chait BT, Roeder RG. The N‐CoR‐HDAC3 nuclear receptor corepressor complex inhibits the JNK pathway through the integral subunit GPS2. Mol Cell 2002;9:611–23. 71. Perissi V, Aggarwal A, Glass CK, Rose DW, Rosenfeld MG. A corepressor/coactivator exchange complex required for transcriptional activation by nuclear receptors and other regulated transcription factors. Cell 2004;116:511–26. 72. Heinzel T, Lavinsky RM, Mullen TM, Soderstrom M, Laherty CD, Torchia J, et al. A complex containing N‐CoR, mSin3 and histone deacetylase mediates transcriptional repression. Nature 1997;387:43–8. 73. Nagy L, Kao HY, Chakravarti D, Lin RJ, Hassig CA, Ayer DE, et al. Nuclear receptor repression mediated by a complex containing SMRT, mSin3A, and histone deacetylase. Cell 1997;89:373–80. 74. Guenther MG, Lane WS, Fischle W, Verdin E, Lazar MA, Shiekhattar R. A core SMRT corepressor complex containing HDAC3 and TBL1, a WD40‐repeat protein linked to deafness. Genes Dev 2000;14:1048–57. 75. Li J, Lin Q, Wang W, Wade P, Wong J. Specific targeting and constitutive association of histone deacetylase complexes during transcriptional repression. Genes Dev 2002;16:687–92. 76. Jones PL, Sachs LM, Rouse N, Wade PA, Shi YB. Multiple N‐CoR complexes contain distinct histone deacetylases. J Biol Chem 2001;276:8807–11. 77. Huang EY, Zhang J, Miska EA, Guenther MG, Kouzarides T, Lazar MA. Nuclear receptor corepressors partner with class II histone deacetylases in a Sin3‐independent repression pathway. Genes Dev 2000;14:45–54. 78. Kao HY, Downes M, Ordentlich P, Evans RM. Isolation of a novel histone deacetylase reveals that class I and class II deacetylases promote SMRT‐mediated repression. Genes Dev 2000;14:55–66. 79. Farhana L, Dawson MI, Leid M, Wang L, Moore DD, Liu G, et al. Adamantyl‐substituted retinoid‐related molecules bind small heterodimer partner and modulate the Sin3A repressor. Cancer Res 2007;67:318–25. 80. Hu X, Lazar MA. Transcriptional repression by nuclear hormone receptors. Trends Endocrinol Metab 2000;11:6–10. 81. Yoon HG, Chan DW, Reynolds AB, Qin J, Wong J. N‐CoR mediates DNA methylation‐ dependent repression through a methyl CpG binding protein Kaiso. Mol Cell 2003;12:723–34.

CHROMATIN REMODELING AND NUCLEAR RECEPTOR SIGNALING

229

82. Underhill C, Qutob MS, Yee SP, Torchia J. A novel nuclear receptor corepressor complex, N‐CoR, contains components of the mammalian SWI/SNF complex and the corepressor KAP‐1. J Biol Chem 2000;275:40463–70. 83. Martens JA, Winston F. Evidence that Swi/Snf directly represses transcription in S. cerevisiae. Genes Dev 2002;16:2231–6. 84. Alenghat T, Yu J, Lazar MA. The N‐CoR complex enables chromatin remodeler SNF2H to enhance repression by thyroid hormone receptor. EMBO J 2006;25:3966–74. 85. Ewing AK, Attner M, Chakravarti D. Novel regulatory role for human Acf1 in transcriptional repression of vitamin D3 receptor‐regulated genes. Mol Endocrinol 2007;21:1791–806. 86. Jepsen K, Hermanson O, Onami TM, Gleiberman AS, Lunyak V, McEvilly RJ, et al. Combinatorial roles of the nuclear receptor corepressor in transcription and development. Cell 2000;102:753–63. 87. Eickbush TH, Moudrianakis EN. The histone core complex: an octamer assembled by two sets of protein–protein interactions. Biochemistry 1978;17:4955–64. 88. Luger K, Mader AW, Richmond RK, Sargent DF, Richmond TJ. Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 1997;389:251–60. 89. Carruthers LM, Hansen JC. The core histone N termini function independently of linker histones during chromatin condensation. J Biol Chem 2000;275:37285–90. 90. Jiang C, Pugh BF. Nucleosome positioning and gene regulation: advances through genomics. Nat Rev Genet 2009;10:161–72. 91. Khorasanizadeh S. The nucleosome: from genomic organization to genomic regulation. Cell 2004;116:259–72. 92. Dorigo B, Schalch T, Bystricky K, Richmond TJ. Chromatin fiber folding: requirement for the histone H4 N‐terminal tail. J Mol Biol 2003;327:85–96. 93. Zeng L, Zhou MM. Bromodomain: an acetyl‐lysine binding domain. FEBS Lett 2002;513:124–8. 94. Brehm A, Tufteland KR, Aasland R, Becker PB. The many colours of chromodomains. Bioessays 2004;26:133–40. 95. Bannister AJ, Kouzarides T. The CBP co‐activator is a histone acetyltransferase. Nature 1996;384:641–3. 96. Mizzen CA, Yang XJ, Kokubo T, Brownell JE, Bannister AJ, Owen‐Hughes T, et al. The TAF (II)250 subunit of TFIID has histone acetyltransferase activity. Cell 1996;87:1261–70. 97. Ogryzko VV, Schiltz RL, Russanova V, Howard BH, Nakatani Y. The transcriptional coactivators p300 and CBP are histone acetyltransferases. Cell 1996;87:953–9. 98. Yang XJ, Ogryzko VV, Nishikawa J, Howard BH, Nakatani Y. A p300/CBP‐associated factor that competes with the adenoviral oncoprotein E1A. Nature 1996;382:319–24. 99. Pokholok DK, Harbison CT, Levine S, Cole M, Hannett NM, Lee TI, et al. Genome‐wide map of nucleosome acetylation and methylation in yeast. Cell 2005;122:517–27. 100. Roth SY, Denu JM, Allis CD. Histone acetyltransferases. Annu Rev Biochem 2001;70:81–120. 101. Hassig CA, Fleischer TC, Billin AN, Schreiber SL, Ayer DE. Histone deacetylase activity is required for full transcriptional repression by mSin3A. Cell 1997;89:341–7. 102. Yang XJ. Lysine acetylation and the bromodomain: a new partnership for signaling. Bioessays 2004;26:1076–87. 103. Hassan AH, Prochasson P, Neely KE, Galasinski SC, Chandy M, Carrozza MJ, et al. Function and selectivity of bromodomains in anchoring chromatin‐modifying complexes to promoter nucleosomes. Cell 2002;111:369–79. 104. de la Cruz X, Lois S, Sanchez‐Molina S, Martinez‐Balbas MA. Do protein motifs read the histone code? Bioessays 2005;27:164–75. 105. Kutney SN, Hong R, Macfarlan T, Chakravarti D. A signaling role of histone‐binding proteins and INHAT subunits pp32 and Set/TAF‐Ibeta in integrating chromatin hypoacetylation and transcriptional repression. J Biol Chem 2004;279:30850–5.

230

BURANAPRAMEST AND CHAKRAVARTI

106. Martin C, Zhang Y. The diverse functions of histone lysine methylation. Nat Rev Mol Cell Biol 2005;6:838–49. 107. Jenuwein T, Laible G, Dorn R, Reuter G. SET domain proteins modulate chromatin domains in eu‐ and heterochromatin. Cell Mol Life Sci 1998;54:80–93. 108. Rea S, Eisenhaber F, O’Carroll D, Strahl BD, Sun ZW, Schmid M, et al. Regulation of chromatin structure by site‐specific histone H3 methyltransferases. Nature 2000;406:593–9. 109. Aagaard L, Schmid M, Warburton P, Jenuwein T. Mitotic phosphorylation of SUV39H1, a novel component of active centromeres, coincides with transient accumulation at mammalian centromeres. J Cell Sci 2000;113(Pt 5): 817–29. 110. Bannister AJ, Zegerman P, Partridge JF, Miska EA, Thomas JO, Allshire RC, et al. Selective recognition of methylated lysine 9 on histone H3 by the HP1 chromo domain. Nature 2001;410:120–4. 111. Lachner M, O’Carroll D, Rea S, Mechtler K, Jenuwein T. Methylation of histone H3 lysine 9 creates a binding site for HP1 proteins. Nature 2001;410:116–20. 112. Nakayama J, Rice JC, Strahl BD, Allis CD, Grewal SI. Role of histone H3 lysine 9 methylation in epigenetic control of heterochromatin assembly. Science 2001;292:110–3. 113. Cao R, Wang L, Wang H, Xia L, Erdjument‐Bromage H, Tempst P, et al. Role of histone H3 lysine 27 methylation in Polycomb‐group silencing. Science 2002;298:1039–43. 114. Tachibana M, Sugimoto K, Nozaki M, Ueda J, Ohta T, Ohki M, et al. G9a histone methyltransferase plays a dominant role in euchromatic histone H3 lysine 9 methylation and is essential for early embryogenesis. Genes Dev 2002;16:1779–91. 115. Chakravarti D, Ogryzko V, Kao HY, Nash A, Chen H, Nakatani Y, et al. A viral mechanism for inhibition of p300 and PCAF acetyltransferase activity. Cell 1999;96:393–403. 116. Hamamori Y, Sartorelli V, Ogryzko V, Puri PL, Wu HY, Wang JY, et al. Regulation of histone acetyltransferases p300 and PCAF by the bHLH protein twist and adenoviral oncoprotein E1A. Cell 1999;96:405–13. 117. Ait‐Si‐Ali S, Ramirez S, Barre FX, Dkhissi F, Magnaghi‐Jaulin L, Girault JA, et al. Histone acetyltransferase activity of CBP is controlled by cycle‐dependent kinases and oncoprotein E1A. Nature 1998;396:184–6. 118. Perissi V, Dasen JS, Kurokawa R, Wang Z, Korzus E, Rose DW, et al. Factor‐specific modulation of CREB‐binding protein acetyltransferase activity. Proc Natl Acad Sci USA 1999;96:3652–7. 119. Seo SB, McNamara P, Heo S, Turner A, Lane WS, Chakravarti D. Regulation of histone acetylation and transcription by INHAT, a human cellular complex containing the set oncoprotein. Cell 2001;104:119–30. 120. Seo SB, Macfarlan T, McNamara P, Hong R, Mukai Y, Heo S, et al. Regulation of histone acetylation and transcription by nuclear protein pp32, a subunit of the INHAT complex. J Biol Chem 2002;277:14005–10. 121. Loven MA, Muster N, Yates JR, Nardulli AM. A novel estrogen receptor alpha‐associated protein, template‐activating factor Ibeta, inhibits acetylation and transactivation. Mol Endocrinol 2003;17:67–78. 122. Loven MA, Davis RE, Curtis CD, Muster N, Yates JR, Nardulli AM. A novel estrogen receptor alpha‐associated protein alters receptor‐deoxyribonucleic acid interactions and represses receptor‐mediated transcription. Mol Endocrinol 2004;18:2649–59. 123. Chakravarti D, Hong R. SET‐ting the stage for life and death. Cell 2003;112:589–91. 124. Schultz‐Norton JR, Ziegler YS, Likhite VS, Yates JR, Nardulli AM. Isolation of novel coregulatory protein networks associated with DNA‐bound estrogen receptor alpha. BMC Mol Biol 2008;9:97. 125. Schneider R, Bannister AJ, Weise C, Kouzarides T. Direct binding of INHAT to H3 tails disrupted by modifications. J Biol Chem 2004;279:23859–62.

CHROMATIN REMODELING AND NUCLEAR RECEPTOR SIGNALING

231

126. Ichijo T, Chrousos GP, Kino T. Activated glucocorticoid receptor interacts with the INHAT component Set/TAF‐Ibeta and releases it from a glucocorticoid‐responsive gene promoter, relieving repression: implications for the pathogenesis of glucocorticoid resistance in acute undifferentiated leukemia with Set‐Can translocation. Mol Cell Endocrinol 2008;283:19–31. 127. Adachi Y, Pavlakis GN, Copeland TD. Identification and characterization of SET, a nuclear phosphoprotein encoded by the translocation break point in acute undifferentiated leukemia. J Biol Chem 1994;269:2258–62. 128. Kular RK, Cvetanovic M, Siferd S, Kini AR, Opal P. Neuronal differentiation is regulated by leucine‐rich acidic nuclear protein (LANP), a member of the inhibitor of histone acetyltransferase complex. J Biol Chem 2009;284:7783–92. 129. Fan Z, Zhang H, Zhang Q. Tumor suppressor pp32 represses cell growth through inhibition of transcription by blocking acetylation and phosphorylation of histone H3 and initiating its proapoptotic activity. Cell Death Differ 2006;13:1485–94. 130. Adegbola O, Pasternack GR. A pp32‐retinoblastoma protein complex modulates androgen receptor‐mediated transcription and associates with components of the splicing machinery. Biochem Biophys Res Commun 2005;334:702–8. 131. Gamble MJ, Erdjument‐Bromage H, Tempst P, Freedman LP, Fisher RP. The histone chaperone TAF‐I/SET/INHAT is required for transcription in vitro of chromatin templates. Mol Cell Biol 2005;25:797–807. 132. Nagata K, Kawase H, Handa H, Yano K, Yamasaki M, Ishimi Y, et al. Replication factor encoded by a putative oncogene, set, associated with myeloid leukemogenesis. Proc Natl Acad Sci USA 1995;92:4279–83. 133. Muto S, Senda M, Akai Y, Sato L, Suzuki T, Nagai R, et al. Relationship between the structure of SET/TAF‐Ibeta/INHAT and its histone chaperone activity. Proc Natl Acad Sci USA 2007;104:4285–90. 134. Fan HY, He X, Kingston RE, Narlikar GJ. Distinct strategies to make nucleosomal DNA accessible. Mol Cell 2003;11:1311–22. 135. Jiang X, Kim HE, Shu H, Zhao Y, Zhang H, Kofron J, et al. Distinctive roles of PHAP proteins and prothymosin‐alpha in a death regulatory pathway. Science 2003;299:223–6. 136. Fan Z, Beresford PJ, Oh DY, Zhang D, Lieberman J. Tumor suppressor NM23‐H1 is a granzyme A‐activated DNase during CTL‐mediated apoptosis, and the nucleosome assembly protein SET is its inhibitor. Cell 2003;112:659–72. 137. Dirksen EH, Cloos J, Braakhuis BJ, Brakenhoff RH, Heck AJ, Slijper M. Human lymphoblastoid proteome analysis reveals a role for the inhibitor of acetyltransferases complex in DNA double‐strand break response. Cancer Res 2006;66:1473–80. 138. Hong W, Kim AY, Ky S, Rakowski C, Seo SB, Chakravarti D, et al. Inhibition of CBP‐mediated protein acetylation by the Ets family oncoprotein PU.1. Mol Cell Biol 2002;22:3729–43. 139. Ko SI, Lee IS, Kim JY, Kim SM, Kim DW, Lee KS, et al. Regulation of histone acetyltransferase activity of p300 and PCAF by proto‐oncogene protein DEK. FEBS Lett 2006;580:3217–22. 140. Jin C, Li H, Murata T, Sun K, Horikoshi M, Chiu R, et al. JDP2, a repressor of AP‐1, recruits a histone deacetylase 3 complex to inhibit the retinoic acid‐induced differentiation of F9 cells. Mol Cell Biol 2002;22:4815–26. 141. Wardell SE, Boonyaratanakornkit V, Adelman JS, Aronheim A, Edwards DP. Jun dimerization protein 2 functions as a progesterone receptor N‐terminal domain coactivator. Mol Cell Biol 2002;22:5451–66. 142. Nakade K, Pan J, Yoshiki A, Ugai H, Kimura M, Liu B, et al. JDP2 suppresses adipocyte differentiation by regulating histone acetylation. Cell Death Differ 2007;14:1398–405. 143. Hublitz P, Kunowska N, Mayer UP, Muller JM, Heyne K, Yin N, et al. NIR is a novel INHAT repressor that modulates the transcriptional activity of p53. Genes Dev 2005;19:2912–24.

232

BURANAPRAMEST AND CHAKRAVARTI

144. Elgin SC. DNAase I‐hypersensitive sites of chromatin. Cell 1981;27:413–5. 145. Gross DS, Garrard WT. Nuclease hypersensitive sites in chromatin. Annu Rev Biochem 1988;57:159–97. 146. McGhee JD, Wood WI, Dolan M, Engel JD, Felsenfeld G. A 200 base pair region at the 50 end of the chicken adult beta‐globin gene is accessible to nuclease digestion. Cell 1981;27:45–55. 147. Yuan GC, Liu YJ, Dion MF, Slack MD, Wu LF, Altschuler SJ, et al. Genome‐scale identification of nucleosome positions in S. cerevisiae. Science 2005;309:626–30. 148. Bernstein BE, Liu CL, Humphrey EL, Perlstein EO, Schreiber SL. Global nucleosome occupancy in yeast. Genome Biol 2004;5:R62. 149. Lee CK, Shibata Y, Rao B, Strahl BD, Lieb JD. Evidence for nucleosome depletion at active regulatory regions genome‐wide. Nat Genet 2004;36:900–5. 150. Chen X, Wang J, Woltring D, Gerondakis S, Shannon MF. Histone dynamics on the interleukin‐2 gene in response to T‐cell activation. Mol Cell Biol 2005;25:3209–19. 151. Vicent GP, Nacht AS, Smith CL, Peterson CL, Dimitrov S, Beato M. DNA instructed displacement of histones H2A and H2B at an inducible promoter. Mol Cell 2004;16:439–52. 152. Brown DD. The role of stable complexes that repress and activate eucaryotic genes. Cell 1984;37:359–65. 153. Segal E, Fondufe‐Mittendorf Y, Chen L, Thastrom A, Field Y, Moore IK, et al. A genomic code for nucleosome positioning. Nature 2006;442:772–8. 154. Boeger H, Griesenbeck J, Strattan JS, Kornberg RD. Nucleosomes unfold completely at a transcriptionally active promoter. Mol Cell 2003;11:1587–98. 155. Reinke H, Horz W. Histones are first hyperacetylated and then lose contact with the activated PHO5 promoter. Mol Cell 2003;11:1599–607. 156. Chandy M, Gutierrez JL, Prochasson P, Workman JL. SWI/SNF displaces SAGA‐acetylated nucleosomes. Eukaryot Cell 2006;5:1738–47. 157. Zhang Y, Shin H, Song JS, Lei Y, Liu XS. Identifying positioned nucleosomes with epigenetic marks in human from ChIP‐Seq. BMC Genomics 2008;9:537. 158. Schones DE, Cui K, Cuddapah S, Roh TY, Barski A, Wang Z, et al. Dynamic regulation of nucleosome positioning in the human genome. Cell 2008;132:887–98. 159. Schones DE, Zhao K. Genome‐wide approaches to studying chromatin modifications. Nat Rev Genet 2008;9:179–91. 160. Dressel U, Thormeyer D, Altincicek B, Paululat A, Eggert M, Schneider S, et al. Alien, a highly conserved protein with characteristics of a corepressor for members of the nuclear hormone receptor superfamily. Mol Cell Biol 1999;19:3383–94. 161. Eckey M, Hong W, Papaioannou M, Baniahmad A. The nucleosome assembly activity of NAP1 is enhanced by Alien. Mol Cell Biol 2007;27:3557–68. 162. Kingston RE, Narlikar GJ. ATP‐dependent remodeling and acetylation as regulators of chromatin fluidity. Genes Dev 1999;13:2339–52. 163. Saha A, Wittmeyer J, Cairns BR. Chromatin remodelling: the industrial revolution of DNA around histones. Nat Rev Mol Cell Biol 2006;7:437–47. 164. Eberharter A, Becker PB. ATP‐dependent nucleosome remodelling: factors and functions. J Cell Sci 2004;117:3707–11. 165. Becker PB, Horz W. ATP‐dependent nucleosome remodeling. Annu Rev Biochem 2002;71:247–73. 166. Peterson CL, Logie C. Recruitment of chromatin remodeling machines. J Cell Biochem 2000;78:179–85. 167. Neigeborn L, Carlson M. Genes affecting the regulation of SUC2 gene expression by glucose repression in Saccharomyces cerevisiae. Genetics 1984;108:845–58.

CHROMATIN REMODELING AND NUCLEAR RECEPTOR SIGNALING

233

168. Stern M, Jensen R, Herskowitz I. Five SWI genes are required for expression of the HO gene in yeast. J Mol Biol 1984;178:853–68. 169. Peterson CL, Herskowitz I. Characterization of the yeast SWI1, SWI2, and SWI3 genes, which encode a global activator of transcription. Cell 1992;68:573–83. 170. Peterson CL, Tamkun JW. The SWI‐SNF complex: a chromatin remodeling machine? Trends Biochem Sci 1995;20:143–6. 171. Martens JA, Winston F. Recent advances in understanding chromatin remodeling by Swi/Snf complexes. Curr Opin Genet Dev 2003;13:136–42. 172. Reyes JC, Muchardt C, Yaniv M. Components of the human SWI/SNF complex are enriched in active chromatin and are associated with the nuclear matrix. J Cell Biol 1997;137:263–74. 173. Aoyagi S, Archer TK. Dynamics of coactivator recruitment and chromatin modifications during nuclear receptor mediated transcription. Mol Cell Endocrinol 2008;280:1–5. 174. Hebbar PB, Archer TK. Chromatin remodeling by nuclear receptors. Chromosoma 2003;111:495–504. 175. Trotter KW, Archer TK. Nuclear receptors and chromatin remodeling machinery. Mol Cell Endocrinol 2007;265–266:162–7. 176. Muchardt C, Yaniv M. A human homologue of Saccharomyces cerevisiae SNF2/SWI2 and Drosophila brm genes potentiates transcriptional activation by the glucocorticoid receptor. EMBO J 1993;12:4279–90. 177. Acevedo ML, Kraus WL. Transcriptional activation by nuclear receptors. Essays Biochem 2004;40:73–88. 178. Archer TK, Lefebvre P, Wolford RG, Hager GL. Transcription factor loading on the MMTV promoter: a bimodal mechanism for promoter activation. Science 1992;255:1573–6. 179. Beato M, Eisfeld K. Transcription factor access to chromatin. Nucleic Acids Res 1997;25:3559–63. 180. Ichinose H, Garnier JM, Chambon P, Losson R. Ligand‐dependent interaction between the estrogen receptor and the human homologues of SWI2/SNF2. Gene 1997;188:95–100. 181. Trotter KW, Archer TK. Reconstitution of glucocorticoid receptor‐dependent transcription in vivo. Mol Cell Biol 2004;24:3347–58. 182. Belandia B, Parker MG. Nuclear receptors: a rendezvous for chromatin remodeling factors. Cell 2003;114:277–80. 183. Chiba H, Muramatsu M, Nomoto A, Kato H. Two human homologues of Saccharomyces cerevisiae SWI2/SNF2 and Drosophila brahma are transcriptional coactivators cooperating with the estrogen receptor and the retinoic acid receptor. Nucleic Acids Res 1994;22:1815–20. 184. Kitagawa H, Fujiki R, Yoshimura K, Mezaki Y, Uematsu Y, Matsui D, et al. The chromatin‐ remodeling complex WINAC targets a nuclear receptor to promoters and is impaired in Williams syndrome. Cell 2003;113:905–17. 185. Fujiki R, Kim MS, Sasaki Y, Yoshimura K, Kitagawa H, Kato S. Ligand‐induced transrepression by VDR through association of WSTF with acetylated histones. EMBO J 2005;24:3881–94. 186. Ito T, Bulger M, Pazin MJ, Kobayashi R, Kadonaga JT. ACF, an ISWI‐containing and ATP‐ utilizing chromatin assembly and remodeling factor. Cell 1997;90:145–55. 187. Varga‐Weisz PD, Wilm M, Bonte E, Dumas K, Mann M, Becker PB. Chromatin‐remodelling factor CHRAC contains the ATPases ISWI and topoisomerase II. Nature 1997;388:598–602. 188. Bochar DA, Savard J, Wang W, Lafleur DW, Moore P, Cote J, et al. A family of chromatin remodeling factors related to Williams syndrome transcription factor. Proc Natl Acad Sci USA 2000;97:1038–43. 189. Fyodorov DV, Blower MD, Karpen GH, Kadonaga JT. Acf1 confers unique activities to ACF/ CHRAC and promotes the formation rather than disruption of chromatin in vivo. Genes Dev 2004;18:170–83.

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190. Goldmark JP, Fazzio TG, Estep PW, Church GM, Tsukiyama T. The Isw2 chromatin remodeling complex represses early meiotic genes upon recruitment by Ume6p. Cell 2000;103:423–33. 191. Fazzio TG, Tsukiyama T. Chromatin remodeling in vivo: evidence for a nucleosome sliding mechanism. Mol Cell 2003;12:1333–40. 192. Moreau JL, Lee M, Mahachi N, Vary J, Mellor J, Tsukiyama T, et al. Regulated displacement of TBP from the PHO8 promoter in vivo requires Cbf1 and the Isw1 chromatin remodeling complex. Mol Cell 2003;11:1609–20. 193. Whitehouse I, Tsukiyama T. Antagonistic forces that position nucleosomes in vivo. Nat Struct Mol Biol 2006;13:633–40. 194. Stopka T, Skoultchi AI. The ISWI ATPase Snf2h is required for early mouse development. Proc Natl Acad Sci USA 2003;100:14097–102. 195. Liu YI, Chang MV, Li HE, Barolo S, Chang JL, Blauwkamp TA, et al. The chromatin remodelers ISWI and ACF1 directly repress Wingless transcriptional targets. Dev Biol 2008;323:41–52. 196. Di Croce L, Koop R, Venditti P, Westphal HM, Nightingale KP, Corona DF, et al. Two‐step synergism between the progesterone receptor and the DNA‐binding domain of nuclear factor 1 on MMTV minichromosomes. Mol Cell 1999;4:45–54. 197. Badenhorst P, Xiao H, Cherbas L, Kwon SY, Voas M, Rebay I, et al. The Drosophila nucleosome remodeling factor NURF is required for Ecdysteroid signaling and metamorphosis. Genes Dev 2005;19:2540–5. 198. Eberharter A, Ferrari S, Langst G, Straub T, Imhof A, Varga‐Weisz P, et al. Acf1, the largest subunit of CHRAC, regulates ISWI‐induced nucleosome remodelling. EMBO J 2001;20:3781–8. 199. Eberharter A, Vetter I, Ferreira R, Becker PB. ACF1 improves the effectiveness of nucleosome mobilization by ISWI through PHD‐histone contacts. EMBO J 2004;23:4029–39. 200. Collins N, Poot RA, Kukimoto I, Garcia‐Jimenez C, Dellaire G, Varga‐Weisz PD. An ACF1‐ ISWI chromatin‐remodeling complex is required for DNA replication through heterochromatin. Nat Genet 2002;32:627–32. 201. Zhou Y, Grummt I. The PHD finger/bromodomain of NoRC interacts with acetylated histone H4K16 and is sufficient for rDNA silencing. Curr Biol 2005;15:1434–8. 202. Zhou Y, Santoro R, Grummt I. The chromatin remodeling complex NoRC targets HDAC1 to the ribosomal gene promoter and represses RNA polymerase I transcription. EMBO J 2002;21:4632–40. 203. Wang X, Arai S, Song X, Reichart D, Du K, Pascual G, Tempst P, Rosenfeld MG, Glass CK, Kurokawa R. Induced ncRNAs allosterically modify RNA-binding proteins in cis to inhibit transcription. Nature 2008;454:126–30.

Nuclear Receptor Repression: Regulatory Mechanisms and Physiological Implications M. David Stewart* and Jiemin Wong{ *Department of Genetics, University of Texas M.D. Anderson Cancer Center, Houston, Texas 77030, USA {

Institute of Biomedical Sciences and School of Life Sciences, East China Normal University, Shanghai 200241, China

I. Introduction ................................................................................. II. Corepressors ................................................................................ A. Ligand‐Independent Corepressors ................................................ B. Ligand‐Dependent Corepressors .................................................. III. Types of NR Repression.................................................................. A. Repression by Unliganded Receptors............................................. B. Repression by Antagonist‐Bound Steroid Receptors .......................... C. Repression by Agonist‐Bound Receptors ........................................ IV. Molecular Mechanisms of Transcriptional Repression ............................ A. Histone Deacetylation................................................................ B. Histone Methylation .................................................................. C. Chromatin Assembly/Remodeling ................................................. D. DNA Methylation ..................................................................... V. Physiological Functions of NR‐Mediated Repression.............................. A. Physiological Function of Repression by Unliganded TR .................... B. TR‐Mediated Repression in the Regulation of Amphibian Metamorphosis ................................................... C. Repression by PPARg ................................................................ D. Repression Mediated by NCoR and SMRT ..................................... E. Repression Mediated by RIP140................................................... VI. Concluding Remarks ...................................................................... References...................................................................................

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The ability to associate with corepressors and to inhibit transcription is an intrinsic property of most members of the nuclear receptor (NR) superfamily. NRs induce transcriptional repression by recruiting multiprotein corepressor complexes. Nuclear receptor corepressor (NCoR) and silencing mediator of retinoic and thyroid receptors (SMRT) are the most well characterized corepressor complexes and mediate repression for virtually all NRs. In turn, corepressor complexes repress transcription because they either contain or associate with chromatin modifying enzymes. These include histone deacetylases, histone Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87007-5

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H3K4 demethylases, histone H3K9 or H3K27 methyltransferases, and ATP‐dependent chromatin remodeling factors. Two types of NR‐interacting corepressors exist. Ligand‐independent corepressors, like NCoR/SMRT, bind to unliganded or antagonist‐bound NRs, whereas ligand‐dependent corepressors (LCoRs) associate with NRs in the presence of agonist. Therefore, LCoRs may serve to attenuate NR‐mediated transcriptional activation. Somewhat unexpectedly, classical coactivators may also function as ‘‘corepressors’’ to mediate repression by agonist‐bound NRs. In this chapter, we will discuss the various modes and mechanisms of repression by NRs as well as discuss the known physiological functions of NR‐mediated repression.

I. Introduction Nuclear receptors (NRs) are ligand‐dependent transcription factors that play pivotal roles in a variety of key metabolic and developmental processes. Their ligands are lipophilic molecules such as steroid hormones, thyroid hormone, retinoic acid, vitamin D, fatty acids, fatty acid derivatives, and xenobiotics. These molecules can diffuse through the plasma membrane and interact with NRs located in either the nuclear or cytosolic compartments. NRs are composed of five functionally distinct domains referred to as A/B, C, D, E, and F. The A/B N‐terminal domain varies greatly between NRs and contains the activation function 1 (AF1). AF1 mediates ligand‐independent transactivation for some NRs. The C region contains a sequence‐specific DNA‐binding domain (DBD) consisting of two zinc fingers. The D domain is a variable hinge region separating the DBD from the ligand‐binding domain (LBD) which comprises region E. The D domain usually contains nuclear import/export sequences. Region E also contains the ligand‐dependent AF2. Finally, the sequence and function of the C‐terminal F domain varies greatly among NRs. NRs are well known for their hormone‐dependent transcriptional activation function. However, NRs also have capacity to repress transcription both in the absence or presence of their ligands. Studies over last decade or two have begun to reveal the molecular mechanisms and physiological relevance of the NR repression function. Important for this discussion of repression by NRs, a subset of NRs including thyroid hormone receptors (TR) and retinoic acid receptors (RAR) are constitutively nuclear and bind to their target genes in the absence of their cognate hormones. These receptors exhibit a potent transcriptional repression function in the absence of hormones. The repression activity could be mapped to their LBD and is transferable.1 It is this property that led to the identification of various proteins termed ‘‘corepressors’’ that preferentially interact with and participate in repression by unliganded receptors.

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The most notable corepressors for NRs are nuclear receptor corepressor (NCoR) and the highly similar silencing mediator of retinoic and thyroid receptors (SMRT). Subsequent molecular and structural analyses have revealed that NCoR and SMRT interact with the NR LBD through one or more corepressor nuclear receptor (CoRNR) boxes.2–4 The CoRNR box consists of the sequence (L/I)XX(I/V)I or LXXX(I/L)XXX(I/L) where X is any amino acid. This interaction is regulated via ligand‐induced conformational change of the LBD. In the absence of ligand, the LBD conformation is such that it prefers interaction with the corepressor proteins containing CoRNR boxes. Upon interaction with agonist, the LBD alters its conformation such that it prefers association with coactivator proteins containing the sequence LXXLL termed the NR box.5 In addition to the corepressors that associate with and mediate repression by unliganded NRs, there is a class of corepressors that contain one or more NR boxes and therefore specifically associate with agonist‐bound receptors. A well studied example is RIP140, which was first identified as an agonist‐ dependent estrogen receptor alpha (ERa) AF2 domain‐interacting protein and thought to be a coactivator.6 These ‘‘ligand‐dependent’’ corepressors may serve to attenuate NR‐mediated transcriptional activity. Given that transcription is a complex multistep process involving a large number of general transcriptional factors and cofactors, in principle a corepressor protein may repress transcription by interfering with any of the steps. This idea is consistent with the large number of corepressors identified so far. Noticeably, corepressors often contain or associate with one or more histone modifying activities that actively inhibit transcription though covalent modifications to the core histones. These modifications include deacetylation, demethylation of H3K4, and methylation of H3K9 or H3K27. Repressive histone modifications can attenuate transcriptional activity either by creating a local repressive heterochromatin environment or by preventing the binding of coactivators or basal machinery. In addition to repression by unliganded NRs, it has become clear that agonist‐bound NRs can also repress transcription of target genes. This phenomenon is in general termed transrepression. The best example of this is the ability of the glucocorticoid receptor (GR) to inhibit expression of proinflammatory genes via direct interaction with NFkB.7,8 We will focus this chapter on the molecular mechanisms of corepressor function. Since recent interesting new work using mouse genetics has shed light on the role of NR‐corepressor interactions in the regulation of gene expression by NRs, we will also discuss the physiological functions of NR repression as well as their disease associations.

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II. Corepressors Transcriptional repression by NRs is mediated by a class of coregulatory proteins termed corepressors. Thus, we will first discuss corepressors in detail and then describe how they are utilized in the various modes of NR‐mediated repression. Simply put, corepressors are molecules that have the ability to inhibit transcription and also interact with NRs. They usually exist in large multiprotein complexes that couple multiple enzymatic, NR binding, chromatin binding, and regulatory activities into a single functional unit. The best studied NR‐corepressor complexes are NCoR and SMRT. Other corepressors exist and may interact with NRs in a cell‐specific or gene‐specific manner. These include Sin3A, Alien, and SUN‐CoR. Additionally, there exists an expanding class of agonist‐bound corepressors, including Hairless, LCoR, MTA1, PRAME, REA, and RIP140.

A. Ligand‐Independent Corepressors Ligand‐independent corepressors are those that bind NRs in the absence of agonist. They contain one or more CoRNR boxes, which interact with the AF2 of the LBD. Upon binding of agonist, ligand‐independent corepressors dissociate from the NR allowing association of NR box‐containing coregulators, typically coactivators. This is the ‘‘canonical’’ molecular mechanism of gene regulation by type II NRs, which bind DNA constitutively, repressing in the absence and activating transcription in the presence of ligand. NCoR and SMRT are the most well characterized ligand‐independent corepressors. They are ubiquitously expressed proteins that mediate repression by numerous NRs as well as other transcription factors. Their molecular mechanisms of action have been extensively studied for the NRs for which they are named: TR and RAR, but their mechanism of action is applicable to all NRs with which they interact. The core NCoR/SMRT protein complex consists of NCoR/SMRT, Transducin b‐like 1/related 1 (TBL1/TBLR1), Histone deacetylase 3 (HDAC3), and G‐protein pathway suppressor 2 (GPS2).9,10 NCoR and SMRT essentially serve as platforms for complex assembly. They bind NRs and associate with each of the other complex subunits. Interaction with NRs is mediated by three CoRNR box‐containing receptor interaction domains (RIDs) located in their C‐termini. HDAC3, as the name implies, is a histone deacetylase and the only deacetylase identified in the core NCoR/SMRT complex. It has activity toward acetylated lysines on all core histones.11 HDAC3 requires interaction with NCoR/SMRT for potent enzymatic activity; thus, the region of NCoR/SMRT that binds and

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activates HDAC3 is referred to as the deacetylase activating domain (DAD).12 TBL1 and TBLR1 are highly related proteins and members of the WD40 superfamily. They directly interact with the N‐terminal repression domain of NCoR/SMRT (RD1) and bind deacetylated histone H4.13 In this manner, they may function to stabilize the corepressor complex on chromatin and thus facilitate repression. Interestingly, TBL1/TBLR1 has also been reported to mediate degradation of NCoR/SMRT by the proteasome during hormone‐ dependent transcriptional activation.14 Little is known of the function of GPS2 other than its ability to inhibit the Jun N‐terminal kinase (JNK) pathway. Direct association with NCoR/SMRT is required for this activity.10 It may function in NR‐mediated transrepression, which will be discussed later in this chapter. While purification of NCoR/SMRT core complex revealed the absence of Sin3A corepressor complex, both NcoR and SMRT have been reported to interact with Sin3A or a component of the Sin3A corepressor complex.15–17 The mammalian SIN3 complex consists of SIN3A/B, HDAC1/2, RbAP46/48, SAP18/30, and SDS3.18–22 The SIN3 complex may be recruited to target genes either by direct interaction with a transcription factor (i.e., Mad1) or indirectly through interaction with NCoR or SMRT.23 NCoR associates with SIN3A/B complexes via the adapter protein SAP30. SMRT only associates with SIN3A complexes and does so through direct interaction with SIN3A.17 Thus, SIN3 corepressor complexes may also contribute to NR‐mediated repression through interaction with NCoR/SMRT. SIN3 complexes are very similar in composition to that of NCoR/SMRT. They both have large proteins that form the molecular scaffold (SIN3A/B, NCoR/SMRT). They both have WD40 superfamily members that stabilize the complexes on chromatin via interaction with core histones (RbAP46/48, TBL1/TBLR1). And they both repress transcription by histone deacetylase subunits (HDAC1/2, HDAC3). Although not components of the core complex, SIN3 also associates with the histone H3K9 methyltransferase ESET/SETDB1 and the ATP‐dependent chromatin remodeling complex SWI/SNF.24,25 Therefore, the ability of NCoR/SMRT to associate with SIN3 complexes may draw additional chromatin modifying enzymes to participate in NR‐mediated transcriptional repression. The corepressor Alien was originally identified as a corepressor for TR, but has since been found to associate with DAX1, COUP‐TFI, COUP‐TFII, AR, and VDR.26–29 Alien is a highly conserved and widely expressed protein. A gene knockout study indicates that it is required for mouse early embryonic development.30 It contains autonomous transcriptional repression activity and represses transcription through both HDAC‐dependent and ‐independent pathways. The HDAC‐dependent pathway involves recruitment of the HDAC1/2‐containing SIN3A complex.31 The HDAC‐independent pathway consists of repression via creation of a compact nucleosome environment and is mediated though recruitment of the nucleosome assembly factor NAP1.32

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In addition to aforementioned histone deacetylase‐dependent mechanism, recent studies indicate that histone methylation and demethylation also contribute to repression by NRs. SUV39H1, a histone H3K9 methyltransferase, has been reported to interact with TR.33 In vitro, this interaction is insensitive to hormone. However, addition of T3 led to dissociation of SUV39H1 from TR in vivo. SUV39H1 enhances TR repression in a H3K9 methyltransferase activity‐dependent manner. Another H3K9 methyltransferase G9a was shown to interact with and participate in repression of CYP7A1 gene by the small heterodimer partner (SHP).34 LSD1 is the first identified histone lysine demethylase that specifically demethylates mono‐ and dimethyl‐H3K4.35 Interestingly, while LSD1 was shown to function as a coactivator for AR and ER, it seems to serve as a corepressor for orphan receptor TLX (also called NR2E1).36,37 LSD1 interacts directly with TLX and potentiates TLX repression through its H3K4 demethylase activity.

B. Ligand‐Dependent Corepressors Unlike ligand‐independent corepressors, ligand‐dependent corepressors (LCoRs) interact with the LBD of agonist‐bound NRs and may contain one or more NR boxes typically found in coactivators. Therefore, this class of corepressors may function to attenuate the activity of agonist‐bound NRs. The list of LCoRs includes hairless (Hr), LCoR, receptor‐interacting protein 140 (RIP140), metastasis associated factor 1 (MTA1), preferentially expressed antigen in melanoma (PRAME), and repressor of estrogen activity (REA). We will briefly mention the best studied of these factors as they have been extensively reviewed elsewhere. RIP140 was the first LCoR identified. It was first identified as an agonist‐ dependent ERa AF2 domain‐interacting protein and thought to be a coactivator.6 It was subsequently found to associate with a number of other NRs.38 Later it was found to have transrepression activity and could negatively regulate agonist‐bound NRs by competing with coactivators for NR binding.39 In addition, RIP140 has been shown to inhibit transcription by means of distinct repression domains that function by both HDAC‐dependent and ‐independent mechanisms.40,41 RIP140 is widely expressed and has critical roles in adipose biology and female reproductive physiology, specifically oocyte release during ovulation.42,43 Similar to RIP140, LCoR was first identified for its ligand‐dependent interaction with ERa, but subsequently was found to associate with many NRs.44 The LCoR gene is broadly expressed in embryonic and adult tissues. LCoR also exhibits both HDAC‐dependent and ‐independent repression. Like most corepressors, LCoR associates with HDAC activity. Specifically, it directly interacts with HDAC3, 6, and 10. Interestingly, its transrepression activity for

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ERa and GR is sensitive to deacetylase inhibitors; whereas, its transrepression activity for progesterone receptor (PR) and vitamin D receptor (VDR) is not. Thus, LCoR exhibits an example of HDAC‐independent repression. The HDAC‐independent repression of LCoR is mediated through its association with CtBP1/2.44 CtBP associates with many transcription factors and is a component of large multiprotein complexes containing histone modifying enzymes. Specifically, CtBP directly interacts with polycomb group (PcG) proteins containing methyltransferase activity for histone H3K27.45 Methylation of H3K27 is well known to be a repressive histone modification. Hr is unique in that it contains both NR and CoRNR boxes; therefore it functions as both a ligand‐independent and LCoR. TR was the first NR found to interact with Hr and for TR, Hr functions as a typical ligand‐independent corepressor.46 Hr associates with the TR LBD in the absence of ligand via its CoRNR box and represses transcription. It also brings HDAC activity via direct interaction with HDAC1, 3, and 5. In the presence of thyroid hormone, Hr dissociates from TR. Unlike NCoR and SMRT, which are ubiquitously expressed, Hr is selectively expressed mainly in the skin and brain.47 Therefore, it may function as a tissue‐specific corepressor for TR. In contrast, Hr is a LCoR for RAR‐related orphan receptor (ROR) and VDR.48,49 It associates with the AF2 domain of these receptors via its two NR boxes. Hr is unable to bind these receptors in the absence of agonist, because its CoRNR motifs are selective for TR. Thus, in tissues where it is expressed, like the brain and skin, Hr attenuates the transcriptional activity of ROR and VDR in response to ligand.

III. Types of NR Repression A. Repression by Unliganded Receptors Type II NRs are those that constitutively reside in the nucleus and bind to their DNA recognition sequences. These NRs include all the isoforms of TR, RAR, VDR, and PPAR. In the absence of ligand, these receptors interact with corepressor complexes and actively repress expression of their target genes. In the presence of ligand, the corepressors are displaced and coactivators are recruited. Repression by type II NRs is generally mediated by ligand‐ independent corepressors, like NCoR and SMRT, which interact with the unliganded NR LBD via CoRNR boxes (Fig. 1A). Certain orphan NRs are constitutive repressors. Therefore, in any tissue in which they are expressed, they bind to response elements and suppress expression of their target genes. Examples include COUP‐TFI, COUP‐TFII, DAX1, EAR2, germ cell nuclear factor (GCNF), SHP, TLX, TR2, and TR4. Like type II NRs, the repression activity of these orphan receptors is mediated by

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A HDAC3

CoRNR box-containing corepressors recruited by unliganded NR

NCoR/SMRT

TR RXR

B HDAC3

CoRNR box-containing corepressors recruited by antagonist-bound NR

NCoR/SMRT 4-HT ER

4-HT ER

C HDAC 1,3,5

NR box-containing corepressors recruited by agonist-bound NR

Hr

GRIP1

Agonist-bound NR represses trancription via classical coactivators

E2 ERa

Vit D VDR RXR

cJun NFκB

FIG. 1. Types of nuclear receptor repression. (A) Repression by unliganded NRs (typical of type II NRs). (B) Repression by antagonist‐bound NRs. The antagonist‐bound LBD favors interaction with corepressors. (C) Repression by agonist‐bound NRs. Two examples are given. Ligand‐dependent corepressors may be recruited by agonist‐bound NRs. Alternatively, classical coactivators may repress transcription by agonist‐bound NRs.

corepressor complexes like NCoR and SMRT. However, since these receptors have no known ligand, they are always maintained in the repressive conformation leading to constitutive repression.

B. Repression by Antagonist‐Bound Steroid Receptors Steroid hormone receptors in general reside in cytoplasm and associate with chaperones in the absence of hormones. Binding of hormones triggers conformational changes that lead to dissociation of cytoplasmic chaperones, nuclear translocation, DNA binding, and ultimately activation of target genes. However, it is well established that antagonist‐bound steroid receptors can lead

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to repression of gene expression (Fig. 1B). Indeed, various antagonists for ER, GR, AR, and PR have been explored for various therapeutic purposes. It is generally believed that different antagonists or partial antagonists can induce the LBD of a NR to adopt different conformations. This different conformation in turn dictates the association of a spectrum of coactivators and corepressors and subsequent transcriptional activation or repression. Since this type of repression has been extensively reviewed, we will not discuss this in detail here.

C. Repression by Agonist‐Bound Receptors While most studies on agonists focus on transcriptional activation, agonist‐ bound receptors can also induce transcriptional repression. One of the best known cases is the negative feedback repression of thyrotropin (TSH) in the pituitary as part of the endocrine hypothalamus–pituitary–thyroid feedback loop.50 The repression function of agonist‐bound receptors are well supported by recent numerous microarray studies. For example, while 17b‐estradiol treatment of MCF7 cells induces activation of a large number of genes, a similar number of genes are repressed.51 Similar results have been reported for TR, VDR, and GR.52–57 Whereas the mechanism whereby agonist‐bound receptors activate gene expression has been extensively studied, little is known about how agonist‐ bound receptors induce transcriptional repression. In the case of feedback repression of TSH by liganded TRb, several working models have been proposed, including (1) direct binding of TR to a negative TRE that leads to the repressive effect of liganded TR, (2) liganded TR associates with the target gene through protein–protein interactions with other transcription factors or cofactors and interferes with transcriptional activation, or (3) squelching effects by sequestering coactivators essential for transcriptional activation.58–62 Each of these mechanisms is supported by some experimental evidence, yet none of these have fully explained the repressive feedback of TSH gene expression by TR. Similarly, recent studies on vitamin D‐induced transcriptional repression of several negative VDR target genes have begun to reveal diverse modes for repression.63,64 Nevertheless, histone deacetylation induced by histone deacetylase corepressors appears to facilitate vitamin D‐induced transcriptional repression via the VDR. ERa, classically a ligand‐induced transcriptional activator, has been reported to function in an opposite manner to regulate expression of the TNFa gene.65 For TNFa, unliganded ERa acted as a coactivator for c‐Jun and NFkB. In the presence of estradiol, ERa repressed transcription of TNFa in a manner dependent on recruitment of the ‘‘coactivator’’ GRIP1. Thus, the authors conclude that GRIP1 can function as either a coactivator or corepressor depending on the context of the gene promoter. Figure 1C illustrates some

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of the modes of repression by agonist‐bound NRs. Taken together, the mechanisms behind agonist‐induced transcriptional repression are probably more complex than those of agonist‐induced transactivation and will be an important area for future study.

IV. Molecular Mechanisms of Transcriptional Repression In principle, NRs and corepressors that they interact with may repress transcription by interfering with any step(s) along the multistep process of transcription. Indeed, NRs have been reported to inhibit transcription by various mechanisms including interfering with the binding of transcription factors to DNA, competition for heterodimer partner RXR, interaction with general transcriptional factors such as TATA‐binding protein (TBP). However, accumulative evidence points to key roles of histone modifications and chromatin remodeling in mediating repression by NRs. In fact, many corepressor proteins are either histone modifying enzymes or function to recruit histone modifying enzymes that deposit repressive marks on chromatin (Fig. 2). Here, we will elaborate on these chromatin related repression mechanisms.

A. Histone Deacetylation By far the most heavily studied molecular mechanism of gene repression is histone deacetylation. It is well known that hypoacetylated histones are associated with gene silencing and heterochromatin. Furthermore, every NR

Histone H3K9 methyltransferases

Histone deacetylases

DNA methyltransferases

ATP-dep. chromatin remodeling factors

NR NR

Histone H3K27 methyltransferases

Histone H3K4 demethylases

FIG. 2. Molecular mechanisms of inhibiting gene expression by NRs. Regardless of the type of NR‐mediated repression (unliganded, antagonist‐bound, agonist‐bound), specific types of chromatin modifying enzymes are recruited to inhibit transcription.

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corepressor associates with one or more HDACs (i.e., HDAC3 for NCoR/ SMRT). Chromatin immunoprecipitation (ChIP) experiments performed on NR target gene promoters have unanimously found that in the absence of ligand the gene is hypoacetylated and becomes hyperacetylated in the presence of agonist. So, histone acetylation is a major mechanism through which NRs mediate transcriptional repression, but how does histone deacetylation actually repress transcription? First, acetylated histones repel internucleosomal interactions.66–68 Internucleosomal interactions are responsible for higher order chromatin structure, which sterically inhibits assembly of transcription machinery. Thus, histone deacetylation would promote the formation of higher order repressive chromatin. Second, acetylation of lysine neutralizes its positive charge thereby loosening the association between histones and DNA.69,70 This theoretically allows histones to be more easily displaced from promoter regions thereby allowing transcription factors access to the promoter. Third, acetylated lysine is the molecular substrate for the bromodomain.71 Bromodomains are found in numerous coactivators and ATP‐dependent chromatin remodeling complexes and function to stabilize their association with their target chromatin template (i.e., target gene promoter). Thus, recruitment of HDAC‐containing corepressor complexes by NRs inhibits transcription through multiple mechanisms.

B. Histone Methylation Histone methylation plays critical roles in regulating chromatin structure and function. Strong evidence indicates that histone methylation also plays important roles in NRs‐induced transcriptional repression. In this regard, CARM1 and PRMT1 were both identified as coactivators for NRs.72–74 CARM1 methylates histone H3 on R2, R17, and R26, whereas PRMT1 primarily methylates histone H4 on R3. Consistent with methyl‐H3K4 being a mark for transcriptional activity, the H3K4 methyltransferase MLL has been identified as a component of the ASC2 coactivator complex.75,76 In contrast, histone methyltransferases (HMTs) with specific activity for H3K9 or H3K27 are autonomous transcriptional repressors. SUV39H1, an H3K9‐specific HMT, was found to associate with unliganded TR.33 Thus, unliganded TR represses transcription through a combination of histone deacetylation mediated by HDAC3‐containing NCoR/SMRT complexes and H3K9 methylation mediated by SUV39H1. Another H3K9 HMT, G9a, was shown to mediate repression by the small heterodimer partner SHP.34,77 Enhancer of zeste homolog 2 (EZH2) is a polycomb protein with histone H3 K27 methyltransferase activity. Conflicting data have been reported for EZH2 as to its function for ER. In one report, EZH2 was shown to activate ER transcription, whereas in the

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other report EZH2 was identified as an REA‐interacting protein that functioned as an ER corepressor.78,79 Thus, the function of EZH2 for ER may be cell type and context dependent. LSD1 is the first identified histone lysine demethylase.35 In vitro purified LSD1 exhibits histone demethylase activity toward only mono‐ and dimethylated H3K4. LSD1 is part of the large protein complex that also contains CoREST and HDAC1/2.80 Given its association with HDAC1/2 and H3K4 demethylase activity, it is not surprising that knockdown of LSD1 by RNAi caused an increase of H3K4 methylation and concomitant derepression of target genes. Consistent with this notion, LSD1 serves as a corepressor for orphan receptor TLX (also called NR2E1).37 LSD1 interacts directly with TLX and potentiates TLX repression through its H3K4 demethylase activity. Interestingly, LSD1 was also identified as an AR‐interacting protein that facilitated AR transactivation.36 This functional switch is brought by a change in its demethylase specificity. Upon interaction with AR, LSD1 was shown to switch its specificity from methyl‐H3K4 to methyl‐H3K9. At this stage, the molecular mechanism that underlies this specificity switch is not clear. In addition to above specific examples, evidence exists for a general involvement of histone methylation in governing ligand‐dependent activation by NRs. It was shown that different H3K9 methyltransferases bring in repressive epigenetic marks and impose gene‐specific gatekeeper functions that prevent unliganded NRs or other transcription factors from binding to their target genes in the absence of ligands or other stimulating signals.81 Histone demethylases including LSD1 thus are required to reverse the repressive epigenetic marks and allow the ligand‐ and signal‐dependent transcriptional activation to take place. This study highlights the functional significance of histone methylation/demethylation in regulation of gene expression.

C. Chromatin Assembly/Remodeling ATP‐dependent chromatin remodeling factors utilize ATP hydrolysis to alter chromatin structure through mechanisms including nucleosome assembly/disassembly, sliding, and conformational changes. While SWI/SNF, a prototype ATP remodeling complex, generally functions as coactivator for NRs, a SNF2H‐containing remodeling complex was reported to be required for efficient repression by unliganded TR.82 In this case, the NCoR/HDAC3 complex and SNF2H complex function cooperatively. Mechanistically, NCoR/HDAC3 generates deacetylated histone H4 tails to which the SNF2H complex binds and induces nucleosomal reorganization. This mechanism of repression through chromatin reorganization is not unique to SNF2H, as corepressor Alien has also been reported to recruit a histone chaperone NAP1 for chromatin assembly.32 NAP1 can assemble or

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disassemble nucleosomes in a DNA replication‐independent manner. However, when tethered to DNA via the heterologous DBD of yeast Gal4, it preferentially represses transcription. Direct interaction of Alien with NAP1 enhances its nucleosome assembly activity. In addition, Alien can directly bind the core histones H3 and H4. Thus, the combination of stable chromatin association of NAP1 via the histone binding property of Alien combined with enhanced nucleosome assembly activity imparted by direct interaction with Alien provides a perfect environment for NAP1 to assemble nucleosomes in a manner that inhibits transcriptional activity. ATP‐dependent chromatin remodeling activity was also shown to participate in repression by the orphan receptor SHP.83 In this study, SHP directly interacted with Brm‐containing SWI/SNF remodeling complexes. Dominant negative ATPase mutants prevented repression. For the CYP7A1 gene, SHP‐ induced repression was mediated by both mSin3A‐HDAC1/2 and SWI/SNF. This is another example of the use of ATP‐dependent chromatin remodeling factors for transcriptional repression. The role of yet another chromatin assembly/remodeling protein Acf1 in NR repression has recently been demonstrated84 and is discussed in detail in Chapter 6.

D. DNA Methylation DNA methylation is a common mechanism for long‐term gene silencing. Since NRs can induce acute transcriptional repression, DNA methylation in general has not been considered as a mechanism for NR‐mediated repression. However, there is evidence for the involvement of DNA methylation in transcriptional repression induced by NRs. Acute promyelocytic leukemia (APL) is a subtype of AML characterized by excess promyelocytes and deficiency in cells of the myeloid lineage. The most common genetic cause of APL is t(15;17) translocation, which generates the leukemia‐promoting PML–RARa fusion protein.85,86 Contrary to full‐length RAR, PML–RARa maintains its interaction with SMRT and NCoR at physiological levels of RA.87,88 For this reason, the fusion protein functions as a constitutive repressor of RAR target genes, blocking differentiation and resulting in leukemogenesis. In addition to recruiting the SMRT/NCoR corepressor complexes, PML–RAR can induce hypermethylation of target genes by recruiting DNA methyltransferases.89 DNA hypermethylation contributes to the leukemogenic potential of PML–RAR. In another study, the corepressor RIP140 was reported to direct both histone and DNA methylation to silence Ucp1 expression in white adipocytes.90 As a recent genome wide study revealed an intimate correlation between histone methylation and DNA methylation,91 perhaps DNA methylation is more commonly involved in transcriptional repression than we previously thought.

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V. Physiological Functions of NR‐Mediated Repression Regardless whether repression is mediated by unliganded or liganded receptors, transcriptional repression is an integral function of all NRs. However, the physiological function of repression is difficult to assess and underappreciated. For example, while it is well established that unliganded TR is a potent repressor, whether this repression activity is physiologically relevant is still a subject of debate. Much of our knowledge concerning the physiological function of NR‐mediated repression comes from knockout studies of corepressor proteins. However, corepressors can mediate repression for both NR and non‐NR transcription factors. Here, we will summarize some recent progress in this area.

A. Physiological Function of Repression by Unliganded TR For most NRs, it is difficult to discern the physiological roles of repression from activation. For TRb, the mutations in the LBD that result in reduced hormone binding are responsible for the syndrome of resistance to thyroid hormone (RTH). RTH is characterized by elevated serum levels of T4 and T3, inappropriately ‘‘normal’’ or elevated serum TSH concentrations, diffuse goiter and varying manifestations of hypothyroidism.92 The RTH‐associated TRb mutations identified so far represent a range of reduced hormone‐binding activities. These TRb mutants maintain association with corepressors SMRT and NCoR at physiological levels of T3 and therefore behave as constitutive repressors. In a sense, the RTH syndrome exemplifies the pathological effects of transcriptional repression by unliganded TRb. Another example of a pathological link to unliganded TR is the v‐Erb A oncoprotein from the avian erythroblastosis virus (AEV). The v‐Erb A protein is a derivative of c‐Erb A, an avian TRa. Compared to c‐Erb A, v‐Erb A was found to sustain a series of alterations including an N‐terminal fusion of viral ‘‘gag’’ sequence, deletion of the C‐terminal AF2 and 13 internal point mutations.93,94 As a consequence, v‐Erb A binds constitutively and with higher affinity to the corepressors SMRT and NCoR.95 This repression function of v‐ Erb A in turn blocks the terminal differentiation of erythroid cells and contributes to its oncogenic activity. Despite the findings described above, it is still unclear if repression by unliganded TR is of physiological significance under normal developmental and growth conditions. In fact, mice with null mutations in all TR isoforms actually exhibit far less severe phenotypes than mice made congenitally

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hypothyroidism by treatment with thyrostatic drugs. These results imply that transcriptional repression by apo‐ or unliganded TRs are responsible for the detrimental effects of hypothyroidism.

B. TR‐Mediated Repression in the Regulation of Amphibian Metamorphosis Xenopus laevis, the African clawed frog, is a model for amphibian development. Frogs begin life as tadpoles and undergo metamorphosis to become air‐ breathing adults. Metamorphosis in frogs is under the master control of thyroid hormone. Although it is unclear if repression by unliganded TR is of physiological relevance in mammals, elegant studies in amphibian frogs have demonstrated a critical role for repression by unliganded TRs in regulating the timing and precise programs of metamorphosis. A dual function model has been proposed for regulation of amphibian development.96 Tadpoles express TRa in virtually all cell types, but do not produce thyroid hormone. Thus, TRa is bound by NCoR/SMRT complexes and mediates silencing of genes involved in metamorphosis.97,98 During metamorphosis, thyroid hormone is produced and TRb becomes expressed at high levels. In this state, TRa and TRb are bound by coactivators and activate expression of prometamorphic genes. As expected, gene activation correlates with dissociation of NCoR/SMRT, binding of coactivators, and histone hyperacetylation. This system not only represses genes that might adversely affect premetamorphic development, but also primes cells to be responsive to thyroid hormone so that they can induce metamorphosis at the appropriate time. The above dual function model is further supported by studies using dominant negative or positive transgenes. First, tadpoles expressing a dominant negative TR, which can bind thyroid hormone but cannot recruit coactivators, are unable to undergo metamorphosis even upon thyroid hormone treatment.99 Second, tadpoles expressing a dominant negative SRC3 also cannot undergo metamorphosis.100 Thus, both corepressor release and coactivator recruitment are required to induce the metamorphic program. Furthermore, expression of a dominant positive TR, which activates TR reporter genes independently of thyroid hormone, causes precocious metamorphosis.101 Additionally, expression of a dominant negative NCoR consisting of only its RID in tadpoles causes derepression of TR target genes and accelerated development.102 Thus, TR‐mediated repression is essential for proper premetamorphic development and regulating the timing of metamorphosis.

C. Repression by PPARg PPARg is a master regulator of adipogenesis. Its repression activity has been linked to maintenance of the preadipocyte cell type or in other words repression by PPARg prevents differentiation of preadipocytes to adipocytes.

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PPARg homozygous null mice die during embryogenesis due to placental defects. Following rescue of the placental phenotype by tetraploid complementation, PPARg null mice survive to term but then manifest defects in adipogenesis.103 White adipose tissue‐specific reduction of PPARg (via genetic deletion of exon 2) results in growth retardation, lipodystrophy, and hyperlipidemia.104,105 Under normal dietary conditions, PPARg activity prevents mobilization of fatty acids and stimulates adipogenesis. However, upon caloric restriction, repression of PPARg target genes prevents fat storage and promotes lipolysis. Thus, at least one of the physiological functions of PPARg‐mediated repression is to allow for the mobilization of fat stores during caloric restriction. In mice, mobilization of fatty acids from white adipose tissue is dependent on the PPARg corepressors, NCoR, and Sirt1 (mammalian ortholog of yeast Sir2).106 Sirt1 represses PPARg target genes, including those involved in fat storage, via direct interaction with NCoR. Caloric restriction induces expression of Sirt1.107 Thus, upon food withdrawal, Sirt1 interacts with PPARg to repress target genes involved in lipogenesis. Accordingly, mice heterozygous for a null mutation in Sirt1 have a limited ability to mobilize fatty acids upon food withdrawal. NIH 3T3‐L1 fibroblasts are a well characterized model of adipogenesis. In the absence of ligand, PPARg represses the adipogenic program in these cells in an NCoR‐dependent manner.108 Additionally, Sirt1 overexpression prevents and Sirt1 knockdown enhances adipogenesis in this model system. In primary adipocytes, increased expression of Sirt1 stimulates lipolysis. Thus, PPARg is a master regulator of adipogenesis and fatty acid metabolism, influencing both the commitment to adipocyte differentiation and the balance between lipogenesis and lipolysis. In this manner, the physiological function of PPARg‐mediated repression is to maintain the preadipocyte cell type and stimulate fatty acid breakdown.

D. Repression Mediated by NCoR and SMRT In general, the physiological functions of NR‐mediated repression have been difficult to address, although some recent studies have begun to shed light on this topic. As discussed above, repression by NRs is mediated by corepressors. Thus, corepressor knockout models have been generated to assess the functional significance of repression. However, most, if not all, NR corepressors also mediate the repressive actions of non‐NR transcription factors. These experiments provide excellent in vivo demonstration of the physiological function of individual corepressors, but fail to precisely address the role of NR‐mediated gene repression.

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Two recent studies have used gene targeting in mouse embryonic stem cells to create alleles of NCoR or SMRT with mutations in their NR interaction domains.109,110 The gene products of these alleles cannot interact with NRs, but it is assumed that they retain their ability to interact with non‐NR transcription factors. The phenotype of the SMRT RID mutant mice was interesting in that homozygous animals did not exhibit embryonic lethality, which was the phenotype of the SMRT null mutation. Instead animals appeared grossly normal and fertile; however, they did exhibit metabolic defects. Homozygous mutants exhibited a 20% decrease in respiration, a 70% increase in adiposity, increased blood glucose, glucose intolerance, and insulin resistance. For NCoR, the RID was conditionally deleted in the liver and was used to test the physiological importance of the NCoR–TR interaction. These mice also appeared grossly normal and fertile. When these mice were pharmacologically induced to be hypothyroid, repression of TR target genes was not observed. Thus, NCoR is critical for TR‐mediated repression in the liver. Interestingly, the authors also found increased expression of many TR target genes compared with wild type controls. These data suggest that NCoR regulates the transcriptional activity of liganded TR and supports an equilibrium model in which corepressors and coactivators compete for the NR LBD.

E. Repression Mediated by RIP140 RIP140 is a corepressor for most, if not all, NRs. The gene encoding RIP140 is expressed in virtually all tissues and subject to regulation by steroid hormones, retinoic acid, and vitamin D. It appears to primarily function in the regulation of energy homeostasis. Accordingly, highest gene expression is found in adipose, muscle, and liver. Studies of homozygous null RIP140 mice (Nrip1 / ) revealed the major function of this corepressor is to regulate metabolic gene expression.42 Nrip1 / mice exhibit a 70% reduction in total body fat, including almost complete absence of subcutaneous fat. White adipocytes were approximately 70% smaller; therefore, these mice have a limited capacity to store triglycerides and cholesterol esters. Additionally, these mice exhibit a 30% increase in respiration owing to an increase in the ratio of slow oxidative to fast twitch muscle fibers. These alterations in adipogenesis and muscle physiology have the ‘‘positive’’ effect of resistance to diet‐induced obesity. Unfortunately, Nrip1 / females are infertile due to an ovulation defect.43 The Nrip1 / phenotype is similar to that of PGC1a overexpression, indicating the main physiological function of this corepressor is to mediate the repression function of PPARg.

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VI. Concluding Remarks To conclude we would like to emphasize the fact that the ability to associate with corepressors and inhibit transcription is an intrinsic property of most members of the NR superfamily. While this is quite apparent for type II and orphan NRs, which constitutively bind DNA and repress transcription in the absence of ligand, it is also true for steroid receptors. Generally speaking, in the absence of hormone, steroid receptors (AR, ER, PR, MR, and GR) reside in the cytoplasm. Upon hormone binding, they translocate to the nucleus, bind DNA, and activate transcription. However, steroid receptors will also associate with corepressors upon binding antagonists. This highlights the fact that CoRNR box‐containing corepressors and NR box‐containing coactivators compete for the LBD of NRs. The equilibrium is shifted in either direction depending on the nature of the ligand. A prime example of this is the cell‐ type specific effects of the mixed agonist/antagonist tamoxifen on association of ERa with coactivators or corepressors.111 NRs mediate transcriptional repression by recruiting multiprotein corepressor complexes. In turn, corepressor complexes repress transcription because they either contain or associate with chromatin modifying enzymes. The best known repressive chromatin modifying enzymes are HDACs. However, we tried in this chapter to emphasize other repressive chromatin modifications brought about by corepressors. These include methylation of histone H3K9 and H3K27 and demethylation of H3K4. Additionally, several corepressor complexes associate with ATP‐dependent chromatin remodeling factors, which in this setting serve to assemble nucleosomes and compact local chromatin. Finally, it is important to note that we are far from understanding the physiological functions of NR‐mediated repression. We have reviewed the few studies that have used mouse models to address this topic and pointed out that it is inherently difficult to separate the effects of NR‐mediated repression from activation. In the future, we expect to see more attempts to determine the precise role of NR‐mediated repression using mouse genetics. Acknowledgments We apologize to colleagues whose original work could not be cited owing to space constraints. The work in J.W.’s laboratory is supported by a grant from National Science Foundation (90919025) and a grant from The Ministry of Science and Technology (2009CB918402). M.D.S. is supported by NCI training grant CA009299.

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References 1. Baniahmad A, Kohne AC, Renkawitz R. A transferable silencing domain is present in the thyroid hormone receptor, in the v‐erbA oncogene product and in the retinoic acid receptor. EMBO J 1992;11:1015–23. 2. Hu X, Lazar MA. The CoRNR motif controls the recruitment of corepressors by nuclear hormone receptors. Nature 1999;402:93–6. 3. Nagy L, Kao HY, Love JD, Li C, Banayo E, Gooch JT, et al. Mechanism of corepressor binding and release from nuclear hormone receptors. Genes Dev 1999;13:3209–16. 4. Perissi V, Staszewski LM, McInerney EM, Kurokawa R, Krones A, Rose DW, et al. Molecular determinants of nuclear receptor‐corepressor interaction. Genes Dev 1999;13:3198–208. 5. Heery DM, Kalkhoven E, Hoare S, Parker MG. A signature motif in transcriptional co‐activators mediates binding to nuclear receptors. Nature 1997;387:733–6. 6. Cavailles V, Dauvois S, L’Horset F, Lopez G, Hoare S, Kushner PJ, et al. Nuclear factor RIP140 modulates transcriptional activation by the estrogen receptor. EMBO J 1995;14:3741–51. 7. Ray A, Prefontaine KE. Physical association and functional antagonism between the p65 subunit of transcription factor NF‐kappa B and the glucocorticoid receptor. Proc Natl Acad Sci USA 1994;91:752–6. 8. Scheinman RI, Gualberto A, Jewell CM, Cidlowski JA, Baldwin AS, Jr. Characterization of mechanisms involved in transrepression of NF‐kappa B by activated glucocorticoid receptors. Mol Cell Biol 1995;15:943–53. 9. Li J, Wang J, Wang J, Nawaz Z, Liu JM, Qin J, et al. Both corepressor proteins SMRT and N‐CoR exist in large protein complexes containing HDAC3. EMBO J 2000;19:4342–50. 10. Zhang J, Kalkum M, Chait BT, Roeder RG. The N‐CoR‐HDAC3 nuclear receptor corepressor complex inhibits the JNK pathway through the integral subunit GPS2. Mol Cell 2002;9:611–23. 11. Emiliani S, Fischle W, Van Lint C, Al‐Abed Y, Verdin E. Characterization of a human RPD3 ortholog HDAC3. Proc Natl Acad Sci USA 1998;95:2795–800. 12. Guenther MG, Barak O, Lazar MA. The SMRT and N‐CoR corepressors are activating cofactors for histone deacetylase 3. Mol Cell Biol 2001;21:6091–101. 13. Yoon HG, Chan DW, Huang ZQ, Li J, Fondell JD, Qin J, et al. Purification and functional characterization of the human N‐CoR complex: the roles of HDAC3, TBL1 and TBLR1. EMBO J 2003;22:1336–46. 14. Perissi V, Aggarwal A, Glass CK, Rose DW, Rosenfeld MG. A corepressor/coactivator exchange complex required for transcriptional activation by nuclear receptors and other regulated transcription factors. Cell 2004;116:511–26. 15. Alland L, Muhle R, Hou H, Jr, Potes J, Chin L, Schreiber‐Agus N, et al. Role for N‐CoR and histone deacetylase in Sin3‐mediated transcriptional repression. Nature 1997;387:49–55. 16. Heinzel T, Lavinsky RM, Mullen TM, Soderstrom M, Laherty CD, Torchia J, et al. A complex containing N‐CoR, mSin3 and histone deacetylase mediates transcriptional repression. Nature 1997;387:43–8. 17. Nagy L, Kao HY, Chakravarti D, Lin RJ, Hassig CA, Ayer DE, et al. Nuclear receptor repression mediated by a complex containing SMRT, mSin3A, and histone deacetylase. Cell 1997;89:373–80. 18. Hassig CA, Fleischer TC, Billin AN, Schreiber SL, Ayer DE. Histone deacetylase activity is required for full transcriptional repression by mSin3A. Cell 1997;89:341–7.

254

STEWART AND WONG

19. Laherty CD, Yang WM, Sun JM, Davie JR, Seto E, Eisenman RN. Histone deacetylases associated with the mSin3 corepressor mediate mad transcriptional repression. Cell 1997;89:349–56. 20. Zhang Y, Iratni R, Erdjument‐Bromage H, Tempst P, Reinberg D. Histone deacetylases and SAP18, a novel polypeptide, are components of a human Sin3 complex. Cell 1997;89:357–64. 21. Zhang Y, Sun ZW, Iratni R, Erdjument‐Bromage H, Tempst P, Hampsey M, et al. SAP30, a novel protein conserved between human and yeast, is a component of a histone deacetylase complex. Mol Cell 1998;1:1021–31. 22. Alland L, David G, Shen‐Li H, Potes J, Muhle R, Lee HC, et al. Identification of mammalian Sds3 as an integral component of the Sin3/histone deacetylase corepressor complex. Mol Cell Biol 2002;22:2743–50. 23. Kasten MM, Ayer DE, Stillman DJ. SIN3‐dependent transcriptional repression by interaction with the Mad1 DNA‐binding protein. Mol Cell Biol 1996;16:4215–21. 24. Yang L, Mei Q, Zielinska‐Kwiatkowska A, Matsui Y, Blackburn ML, Benedetti D, et al. An ERG (ets‐related gene)‐associated histone methyltransferase interacts with histone deacetylases 1/2 and transcription co‐repressors mSin3A/B. Biochem J 2003;369:651–7. 25. Underhill C, Qutob MS, Yee SP, Torchia J. A novel nuclear receptor corepressor complex, N‐CoR, contains components of the mammalian SWI/SNF complex and the corepressor KAP‐1. J Biol Chem 2000;275:40463–70. 26. Dressel U, Thormeyer D, Altincicek B, Paululat A, Eggert M, Schneider S, et al. Alien, a highly conserved protein with characteristics of a corepressor for members of the nuclear hormone receptor superfamily. Mol Cell Biol 1999;19:3383–94. 27. Altincicek B, Tenbaum SP, Dressel U, Thormeyer D, Renkawitz R, Baniahmad A. Interaction of the corepressor Alien with DAX‐1 is abrogated by mutations of DAX‐1 involved in adrenal hypoplasia congenita. J Biol Chem 2000;275:7662–7. 28. Polly P, Herdick M, Moehren U, Baniahmad A, Heinzel T, Carlberg C. VDR‐Alien: a novel, DNA‐selective vitamin D(3) receptor‐corepressor partnership. FASEB J 2000;14:1455–63. 29. Moehren U, Papaioannou M, Reeb CA, Hong W, Baniahmad A. Alien interacts with the human androgen receptor and inhibits prostate cancer cell growth. Mol Endocrinol 2007;21:1039–48. 30. Lykke‐Andersen K, Schaefer L, Menon S, Deng XW, Miller JB, Wei N. Disruption of the COP9 signalosome Csn2 subunit in mice causes deficient cell proliferation, accumulation of p53 and cyclin E, and early embryonic death. Mol Cell Biol 2003;23:6790–7. 31. Moehren U, Dressel U, Reeb CA, Vaisanen S, Dunlop TW, Carlberg C, et al. The highly conserved region of the co‐repressor Sin3A functionally interacts with the co‐repressor Alien. Nucleic Acids Res 2004;32:2995–3004. 32. Eckey M, Hong W, Papaioannou M, Baniahmad A. The nucleosome assembly activity of NAP1 is enhanced by Alien. Mol Cell Biol 2007;27:3557–68. 33. Li J, Lin Q, Yoon HG, Huang ZQ, Strahl BD, Allis CD, et al. Involvement of histone methylation and phosphorylation in regulation of transcription by thyroid hormone receptor. Mol Cell Biol 2002;22:5688–97. 34. Fang S, Miao J, Xiang L, Ponugoti B, Treuter E, Kemper JK. Coordinated recruitment of histone methyltransferase G9a and other chromatin‐modifying enzymes in SHP‐mediated regulation of hepatic bile acid metabolism. Mol Cell Biol 2007;27:1407–24. 35. Shi Y, Lan F, Matson C, Mulligan P, Whetstine JR, Cole PA, et al. Histone demethylation mediated by the nuclear amine oxidase homolog LSD1. Cell 2004;119:941–53. 36. Metzger E, Wissmann M, Yin N, Muller JM, Schneider R, Peters AH, et al. LSD1 demethylates repressive histone marks to promote androgen‐receptor‐dependent transcription. Nature 2005;437:436–9.

NUCLEAR RECEPTOR REPRESSION

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37. Yokoyama A, Takezawa S, Schule R, Kitagawa H, Kato S. Transrepressive function of TLX requires the histone demethylase LSD1. Mol Cell Biol 2008;28:3995–4003. 38. Augereau P, Badia E, Balaguer P, Carascossa S, Castet A, Jalaguier S, et al. Negative regulation of hormone signaling by RIP140. J Steroid Biochem Mol Biol 2006;102:51–9. 39. Lopez GN, Webb P, Shinsako JH, Baxter JD, Greene GL, Kushner PJ. Titration by estrogen receptor activation function‐2 of targets that are downstream from coactivators. Mol Endocrinol 1999;13:897–909. 40. Wei LN, Hu X, Chandra D, Seto E, Farooqui M. Receptor‐interacting protein 140 directly recruits histone deacetylases for gene silencing. J Biol Chem 2000;275:40782–7. 41. Christian M, Tullet JM, Parker MG. Characterization of four autonomous repression domains in the corepressor receptor interacting protein 140. J Biol Chem 2004;279:15645–51. 42. Leonardsson G, Steel JH, Christian M, Pocock V, Milligan S, Bell J, et al. Nuclear receptor corepressor RIP140 regulates fat accumulation. Proc Natl Acad Sci USA 2004;101:8437–42. 43. White R, Leonardsson G, Rosewell I, Ann Jacobs M, Milligan S, Parker M. The nuclear receptor co‐repressor nrip1 (RIP140) is essential for female fertility. Nat Med 2000;6:1368–74. 44. Fernandes I, Bastien Y, Wai T, Nygard K, Lin R, Cormier O, et al. Ligand‐dependent nuclear receptor corepressor LCoR functions by histone deacetylase‐dependent and ‐independent mechanisms. Mol Cell 2003;11:139–50. 45. Sewalt RG, Gunster MJ, van der Vlag J, Satijn DP, Otte AP. C‐Terminal binding protein is a transcriptional repressor that interacts with a specific class of vertebrate Polycomb proteins. Mol Cell Biol 1999;19:777–87. 46. Potter GB, Beaudoin GM, III, DeRenzo CL, Zarach JM, Chen SH, Thompson CC. The hairless gene mutated in congenital hair loss disorders encodes a novel nuclear receptor corepressor. Genes Dev 2001;15:2687–701. 47. Potter GB, Zarach JM, Sisk JM, Thompson CC. The thyroid hormone‐regulated corepressor hairless associates with histone deacetylases in neonatal rat brain. Mol Endocrinol 2002;16:2547–60. 48. Hsieh JC, Sisk JM, Jurutka PW, Haussler CA, Slater SA, Haussler MR, et al. Physical and functional interaction between the vitamin D receptor and hairless corepressor, two proteins required for hair cycling. J Biol Chem 2003;278:38665–74. 49. Moraitis AN, Giguere V, Thompson CC. Novel mechanism of nuclear receptor corepressor interaction dictated by activation function 2 helix determinants. Mol Cell Biol 2002;22:6831–41. 50. Chatterjee VK, Lee JK, Rentoumis A, Jameson JL. Negative regulation of the thyroid‐ stimulating hormone alpha gene by thyroid hormone: receptor interaction adjacent to the TATA box. Proc Natl Acad Sci USA 1989;86:9114–8. 51. Coser KR, Chesnes J, Hur J, Ray S, Isselbacher KJ, Shioda T. Global analysis of ligand sensitivity of estrogen inducible and suppressible genes in MCF7/BUS breast cancer cells by DNA microarray. Proc Natl Acad Sci USA 2003;100:13994–9. 52. Yen PM, Feng X, Flamant F, Chen Y, Walker RL, Weiss RE, et al. Effects of ligand and thyroid hormone receptor isoforms on hepatic gene expression profiles of thyroid hormone receptor knockout mice. EMBO Rep 2003;4:581–7. 53. Liu YY, Brent GA. Thyroid hormone‐dependent gene expression in differentiated embryonic stem cells and embryonal carcinoma cells: identification of novel thyroid hormone target genes by deoxyribonucleic acid microarray analysis. Endocrinology 2005;146:776–83. 54. Galon J, Franchimont D, Hiroi N, Frey G, Boettner A, Ehrhart‐Bornstein M, et al. Gene profiling reveals unknown enhancing and suppressive actions of glucocorticoids on immune cells. FASEB J 2002;16:61–71.

256

STEWART AND WONG

55. James CG, Ulici V, Tuckermann J, Underhill TM, Beier F. Expression profiling of Dexamethasone‐treated primary chondrocytes identifies targets of glucocorticoid signalling in endochondral bone development. BMC Genomics 2007;8:205. 56. Swami S, Raghavachari N, Muller UR, Bao YP, Feldman D. Vitamin D growth inhibition of breast cancer cells: gene expression patterns assessed by cDNA microarray. Breast Cancer Res Treat 2003;80:49–62. 57. Wang TT, Tavera‐Mendoza LE, Laperriere D, Libby E, MacLeod NB, Nagai Y, et al. Large‐ scale in silico and microarray‐based identification of direct 1,25‐dihydroxyvitamin D3 target genes. Mol Endocrinol 2005;19:2685–95. 58. Hollenberg AN, Monden T, Flynn TR, Boers ME, Cohen O, Wondisford FE. The human thyrotropin‐releasing hormone gene is regulated by thyroid hormone through two distinct classes of negative thyroid hormone response elements. Mol Endocrinol 1995;9:540–50. 59. Shibusawa N, Hollenberg AN, Wondisford FE. Thyroid hormone receptor DNA binding is required for both positive and negative gene regulation. J Biol Chem 2003;278:732–8. 60. Ortiga‐Carvalho TM, Shibusawa N, Nikrodhanond A, Oliveira KJ, Machado DS, Liao XH, et al. Negative regulation by thyroid hormone receptor requires an intact coactivator‐binding surface. J Clin Invest 2005;115:2517–23. 61. Wulf A, Wetzel MG, Kebenko M, Kroger M, Harneit A, Merz J, et al. The role of thyroid hormone receptor DNA binding in negative thyroid hormone‐mediated gene transcription. J Mol Endocrinol 2008;41:25–34. 62. Wang D, Xia X, Liu Y, Oetting A, Walker RL, Zhu Y, et al. Negative regulation of TSH{alpha} target gene by thyroid hormone involves histone acetylation and corepressor complex dissociation. Mol Endocrinol 2009;23:600–9. 63. Murayama A, Kim MS, Yanagisawa J, Takeyama K, Kato S. Transrepression by a liganded nuclear receptor via a BHLH activator through co‐regulator switching. EMBO J 2004;23:1598–608. 64. Kato S, Fujiki R, Kim MS, Kitagawa H. Ligand‐induced transrepressive function of VDR requires a chromatin remodeling complex, WINAC. J Steroid Biochem Mol Biol 2007;103:372–80. 65. Cvoro A, Tzagarakis‐Foster C, Tatomer D, Paruthiyil S, Fox MS, Leitman DC. Distinct roles of unliganded and liganded estrogen receptors in transcriptional repression. Mol Cell 2006;21:555–64. 66. Garcia‐Ramirez M, Rocchini C, Ausio J. Modulation of chromatin folding by histone acetylation. J Biol Chem 1995;270:17923–8. 67. Tse C, Sera T, Wolffe AP, Hansen JC. Disruption of higher‐order folding by core histone acetylation dramatically enhances transcription of nucleosomal arrays by RNA polymerase III. Mol Cell Biol 1998;18:4629–38. 68. Wang X, Hayes JJ. Acetylation mimics within individual core histone tail domains indicate distinct roles in regulating the stability of higher‐order chromatin structure. Mol Cell Biol 2008;28:227–36. 69. Cary PD, Crane‐Robinson C, Bradbury EM, Dixon GH. Effect of acetylation on the binding of N‐terminal peptides of histone H4 to DNA. Eur J Biochem 1982;127:137–43. 70. Hong L, Schroth GP, Matthews HR, Yau P, Bradbury EM. Studies of the DNA binding properties of histone H4 amino terminus. Thermal denaturation studies reveal that acetylation markedly reduces the binding constant of the H4 ‘‘tail’’ to DNA. J Biol Chem 1993;268:305–14. 71. Dhalluin C, Carlson JE, Zeng L, He C, Aggarwal AK, Zhou MM. Structure and ligand of a histone acetyltransferase bromodomain. Nature 1999;399:491–6.

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72. Chen D, Ma H, Hong H, Koh SS, Huang SM, Schurter BT, et al. Regulation of transcription by a protein methyltransferase. Science 1999;284:2174–7. 73. Koh SS, Chen D, Lee YH, Stallcup MR. Synergistic enhancement of nuclear receptor function by p160 coactivators and two coactivators with protein methyltransferase activities. J Biol Chem 2001;276:1089–98. 74. Wang H, Huang ZQ, Xia L, Feng Q, Erdjument‐Bromage H, Strahl BD, et al. Methylation of histone H4 at arginine 3 facilitating transcriptional activation by nuclear hormone receptor. Science 2001;293:853–7. 75. Goo YH, Sohn YC, Kim DH, Kim SW, Kang MJ, Jung DJ, et al. Activating signal cointegrator 2 belongs to a novel steady‐state complex that contains a subset of trithorax group proteins. Mol Cell Biol 2003;23:140–9. 76. Lee S, Lee DK, Dou Y, Lee J, Lee B, Kwak E, et al. Coactivator as a target gene specificity determinant for histone H3 lysine 4 methyltransferases. Proc Natl Acad Sci USA 2006;103:15392–7. 77. Boulias K, Talianidis I. Functional role of G9a‐induced histone methylation in small heterodimer partner‐mediated transcriptional repression. Nucleic Acids Res 2004;32:6096–103. 78. Hwang C, Giri VN, Wilkinson JC, Wright CW, Wilkinson AS, Cooney KA, et al. EZH2 regulates the transcription of estrogen‐responsive genes through association with REA, an estrogen receptor corepressor. Breast Cancer Res Treat 2008;107:235–42. 79. Shi B, Liang J, Yang X, Wang Y, Zhao Y, Wu H, et al. Integration of estrogen and Wnt signaling circuits by the polycomb group protein EZH2 in breast cancer cells. Mol Cell Biol 2007;27:5105–19. 80. Shi YJ, Matson C, Lan F, Iwase S, Baba T, Shi Y. Regulation of LSD1 histone demethylase activity by its associated factors. Mol Cell 2005;19:857–64. 81. Garcia‐Bassets I, Kwon YS, Telese F, Prefontaine GG, Hutt KR, Cheng CS, et al. Histone methylation‐dependent mechanisms impose ligand dependency for gene activation by nuclear receptors. Cell 2007;128:505–18. 82. Alenghat T, Yu J, Lazar MA. The N‐CoR complex enables chromatin remodeler SNF2H to enhance repression by thyroid hormone receptor. EMBO J 2006;25:3966–74. 83. Kemper JK, Kim H, Miao J, Bhalla S, Bae Y. Role of an mSin3A‐Swi/Snf chromatin remodeling complex in the feedback repression of bile acid biosynthesis by SHP. Mol Cell Biol 2004;24:7707–19. 84. Ewing AK, Attner M, Chakravarti D. Novel regulatory role for human Acf1 in transcriptional repression of vitamin D3 receptor‐regulated genes. Mol Endocrinol 2007;21:1791–806. 85. de The H, Lavau C, Marchio A, Chomienne C, Degos L, Dejean A. The PML‐RAR alpha fusion mRNA generated by the t(15;17) translocation in acute promyelocytic leukemia encodes a functionally altered RAR. Cell 1991;66:675–84. 86. Kakizuka A, Miller WH, Jr, Umesono K, Warrell RP, Jr, Frankel SR, Murty V, et al. Chromosomal translocation t(15;17) in human acute promyelocytic leukemia fuses RAR alpha with a novel putative transcription factor, PML. Cell 1991;66:663–74. 87. Hong SH, David G, Wong CW, Dejean A, Privalsky ML. SMRT corepressor interacts with PLZF and with the PML‐retinoic acid receptor alpha (RARalpha) and PLZF‐RARalpha oncoproteins associated with acute promyelocytic leukemia. Proc Natl Acad Sci USA 1997;94:9028–33. 88. Guidez F, Ivins S, Zhu J, Soderstrom M, Waxman S, Zelent A. Reduced retinoic acid‐sensitivities of nuclear receptor corepressor binding to PML‐ and PLZF‐RARalpha underlie molecular pathogenesis and treatment of acute promyelocytic leukemia. Blood 1998;91:2634–42.

258

STEWART AND WONG

89. Di Croce L, Raker VA, Corsaro M, Fazi F, Fanelli M, Faretta M, et al. Methyltransferase recruitment and DNA hypermethylation of target promoters by an oncogenic transcription factor. Science 2002;295:1079–82. 90. Kiskinis E, Hallberg M, Christian M, Olofsson M, Dilworth SM, White R, et al. RIP140 directs histone and DNA methylation to silence Ucp1 expression in white adipocytes. EMBO J 2007;26:4831–40. 91. Meissner A, Mikkelsen TS, Gu H, Wernig M, Hanna J, Sivachenko A, et al. Genome‐scale DNA methylation maps of pluripotent and differentiated cells. Nature 2008;454:766–70. 92. Refetoff S. Resistance to thyroid hormone. Clin Lab Med 1993;13:563–81. 93. Sap J, Munoz A, Damm K, Goldberg Y, Ghysdael J, Leutz A, et al. The c‐erb‐A protein is a high‐affinity receptor for thyroid hormone. Nature 1986;324:635–40. 94. Weinberger C, Thompson CC, Ong ES, Lebo R, Gruol DJ, Evans RM. The c‐erb‐A gene encodes a thyroid hormone receptor. Nature 1986;324:641–6. 95. Lee S, Privalsky ML. Multiple mutations contribute to repression by the v‐Erb A oncoprotein. Oncogene 2005;24:6737–52. 96. Sachs LM, Damjanovski S, Jones PL, Li Q, Amano T, Ueda S, et al. Dual functions of thyroid hormone receptors during Xenopus development. Comp Biochem Physiol B Biochem Mol Biol 2000;126:199–211. 97. Sachs LM, Jones PL, Havis E, Rouse N, Demeneix BA, Shi YB. Nuclear receptor corepressor recruitment by unliganded thyroid hormone receptor in gene repression during Xenopus laevis development. Mol Cell Biol 2002;22:8527–38. 98. Tomita A, Buchholz DR, Shi YB. Recruitment of N‐CoR/SMRT‐TBLR1 corepressor complex by unliganded thyroid hormone receptor for gene repression during frog development. Mol Cell Biol 2004;24:3337–46. 99. Buchholz DR, Hsia SC, Fu L, Shi YB. A dominant‐negative thyroid hormone receptor blocks amphibian metamorphosis by retaining corepressors at target genes. Mol Cell Biol 2003;23:6750–8. 100. Paul BD, Fu L, Buchholz DR, Shi YB. Coactivator recruitment is essential for liganded thyroid hormone receptor to initiate amphibian metamorphosis. Mol Cell Biol 2005;25:5712–24. 101. Buchholz DR, Tomita A, Fu L, Paul BD, Shi YB. Transgenic analysis reveals that thyroid hormone receptor is sufficient to mediate the thyroid hormone signal in frog metamorphosis. Mol Cell Biol 2004;24:9026–37. 102. Sato Y, Buchholz DR, Paul BD, Shi YB. A role of unliganded thyroid hormone receptor in postembryonic development in Xenopus laevis. Mech Dev 2007;124:476–88. 103. Barak Y, Nelson MC, Ong ES, Jones YZ, Ruiz‐Lozano P, Chien KR, et al. PPAR gamma is required for placental, cardiac, and adipose tissue development. Mol Cell 1999;4:585–95. 104. Akiyama TE, Sakai S, Lambert G, Nicol CJ, Matsusue K, Pimprale S, et al. Conditional disruption of the peroxisome proliferator‐activated receptor gamma gene in mice results in lowered expression of ABCA1, ABCG1, and apoE in macrophages and reduced cholesterol efflux. Mol Cell Biol 2002;22:2607–19. 105. Koutnikova H, Cock TA, Watanabe M, Houten SM, Champy MF, Dierich A, et al. Compensation by the muscle limits the metabolic consequences of lipodystrophy in PPAR gamma hypomorphic mice. Proc Natl Acad Sci USA 2003;100:14457–62. 106. Picard F, Kurtev M, Chung N, Topark‐Ngarm A, Senawong T, Machado De Oliveira R, et al. Sirt1 promotes fat mobilization in white adipocytes by repressing PPAR‐gamma. Nature 2004;429:771–6. 107. Chen D, Bruno J, Easlon E, Lin SJ, Cheng HL, Alt FW, et al. Tissue‐specific regulation of SIRT1 by calorie restriction. Genes Dev 2008;22:1753–7.

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108. Yu C, Markan K, Temple KA, Deplewski D, Brady MJ, Cohen RN. The nuclear receptor corepressors NCoR and SMRT decrease peroxisome proliferator‐activated receptor gamma transcriptional activity and repress 3T3‐L1 adipogenesis. J Biol Chem 2005;280:13600–5. 109. Astapova I, Lee LJ, Morales C, Tauber S, Bilban M, Hollenberg AN. The nuclear corepressor, NCoR, regulates thyroid hormone action in vivo. Proc Natl Acad Sci USA 2008;105:19544–9. 110. Nofsinger RR, Li P, Hong SH, Jonker JW, Barish GD, Ying H, et al. SMRT repression of nuclear receptors controls the adipogenic set point and metabolic homeostasis. Proc Natl Acad Sci USA 2008;105:20021–6. 111. Smith CL, Nawaz Z, O’Malley BW. Coactivator and corepressor regulation of the agonist/ antagonist activity of the mixed antiestrogen, 4‐hydroxytamoxifen. Mol Endocrinol 1997;11:657–66.

The Roles and Action Mechanisms of p160/SRC Coactivators and the ANCCA Coregulator in Cancer Elaine Y.C. Hsia,*,{ June X. Zou,{,z and Hong‐Wu Chen*,z *Department of Biochemistry and Molecular Medicine, University of California at Davis, Sacramento, California 95817 {

Department of Internal Medicine, School of Medicine, University of California at Davis, Sacramento, California 95817

z

UC Davis Cancer Center/Basic Sciences, University of California at Davis, Sacramento, California 95817

I. Introduction: The Discovery of AIB1/ACTR/SRC‐3 as a Nuclear Hormone Receptor Coactivator and a Gene Amplified in Cancer ........................... II. Aberrant Genetic Regulation of p160/SRC Expression in Cancers ............. A. Chromosomal Alterations............................................................ B. Polymorphisms......................................................................... C. Aberrant Expression of SRCs in Cancers and the Potential Underlying Mechanisms ............................................................................ III. The p160/SRCs Functions and Their Action Mechanisms in Cancer Cells... A. Cell Proliferation, Survival, Invasion, and Metastasis ......................... B. Tamoxifen Resistance in Breast Cancer .......................................... C. Androgen Independence in Prostate Cancer.................................... IV. Functions of p160/SRCs in Tumorigenesis Revealed in Animal Models ...... A. SRCs in Mouse Mammary Tumorigenesis and Tumor Metastasis.......... B. SRC‐3 in the Mouse Prostate Cancer Model ................................... V. The Coregulator ANCCA, a Unique Target of AIB1/ACTR and a Potential Key Player in Cancer...................................................................... A. ANCCA is a Hormone‐Induced Nuclear Receptor Coregulator and a Target of AIB1/ACTR ................................................................ B. ANCCA, a Potential Key Player in Tumorigenesis, is Frequently Overexpressed in Different Cancers .............................................. VI. Concluding Remarks ...................................................................... References...................................................................................

Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87008-7

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Chromosomal aberrations involving genes encoding members of the p160/SRC transcriptional coactivator family such as AIB1/ACTR and TIF2 implicated the coactivators in malignancy of human cells. Significant progress has been made in the last decade toward uncovering their roles in the development and progression of solid tissue tumors as well as leukemia and understanding of the underlying molecular mechanisms. Here, we review their genetic aberrations and dysregulation in expression in breast cancer, prostate cancer, and other nonhormone‐responsive cancers. The experimental evidence gathered from studies using cell culture and animal models strongly supports a critical and, in some circumstances, their oncogenic function. We summarize results that the SRCs may contribute to tumorigenesis and disease progression through transcription factors such as E2F, PEA3, and AP‐1 and through an intimate control of signaling pathways of growth factors‐Akt and the receptor tyrosine kinases. The finding that a recently identified nuclear receptor coregulator ANCCA, like the SRCs, is frequently overexpressed in many types of cancers again underscores their broader roles in cancer.

I. Introduction: The Discovery of AIB1/ACTR/SRC‐3 as a Nuclear Hormone Receptor Coactivator and a Gene Amplified in Cancer It is generally conceived that different members of the nuclear hormone receptor superfamily play distinct roles in cancer.1 For instance, estrogen receptor (ER)‐alpha and androgen receptor (AR) are well known for their aberrant function in the development and progression of breast cancers and prostate cancers, respectively. However, loss or impairment of normal function of other receptors such as retinoic acid receptor (RAR)‐alpha was strongly implicated in the development or progression of other malignancies such as leukemia.2,3 Although the p160/SRCs were initially thought to function as coactivators for a specific receptor or a group of receptors (such as steroid receptors), it quickly became clear that each of the three SRCs can mediate transcriptional activation for multiple members of the receptor superfamily. For instance, SRC‐1 can serve as a coactivator for estrogen receptors as well as for RARs. Indeed, SRC family of proteins shares several functional domains that mediate interaction with the different receptors and other transcriptional coregulators such as p300, CBP, and CARM1 (Fig. 1). Thus, whether and how the SRCs are involved in tumorigenesis was not immediately apparent. However, the identification of a third member of the SRC family changed this perception.

ACTION MECHANISMS OF

p160/SRC AND THE ANCCA IN CANCER PAS

bHLH

A

B S/T

263

RID

AD1

AD2

Nuclear receptors

CBP/ p300

CARM1/ PRMT1

SRC-1 1

Q L1 L2 L3

L7

L4 L5 L6

TIF2 1

1441

HAT

1464

Q EID

ACTR/AIB1 1

Q

Q

HAT

1412

FIG. 1. The functional domains of the p160/SRC coactivator family. The locations of conserved structural and functional domains for full‐length human SRC proteins are indicated by filled or textured boxes and bars. bHLH, basic helix‐loop‐helix; PAS, Per/ARNT/Sim domain; S/T, serine/ threonine‐rich regions; RID, receptor interaction domain; L1–L6 (L7), LXXLL alpha‐helix motifs; Q, glutamine‐rich regions; HAT, histone acetyltransferase domains; AD1 and AD2, transcriptional activation domains; EID, E2F1 interaction domain. Interaction partners for RID, AD1, and AD2 are indicated under the lines. The number of amino acid residues for different human SRCs are based on NCBI protein database entries: NP_003734 (SRC‐1), NP_006531 (TIF2), AAB92368 (AIB1/ACTR).

While many reported the third SRC identification using yeast‐based protein–protein interaction methods (hence the different names given, including ACTR, RAC3, and TRAM1. Here, we will use AIB1/ACTR for the human gene and SRC‐3 for the mouse gene),4–6 one study took a cancer genetic approach. Previous molecular cytogenetic studies had indicated that chromosomal region 20q is one of the common locations frequently amplified in breast cancer.7–9 To identify the target genes involved, a chromosome microdissection‐ hybrid selection technique was employed to clone the cDNA sequences expressed from the amplified region. The partial cDNA sequences were found to be encoded by different genes with amplified in breast cancer 1 (AIB1) located at 20q12 and AIB3 at 20q11.10 Upon cloning of the full‐length cDNA sequence, it became apparent that AIB1/ACTR was a third member of human SRC family of coactivators.11 Analysis of AIB1/ACTR transcript level in human breast cancer specimens and cell lines revealed that tumors and cell lines with its gene amplification displayed high levels of expression, indicating that AIB1/ACTR gene amplification resulted in its overexpression. Thus, together with the demonstration of AIB1/ACTR as an ER coactivator, the findings strongly suggested that this third SRC plays an important role in breast cancer. As reviewed in the following sections, multiple lines of evidence from many studies conducted since its initial identification now clearly indicate that aberrant AIB1/ACTR/SRC‐3 is indeed critical for tumorigenesis in multiple types of cancers and that the mechanisms underlying its oncogenicity go well

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beyond its function as an ER coactivator. Studies on the other two SRCs also strongly implicated their involvement in different malignancies with complex mechanisms.

II. Aberrant Genetic Regulation of p160/SRC Expression in Cancers Much research has established a strong link between altered p160/SRC coregulator gene expression and disease, particularly cancer. Mutations in p160 coactivator genes appear to be rare events in cancers.12–14 Thus, aberrations of p160 genes identified so far in cancers are mainly in their overexpression at both transcript and protein levels and, to a much lesser extent, in alterations of their structure and function. Here, we review evidence for the roles of various genetic aberrations in cancer‐associated p160 coactivator expression and the potential underlying mechanisms that link altered expression to cancer.

A. Chromosomal Alterations 1. GENE AMPLIFICATION OF AIB1 AND SRC‐1 Gene amplification and the consequent overexpression of AIB1/ACTR have been frequently detected in various types of solid tumors (Table I). One of the first groups to discover AIB1/ACTR identified the gene as an expressed sequence from chromosome 20q12 that is amplified in approximately 10% of primary breast cancer biopsies (hence, AIB1) and in a panel of ER positive breast and ovarian cancer cell lines.11 Elevation of AIB1/ACTR mRNA in 31–64% of breast cancer specimens compared to normal mammary epithelium suggests that other mechanisms besides gene amplification significantly contribute to overexpression.11,15 Indeed, similar statistics for AIB1/ACTR amplification and overexpression were observed in clinical samples of primary gastric, urothelial, colorectal, and esophageal squamous cell carcinomas examined with FISH and immunohistochemistry,16–19 although one study found AIB1/ACTR amplification in 37% of pancreatic adenocarcinoma (n ¼ 78).20 While gene amplification consistently correlated with high protein levels, AIB1/ACTR overexpression was also observed in a significant proportion of tumor samples with normal gene copy number. AIB1/ACTR amplification and accompanied overexpression have been suggested as potential markers for disease progression and prognosis with varying power for different types of tumors. Although there are some conflicting reports on whether AIB1/ACTR is expressed exclusively in mammary tumor tissue, it is consistently found to be significantly overexpressed in tumors relative to normal mammary epithelium,21,22 and AIB1/ACTR

TABLE I GENETIC ALTERATIONS DETECTED FOR THE HUMAN p160/SRC COACTIVATORS Genetic alteration

SRC

Amplification

AIB1/ACTR

Type

Locus or position

Phenotype

Cancer

References

20q12

Overexpression

Breast

11, 57

Ovarian

11, 25, 57

Gastric

16

Urothelial

17

Colorectal

18

Esophageal

19

Pancreatic

20

SRC‐1 Chromosomal rearrangement

Hepatocellular

23

(Prostate)

28–30

(Endometrial)

24

(Breast)

55, 61, 100

Prostate

12, 26

TIF2

Inversion

inv(8)(p11q13)

MOZ–TIF2

Acute myeloid leukemia (M4, M5 subtypes)

33, 36, 37

TIF2

Translocation

t(8;12)(q13;p13)

ETV6–TIF2

Acute lymphoblastic leukemia

40

SRC‐1

t(2;2)(q35;p23)

PAX3–SRC‐1

Pediatric rhabdomyosarcoma

42

AIB1/ACTR

t(8;20)(p11;q13)

MOZ–ACTR

Acute myeloid leukemia (M5 subtype)

43

(Continues)

TABLE I (Continued) Genetic alteration

SRC

Type

Locus or position

Phenotype

Cancer

References

Polymorphism

AIB1/ACTR

SNP

1758G > C

Q586H

Breast cancer risk

13

2880A > G

T960T

N/A

A407S

Amino acid residues 1244–1272

Variable poly‐Q length

TIF2 AIB1/ACTR

VNTR (CAG)n

14 Breast cancer risk

44–49, 52

Prostate cancer risk

50

Ovarian

51

The alterations are listed according to type, location, resulting phenotype, and associated tissues for which the alteration was observed in or linked to risk for malignant disease. For cancers depicted in italicized letters inside parentheses, SRC overexpression, but not gene amplification, has been observed; or information regarding amplification in the clinical studies is unavailable. The symbol > indicates that the left allele is the more common variant.

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overexpression correlates with high tumor grade.15,22 In breast and bladder cancers, frequency of AIB1/ACTR overexpression does not change significantly with different histological stages, suggesting that overexpression may be an early event that is maintained throughout tumor progression.15,17 By contrast, high AIB1/ACTR expression or amplification was detected more frequently in the later stages of gastric, colorectal, pancreatic, esophageal, and endometrial cancer, and in metastatic or recurrent hepatocellular carcinoma when compared to primary disease.16,18–20,23,24 Significant correlation between overexpressed AIB1/ACTR and increased expression of Ki‐67 (a cell proliferation marker) suggests that high levels of AIB1/ACTR confer a growth advantage during tumor progression of urothelial and esophageal squamous cells.17,19 Poor prognosis (lower disease‐free survival or DFS) also correlated with AIB1/ACTR overexpression in bladder cancer and endometrial cancer patients, and with gene amplification in gastric cancer patients.16,17,24 In a study with limited number of sporadic ovarian tumor samples, AIB1/ACTR amplification was linked to a tendency towards lower DFS.25 In summary, these clinical studies suggest that AIB1/ACTR amplification and overexpression can be useful markers for prognostic evaluation of patients with various malignant diseases. The notable discrepancies between percentage of cases with increased gene copy number and high RNA or protein levels, however, also suggest that genetic regulation of AIB1/ACTR expression is complex and most certainly involves transcriptional and posttranscriptional mechanisms. In contrast to AIB1/ACTR amplification, there is little information regarding gene copy number change in SRC‐1 and SRC‐2/TIF2 genes. One study thus far reported a very low frequency (2 out of 70 tumors examined) of SRC‐1 gene amplification in androgen ablation‐resistant prostate tumors and xenografts.12 Interestingly, studies using matched patient specimens often show comparable SRC‐1, TIF2, and AIB1/ACTR expression levels in tumor and normal prostate tissue or BPH.26,27 Other studies show variable levels of AIB1/ACTR expression in normal, BPH and malignant prostate tissues. Nevertheless, most clinical studies reveal significant association of higher protein expression of the p160 coactivators and markers of more aggressive prostate cancer and disease progression.26,28–31 2. CHROMOSOMAL TRANSLOCATION AND INVERSION INVOLVING THE SRCS Chromosomal rearrangements involving the p160 coactivators AIB1/ACTR and TIF2 (also named as SRC‐2 and GRIP1) have been linked to the development of acute leukemias, of which the most extensively characterized is the MOZ–TIF2 fusion resulting from inversion of chromosome 8 at inv(8) (p11q13). MOZ is a member of the MYST family of HATs and is involved in proper development of hematopoietic progenitors.32 Fusion proteins resulting

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from chromosomal translocation of the MOZ gene with CBP/p300 and inversion with TIF2 are associated with M4/M5 subtypes of acute myeloid leukemia (AML) in humans.33 When retrovirally expressed, MOZ–TIF2 immortalizes myeloid progenitors in vitro and recapitulates AML in bone marrow transplant assays.34 Similar approaches were used to demonstrate that MOZ–TIF2 but not BCR–ABL oncogene induces self‐renewal capacity in committed myeloid progenitors.35 As a result of the inversion, the fusion product retains the conserved MYST and HAT domains of MOZ and the CBP‐interaction (AD1) and activation 2 domains (AD2) of TIF2 (Fig. 2).36,37

Exon 16 1117

Exon 13 869

MYST Zn

PHD MOZ – TIF2 inv(8)(p11q13)

1

209

310

HAT

522

AD1

Acidic 703

1057 1121

Exon 16 1117

Exon 13 918

Acidic

AD1

AD2 1305

1464

MYST PHD MOZ – ACTR/NCOA3 t(8;20)(p11q13)

1

209

Exon 4 154

Zn

310

HAT

522

703

1

AD2

1057 1121

1305

Exon 6 338

Paired Homeodomain Box PAX3 – SRC-1 t(2;2)(q35;p23)

1

1412

Exon 15 1010

AD1 ETV6 – TIF2 t(8;12)(q13;p13)

1017

AD2

1464

Exon 13 843

AD1 920 977

AD2 1172

1441

FIG. 2. p160/SRC protein fusions derived from chromosomal rearrangements. Schematic of chimeric proteins resulting from the chromosomal rearrangements indicated on left. The break point exon and amino acid residue for each fusion partner are indicated by the forked lines. Conserved structural and functional domains are labeled above each diagram. Numbers below each diagram represent amino acid residues. PHD, plant homeodomain‐linked zinc fingers; MYST, MOZ, Ybf2/Sas3, Sas2, and Tip60 domain; Zn, C2HC zinc finger.

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Notably, reporter gene analyses show that MOZ–TIF2 displays a dominant‐ negative effect on transcription by nuclear hormone receptors and other CBP‐ dependent activators such as p53.38 Further examination using deletion mutants to probe the structure–function of MOZ–TIF2 showed that both the dominant‐negative effect and the transforming capacity of the fusion protein depend on a nucleosome recognition motif in MOZ and the CBP‐interaction domain of TIF2, while interaction with CBP involves both the MOZ and TIF2 portions.34,38 In an inducible cell line, chromatin immunoprecipitation demonstrated that expression of MOZ–TIF2 alters RAR coregulator recruitment and histone modification at the RARbeta2 promoter. Specifically, MOZ–TIF2 increased promoter histone acetylation and CBP recruitment to unliganded RAR corepressor complex, and decreased CBP recruitment in the presence of ligand without affecting histone acetylation.39 Globally, MOZ–TIF2 expression correlated with depletion of cellular CBP levels and dissociation of CBP from subnuclear PML bodies, an effect that required the TIF2 CBP-interaction domain.38 These studies suggest that global and promoter‐specific perturbation of CBP localization and recruitment to chromatin by MOZ–TIF2 may represent an aberrant process through which transcriptional programs are epigenetically disrupted to subvert normal differentiation and thus promote transformation of hemataopoietic progenitor cells. More recently, a novel ETV6–TIF2 fusion gene resulting from t(8;12)(q13; p13) translocation was identified in six cases of childhood acute lymphoblastic leukemia (ALL) involving both T‐lymphoid and myeloid lineages. The fusion protein retains the PNT protein dimerization domain of ETV6 and the CBP‐ interaction and AD2 domains of TIF2 (Fig. 2).40 ETV6 is a repressor of the Ets family of transcription factors and has been shown to possess tumor suppressor properties in vitro. ETV6 is a common target of chromosomal rearrangements to form fusion proteins with more than 20 partners in diverse types of leukemia.41 Transformative properties attributed to ETV6–TIF2 have not yet been demonstrated, although presence of the fusion correlated with frequent heterozygous activating mutations in NOTCH1 gene (four out of six), which had previously been linked to T‐cell ALL and rare cases of AML.40 It was speculated that the ETV6 translocation and NOTCH1 mutation represent two genetic events leading to progenitor transformation and specific expansion of T/myeloid lineages, although further studies utilizing ectopic expression of ETV6–TIF2 will be needed to test this model. It is likely that cancer‐associated fusion proteins resulting from genetic rearrangements with other p160 coactivators will be discovered as gene expression signatures that are identified to distinguish subtypes of malignant disease. Gene expression profiling to classify subtypes of pediatric rhabdomyosarcoma led to the discovery of a PAX3–SRC‐1 fusion resulting from t(2;2)(q35;p23) translocation, composed of paired box and homeodomain DNA-binding

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domains from PAX3 and the CBP‐interaction and AD2 domains of SRC‐1 (Fig. 2). Functional analysis of the fusion demonstrated that, like other PAX3 (7) translocation fusions associated with the alveolar subgroup of rhabdomyosarcoma, the PAX3–SRC‐1 is a more potent transcriptional activator than wild‐ type PAX3.42 Recently, a single case of M5 type AML revealed AIB1/ACTR to be another fusion partner for MOZ resulting from t(8;20)(p11;q13) translocation to produce MOZ–AIB1/ACTR chimeric protein (Fig. 2).43 Finally, it is noteworthy that only the MOZ–TIF2, ETV6–TIF2, MOZ– AIB1/ACTR, and PAX3–SRC‐1 fusion proteins, but not the reciprocal fusions stemming from each chromosome rearrangement, have been detected in malignant diseases (Fig. 2 and Table I). The conservation of the CBP‐interaction and AD2 domains in all of the relevant p160/SRC gene fusion products also suggest that the fusion proteins use a similar or common mechanism to promote oncogenesis by targeting the transcriptional machinery.

B. Polymorphisms 1. SINGLE NUCLEOTIDE POLYMORPHISMS (SNPS) A number of population genotyping studies have revealed that SNPs or variable number of tandem repeats (VNTRs) in AIB1 may be associated with risk for cancers in various hormone‐responsive tissues. For example, SNPs in the AIB1/ACTR gene resulting in Q586H (1758G > C) and T960T (2880A > G), where > designates the more frequent variant, have been found to be a protective factor against breast cancer when cohorts of healthy women and women with familial breast cancer were compared in a population of German and Polish nationals.13 In both populations, genotypes containing the rare variant for each SNP were more frequent in the control groups compared with the diseased cases. The two rare variants were not linked to each other, but haplotype analysis showed that having the rare protective variant in just one SNP site was significantly more frequent in the controls than in diseased cases when compared to having the common variant in both SNP sites. Due to the lack of functional studies, it is unclear how the rare SNP variants might influence breast cancer risk, although the authors speculate that Q586H could alter protein structure and that the silent T960T polymorphism may affect posttranscriptional regulation through altered codon usage. A more recent study identified SNPs in the p160 coactivator genes from a multiethnic panel of women with advanced breast cancer. In subsequent allelic frequency analyses with a larger multiethnic cohort (1612 cases and 1961 controls), all but one of the identified SNPs (AIB1/ACTR M391V) were found across the different populations. Notably, a A407S SNP in TIF2 showed significant positive correlation with breast cancer (OR 2.25) and advanced disease stage (OR 4.21 in a subgroup of 420 cases).14 Taken together, these

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two studies suggest that the link between a particular coactivator SNP and breast cancer risk can vary depending on the ethnic heterogeneity of the observed group, and that other genetic or environmental agents may affect the risk factor associated with a SNP for a particular ethnic group. 2. VARIABLE NUMBER OF POLYGLUTAMINE REPEATS The potential association of polyglutamine repeat number in AIB1/ACTR gene with breast cancer risk has also been explored. The polymorphic stretch of glutamines lies between residues 1244 and 1272 in the carboxy‐terminal region of AIB1/ACTR and is encoded by a variable number of CAG repeats (Fig. 1).44 The first correlation between longer polyQ tract and breast cancer risk was found in a case–control study involving 448 women (278 cases and 170 controls) with germ‐line mutations in BRCA1/2 genes. Women carrying alleles with more than 28 CAG repeats had significantly higher breast cancer risk compared to women with fewer repeats (OR 1.96), and those at highest risk additionally had either never given birth or had given birth over the age of 30 years (OR 4.62–6.97).45 However, in a subsequent larger study of unselected 1175 BRCA1/2 mutation carriers, no correlation between polyQ length and breast cancer risk was observed,46 refuting results from an earlier and smaller study of 311 unselected BRCA1/2 mutation carriers.47 Another large, case– control study that compared women with familial breast cancer (n ¼ 591) with healthy women (n ¼ 536) demonstrated no effect of polyQ length in AIB1/ ACTR on breast cancer risk, even in the absence of BRCA1/2 mutations.48 Most recently, a retrospective cohort study of women with BRCA1 mutation (176 cases and 140 controls) showed a correlation of breast cancer risk with smoking history and AIB1 polyQ length. Specifically, the authors found significant and additive protective effects from having positive smoking history and having at least one polyQ allele of 28 repeats,49 the most common low‐risk allele corroborated by this and other earlier studies.45,47,48 Confounding further the potential role of AIB1/ACTR polyQ length in breast cancer risk, studies examining the impact of shorter than average polyQ alleles have consistently demonstrated an inverse correlation between repeat length and risk for other types of cancer. In a case–control study of Chinese men (189 cases and 301 controls), those homozygous for an AIB1/ACTR allele with less than 29 repeats were at significantly higher risk for prostate cancer compared to those homozygous for 29 repeats (the most common length).50 In a small study of ovarian cancer patients (n ¼ 89), those homozygous for having less than 29 repeats in each allele had significantly lower DFS and shorter time to disease recurrence.51 Another study looking at both repeat length and sequence found that having at least one allele with less than 27 repeats was significantly associated with 32 sporadic breast cancer cases when compared to

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16 familial breast cancer cases with BRCA1/2 mutation or 43 controls. Additionally, sequence comparison between the groups showed higher sequence variability indicative of somatic instability in the cases compared to controls.52 Given the discrepancies among the studies, it remains unclear whether the VNTR in the AIB1/ACTR polyQ tract impacts breast cancer risk and, if it does, how this variation interacts with other genetic or environmental risk factors such as BRCA1/2 mutation. The studies do, however, point out the need for molecular examination of polyQ length on AIB1/ACTR functions such as transactivation or protein–protein interactions. Progress in this area could reveal pathways and targets that are impacted by the AIB1/ACTR polyQ repeat variation in cancer. Shorter polyQ length in AR has been shown to enhance its transcriptional activity and ligand‐dependent interaction with p160/SRC coactivators in prostate cancer cells.53 It is possible that polyQ length in AIB1/ ACTR modulates its association with other factors implicated in AIB1/ACTR‐ mediated tumorigenesis such as ER, AR, and E2F transcription factors (as described in Section III). In support of this hypothesis, the analogous polyQ region of SRC‐1 interacts directly with AR and is required to enhance its signaling.54

C. Aberrant Expression of SRCs in Cancers and the Potential Underlying Mechanisms 1. OVEREXPRESSION IN CANCERS AND THE DISEASE IMPLICATIONS High levels of SRC‐1, TIF2, or AIB1/ACTR expression have been correlated with clinical parameters in breast and prostate cancers such as higher recurrence rate after hormone deprivation therapies and lower disease‐free survival.29,55,56 In prostate tumors, increased SRC‐1 or AIB1/ACTR protein expression correlates with more aggressive pathology including high grade, advanced stage, and lymph node metastasis.26,28,30 High level expression of AIB1/ACTR was also found to correlate with biochemical markers of increased prostate cancer cell proliferation and survival, as well as PSA recurrence and reduced DFS.28–30 High SRC‐1 and TIF2 expression have also been found to correlate with PSA recurrence after androgen ablation therapy.31 Although the p160 coactivators are expressed in both AR‐positive and AR‐negative prostate tumors and xenografts,27,56 any prognostic significance associated with their coexpression or lack thereof has not yet been established. In breast cancer, elevated expression of AIB1/ACTR was frequently correlated with high tumor grade.15,22 Immunohistochemical analysis showed that AIB1/ACTR overexpression correlated with absence of ER or progesterone receptor (PR) expression in invasive breast tumor specimens.15 Although an earlier report documented a correlation with ER and PR positivity, gene amplification was the only parameter analyzed in a relatively small‐scale study.57

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Similarly, no association between SRC‐1 and ER expression was observed in another clinical study using breast tumor samples.55 Notably, both SRC‐1 and AIB1/ACTR were found to be frequently coexpressed with the growth factor receptor HER2/Neu, and their overexpression in HER2‐positive tumors was associated with poor outcome in response to adjuvant tamoxifen therapy.15,55,58 Interestingly, AIB1/ACTR gene amplification or overexpression had no unilateral effect on DFS58–60 whereas SRC‐1 significantly associated with lower DFS.55,61 Thus, resistance to tamoxifen therapy in AIB1/ACTR overexpressing breast tumors may be specific to the subset that coexpresses HER2. More recently, the scope of partnership between AIB1/ACTR and the epidermal growth factor receptor family as a prognostic indicator for tamoxifen resistance has broadened to include HER1–3.59 Finally, it is worthwhile to note that AIB1/ACTR overexpression has been observed in a wide spectrum of human cancers—many of which are not expected to be hormone‐responsive. These include bladder, colorectal, gastric, pancreatic, liver, and endometrial cancers.16–18,23,62,63 AIB1/ACTR overexpression also correlated with poor patient survival in subsets of bladder, colorectal, and endometrial cancer patients, in addition to high grade or disease stage (see Section II.A). Collectively, these studies established p160 coactivators as critical mediators of tumor progression and pathology at different stages of disease. Furthermore, the above studies imply that this tumor‐promotion function of SRCs does not always involve the coactivators’ role in hormone receptor activity and hormone‐regulated gene expression. 2. TRANSCRIPTIONAL DEREGULATION OF AIB1/ACTR GENE Recent studies have identified potential mechanisms that control AIB1/ ACTR expression at the transcriptional and posttranscriptional levels, which may be deregulated in cancer.64–68 Although the exact contribution of transcriptional versus posttranscriptional pathways to AIB1/ACTR gene deregulation in cancer is unclear at this point, these studies nonetheless indicate that both mechanisms are likely involved in mediating changes of AIB1/ACTR expression depending on cellular context and extracellular signals. An early study with MCF‐7 breast cancer cells showed that endogenous AIB1/ACTR expression is primarily regulated at the transcriptional level in response to ER ligands.69 Whereas hormone depletion increased AIB1/ACTR RNA and protein levels, subsequent estrogen treatment suppressed the expression back to basal levels. Antiestrogens such as tamoxifen and all‐trans retinoic acid (atRA), on the other hand, had the opposite effect and induced AIB1/ACTR expression. Cotreatment with transcriptional and protein synthesis inhibitors confirmed that AIB1/ACTR upregulation occurred primarily at the transcriptional level and demonstrated that new protein synthesis was required for part of the induction. Although the secondary protein that is

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synthesized in response to the antiestrogens was not clearly identified, both treatment and antibody blocking experiments suggested that the growth factor TGF‐beta contributes to the AIB1/ACTR induction response. It is known that the AIB1/ACTR gene is amplified and overexpressed in MCF‐7 cells compared to other breast cancer cell lines,70 and it should be noted that no changes in cell cycle indicative of an antiproliferative effect from the antiestrogen concentrations used in this study were observed.69 Thus, it is unclear whether the negative feedback mechanism of hormones on AIB1/ACTR expression is relevant to the regulation of AIB1/ACTR in a normal physiological context. Nonetheless, in conjunction with the finding that AIB1/ACTR is involved in agonist but not antagonist‐mediated ER degradation,71 it is tempting to speculate that accompanying fluctuations in AIB1/ACTR levels may influence the extent of ER signaling attenuation in response to steroidal stimulation and antiestrogen therapy. In contrast to the effect observed with estrogen, a recent study showed that the androgen dihydroxytestosterone (DHT) upregulated AIB1/ACTR expression in LNCaP prostate cancer cells. Ectopically overexpressed AR enhanced DHT induction of AIB1/ACTR mRNA, which was observed by 4 h after DHT treatment. The findings suggest that androgens can directly regulate the AIB1/ACTR gene to potentiate AR signaling through a positive feedback mechanism.72 Two independent studies identifying the AIB1/ACTR promoter determined that it is directly regulated by E2F transcription factors through an autoregulatory loop.64,65 AIB1/ACTR had been previously identified as a coactivator for E2F73 and is required in both normal and malignant human cells for cell cycle progression from G1 to S phase, and for the expression of E2F target genes involved in the transition including cyclin E, cdk2, and E2F1. Remarkably, AIB1/ACTR RNA and protein levels followed a similar cell cycle dependent expression pattern, peaking at G1/S, indicating that it is also an E2F target gene.64 Truncation reporter gene analysis with genomic DNA fragments identified a minimal 0.6 kb E2F‐responsive region encompassing the first exon and promoter of AIB1/ACTR.64,65 Furthermore, ChIP analysis showed that dynamic recruitment of both E2F1 and AIB1/ACTR to the promoters of AIB1/ACTR and other E2F target genes followed their pattern of expression during cell cycle progression.64 These findings strongly support a general model for regulation of AIB1/ACTR expression that is primarily determined at the transcriptional level and is cell cycle dependent. Furthermore, it is tempting to speculate that such a mechanism might be deregulated in cancer cells with hyperactive E2F or AIB1/ACTR, and that the positive regulatory feedback loop between the two might exacerbate AIB1/ACTR overexpression. Much work remains in terms of elucidating the full range of conditions and factors that regulate AIB1/ACTR at the transcriptional level, and determining how perturbations in these mechanisms might lead to aberrant AIB1/ACTR

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expression. Because E2Fs represent the hub through which all growth promoting signals converge, further examination of the cis and trans factors underlying E2F‐mediated regulation of the AIB1/ACTR promoter could reveal insights as to how AIB1/ACTR expression might be deregulated in different cell types.

III. The p160/SRCs Functions and Their Action Mechanisms in Cancer Cells The relevance of p160 coactivators in malignant disease prognosis, particularly in patient(s) response to antihormone therapies for breast and prostate cancer, has spurred much progress in elucidating the role that these coactivators play in cancer cell behavior. Importantly, more recent studies underscore the important functions that these coactivators may have in terms of controlling cancer cell proliferation, survival, invasion, and metastasis, as well as tumor resistance to therapeutics.

A. Cell Proliferation, Survival, Invasion, and Metastasis 1. HORMONE‐DEPENDENT AND ‐INDEPENDENT PROLIFERATION OF PROSTATE AND BREAST CANCER CELLS: P160 COACTIVATORS AS MEDIATORS OF ETS, STAT, AND E2F TRANSCRIPTIONAL PROGRAMS Since the p160/SRCs were initially characterized as coactivators for nuclear hormone receptors including ER and AR, initial studies on their potential function in cancer cell proliferation were focused on their role in hormone‐ responsive or receptor‐positive cancer cells and tumors. For instance, an early study using ribozyme targeting AIB1/ACTR demonstrated that the high level of AIB1/ACTR in MCF7 cells is required for estrogen‐stimulated proliferation.74 RNAi‐mediated knocking down of SRC‐1 demonstrated that SRC‐1 is important for AR target gene expression and for the proliferation of AR positive androgen‐dependent LNCaP and androgen‐independent but AR‐positive C4–2 prostate cancer cell lines, but not for the proliferation of AR negative PC‐3 and DU145 prostate cancer cells.26 Likewise, in another study, specific depletion of AIB1/ACTR strongly inhibited proliferation of LNCaP and AR‐ positive but androgen‐independent C4–2B cells as well as the androgen‐ stimulated growth of CWR22 xenograft tumors.75 However, in subsequent studies where both receptor‐positive and ‐negative cancer cells were rigorously examined, it was found that AIB1/ACTR and TIF2 are required for the proliferation and survival of not only the receptor‐positive and hormone‐ stimulated cancer cells but also the ER‐ or AR‐negative cancer cells.29,64,73,76 These later findings strongly suggest that the function of SRCs in cancer is not limited to serving as coactivators for ER or AR.

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Well characterized ER target genes involved in control of cell proliferation, such as cyclin D1 and c‐myc, are directly regulated by p160 coactivators as assessed by ChIP analysis of estrogen‐stimulated breast cancer cells.77–79 The chemokine SDF‐1 is a more recently identified ER target gene that is required for estrogen‐induced cell proliferation through binding and activating its cognate receptor CXCR4 in ER‐alpha positive breast and ovarian cancer cells.80 Subsequent study using MCF‐7 breast cancer sublines expressing variable levels of SRC‐1 suggests that this coactivator is involved in both the basal and estrogen‐inducible expression of SDF‐1, as well as estrogen‐stimulated proliferation. Limited evidence from this study using ectopic expression showed that TIF2 and AIB1/ACTR can similarly modulate SDF‐1 basal expression,81 although AIB1/ACTR depletion specifically blocked estrogen‐stimulated cyclin D1 expression and proliferation of MCF‐7 cells in a recent study.82 In neither study did depletion of AIB1/ACTR (or other p160 coactivators) affect estrogen induction of c‐Myc, suggesting some functional redundancy in the coactivator’s roles in mediating expression of subsets of ER target genes. Independently of ER, the p160 coactivators have been implicated in regulating the expression of target genes in response to growth factor signaling to promote cell proliferation. The c‐Myc gene is a direct target of Ets‐2 and STAT3 transcription factors which activate gene expression in response to MAPK and leptin signaling pathways, respectively. RNAi depletion analysis showed that Ets2‐mediated transactivation of c‐Myc reporter gene and induction of endogenous c‐Myc in hormone resistant/insensitive breast cancer cells were dependent on SRC‐1 and AIB1/ACTR expression.61 Depletion of AIB1/ ACTR but not SRC‐1 also resulted in lower Ets‐2 transcription factor protein levels, suggesting that the coactivators may differentially mediate their effects on c‐myc expression by regulating Ets2 activity and/or expression. ChIP and reporter gene analysis demonstrated that SRC‐1 is specifically recruited to and transactivates c‐myc promoter in response to leptin treatment of MCF‐7 cells. In support of a critical role for SRC‐1 in leptin‐stimulated proliferation through STAT3, SRC‐1 directly interacts with STAT3 in vitro and in vivo, and specific inhibition of STAT3 signaling or depletion of SRC‐1 blocked leptin‐induced proliferation.83 Because Ets2 and STAT3 regulate a number of genes critically involved in cell growth and proliferation,84,85 these studies suggest distinct and combinatorial functions for p160 coactivators in mediating response of diverse gene regulatory networks to various nonsteroid hormonal signaling pathways. In an attempt to determine the molecular mechanism by which AIB1/ ACTR functions to promote hormone receptor independent cancer cell proliferation, our laboratory has focused on the role of AIB1/ACTR in controlling expression of genes involved in cell cycle progression. Using RNAi‐mediated depletion of AIB1/ACTR in hormone dependent and independent breast and prostate cancer cell lines, we showed that AIB1/ACTR is required for cell cycle

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progression from G1 to S phase, and for the expression of E2F target genes involved in the transition including cyclins, cdks, and E2F1.64,73,75 Substantiating the importance of AIB1/ACTR involvement in E2F function, mutational analysis of AIB1/ACTR structure and function showed that hormone‐ independent proliferation did not require association with ER, but depended instead on its direct interaction with E2F1.73 By acting as an E2F coactivator, AIB1/ACTR would be predicted to aberrantly enhance cell proliferation when overexpressed, a condition that could drive tumorigenesis and tumor progression. In agreement with this hypothesis, deletion of the E2F interaction domain in AIB1/ACTR abolished its transformative properties when ectopically overexpressed in HMECs (Fig. 1).64 In summary, these studies indicate that AIB1/ ACTR predominantly acts through cell cycle regulator E2Fs to promote cell proliferation regardless of ER status in breast cancer cells and androgen dependence in prostate cancer cells. 2. IGF/AKT SIGNALING AND PATHWAY COMPONENTS—DIRECT AND INDIRECT TARGETS FOR AIB1/ACTR‐MEDIATED SURVIVAL One of the major protein kinase signaling pathways modulated by AIB1/ ACTR appears to be the IGF‐1/PI3K/Akt pathway. IGF stimulates cellular mitogenesis, transformation, and differentiation, and inhibits apoptosis or anoikis through signaling by the IGF‐1 receptor, leading to sequential activation of intracellular substrates or kinase proteins such as IRS‐1 and IRS‐2, PI3‐kinase, and Akt. SRC‐3 deficiency in otherwise normal mice resulted in dwarfism and decrease of IGF‐1 gene expression.86,87 In line with the findings from these knockout (KO) studies, IGF‐1 mRNA levels in the mammary epithelial cells and serum were significantly elevated in the MMTV–SRC‐3 transgenic (Tg) mice.88 Markedly elevated activation of IGF‐1 receptor, Akt, GSK3, and mTOR as well as the downstream effectors eIF4G and p70S6K were detected in cell cultures derived from the AIB1/ACTR Tg compared to wild‐type mammary glands from 10‐week-old mice by Western blot analysis with antibodies for phosphorylated forms of the proteins (Fig. 3).88 In the MMTV‐ras Tg model, SRC‐3 deficiency dampened the ras‐induced expression of IRS‐1 and IRS‐2 in mammary glands and tumors, as well as IGF‐1 stimulated mitogenic response in the mammary tumor cells, supporting the notion that AIB1/ACTR/SRC‐3 plays a major role in activated ras‐induced mammary tumorigenesis by mediating elevated IGF‐1 signaling.89 A critical role of AIB1/ACTR in mediating IGF‐1 signaling has also been demonstrated in human cancer cell lines. AIB1/ACTR depletion in MCF‐7 breast cancer cells abrogated IGF‐1 stimulated colony formation by increasing apoptosis under anchorage‐independent conditions,76 while knocking down of AIB1/ACTR expression was sufficient to induce apoptosis in adherent LNCaP and PC3 prostate cancer cells.29 These effects were notably accompanied by

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IGF-1

P-

IGF1R

IGF-2

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SGK

P- p70 S6K P-

P- mTOR

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FoxO Tx factors FIG. 3. IGF‐1/PI3K/Akt pathway components regulated by AIB1/ACTR/SRC‐3. Diagram of the IGF‐1 receptor signaling pathway is shown with arrowhead lines indicating direction of sequential activation for downstream intracellular substrates, adapted from Ref. 90. Hammer head lines indicate inhibitory activities. Components whose expression is modulated by AIB1/ ACTR/SRC‐3 in various experimental systems are designated in rectangular boxes with fill pattern indicating the observed changes: protein (black), RNA (gray), both protein and RNA (unfilled). Components whose protein expression is unaltered by elevated AIB1/ACTR/SRC388 are designated in unfilled ovals. Components marked with a P‐attachment indicate their elevated phosphorylation associated with AIB1/ACTR expression.88 Double dashed horizontal line represents extracellular membrane, and the arc depicts the nucleus.

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decreases in protein levels of IGF/Akt pathway components including IGF‐1, IGF‐1 receptor, IRS‐1, IRS2, and the downstream antiapoptotic gene Bcl2, as well as total or phosphorylated/activated Akt.76,90 The molecular mechanisms by which AIB1/ACTR regulates the level of IGF/Akt pathway components appear to be complex. The involvement of AIB1/ACTR in direct transcriptional regulation of their expression has been established in models of prostate cancer.90 Upregulation of expression for these components was observed at both mRNA and protein levels in AIB1/ ACTR overexpressing LNCaP cells. Concordantly, their expression was reduced in the prostate of AIB1/ACTR knockout (KO) mice or siRNA‐AIB1/ ACTR treated PC‐3 cells compared to controls (Fig. 3, white boxes). ChIP assay confirmed at least some of the components are direct target genes of AP‐1 and AIB1/ACTR, and transactivation of IRS‐2 promoter through AP‐1 was synergistically enhanced by AIB1/ACTR. Notably, depletion of c‐Fos showed that AP‐1 is required for AIB1/ACTR recruitment to the target gene promoters, suggesting that AIB1/ACTR and AP‐1 coordinately regulate expression of IGF/Akt signaling components at the transcriptional level in prostate cancer.90 In contrast, the mechanism through which AIB1/ACTR controls expression of IGF/Akt components in breast cancer cells is less clear. Except for IGF‐1, no data have been reported on whether AIB1/ACTR controls the transcription of IGF‐1 receptors, IRS, and the other downstream signaling genes. Intriguingly, a novel mechanism by which AIB1/ACTR indirectly controls the expression of IGF‐1 was recently identified using the SRC‐3 KO mouse model.91 It is well known that IGF‐I in circulation associates with IGF‐binding proteins such as IGFBP‐3, which stabilizes IGF‐1 and enhances its cellular signaling. Remarkably, SRC‐3 KO mice exhibit lower serum levels of both IGF‐1 and IGFBP‐3. Further analysis revealed that the transcription of IGFBP‐3, but not IGF‐1 was impaired in the cells of SRC‐3 KO mice. Ligand‐induced transactivation of IGFBP‐3 gene through vitamin‐D receptor was significantly attenuated in KO compared to wild‐type MEFs.91 Thus, at least in the animal model, SRC‐3 may function to mediate IGF‐1 signaling by controlling expression of a factor(s) that maintain(s) circulatory levels of the growth factor. 3. INVASION AND METASTASIS MEDIATED BY AIB1/ACTR, AP‐1, AND PEA3 High levels of SRC‐1 and AIB1/ACTR expression have been shown to promote the invasive capabilities of breast cancer cells in vitro, and several studies suggest that the coactivators, particularly AIB1/ACTR, function in regulating a subset of invasion and metastasis‐associated genes.81,92,93 Matrix metalloproteinases (MMPs) participate in the invasion process by degrading extracellular matrix (ECM) during cancer metastasis. In one study addressing

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its possible involvement in tumor metastasis, AIB1/ACTR knockdown strongly inhibited the invasiveness of breast cancer cells (MDA‐MB‐231 or T‐47D) while its ectopic expression decreased their adhesion to ECMs such as fibronectin and collagen IV and promoted their invasion in the Matrigel assay. Further molecular analysis indicates that AIB1/ACTR functions as a coactivator of AP‐1 to mediate the expression of specific members of MMPs such as MMP‐7 and MMP‐10.93 This finding is consistent with the notion that MMP‐7 is one of the few MMPs that are synthesized and secreted primarily by the tumor cells and play key roles in major aspects of cancer metastasis. In another study, cell lines derived from mammary tumors of SRC‐3 KO mice carrying MMTV‐driven PyMT transgene were significantly less motile and invasive, and expressed lower MMP‐2 and MMP‐9 levels than did their wild‐type/PyMT counterparts.92 AIB1/ACTR/SRC‐3 knockdown and rescue experiments using mouse and human mammary tumor cells demonstrated that AIB1/ACTR directly controls the MMP expression. Reporter gene assay and regular coimmunoprecipitation and ChIP experiments detected the involvement of AIB1/ ACTR–PEA3 protein complex in control of the MMP transcription in the cancer cells. Importantly, IHC analysis of over 500 human breast tumors indicated a statistically significant association between the elevated levels of AIB1/ACTR, PEA3, and the MMPs,92 further supporting the notion that AIB1/ ACTR promotes PEA3‐mediated expression of a subset of MMPs in human breast cancer cells. Recently, a study dissected the role of AIB1/ACTR in prostate cancer cell migration and invasion.30 It was found that AIB1/ACTR depletion decreased cell motility by attenuating FAK signaling and a significant correlation was observed between levels of AIB1/ACTR and the phosphorylated/activated form of FAK in a cohort of prostate tumors. AIB1/ACTR depletion additionally decreased prostate cancer cell invasion and expression of MMP‐2 and MMP‐13, while AIB1/ACTR overexpression had the opposite effects. ChIP and reporter gene assay demonstrated that AIB1/ACTR directly regulates MMP‐2 and MMP‐13 expression, while binding site mutation analysis of the MMP‐13 promoter revealed that AIB1/ACTR recruitment and transactivation involves both AP‐1 and PEA3. Clinically, higher levels of AIB1/ACTR transcript were observed in prostate cancer specimens from patients positive for seminal vesicle invasion, lymph node metastasis, and PSA recurrence compared to patients without these clinicopathogical parameters.30 These studies collectively suggest that AIB1/ACTR promotes metastatic properties in breast and prostate cancer cells at least in part by regulating MMP family genes that are involved in invasion, although the mechanism by which AIB1/ACTR is involved in FAK‐mediated cell motility is less well understood.

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B. Tamoxifen Resistance in Breast Cancer Tamoxifen displays partial ER agonist or antagonist activity depending on tissue type. Cellular factors that influence the agonistic versus antagonistic properties of tamoxifen and other SERMs are beginning to be understood. Studies on nuclear receptor coregulators suggest that one mechanism for the tissue‐specific effect of tamoxifen is through selective recruitment of coactivator or corepressor complexes on promoters of ER target genes. It was demonstrated that in Ishikawa endometrial cancer cells, but not in MCF‐7 breast cancer cells, that high levels of SRC‐1 are important for tamoxifen‐stimulated ER target gene expression and cell proliferation.94 However, the mechanisms underlying the development of tamoxifen resistance in breast cancer appear to be far more complex than a simple balance of relative levels of individual coregulators. Nevertheless, recent studies clearly indicate important roles played by the aberrant SRCs.

1. AIB1/ACTR AND HER2 OVEREXPRESSION PROMOTE TAMOXIFEN RESISTANCE In vitro studies in breast cancer cells have provided some insight to the mechanism by which AIB1/ACTR cross talk with HER2 disrupts tamoxifen‐ mediated inhibition of cell proliferation. In HER2 positive endocrine resistant breast cancer cells, siRNA‐mediated disruption of AIB1/ACTR expression was sufficient to switch tamoxifen from partial agonist to antagonist, as displayed by changes in cell proliferation and the induction of ER target gene pS2.95,96 On the other hand, stable expression of HER2 was sufficient to switch tamoxifen to having agonist effect in MCF‐7 cells (in which AIB1/ACTR is overexpressed), as indicated by increased cell proliferation, tumor growth, and coactivator recruitment to and induction of ER target genes. Notably, this finding was accompanied by phosphorylation of ER, AIB1/ACTR, HER2, and HER downstream effectors such as MAPK and Akt. The in vitro and in vivo agonist effects of tamoxifen in these HER2/AIB1/ACTR overexpressing cells was dependent on growth factor receptor activity—that is, they were abolished by the inhibitor gefitinib—indicating that ER and AIB1/ACTR are phosphorylated through growth factor signaling that is activated by tamoxifen via nongenomic activities.97 Together, these studies suggest that nongenomic activation of ER by bidirectional ER/HER2 cross talk depends on p160 coactivator expression, and represents an important mechanism for bypassing antagonist effect of tamoxifen through recruitment and sustained activation of alternative kinase‐ mediated proliferation pathways. Furthermore, they indicate that inhibiting HER family member activation could enhance antiproliferative effects of

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tamoxifen in HER2/AIB1/ACTR overexpressing breast cancer cells. Indeed, a recent study showed improved sensitivity to tamoxifen in BT474 cells that were initially treated with the HER kinase inhibitor Herceptin.98 2. AIB1/ACTR COOPERATES WITH ETS FAMILY MEMBERS TO POTENTIATE HER2 SIGNALING Members of the Ets family of MAPK‐dependent transcription factors are known downstream targets of HER2 signaling, and have been implicated to function with p160 coactivators to promote tamoxifen resistance. All three p160 coactivators are capable of enhancing transactivation of Ets2 and members of the PEA3 subfamily including ER81.61,99 Coimmunoprecipitation and electrophoretic mobility shift assay using extracts from ER negative SKBR3 or primary breast cancer cultures show that growth factor stimulation enhances coactivator interaction with Ets2 and enhances their recruitment to Ets response element.100 Clinically, Ets2 and PEA3 expression correlated with that of SRC‐1,55,61 while coexpression of SRC‐1 and Ets2 or PEA3 significantly correlated with decreased DFS in HER2 positive patients receiving adjuvant therapy.55,100 These studies suggest that, in addition to ER/HER2 cross talk, p160 coactivators can potentiate HER2 signaling selectively by enhancing the activity of downstream Ets family transcription factors. 3. AIB1/ACTR ACTIVATES E2F‐REGULATED CELL PROLIFERATIVE PROGRAM IN THE PRESENCE OF TAMOXIFEN Aside from involvement in growth factor‐mediated ER or HER2 activation, studies from our laboratory reveal that AIB1/ACTR overexpression may disrupt antagonist response to tamoxifen by directly potentiating E2F transcription factor‐mediated expression of cell cycle regulator genes.73 The AIB1/ACTR‐ E2F pathway may represent an alternative route toward cell proliferation in breast cancer cells with disrupted ER signaling as in the case of antiestrogen treatment. In ER positive breast cancer cells, ectopic AIB1/ACTR overexpression was sufficient to confer tamoxifen resistance and promote cell proliferation with concomitant upregulation of E2F target genes involved in cell cycle progression including cyclin A, cyclin E, cdk2, and E2F1. In agreement with a hormone‐ independent pathway, mutational analysis of AIB1/ACTR structure and function indicated that the effect on cell proliferation does not require association with ER, but depends instead on direct interaction with E2F1 (Fig. 1).73 4. AN AIB1/ACTR ISOFORM MAY PROMOTE AGONIST ACTIVITY OF TAMOXIFEN Reiter et al. identified an N‐terminally truncated AIB1/ACTR isoform that lacks the bHLH and part of the PAS domain and comprises up to 5% of total expressed AIB1/ACTR mRNA in breast cancer and human mammary

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epithelial cells.101 In a small panel of tissue specimens, the relative expression of this AIB1/ACTR isoform was significantly higher in tumors compared to normal samples. The isoform was more efficient than the full-length AIB1/ACTR in enhancing ligand‐dependent transactivation of ER‐alpha and PR‐beta, as well as potentiating EGF‐induced transcription.101 Its ectopic expression in mice driven by the human cytomegalovirus immediate early gene 1 (hCMVIE1) promoter increased proliferation of mammary epithelial cells but was insufficient to induce tumor formation in the transgenic mice.102 However, results from reporter gene assays suggest that the isoform can more effectively enhance the agonist effect of tamoxifen in breast and endometrial cancer cells.103 Further studies will be needed to address the functional significance of the AIB1/ACTR isoform in tamoxifen resistance of human breast cancer and the underlying mechanism. 5. SRC‐1 IS INVOLVED IN AGONISTIC ACTIVITY OF TAMOXIFEN A recent study examining the mechanism of ER‐alpha and SRC‐1 interaction in tamoxifen agonist activity suggests that functional interaction between the two in the presence of tamoxifen is dependent on protein kinase A (PKA)‐ mediated phosphorylation of serine‐305 in the hinge region of ER‐alpha.104 The modification triggers the orientation of tamoxifen‐bound ER‐alpha C‐terminus toward SRC‐1, which renders the receptor–coactivator complex competent for transcriptional activation. While the selective nuclear receptor coactivator PGC‐1beta has also been shown to enhance the agonist activity of tamoxifen,105 a more recent study demonstrated that the molecular basis for this function involves cooperative interaction with SRC‐1.106 Structure– function analysis using in vitro binding and yeast‐two hybrid approaches revealed that PGC‐1beta directly binds to and synergizes with SRC‐1 to efficiently coactivate tamoxifen‐bound ER‐alpha. Studies with breast cancer specimens found that SRC‐1 expression correlates with the disease recurrence, supporting the notion of SRC‐1 involvement in tamoxifen resistance.61,100 However, as described above, an alternative mechanism of SRC‐1 in tamoxifen resistance might be its coactivator action for the Ets transcription factor.

C. Androgen Independence in Prostate Cancer Although androgen ablation is an initial treatment of choice for prostate cancer, much evidence highlights an important role for AR in subsequent development and progression of the disease to a hormone‐refractory state. The role of AR in mediating prostate cancer recurrence is under heavy investigation. AR reactivation under androgen deprivation appears to involve several possible mechanisms including AR mutations and/or overexpression and posttranslational modifications through aberrant growth factor/kinase‐mediated

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signaling. Many studies also indicate an important role of p160 coactivators in enhancing AR activation by low androgen levels or alternative growth factor kinase‐mediated signaling in prostate cancer. 1. P160 COACTIVATOR LEVELS ARE ALTERED BY ANDROGEN ABLATION TO FACILITATE LIGAND‐DEPENDENT AND ‐INDEPENDENT AR ACTIVATION Specific RNAi‐mediated depletion of the individual p160 coactivators has demonstrated that their expression is important for the proliferation of AR positive androgen‐dependent and ‐independent prostate cancer cell lines.26,29,31,75 Higher protein expression of SRC‐1 and TIF2 was observed in a CWR22 xenograft mouse model of recurrent prostate cancer after androgen ablation and in a small number of clinical samples (n ¼ 8) of recurrent tumors compared to BPH and androgen‐dependent primary tumors.56 TIF2 overexpression enhanced AR transactivation by low concentrations of adrenal androgens and by low‐affinity steroids in reporter gene analysis.107 Another study using AR–LBD mutants suggests that ligand binding is required for hormone‐refractory growth in vitro and in vivo.108 These findings suggest that changes in p160 coactivator levels during disease recurrence can influence AR sensitivity to the growth stimulatory effects of low androgen concentrations and other circulating hormones. Although the mechanisms by which androgen deprivation modulates coactivator levels are not yet fully elucidated, one study showed that TIF2 expression in androgen‐dependent prostate cancer cells is significantly reduced by androgen treatment, while molecular analysis of TIF2 promoter revealed that it is directly targeted and repressed by agonist‐ or antagonist‐bound AR.31 Along with the consistent finding that AR expression also increases in models of hormone‐refractory recurrence, this suggests that upregulation of TIF2 is a consequence of androgen deprivation that elicits a hypersensitive AR activation response to residual hormones. Interestingly, several studies have shown that the p160 coactivators are involved in androgen‐independent AR activation and expression of target genes. AR target genes are upregulated in response to androgen treatment in androgen‐dependent prostate cancer cells, whereas they are constitutively expressed in androgen‐independent derivatives. Ectopic AIB1/ACTR expression increased mRNA level of AR target gene PSA in LNCaP cells in the absence of DHT,109 and TIF2 depletion by shRNA lowered basal PSA levels in the androgen‐independent derivative of LNCaP cells.110 These findings suggest that elevated levels of p160 coactivators can elicit AR loading onto chromatin independently of ligand to nucleate an activated transcription complex on PSA and perhaps other AR target genes that control cell proliferation. Another study demonstrated that overexpression of AR in LNCaP cells is sufficient to confer growth in low‐androgen conditions and resistance to antagonists such as bicalutamide. ChIP analysis of target genes showed that

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bicalutimide induced recruitment of activated AR transcription complex components including the SRC coactivators and RNAPII in AR‐infected LNCaP cells but not in vector‐infected cells.108 2. AR MUTATIONS AND KINASE SIGNALING CAN INCREASE LIGAND AND P160 COACTIVATOR BINDING AFFINITY Aside from their overexpression, the p160 coactivators may cooperate with other mechanisms that enhance AR activation under androgen‐deprived conditions. Gain of function mutations in the AR have been implicated to confer selective advantage for hormone‐refractory growth of prostate tumor cells and are associated with advanced stages of disease and antiandrogen therapy.111 Some of these AR mutants display promiscuous activation by AR antagonists, adrenal androgens such as DHEA, and nonandrogen hormones such as estradiol and progesterone. One such mutation, H874Y located in helix 10/11 of the LBD, has been shown to increase the binding affinity for and transactivation sensitivity to the p160 coactivators in the presence of androgen.112 Shorter length of polyglutamine tract located in the AR N‐terminal domain is associated with higher AR transcriptional activity and prostate cancer risk. The polyglutamine length inversely correlated with ligand binding affinity and ligand‐dependent interaction between AR and p160 coactivators.53 Another adaptive change implicated in recurrent prostate cancer is increased ligand‐independent AR activation through kinase signaling pathways that elicit phosphorylation of the receptor and its coactivators. IGF‐1, EGF, keratinocyte growth factor, IL‐6, neuropeptides, and HER2 overexpression have all been reported to induce AR transactivation in the absence of androgen.113–115 Mutational analysis revealed that MAPK phosphorylation of SRC‐1 on Ser1185 and Thr1179 is required for optimal ligand‐independent AR activation by IL‐6.114 ChIP assay indicated that neuropeptides can selectively induce the recruitment of SRC‐1 and AIB1/ACTR, but not TIF2, preferentially to the proximal promoter region of AR target gene PSA.115 Interestingly, EGF increased TIF2 expression in cells derived from recurrent human prostate cancer xenografts and was sufficient to reverse the reduction of both AR and TIF2 observed following castration. Unlike androgen treatment, the effect of EGF on TIF2 expression was posttranscriptional as the changes occurred at the protein but not at the mRNA level, and depended on phosphorylation by MAPK of a serine residue located close to one of the LXXLL motifs in the receptor interaction domain of TIF2.116 Thus, aside from direct activation of AR and associated coactivators, MAPK‐mediated phosphorylation induced by various activated growth factor and cytokine signaling pathways may also function to increase p160 coactivator levels in hormone‐refractory prostate cancer cells, and in doing so, elicit a cooperative mechanism to enhance expression of AR target genes.

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IV. Functions of p160/SRCs in Tumorigenesis Revealed in Animal Models A. SRCs in Mouse Mammary Tumorigenesis and Tumor Metastasis Since AIB1/ACTR/SRC‐3 gene was found to be frequently amplified and/ or overexpressed in multiple types of cancers, studies on the function of SRCs in tumorigenesis using animal models have been focused primarily on SRC‐3. In an early study aimed at exploring the oncogenic function of AIB1/ACTR in mouse mammary gland, multiple AIB1/ACTR transgenic mouse lines were made where the expression of human AIB1/ACTR cDNA was controlled by hormone‐responsive MMTV‐promoter.88 Overexpression of AIB1/ACTR resulted in significantly increased size of mammary gland (30–40% larger compared to wild type (WT)) at puberty and in delayed involution postweaning. These phenotypes are likely due to the cellular effects of AIB1/ACTR overexpression such as increased cell proliferation and delayed apoptosis observed in the transgenic animal mammary glands. As the transgenic animals age, an increasing number (over 70%) of them developed tumors and the average latency is about 16 months. Interestingly, only about half of the animals developed adenocarcinomas in the mammary glands and the other half developed tumors in other tissues including pituitary, uterus, lung, and tumors of mesenchymal origin. Different from tumors developed in many other transgenic models for breast cancer,117 the majority of mammary adenocarincomas induced by AIB1/ACTR transgene are ERa‐positive, suggesting that the receptor is an important mediator of AIB1/ACTR mammary tumorigenesis. Therefore, this study clearly demonstrated that aberrantly expressed AIB1/ ACTR can act as a potent oncogene. The role of AIB1/ACTR/SRC‐3 in mammary tumorigenesis was also examined in other studies using different approaches including oncogene and carcinogen‐induced tumorigenesis. MMTV‐driven, v‐Ha‐ras transgenic mice are highly susceptible to mammary tumorigenesis.118 Interestingly, elevated levels of SRC‐3 protein were detected in the mammary tumors of SRC‐3þ/þ‐ras mice.89 By cross‐breeding SRC‐3 KO mice with the H‐ras mice, it was found that ras‐ induced mammary tumor onset was significantly delayed and the tumor incidence was also dramatically reduced in SRC‐3 / ‐ras mice. Intriguingly, SRC‐3 gene inactivation did not affect the tumorigenesis‐stimulatory role of ovarian hormones. This is based on the results that SRC‐3 deficiency prolonged the tumor latency similarly in mice with normal, elevated, or depleted ovarian hormones. Recently, a crucial role of SRC‐3 was also revealed in HER2/Neu‐induced mammary tumorigenesis.119 One allele inactivation of SRC‐3 was sufficient to significantly delay Neu‐induced tumor development while its total deletion

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completely prevented the oncogene‐induced tumorigenesis in mouse mammary glands. In a carcinogen‐induced tumorigenesis model, SRC‐3 deficiency was found to suppress DMBA‐induced tumor development in mammary glands but not in the skin,120 consistent with the critical role of SRC‐3 in normal mammary gland development, as SRC‐3 / mice displayed significant delay in mammary ductal morphogenesis.86 However, the tumor stimulatory effect of increased levels of hormones (via pituitary isografts) was not compromised in the DMBA‐ tumor model. Together, these studies lent strong genetic evidence for the crucial role of SRC‐3 in mammary tumorigenesis. They also suggest that SRC‐3 in tumorigenesis may act, in part, through a mechanism independent of the ovarian hormones and their receptors. The role of SRC‐3 in tumor metastasis has also been examined in animal models.92 Polyomavirus middle T antigen (PyMT), activating via major kinase signaling pathways including PI3K/Akt and ras/MAPK, can rapidly induce mammary tumorigenesis and extensive lung metastasis.121 When the MMTV‐driven PyMT transgenic mice were crossed with the SRC‐3 KO mice, a marked inhibition of tumor metastasis to the lung was observed in SRC‐3 / /PyMT mice, as the SRC‐3 KO mice showed much decreased metastatic tumor areas in the lung (from 8.5% of lung section in WT to 2.5% in SRC‐3 / /PyMT mice). Transplantation of SRC‐3 / /PyMT tumors to the mammary fat pads of nontransgenic normal mice showed similar defects of lung metastasis, indicating that SRC‐3 function in the primary tumor cells is required for effective lung metastasis.92 Further analysis revealed that the tumor cells from SRC‐3 / /PyMT mice had highly reduced migration and invasion activities, which correlates with their more differentiated phenotype including the retention of the expression of epithelial cell markers (E‐cadherin and b‐catenin) in SRC‐3 KO tumors. The SRC‐1 gene function in mammary tumorigenesis has also recently been examined in the PyMT model.122 Interestingly, unlike the observation of a significant delay of tumor onset in SRC‐3 KO mice, SRC‐1 deficiency showed little effect on PyMT‐induced tumor initiation and growth. However, SRC‐1 gene inactivation caused a drastic decrease of tumor lung metastasis. The occurrence of lung tumor foci was markedly reduced from 85% in WT/PyMT mice to about 10% in SRC‐1 / /PyMT mice. Moreover, tumor cells in the blood were much less frequently detected in SRC‐1 KO mice, suggesting that SRC‐1 plays a role in tumor cell intravasation. Like SRC‐3, tumor transplantation experiments demonstrated an intrinsic role of SRC‐1 in the tumor metastasis.

B. SRC‐3 in the Mouse Prostate Cancer Model All three SRCs have been found to act as strong coactivators for AR and play critical roles in androgen‐stimulated expression of genes such as PSA in human prostate cancer cells.123,124 Studies with human tumor specimens and

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cancer cell lines strongly suggest important roles of the SRCs in prostate cancer. However, thus far only one study on SRC‐3 in prostate cancer using animal model was reported.125 The transgenic adenocarcinoma of the mouse prostate (TRAMP) model was one of the first animal models developed for prostate cancer. The model uses rat probasin promoter to express the SV40 T and t antigens specifically in the terminally differentiated prostate epithelial cells.126 The large T antigen functions as an oncoprotein through interaction with pRb and p53 tumor suppressors and the small t antigen can also disrupt cell cycle control when interacting with a protein phosphatase. TRAMP males develop locally invasive tumors that are metastatic and poorly differentiated. When SRC‐3 was inactivated, tumor progression was strongly inhibited as most of SRC‐3 / /TRAMP mice only developed early stage, well‐ differentiated adenocarcinomas that retain expression of epithelial markers such as E‐cadherin and cytokeratin 8. Another intriguing finding was that SRC‐3 expression, which was hardly detectable in normal prostate epithelium, was highly elevated in the malignant prostate of TRAMP mice during the tumor progression, peaking in the late stage, poorly differentiated tumors.125 Together with the lack of evidence that SRC‐3 deficiency affected tumor onset and/or incidence, the results support the conclusion that high levels of SRC‐3 are required specifically for prostate cancer progression to a more advanced stage.

V. The Coregulator ANCCA, a Unique Target of AIB1/ACTR and a Potential Key Player in Cancer A. ANCCA is a Hormone‐Induced Nuclear Receptor Coregulator and a Target of AIB1/ACTR The important role of AIB1/ACTR in cancer development and progression prompted many to search for key downstream targets that might mediate its oncogenic activity. In one of our studies, we performed gene expression microarray analysis to identify genes with expression altered by overexpressed AIB1/ACTR independent of hormone. One of the cDNA sequences upregulated by the ectopic AIB1/ACTR was predicted to possess two AAAþ ATPase domains (AAA‐D1 and AAA‐D2) and a bromodomain at its C‐terminus, but its function was not characterized at the time. To account for its recognized domains, its biological function and its overexpression in cancers (described below), we designate this gene as ANCCA for AAAþ nuclear cofactor cancer‐ associated. Consistent with the fact that AAAþ (ATPase associated with various cellular activities) domains are evolutionarily highly conserved protein

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modules,127 genes with similar AAAþ and bromodomain composition can be found in the database across the entire eukaryotes (except Drosophila melanogaster) including lex‐1 in Caenorhabditis elegans and yta7 in Saccharomyces cerevisiae. Supporting the conclusion that ANCCA is a direct target of AIB1/ACTR, ectopic AIB1/ACTR induced the expression of ANCCA in ER‐positive breast cancer cells with ER activity inhibited by antiestrogen ICI182,780, while endogenous AIB1/ACTR protein was found to occupy ANCCA promoter.128 Unexpectedly, when the breast cancer cells were treated with estrogen, ANCCA expression was also strongly induced both at its transcript and protein levels. Remarkably, over 10‐fold induction of ANCCA by androgen DHT can also be detected in human prostate cancer cells.129 Together with our unpublished observation that ANCCA expression is stimulated by estrogen in ovarian hormone‐responsive tissues of mouse, these results suggest that ANCCA may function as a nuclear coregulator to mediate hormone responses. Supporting this notion, we found that ANCCA interacts directly with ER‐alpha and AR and, upon hormone stimulation, is recruited to a specific subset of the receptor target genes such as cyclin D1, c‐myc and E2F1 in breast cancer cells, and IRS‐2 and SGK‐1 in prostate cancer cells.128,129 ANCCA depletion strongly inhibited the target gene expression. Remarkably, ANCCA appears not to play a crucial role in hormone induction of a group of genes such as pS2 and PSA/ KLK3, and does not directly interact with other receptors such as RARs. These results suggest that ANCCA as a nuclear receptor coregulator has a functional mechanism distinct from that of the SRCs.

B. ANCCA, a Potential Key Player in Tumorigenesis, is Frequently Overexpressed in Different Cancers The identified ANCCA target genes such as cyclin D1, c‐myc, E2F1, and IRS‐2 are well known for their roles in tumorigenesis. Accordingly, suppression of ANCCA expression in breast cancer and prostate cancer cells resulted in strong inhibition of cell proliferation, cell cycle progression, and their survival.128,129 Supporting the notion that ANCCA may be a key player in cancer, several gene expression profiling studies aimed at identifying specific groups of aberrantly expressed genes as signatures for cancer prognosis listed ANCCA (in the name of pro2000 or atad2) as one of the genes frequently overexpressed in large cohorts of breast cancer specimens. For instance, in one study, ANCCA/pro2000 is one of the 39 genes with increased expression correlating with transition from preinvasive (DCIS) to invasive ductal carcinoma (IDC) of breast cancer.130 In another study, ANCCA/pro2000 is one of the 76 genes identified as a gene signature for prediction of distant metastasis of ER‐positive but lymph‐node‐negative primary breast cancer.131 The clinical significance of

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the 76 genes as prognosis marker was later validated in several multicenter studies.132,133 In an analysis of limited number of high grade osteosarcoma specimens, high transcript level of ANCCA/ATAD2 is associated with decrease in event‐free survival of the patients.134 Our recent IHC study suggests that ANCCA protein is overexpressed in a subset of human breast cancers (unpublished data) and prostate cancers, and that its overexpression in prostate cancer correlates with higher Gleason score.129 Together these findings strongly implicate ANCCA as a key player in tumorigenesis and/or cancer progression.

VI. Concluding Remarks The last decade has witnessed a rapid advance in our understanding of the role of p160/SRC family of transcriptional coregulators in human cancers. The major findings include the oncogenic function of AIB1/ACTR/SRC‐3 revealed in animal models, the crucial roles of different SRCs in control of proliferation and survival of different cancer cells and the several mechanistic pathways deregulated by aberrantly expressed SRCs. However, many important questions remain to be addressed. For example, it remains to be seen whether the other two SRCs and ANCCA possess oncogenic activity and whether overexpressed AIB1/ACTR also acts as an oncogene in other tissues such as prostate and pancreas. Given the results that aged mice with SRC‐3 gene inactivated developed B‐cell lymphoma,135 it will be important to better understand the specific tissue/cellular context where the SRCs exert their protumorigenic roles. Also, although a strong link of AIB1/ACTR overexpression with erbB2/HER2 was revealed in human breast cancer tumors, a direct demonstration of such link in experimental systems has been lacking. Moreover, several clinically relevant issues remain unresolved. For instance, what are the initial triggers for the gene amplification‐independent overexpression of SRCs and ANCCA? To what extent might their action mechanisms identified using the experimental models operate in the human tumors? Finally, how might these coregulators be targeted for development of therapeutics? Continued research on these and other issues will certainly provide valuable insights to the prevention, prognosis, and treatment of the multiple types of malignancy where the coregulators are intimately involved. Acknowledgments This work was partially supported by an NIH grant R01CA113860 (HWC). E.Y.H. was a recipient of The Flloyd & Mary Schwall Medical Fellowship.

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References 1. Conzen SD. Minireview: nuclear receptors and breast cancer. Mol Endocrinol 2008;22:2215–28. 2. Mongan NP, Gudas LJ. Diverse actions of retinoid receptors in cancer prevention and treatment. Differentiation 2007;75:853–70. 3. Scaglioni PP, Pandolfi PP. The theory of APL revisited. Curr Top Microbiol Immunol 2007;313:85–100. 4. Chen H, Lin RJ, Schiltz RL, Chakravarti D, Nash A, Nagy L, et al. Nuclear receptor coactivator ACTR is a novel histone acetyltransferase and forms a multimeric activation complex with P/CAF and CBP/p300. Cell 1997;90:569–80. 5. Li H, Gomes PJ, Chen JD. RAC3, a steroid/nuclear receptor‐associated coactivator that is related to SRC‐1 and TIF2. Proc Natl Acad Sci USA 1997;94:8479–84. 6. Takeshita A, Cardona GR, Koibuchi N, Suen CS, Chin WW. TRAM‐1, a novel 160‐kda thyroid hormone receptor activator molecule, exhibits distinct properties from steroid receptor coactivator‐1. J Biol Chem 1997;272:27629–34. 7. Isola JJ, Kallioniemi OP, Chu LW, Fuqua SA, Hilsenbeck SG, Osborne CK, et al. Genetic aberrations detected by comparative genomic hybridization predict outcome in node‐negative breast cancer. Am J Pathol 1995;147:905–11. 8. Kallioniemi A, Kallioniemi OP, Piper J, Tanner M, Stokke T, Chen L, et al. Detection and mapping of amplified DNA sequences in breast cancer by comparative genomic hybridization. Proc Natl Acad Sci USA 1994;91:2156–60. 9. Tanner MM, Tirkkonen M, Kallioniemi A, Isola J, Kuukasjarvi T, Collins C, et al. Independent amplification and frequent co‐amplification of three nonsyntenic regions on the long arm of chromosome 20 in human breast cancer. Cancer Res 1996;56:3441–5. 10. Guan XY, Xu J, Anzick SL, Zhang H, Trent JM, Meltzer PS. Hybrid selection of transcribed sequences from microdissected DNA: isolation of genes within amplified region at 20q11‐q13.2 in breast cancer. Cancer Res 1996;56:3446–50. 11. Anzick SL, Kononen J, Walker RL, Azorsa DO, Tanner MM, Guan X‐Y, et al. AIB1, a steroid receptor coactivator amplified in breast and ovarian cancer. Science 1997;277:965–8. 12. Maki HE, Waltering KK, Wallen MJ, Martikainen PM, Tammela TL, van Weerden WM, et al. Screening of genetic and expression alterations of SRC1 gene in prostate cancer. Prostate 2006;66:1391–8. 13. Burwinkel B, Wirtenberger M, Klaes R, Schmutzler RK, Grzybowska E, Forsti A, et al. Association of NCOA3 polymorphisms with breast cancer risk. Clin Cancer Res 2005;11:2169–74. 14. Haiman CA, Garcia RR, Hsu C, Xia L, Ha H, Sheng X, et al. Screening and association testing of common coding variation in steroid hormone receptor co‐activator and co‐repressor genes in relation to breast cancer risk: the Multiethnic Cohort. BMC Cancer 2009;9:43. 15. Bouras T, Southey MC, Venter DJ. Overexpression of the steroid receptor coactivator AIB1 in breast cancer correlates with the absence of estrogen and progesterone receptors and positivity for p53 and HER2/neu. Cancer Res 2001;61:903–7. 16. Sakakura C, Hagiwara A, Yasuoka R, Fujita Y, Nakanishi M, Masuda K, et al. Amplification and over‐expression of the AIB1 nuclear receptor co‐activator gene in primary gastric cancers. Int J Cancer 2000;89:217–23. 17. Luo JH, Xie D, Liu MZ, Chen W, Liu YD, Wu GQ, et al. Protein expression and amplification of AIB1 in human urothelial carcinoma of the bladder and overexpression of AIB1 is a new independent prognostic marker of patient survival. Int J Cancer 2008;122:2554–61.

292

HSIA ET AL.

18. Xie D, Sham JST, Zeng W‐F, Lin H‐L, Bi J, Che L‐H, et al. Correlation of AIB1 overexpression with advanced clinical stage of human colorectal carcinoma. Hum Pathol 2005;36:777–83. 19. Xu F‐P, Xie D, Wen J‐M, Wu H‐X, Liu Y‐D, Bi J, et al. SRC‐3/AIB1 protein and gene amplification levels in human esophageal squamous cell carcinomas. Cancer Lett 2007;245:69–74. 20. Henke RT, Haddad BR, Kim SE, Rone JD, Mani A, Jessup JM, et al. Overexpression of the nuclear receptor coactivator AIB1 (SRC‐3) during progression of pancreatic adenocarcinoma. Clin Cancer Res 2004;10:6134–42. 21. List HJ, Reiter R, Singh B, Wellstein A, Riegel AT. Expression of the nuclear coactivator AIB1 in normal and malignant breast tissue. Breast Cancer Res Treat 2001;68:21–8. 22. Hudelist G, Czerwenka K, Kubista E, Marton E, Pischinger K, Singer CF. Expression of sex steroid receptors and their co‐factors in normal and malignant breast tissue: AIB1 is a carcinoma‐specific co‐activator. Breast Cancer Res Treat 2003;78:193–204. 23. Wang Y, Wu MC, Sham JS, Zhang W, Wu WQ, Guan XY. Prognostic significance of c‐myc and AIB1 amplification in hepatocellular carcinoma. A broad survey using high‐throughput tissue microarray. Cancer 2002;95:2346–52. 24. Sakaguchi H, Fujimoto J, Sun WS, Tamaya T. Clinical implications of steroid receptor coactivator (SRC)‐3 in uterine endometrial cancers. J Steroid Biochem Mol Biol 2007;104:237–40. 25. Tanner MM, Grenman S, Koul A, Johannsson O, Meltzer P, Pejovic T, et al. Frequent amplification of chromosomal region 20q12‐q13 in ovarian cancer. Clin Cancer Res 2000;6:1833–9. 26. Agoulnik IU, Vaid A, Bingman WE, III, Erdeme H, Frolov A, Smith CL, et al. Role of SRC‐1 in the promotion of prostate cancer cell growth and tumor progression. Cancer Res 2005;65:7959–67. 27. Linja MJ, Porkka KP, Kang Z, Savinainen KJ, Janne OA, Tammela TL, et al. Expression of androgen receptor coregulators in prostate cancer. Clin Cancer Res 2004;10:1032–40. 28. Gnanapragasam VJ, Leung HY, Pulimood AS, Neal DE, Robson CN. Expression of RAC 3, a steroid hormone receptor co‐activator in prostate cancer. Br J Cancer 2001;85:1928–36. 29. Zhou HJ, Yan J, Luo W, Ayala G, Lin SH, Erdem H, et al. SRC‐3 is required for prostate cancer cell proliferation and survival. Cancer Res 2005;65:7976–83. 30. Yan J, Erdem H, Li R, Cai Y, Ayala G, Ittmann M, et al. Steroid receptor coactivator‐3/AIB1 promotes cell migration and invasiveness through focal adhesion turnover and matrix metalloproteinase expression. Cancer Res 2008;68:5460–8. 31. Agoulnik IU, Vaid A, Nakka M, Alvarado M, Bingman WE, III, Erdem H, et al. Androgens modulate expression of transcription intermediary factor 2, an androgen receptor coactivator whose expression level correlates with early biochemical recurrence in prostate cancer. Cancer Res 2006;66:10594–602. 32. Yang XJ, Ullah M. MOZ and MORF, two large mystic hats in normal and cancer stem cells. Oncogene 2007;26:5408–19. 33. Katsumoto T, Yoshida N, Kitabayashi I. Roles of the histone acetyltransferase monocytic leukemia zinc finger protein in normal and malignant hematopoiesis. Cancer Sci 2008;99:1523–7. 34. Deguchi K, Ayton PM, Carapeti M, Kutok JL, Snyder CS, Williams I.R, et al. MOZ‐TIF2‐ induced acute myeloid leukemia requires the MOZ nucleosome binding motif and TIF2‐ mediated recruitment of CBP. Cancer Cell 2003;3:259–71. 35. Huntly BJ, Shigematsu H, Deguchi K, Lee BH, Mizuno S, Duclos N, et al. MOZ‐TIF2, but not BCR‐ABL, confers properties of leukemic stem cells to committed murine hematopoietic progenitors. Cancer Cell 2004;6:587–96.

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36. Carapeti M, Aguiar RC, Goldman JM, Cross NC. A novel fusion between MOZ and the nuclear receptor coactivator TIF2 in acute myeloid leukemia. Blood 1998;91:3127–33. 37. Liang J, Prouty L, Williams BJ, Dayton MA, Blanchard KL. Acute mixed lineage leukemia with an inv(8)(p11q13) resulting in fusion of the genes for MOZ and TIF2. Blood 1998;92:2118–22. 38. Kindle KB, Troke PJ, Collins HM, Matsuda S, Bossi D, Bellodi C, et al. MOZ‐TIF2 inhibits transcription by nuclear receptors and p53 by impairment of CBP function. Mol Cell Biol 2005;25:988–1002. 39. Collins HM, Kindle KB, Matsuda S, Ryan C, Troke PJ, Kalkhoven E, et al. MOZ‐TIF2 alters cofactor recruitment and histone modification at the RARbeta2 promoter: differential effects of MOZ fusion proteins on CBP‐ and MOZ‐dependent activators. J Biol Chem 2006;281:17124–33. 40. Strehl S, Nebral K, Konig M, Harbott J, Strobl H, Ratei R, et al. ETV6‐NCOA2: a novel fusion gene in acute leukemia associated with coexpression of T‐lymphoid and myeloid markers and frequent NOTCH1 mutations. Clin Cancer Res 2008;14:977–83. 41. Bohlander SK. ETV6: a versatile player in leukemogenesis. Semin Cancer Biol 2005;15:162–74. 42. Wachtel M, Dettling M, Koscielniak E, Stegmaier S, Treuner J, Simon‐Klingenstein K, et al. Gene expression signatures identify rhabdomyosarcoma subtypes and detect a novel t(2;2) (q35;p23) translocation fusing PAX3 to NCOA1. Cancer Res 2004;64:5539–45. 43. Esteyries S, Perot C, Adelaide J, Imbert M, Lagarde A, Pautas C, et al. NCOA3, a new fusion partner for MOZ/MYST3 in M5 acute myeloid leukemia. Leukemia 2008;22:663–5. 44. Hayashi Y, Yamamoto M, Ohmori S, Kikumori T, Imai T, Funahashi H, et al. Polymorphism of homopolymeric glutamines in coactivators for nuclear hormone receptors. Endocr J 1999;46:279–84. 45. Rebbeck TR, Wang Y, Kantoff PW, Krithivas K, Neuhausen SL, Godwin AK, et al. Modification of BRCA1‐ and BRCA2‐associated breast cancer risk by AIB1 genotype and reproductive history. Cancer Res 2001;61:5420–4. 46. Hughes DJ, Ginolhac SM, Coupier I, Barjhoux L, Gaborieau V, Bressac‐de‐Paillerets B, et al. Breast cancer risk in BRCA1 and BRCA2 mutation carriers and polyglutamine repeat length in the AIB1 gene. Int J Cancer 2005;117:230–3. 47. Kadouri L, Kote‐Jarai Z, Easton DF, Hubert A, Hamoudi R, Glaser B, et al. Polyglutamine repeat length in the AIB1 gene modifies breast cancer susceptibility in BRCA1 carriers. Int J Cancer 2004;108:399–403. 48. Wilkening S, Burwinkel B, Grzybowska E, Klaes R, Pamula J, Pekala W, et al. Polyglutamine repeat length in the NCOA3 does not affect risk in familial breast cancer. Cancer Epidemiol Biomarkers Prev 2005;14:291–2. 49. Colilla S, Kantoff PW, Neuhausen SL, Godwin AK, Daly MB, Narod SA, et al. The joint effect of smoking and AIB1 on breast cancer risk in BRCA1 mutation carriers. Carcinogenesis 2006;27:599–605. 50. Hsing AW, Chokkalingam AP, Gao Y‐T, Wu G, Wang X, Deng J, et al. Polymorphic CAG/CAA repeat length in the AIB1/SRC‐3 gene and prostate cancer risk: a population‐based case‐ control study. Cancer Epidemiol Biomarkers Prev 2002;11:337–41. 51. Li AJ, Lerner DL, Gapuzan M‐ER, Karlan BY. AIB1 polymorphisms predict aggressive ovarian cancer phenotype. Cancer Epidemiol Biomarkers Prev 2005;14:2919–22. 52. Dai P, Wong L‐JC. Somatic instability of the DNA sequences encoding the polymorphic polyglutamine tract of the AIB1 gene. J Med Genet 2003;40:885–90. 53. Wang Q, Udayakumar TS, Vasaitis TS, Brodie AM, Fondell JD. Mechanistic relationship between androgen receptor polyglutamine tract truncation and androgen‐dependent transcriptional hyperactivity in prostate cancer cells. J Biol Chem 2004;279:17319–28.

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54. Bevan CL, Hoare S, Claessens F, Heery DM, Parker MG. The AF1 and AF2 domains of the androgen receptor interact with distinct regions of SRC1. Mol Cell Biol 1999;19:8383–92. 55. Fleming FJ, Myers E, Kelly G, Crotty TB, McDermott EW, O’Higgins NJ, et al. Expression of SRC‐1, AIB1, and PEA3 in HER2 mediated endocrine resistant breast cancer; a predictive role for SRC‐1. J Clin Pathol 2004;57:1069–74. 56. Gregory CW, He B, Johnson RT, Ford OH, Mohler JL, French FS, et al. A mechanism for androgen receptor‐mediated prostate cancer recurrence after androgen deprivation therapy. Cancer Res 2001;61:4315–9. 57. Bautista S, Valles H, Walker RL, Anzick S, Zeillinger R, Meltzer P, et al. In breast cancer, amplification of the steroid receptor coactivator gene AIB1 is correlated with estrogen and progesterone receptor positivity. Clin Cancer Res 1998;4:2925–9. 58. Osborne CK, Bardou V, Hopp TA, Chamness GC, Hilsenbeck SG, Fuqua SAW, et al. Role of the estrogen receptor coactivator AIB1 (SRC‐3) and HER‐2/neu in tamoxifen resistance in breast cancer. J Natl Cancer Inst 2003;95:353–61. 59. Kirkegaard T, McGlynn LM, Campbell FM, Muller S, Tovey SM, Dunne B, et al. Amplified in breast cancer 1 in human epidermal growth factor receptor—positive tumors of tamoxifen‐ treated breast cancer patients. Clin Cancer Res 2007;13:1405–11. 60. Dihge L, Bendahl PO, Grabau D, Isola J, Lovgren K, Ryden L, et al. Epidermal growth factor receptor (EGFR) and the estrogen receptor modulator amplified in breast cancer (AIB1) for predicting clinical outcome after adjuvant tamoxifen in breast cancer. Breast Cancer Res Treat 2008;109:255–62. 61. Al‐azawi D, Ilroy MM, Kelly G, Redmond AM, Bane FT, Cocchiglia S, et al. Ets‐2 and p160 proteins collaborate to regulate c‐Myc in endocrine resistant breast cancer. Oncogene 2008;27:3021–31. 62. Balmer NN, Richer JK, Spoelstra NS, Torkko KC, Lyle PL, Singh M. Steroid receptor coactivator AIB1 in endometrial carcinoma, hyperplasia and normal endometrium: correlation with clinicopathologic parameters and biomarkers. Mod Pathol 2006;19:1593–605. 63. Ghadimi BM, Schrock E, Walker RL, Wangsa D, Jauho A, Meltzer PS, et al. Specific chromosomal aberrations and amplification of the AIB1 nuclear receptor coactivator gene in pancreatic carcinomas. Am J Pathol 1999;154:525–36. 64. Louie MC, Revenko AS, Zou JX, Yao J, Chen HW. Direct control of cell cycle gene expression by proto‐oncogene product ACTR, and its autoregulation underlies its transforming activity. Mol Cell Biol 2006;26:3810–23. 65. Mussi P, Yu C, O’Malley BW, Xu J. Stimulation of steroid receptor coactivator‐3 (SRC‐3) gene overexpression by a positive regulatory loop of E2F1 and SRC‐3. Mol Endocrinol 2006;20:3105–19. 66. Li X, Lonard DM, Jung SY, Malovannaya A, Feng Q, Qin J, et al. The SRC‐3/AIB1 coactivator is degraded in a ubiquitin‐ and ATP‐independent manner by the REG[gamma] proteasome. Cell 2006;124:381–92. 67. Wu R‐C, Feng Q, Lonard DM, O’Malley BW. SRC‐3 coactivator functional lifetime is regulated by a phospho‐dependent ubiquitin time clock. Cell 2007;129:1125–40. 68. Yi P, Feng Q, Amazit L, Lonard DM, Tsai SY, Tsai M‐J, et al. Atypical protein kinase C regulates dual pathways for degradation of the oncogenic coactivator SRC‐3/AIB1. Mol Cell 2008;29:465–76. 69. Lauritsen KJ, List HJ, Reiter R, Wellstein A, Riegel AT. A role for TGF‐beta in estrogen and retinoid mediated regulation of the nuclear receptor coactivator AIB1 in MCF‐7 breast cancer cells. Oncogene 2002;21:7147–55. 70. Glaeser M, Floetotto T, Hanstein B, Beckmann MW, Niederacher D. Gene amplification and expression of the steroid receptor coactivator SRC3 (AIB1) in sporadic breast and endometrial carcinomas. Horm Metab Res 2001;33:121–6.

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71. Shao W, Keeton EK, McDonnell DP, Brown M. Coactivator AIB1 links estrogen receptor transcriptional activity and stability. Proc Natl Acad Sci USA 2004;101:11599–604. 72. Urbanucci A, Waltering KK, Suikki HE, Helenius MA, Visakorpi T. Androgen regulation of the androgen receptor coregulators. BMC Cancer 2008;8:219. 73. Louie MC, Zou JX, Rabinovich A, Chen HW. ACTR/AIB1 functions as an E2F1 coactivator to promote breast cancer cell proliferation and antiestrogen resistance. Mol Cell Biol 2004;24:5157–71. 74. List HJ, Lauritsen KJ, Reiter R, Powers C, Wellstein A, Riegel AT. Ribozyme targeting demonstrates that the nuclear receptor coactivator AIB1 is a rate‐limiting factor for estrogen‐dependent growth of human MCF‐7 breast cancer cells. J Biol Chem 2001;276:23763–8. 75. Zou JX, Zhong Z, Shi XB, Tepper CG, deVere White RW, Kung HJ, et al. ACTR/AIB1/SRC‐3 and androgen receptor control prostate cancer cell proliferation and tumor growth through direct control of cell cycle genes. Prostate 2006;66:1474–86. 76. Oh A, List H‐J, Reiter R, Mani A, Zhang Y, Gehan E, et al. The nuclear receptor coactivator AIB1 mediates insulin‐like growth factor I‐induced phenotypic changes in human breast cancer cells. Cancer Res 2004;64:8299–308. 77. Chen H, Lin RJ, Xie W, Wilpitz D, Evans RM. Regulation of hormone‐induced histone hyperacetylation and gene activation via acetylation of an acetylase. Cell 1999;98:675–86. 78. Shang Y, Hu X, DiRenzo J, Lazar MA, Brown M. Cofactor dynamics and sufficiency in estrogen receptor‐regulated transcription. Cell 2000;103:843–52. 79. Planas‐Silva MD, Shang Y, Donaher JL, Brown M, Weinberg RA. AIB1 enhances estrogen‐ dependent induction of cyclin D1 expression. Cancer Res 2001;61:3858–62. 80. Hall JM, Korach KS. Stromal cell‐derived factor 1, a novel target of estrogen receptor action, mediates the mitogenic effects of estradiol in ovarian and breast cancer cells. Mol Endocrinol 2003;17:792–803. 81. Kishimoto H, Wang Z, Bhat‐Nakshatri P, Chang D, Clarke R, Nakshatri H. The p160 family coactivators regulate breast cancer cell proliferation and invasion through autocrine/paracrine activity of SDF‐1alpha/CXCL12. Carcinogenesis 2005;26:1706–15. 82. Karmakar S, Foster EA, Smith CL. Unique roles of p160 coactivators for regulation of breast cancer cell proliferation and estrogen receptor‐{alpha} transcriptional activity. Endocrinology 2009;150:1588–96. 83. Yin N, Wang D, Zhang H, Yi X, Sun X, Shi B, et al. Molecular mechanisms involved in the growth stimulation of breast cancer cells by leptin. Cancer Res 2004;64:5870–5. 84. Hsu T, Trojanowska M, Watson DK. Ets proteins in biological control and cancer. J Cell Biochem 2004;91:896–903. 85. Frank DA. STAT3 as a central mediator of neoplastic cellular transformation. Cancer Lett 2007;251:199–210. 86. Xu J, Liao L, Ning G, Yoshida‐Komiya H, Deng C, O’Malley BW. The steroid receptor coactivator SRC‐3 (p/CIP/RAC3/AIB1/ACTR/TRAM‐1) is required for normal growth, puberty, female reproductive function, and mammary gland development. Proc Natl Acad Sci USA 2000;97:6379–84. 87. Wang Z, Rose DW, Hermanson O, Liu F, Herman T, Wu W, et al. Regulation of somatic growth by the p160 coactivator p/CIP. Proc Natl Acad Sci USA 2000;97:13549–54. 88. Torres‐Arzayus MI, de Mora JF, Yuan J, Vazquez F, Bronson R, Rue M, et al. High tumor incidence and activation of the PI3K/AKT pathway in transgenic mice define AIB1 as an oncogene. Cancer Cell 2004;6:263–74. 89. Kuang SQ, Liao L, Zhang H, Lee AV, O’Malley BW, Xu J. AIB1/SRC‐3 deficiency affects insulin‐like growth factor I signaling pathway and suppresses v‐Ha‐ras‐induced breast cancer initiation and progression in mice. Cancer Res 2004;64:1875–85.

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90. Yan J, Yu C‐T, Ozen M, Ittmann M, Tsai SY, Tsai M‐J. Steroid receptor coactivator‐3 and activator protein‐1 coordinately regulate the transcription of components of the insulin‐like growth factor/AKT signaling pathway. Cancer Res 2006;66:11039–46. 91. Liao L, Chen X, Wang S, Parlow AF, Xu J. Steroid receptor coactivator 3 maintains circulating insulin‐like growth factor I (IGF‐I) by controlling igf‐binding protein 3 expression. Mol Cell Biol 2008;28:2460–9. 92. Qin L, Liao L, Redmond A, Young L, Yuan Y, Chen H, et al. The AIB1 oncogene promotes breast cancer metastasis by activation of PEA3‐mediated matrix metalloproteinase 2 (MMP2) and MMP9 expression. Mol Cell Biol 2008;28:5937–50. 93. Li LB, Louie MC, Chen HW, Zou JX. Proto‐oncogene ACTR/AIB1 promotes cancer cell invasion by up‐regulating specific matrix metalloproteinase expression. Cancer Lett 2008;261:64–73. 94. Shang Y, Brown M. Molecular determinants for the tissue specificity of SERMs. Science 2002;295:2465–8. 95. Su Q, Hu S, Gao H, Ma R, Yang Q, Pan Z, et al. Role of AIB1 for tamoxifen resistance in estrogen receptor‐positive breast cancer cells. Oncology 2008;75:159–68. 96. Mc Ilroy M, Fleming FJ, Buggy Y, Hill ADK, Young LS. Tamoxifen‐induced ER‐{alpha}‐SRC‐ 3 interaction in HER2 positive human breast cancer; a possible mechanism for ER isoform specific recurrence. Endocr Relat Cancer 2006;13:1135–45. 97. Shou J, Massarweh S, Osborne CK, Wakeling AE, Ali S, Weiss H, et al. Mechanisms of tamoxifen resistance: increased estrogen receptor‐HER2/neu cross‐talk in ER/HER2‐positive breast cancer. J Natl Cancer Inst 2004;96:926–35. 98. Chen B, Wang Y, Kane SE, Chen S. Improvement of sensitivity to tamoxifen in estrogen receptor‐positive and Herceptin‐resistant breast cancer cells. J Mol Endocrinol 2008;41: 367–77. 99. Goel A, Janknecht R. Concerted activation of ETS protein ER81 by p160 coactivators, the acetyltransferase p300 and the receptor tyrosine kinase HER2/Neu. J Biol Chem 2004;279:14909–16. 100. Myers E, Hill AD, Kelly G, McDermott EW, O’Higgins NJ, Buggy Y, et al. Associations and interactions between Ets‐1 and Ets‐2 and coregulatory proteins, SRC‐1, AIB1, and NCoR in breast cancer. Clin Cancer Res 2005;11:2111–22. 101. Reiter R, Wellstein A, Riegel AT. An isoform of the coactivator AIB1 that increases hormone and growth factor sensitivity is overexpressed in breast cancer. J Biol Chem 2001;276:39736–41. 102. Tilli MT, Reiter R, Oh AS, Henke RT, McDonnell K, Gallicano GI, et al. Overexpression of an N‐terminally truncated isoform of the nuclear receptor coactivator amplified in breast cancer 1 leads to altered proliferation of mammary epithelial cells in transgenic mice. Mol Endocrinol 2005;19:644–56. 103. Reiter R, Oh AS, Wellstein A, Riegel AT. Impact of the nuclear receptor coactivator AIB1 isoform AIB1‐Delta3 on estrogenic ligands with different intrinsic activity. Oncogene 2004;23:403–9. 104. Zwart W, Griekspoor A, Berno V, Lakeman K, Jalink K, Mancini M, et al. PKA‐induced resistance to tamoxifen is associated with an altered orientation of ERalpha towards co‐activator SRC‐1. EMBO J 2007;26:3534–44. 105. Kressler D, Schreiber SN, Knutti D, Kralli A. The PGC‐1‐related protein PERC is a selective coactivator of estrogen receptor alpha. J Biol Chem 2002;277:13918–25. 106. Kressler D, Hock MB, Kralli A. Coactivators PGC‐1beta and SRC‐1 interact functionally to promote the agonist activity of the selective estrogen receptor modulator tamoxifen. J Biol Chem 2007;282:26897–907.

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107. Fujimoto N, Miyamoto H, Mizokami A, Harada S, Nomura M, Ueta Y, et al. Prostate cancer cells increase androgen sensitivity by increase in nuclear androgen receptor and androgen receptor coactivators; a possible mechanism of hormone‐resistance of prostate cancer cells. Cancer Invest 2007;25:32–7. 108. Chen CD, Welsbie DS, Tran C, Baek SH, Chen R, Vessella R, et al. Molecular determinants of resistance to antiandrogen therapy. Nat Med 2004;10:33–9. 109. Louie MC, Yang HQ, Ma AH, Xu W, Zou JX, Kung HJ, et al. Androgen‐induced recruitment of RNA polymerase II to a nuclear receptor‐p160 coactivator complex. Proc Natl Acad Sci USA 2003;100:2226–30. 110. Shi XB, Xue L, Zou JX, Gandour‐Edwards R, Chen H, deVere White RW. Prolonged androgen receptor loading onto chromatin and the efficient recruitment of p160 coactivators contribute to androgen‐independent growth of prostate cancer cells. Prostate 2008;68:1816–26. 111. Balk SP. Androgen receptor as a target in androgen‐independent prostate cancer. Urology 2002;60:132–8. 112. Duff J, McEwan IJ. Mutation of histidine 874 in the androgen receptor ligand‐binding domain leads to promiscuous ligand activation and altered p160 coactivator interactions. Mol Endocrinol 2005;19:2943–54. 113. Feldman BJ, Feldman D. The development of androgen‐independent prostate cancer. Nat Rev Cancer 2001;1:34–45. 114. Ueda T, Mawji NR, Bruchovsky N, Sadar MD. Ligand‐independent activation of the androgen receptor by interleukin‐6 and the role of steroid receptor coactivator‐1 in prostate cancer cells. J Biol Chem 2002;277:38087–94. 115. Desai SJ, Ma AH, Tepper CG, Chen HW, Kung HJ. Inappropriate activation of the androgen receptor by nonsteroids: involvement of the Src kinase pathway and its therapeutic implications. Cancer Res 2006;66:10449–59. 116. Gregory CW, Fei X, Ponguta LA, He B, Bill HM, French FS, et al. Epidermal growth factor increases coactivation of the androgen receptor in recurrent prostate cancer. J Biol Chem 2004;279:7119–30. 117. Allred DC, Medina D. The relevance of mouse models to understanding the development and progression of human breast cancer. J Mammary Gland Biol Neoplasia 2008;13:279–88. 118. Sinn E, Muller W, Pattengale P, Tepler I, Wallace R, Leder P. Coexpression of MMTV/ v‐Ha‐ras and MMTV/c‐myc genes in transgenic mice: synergistic action of oncogenes in vivo. Cell 1987;49:465–75. 119. Fereshteh MP, Tilli MT, Kim SE, Xu J, O’Malley BW, Wellstein A, et al. The nuclear receptor coactivator amplified in breast cancer‐1 is required for Neu (ErbB2/HER2) activation, signaling, and mammary tumorigenesis in mice. Cancer Res 2008;68:3697–706. 120. Kuang SQ, Liao L, Wang S, Medina D, O’Malley BW, Xu J. Mice lacking the amplified in breast cancer 1/steroid receptor coactivator‐3 are resistant to chemical carcinogen‐induced mammary tumorigenesis. Cancer Res 2005;65:7993–8002. 121. Guy CT, Cardiff RD, Muller WJ. Induction of mammary tumors by expression of polyomavirus middle T oncogene: a transgenic mouse model for metastatic disease. Mol Cell Biol 1992;12:954–61. 122. Wang S, Yuan Y, Liao L, Kuang SQ, Tien JC, O’Malley BW, et al. Disruption of the SRC‐1 gene in mice suppresses breast cancer metastasis without affecting primary tumor formation. Proc Natl Acad Sci USA 2009;106:151–6. 123. Heinlein CA, Chang C. Androgen receptor in prostate cancer. Endocr Rev 2004;25:276–308. 124. Heemers HV, Tindall DJ. Androgen receptor (AR) coregulators: a diversity of functions converging on and regulating the AR transcriptional complex. Endocr Rev 2007;28:778–808.

298

HSIA ET AL.

125. Chung AC, Zhou S, Liao L, Tien JC, Greenberg NM, Xu J. Genetic ablation of the amplified‐ in‐breast cancer 1 inhibits spontaneous prostate cancer progression in mice. Cancer Res 2007;67:5965–75. 126. Greenberg NM, DeMayo F, Finegold MJ, Medina D, Tilley WD, Aspinall JO, et al. Prostate cancer in a transgenic mouse. Proc Natl Acad Sci USA 1995;92:3439–43. 127. Erzberger JP, Berger JM. Evolutionary relationships and structural mechanisms of AAAþ proteins. Annu Rev Biophys Biomol Struct 2006;35:93–114. 128. Zou JX, Revenko AS, Li LB, Gemo AT, Chen HW. ANCCA, an estrogen‐regulated AAAþ ATPase coactivator for eralpha, is required for coregulator occupancy and chromatin modification. Proc Natl Acad Sci USA 2007;104:18067–72. 129. Zou JX, Guo L, Revenko AS, Tepper CG, Gemo AT, Kung H‐J, et al. Androgen‐induced coactivator ANCCA mediates specific androgen receptor signaling in prostate cancer. Cancer Res 2009;69:3339–46. 130. Ma XJ, Salunga R, Tuggle JT, Gaudet J, Enright E, McQuary P, et al. Gene expression profiles of human breast cancer progression. Proc Natl Acad Sci USA 2003;100:5974–9. 131. Wang Y, Klijn JG, Zhang Y, Sieuwerts AM, Look MP, Yang F, et al. Gene‐expression profiles to predict distant metastasis of lymph‐node‐negative primary breast cancer. Lancet 2005;365:671–9. 132. Foekens JA, Atkins D, Zhang Y, Sweep FC, Harbeck N, Paradiso A, et al. Multicenter validation of a gene expression‐based prognostic signature in lymph node‐negative primary breast cancer. J Clin Oncol 2006;24:1665–71. 133. Desmedt C, Piette F, Loi S, Wang Y, Lallemand F, Haibe‐Kains B, et al. Strong time dependence of the 76‐gene prognostic signature for node‐negative breast cancer patients in the TRANSBIG multicenter independent validation series. Clin Cancer Res 2007;13:3207–14. 134. Fellenberg J, Bernd L, Delling G, Witte D, Zahlten‐Hinguranage A. Prognostic significance of drug‐regulated genes in high‐grade osteosarcoma. Mod Pathol 2007;20:1085–94. 135. Coste A, Antal MC, Chan S, Kastner P, Mark M, O’Malley BW, et al. Absence of the steroid receptor coactivator‐3 induces B‐cell lymphoma. EMBO J 2006;25:2453–64.

Protein Arginine Methyltransferases: Nuclear Receptor Coregulators and Beyond Peter Kuhn and Wei Xu McArdle Laboratory for Cancer Research, University of Wisconsin, Madison, Wisconsin 53706

I. Introduction ............................................................................... II. Enzymatic Activity of PRMTs ......................................................... A. PRMTs Catalyzing Asymmetric Dimethylation (Type I) .................... B. PRMTs Catalyzing Symmetric Dimethylation (Type II) .................... C. PRMTs Without Identified Activity ............................................. D. Distributive Versus Processive Mechanism .................................... E. Substrate Specificity and Regulation ............................................ F. Posttranslational Regulation of PRMTs......................................... III. PRMTs in Transcriptional Regulation ............................................... A. PRMTs as Histone Methyltransferases ......................................... B. PRMTs Involved in Nuclear Receptor Regulation ........................... C. PRMTs Regulate Transcription Mediated by Other Transcription Factors ............................................................... D. Interplay of PRMTs During Transcriptional Regulation .................... E. PRMTs in Transcription Elongation............................................. IV. PRMTs in Posttranscriptional Regulation .......................................... V. Structural Analysis of PRMTs ......................................................... VI. Small Molecule Inhibitors for PRMTs............................................... VII. Biological Functions of PRMTs ....................................................... A. PRMTs in Differentiation and Development.................................. B. PRMTs in Cancer ................................................................... C. PRMTs in Viral Infection and Immune Response ........................... D. PRMTs in Metabolism ............................................................. E. PRMTs and DNA Methylation ................................................... F. PRMTs and DNA Repair .......................................................... G. PRMTs and Chromatin Domains ................................................ VIII. Concluding Remarks .................................................................... References.................................................................................

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Protein arginine methyltransferases (PRMTs) are a family of enzymes that play a crucial role in diverse cellular functions. Several PRMTs have been associated with gene expression regulation, in which PRMTs act as histone methyltransferases, secondary coregulators of transcription, or facilitate mRNA splicing and stability. Additional functions include modulation of protein

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localization, ribosomal assembly, and signal transduction. At the organismal level, several PRMTs appear to be important for development and may play an important role in cancer. The relationships between their cellular and organismal functions are poorly understood; at least in part due to the large body of enzymatic substrates for PRMTs and their transcriptional targets that remain to be determined. Specific PRMT inhibitors have been developed in recent years, which should help to shed light on their diverse biological roles. Connecting PRMT cellular functions with their global effects on an organism will facilitate development of novel treatments for human diseases.

I. Introduction Methyltransferases are a diverse group of enzymes that modify DNA, RNA, small molecules, lipids, and proteins.1 The common aspect of this highly variable group is that many share a similar catalytic core structure utilizing a common methyl donor, S‐adenosylmethionine (AdoMet). Five classes of AdoMet dependent methyltransferases have been identified.2–4 Class I enzymes are defined by a seven strand twisted b‐sheet structure. The Class I enzymes contain four short motifs that stabilize AdoMet binding to the enzyme and facilitate methylation.5 Motif I has a highly conserved amino acid sequence VLDxGxGxG3 which is often mutated to create AdoMet binding deficient and inactive enzymes. Description of other general motifs and the protein arginine methyltransferase (PRMT)‐specific ‘‘double E’’ and ‘‘THW’’ loops can be found in previous publications.1,3 These motifs have facilitated the identification of PRMT and related enzymes.5 PRMTs are classified as Type I, Type II, Type III, or Type IV enzymes. Type I enzymes catalyze formation of o‐NG‐monomethylarginine (MMA) and asymmetric o‐NG,NG‐dimethylarginines (aDMA); type II enzymes catalyze the formation of MMA and symmetric o‐NG,NG‐dimethylarginines (sDMA); type III enzymes catalyze the formation of MMA only while type IV enzymes uniquely catalyze the formation of d‐NG‐mono‐methylarginine6 (Fig. 1). From the initial observations of methylarginine in cells,7 it took 30 years to identify the first mammalian PRMT.8 Over the last decade many PRMTs have been identified in a variety of eukaryotic species. Three PRMTs have been described in Saccharomyces cerevisiae, each having different enzymatic activities.9,10 Eleven RMT have been identified in Arabidopsis thaliana11 while nine have been identified in Drosophila.12 Up to 11 enzymes have been reported in mammals2,13 and this family of PRMTs will be the focus of this review. Evolution and phylogenetic analysis of PRMTs have been extensively reviewed.13 The schematic alignment of 10 human PRMTs is shown in Fig. 2.

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CH3 H2N

+

NH

Type I PRMTI PRMT3 CARM1 PRMT6 PRMT8

Type III PRMT7(?) HsI7(?)

+ AdoMet

NH + AdoHcy

+ AdoMet

H2 N

+

NH2

H 2N

+ COO−

NH

MMA(w)

(CH2)3 +

H 2N

+

NH2

COO− H3N Arginine + AdoMet

CH3

N

H 3N

+

+ AdoMet

COO−

CH3

+ AdoHcy

(CH2)3

NH

+ H3N COO− MMA(w)

Type IV RMT2 + AdoHcy

N

+

NH

NH Type I, II + AdoHcy + AdoMet (CH ) 2 3

(CH2)3 +

H2N

CH3

(CH2)3

H3N

CH3

+ H3N COO− aDMA(w) CH3

CH3 HN

+

NH

NH Type ll PRMT5 PRMT7 FBXO11

MMA(d )

(CH2)3 + AdoHcy + H3N COO− sDMA(w)

FIG. 1. Structures of methylated arginines in proteins. The CH3 group (bold) is transferred from AdoMet to arginine forming aDMA (type I), sDMA (type II), MMA (o) (type III), or MMA (d) (type IV). Methyltransferases are classified Type I–IV based on the methylarginine products. (Adapted from Ref. 2 with permission from Mark Bedford.)

II. Enzymatic Activity of PRMTs A. PRMTs Catalyzing Asymmetric Dimethylation (Type I) PRMT1 was the first mammalian RMT identified as the result of a yeast two hybrid screen for proteins that interact with the highly similar TIS21 and BTG1 proteins.8 PRMT1 is found mostly in the nucleus, while some PRMT1 resides in the cytoplasm.14 Recombinant PRMT activity was first demonstrated by incubation of PRMT1 with radiolabeled AdoMet and the R1 peptide (GGFGGRGGFG). This peptide contains one copy of an arginine methylation motif commonly found in the RNA binding domains of proteins involved in mRNA splicing.15,16 These RGG ‘‘boxes’’ are also called glycine arginine rich (GAR) motifs. Additionally found in non‐RNA‐binding proteins, GAR motifs are frequently the sites of PRMT methylation. PRMT1 methylation substrates

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C

D

E

PRMT1/HRMT1L2 PRMT2/HRMT1L1 PRMT3 PRMT4/CARM1 PRMT5/JBP1 PRMT6

AB C

D

E

PRMT7 PRMT8

myr

FBXO11/PRMT9 PRMT10/9 A: Motif 1 VLD/EVGxGxG B: Post 1 V/IxG/AxD/E C: Motif 2 F/I/VDI/L/K D: Motif 3 LR/KxxG E: THW loop

Type I

SH3 domain

Type II No activity

TPR repeat

Zn finger

F box

High similarity Low similarity

FIG. 2. Schematic alignment of 10 human PRMTs. The conserved motifs and THW loop are shown as black box. Other unique structural features are indicated in graph. (Adapted from Ref. 2 with permission from Mark Bedford.)

do include variations of the RGG box,17 and PRMT1 also methylates RxR sequences18 and even highly divergent sequences.19–21 Being the predominant type I enzyme, PRMT1 is responsible for 85% of cellular aDMA formation22 and is involved in diverse cellular processes. PRMT3 was first identified as a PRMT1 binding protein in a yeast two hybrid screen.23 PRMT3 contains a unique extended N‐terminus that is highly acidic and contains a C2H2 zinc finger motif. Deletion of the N‐terminal domain significantly reduces PRMT3‐mediated methylation of the GAR motif and other RNA‐associated PRMT3 substrates.24 Found in the cytoplasm, PRMT3 appears to be primarily important for regulation of ribosomal protein levels25,26 but may also be important in apoptotic signaling.27 Coactivator‐associated RMT 1, CARM1/PRMT4, was first identified in a yeast two hybrid screen in search of glucocorticoid receptor interacting protein 1 (GRIP1) interacting proteins.28 CARM1 is somewhat unique from other type I PRMTs in that it does not methylate substrates containing a GAR motif.29 CARM1 is a transcriptional coactivator for a number of transcription factors, and is also important for mRNA processing. Consistent with these functions, CARM1 is predominantly nuclear, although cytoplasmic CARM1 has been observed in prostate tumors.30

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PRMT6 was first identified by the gapped BLAST method searching for PRMT family members.14 PRMT6 can methylate the GAR motif, exhibits some overlap with PRMT1 substrate specificity, and significant automethylation. PRMT6 is predominantly found in the nucleus, and appears to play a role in chromatin structure and viral infection.31–33 PRMT8 was first identified in a screen for genes selectively expressed in neural progenitor cells.34 Further studies indicate that PRMT8 is exclusively expressed in brain tissue.35 PRMT8 localization is further limited to neurons in specific regions of the brain, including the somatosensory, limbic, and motor systems.36 PRMT8 exhibits type I methylation of the GAR motif and localizes to the plasma membrane bound by an N‐terminal myristoylation.35 The N‐terminus of PRMT8 appears to negatively regulate methyltransferase activity in vitro as well as facilitate binding to SH3 domains through two proline rich (PR) domains.37 PRMT2, containing an SH3 domain, binds to PRMT8 in GST pull‐down assays. PRMT8 is also able to specifically heterodimerize with PRMT1,35 in analogy to PRMT1 and PRMT3 heterodimers.23 PRMT8 binds to numerous common PRMT substrates including RNA binding proteins, hnRNPs, and RNA helicases, as well as some unique proteins including actin, tubulin, and hsp70.38

B. PRMTs Catalyzing Symmetric Dimethylation (Type II) PRMT5 was originally identified by yeast two hybrid as a JAK2 kinase interacting protein and thus originally called Jak binding protein 1(JBP1).39 PRMT5 was the first type II RMT identified. Consistent with other PRMTs, motif I of PRMT5 is necessary for its enzymatic activity.40 PRMT5–GFP fusion proteins initially suggested that PRMT5 resides primarily in the cytoplasm.14 However, biochemical fractionation indicates both the nuclear and cytoplasmic localization.41 Localization may be cell type or stage‐specific as both nuclear and cytoplasmic PRMT5 is observed exclusively at different developmental stages of the testes.42 PRMT5 currently has two major functions which coincide with its locations. First, it contributes to complex assembly of ribonucleoproteins in the cytoplasm.43 Second, PRMT5 plays an important role in transcriptional regulation, either through histone modifications44,45 or interactions with RNA Polymerase II elongation factors.46 PRMT7 was first identified by the gapped‐BLAST scanning of GenBank for protein sequences similar to S‐AdoMet binding domains. PRMT7 contains two PRMT domains, each having four conserved methyltransferase motifs. Deletion mutants containing only one domain do not exhibit independent activity.47 This initial report observed only MMA product, prompting the authors to assign a type III classification to PRMT7. Subsequent work has demonstrated symmetric dimethylation of PRMT7 substrates under lower substrate concentrations.48 If PRMT7 utilizes a distributive methylation mechanism, limited

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incubation with high substrate concentration would result in monomethylation due to the low likelihood of the enzyme finding the same substrate molecule twice. Since PRMT7 has the ability to catalyze a type II reaction, this raises the question whether type III methylation is an intrinsic property of an enzyme or it merely describes an incomplete methylation event. To our knowledge, no mechanism has been described to restrict RMTs to catalyze only MMA. Another proposed type III RMT Hsl7 has also recently been shown to catalyze symmetric dimethylation.49 As the second identified type II PRMT, PRMT7 methylates many similar substrates to PRMT5.48 Interestingly, a small amount of aDMA was also observed when PRMT7 methylates peptide substrates, suggesting that this enzyme may have less specificity for substrate orientation in the substrate binding pocket or the dual PRMT domains may have separate enzymatic activities.48 No Type II PRMT crystal structure has been solved thus far. Two proteins have been assigned as PRMT9 by different research groups. One PRMT9 (located at human chromosome 2p16) is alternatively named F‐box only protein 11 (FBXO11), which was identified as a putative PRMT by a search for proteins that possess motif I, however its primary structure is very different from that of PRMT1–8.50 PRMT9/FBXO11 was found to catalyze formation of MMA and sDMA, as well as a lesser amount of aDMA, and thus is classified as a type II PRMT.

C. PRMTs Without Identified Activity PRMT2 was first identified as HRMT1L1 by chromosomal transcription mapping of Chr.2151,52 and is predominantly found in the nucleus.14 PRMT2 uniquely contains an N‐terminal SH3 domain thought to play a role in protein– protein interactions. While PRMT2 does bind to AdoMet and the AdoMet binding domain is necessary for PRMT2‐mediated coactivation,53 it has not been shown as an active enzyme. Thus, while PRMT2 regulates transcription of nuclear receptors,53,54 the mechanism of activation is not clear. Another PRMT9 (located at human chromosome 4q31) named by Bedford group was described as PRMT10 by Krause et al.13,55 This PRMT encodes a protein with 835 amino acids. PRMT9/10 harbors two putative AdoMet‐binding motifs and a tetratricopeptide repeat domain (TPR) at its N‐terminal end. It resembles PRMT7 phylogenetically and structurally, although its enzymatic activity has not been elucidated.

D. Distributive Versus Processive Mechanism The mechanism by which PRMTs mediate dimethylation on arginine is still under debate. A distributive mechanism is performed by an enzyme that covalently attaches a single modification then releases the substrate. In contrast, a processive mechanism is defined by the enzyme’s ability to maintain

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interaction with the substrate until multiple covalent modifications have been achieved. Initial studies of peptide methylation suggested that PRMT7 produces more monomethylated peptides as substrate concentration increases, supporting a distributive mechanism.48 Recent PRMT1 kinetic studies give evidence for a partial processive mechanism.56,57 PRMT6 exhibits a distributive mechanism on peptide substrates.58 We find that CARM1 automethylation occurs via a processive mechanism (Kuhn and Xu, submitted for publication). The observed mechanism may be affected by the size of the substrate (i.e., proteins offer more docking sites than short peptides, thus are more likely to present a processive mechanism).

E. Substrate Specificity and Regulation The role of surface residues distal to the active site in PRMT substrate specificity remains poorly defined. PRMT1 has exhibited lower activity for peptides in comparison to proteins with similar recognition sites,59 supporting the notion that an extended substrate structure enhances methyltransferase activity. Individual mutation of several surface residues in PRMT1 alters methyltransferase activity, although the specific role of these residues in the enzyme is undefined.60 The methylation of H3R2 and H3R26 by CARM1 was recently attributed to the presence of CARM1 N‐terminal domain.61 This was in contrast to H3R17 methylation, which only requires the central catalytic core. While the mechanism defining PRMT substrate specificity is not completely understood, a number of studies indicate PRMT substrate recognition is highly variable and context dependent. There is some amino acid sequence disparity between type I and type II PRMTs proximal to the active site. In spite of this, GST‐GAR, histone H4R3, and MBP are methylated by both type I and type II enzymes.62–64 Other substrates such as EWS and PABPN1 are highly methylated by multiple type I or type II PRMTs, respectively,65,66 exhibiting partial overlap in specificity. This combination of specificity and redundancy suggests that substrate recognition is not solely determined by the primary substrate structure. Further support comes from the lack of an apparent recognition motif among exclusive CARM1 substrates.61 Thus, the secondary or tertiary structure of recognition sites may be more important for specificity of an enzyme like CARM1. The apparent overlap of substrate specificity within or between type I and type II PRMTs may also be differentiated in vivo by PRMT interacting proteins and subcellular localization. Significant evidence exists for the role of protein binding partners in PRMTs mediate methylation. PRMT1 is regulated by multiple substrate binding partners. TIS21 and BTG1 bind directly to PRMT1 and significantly regulate PRMT1‐mediated methylation of substrates in rat cell lysates.8,23 BTG1 was further shown to regulate PRMT1‐mediated methylation in a substrate dependent manner, facilitating histone H4 but inhibiting methylation of a Sam68 peptide.67 The BTG1

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interacting protein hCAF also binds to PRMT1 and decreases methylation on selective substrates. BTG2 binds to PRMT1 in neurons and knockdown of BTG2 reduces asymmetric dimethylation.68 PRMT3 methylation of a GAR motif is inhibited by interaction with the tumor suppressor DAL‐1/4.1B.25 PRMT3 also forms a complex with rpS2, which increases the methyltransferase activity of PRMT3.69 PRMT7 exhibits independent and specific activity in vitro, but the addition of CTCFL increases PRMT7 activity significantly.70 Finally, while PRMT8 activity in vitro is significantly lower than that of PRMT1, both proteins show significant and similar levels of activity when incubated with cell lysates.38 These results suggest that other cellular factors contribute to the activity of one or both proteins, further supporting at least partial dependency of PRMTs on cofactor binding.

F. Posttranslational Regulation of PRMTs While PRMTs are characterized for their ability to modify proteins, the posttranslational modifications on PRMTs have gone overlooked until very recently. Automethylation has been observed on several PRMTs but the mechanism is poorly understood. Initial experiments with GST‐PRMT6 indicate that it has a high automethylation activity in vitro but the specific site and function of automethylation is undefined.14 PRMT8 also exhibits automethylation activity, asymmetrically dimethylating two residues in the GST tag (raising concerns about PRMT6 automethylation), as well as dimethylating R73 and monomethylating R58.37 The functional role of PRMT8 automethylation has not been determined, although it is interesting to note that the monomethylated R58 residue is located in the PR domain II, which is necessary for PRMT8 association with SH3 domains. The automethylation of other GST‐PRMTs was demonstrated to be minimal, with weak signals for PRMT1 and CARM1. Weak automethylation by CARM1 has been observed previously.71 Recent work in our laboratory suggests that the lack of observed CARM1 automethylation in vitro may be due to near saturated automethylation occurring in vivo prior to analysis (Kuhn and Xu, submitted for publication). Thus, PRMT automethylation may be more widespread than previously thought. Posttranslational modification of PRMTs as a regulatory mechanism in signal transduction has recently emerged. CARM1 was recently shown to be phosphorylated at S229 during mitosis.72 A serine to glutamic acid (Glu or E) mutation mimicking phospho‐CARM1 blocks homodimerization and abrogates both the methyltransferase and estrogen receptor (ER) activating functions of CARM1. This highlights the importance of dimerization in CARM1 function and represents the first example of regulation of PRMT activity by a posttranslational modification.

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III. PRMTs in Transcriptional Regulation A. PRMTs as Histone Methyltransferases Histone modifications were first observed over 45 years ago73 and are a major research focus today. The temporal and conditional nature of many histone modifications indicates that these marks are highly context dependent, where the presence of one histone modification can either inhibit or enhance modification of a different histone residue. This complex interdependency of histone modifications has been termed the ‘‘histone code.’’ Significant work has established that these modifications function like a regulatory cascade, each mark dependent on others for their appropriate function.74,75 This model is well established; however our ability to read this code is still limited. Lysine acetylation and methylation are the predominate focus of those studying chromatin remodeling and epigenetic signaling, whereas the role of arginine methylation has received considerably less attention.76,77 Histone modification by PRMT family members results in both the activation and repression of gene expression. Although the mechanism of dynamic arginine methylation and demethylation is still poorly defined, arginine methylation is dependent on and essential for other types of histone modifications. 1. HISTONE MODIFICATION CATALYZED BY TYPE I PRMTs PRMT1 has been shown to methylate histone H4R3 and loss of PRMT1 leads to a loss of detectable H4R3 methylation.78,79 However, further research of other PRMTs suggests that H4R3 is a target substrate for multiple PRMTs, complicating the function of this mark considerably. In association with PRMT1, H4R3 methylation is an activating mark for transcription, facilitating subsequent acetylation by p300.64,80 However, preacetylation of H4 is inhibitory for R3 methylation by PRMT1. SE translocation (SET) protein protects H4 from acetylation, and this hypoacetylation is essential for PRMT1‐mediated histone methylation.81 Thus, the presence or absence of acetylation is not an absolute determinate of transcriptional activation, as acetylation may be associated with transcriptional activation or repression in the context of other histone modifications and chromatin‐associated proteins. This highlights the need to examine acetylation (and methylation) of specific histone residues, since the mere presence or absence of nonspecific acetylation does not determine gene expression. CARM1 was the first PRMT identified as a histone methyltransferase.28 CARM1 was originally identified to methylate histone H3 at R2, R17, and R26, with R17 and R26 being the predominant sites for methylation.29 Additionally, CARM1 appears to methylate one or more arginine residues in the C‐terminal end of the H3 tail.29 Methylation at H3R17 is often associated with CARM1

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activation of nuclear receptors and other transcription factors.82,83 CARM1‐ mediated histone modifications also interact with other chromatin marks. Acetylation by CBP at K18 precedes acetylation at K23, and these modifications are necessary for CARM1 methylation at R17.84 Kinetic analysis shows that the acetylation of K18 potentiates R17 methylation of H3 tails by increasing the catalytic rate of H3R17 methylation without altering CARM1‐histone peptide binding affinity.61 While histone modifications often alter the binding affinity for effectors and other histone modifiers, the interaction between H3K18 acetylation and CARM1 activity appears to be somewhat unique. Sequential histone modification appears to facilitate a synergistic activation by PRMT1 and CARM1. In vitro transcription has demonstrated that the ordered addition of PRMT1, p300, and CARM1 significantly affects transcriptional activity.85 The acetylation by CBP/p300 acts a bridge between PRMT1 and CARM1 modifications, conveying the activation signal initiated by PRMT1 to direct CARM1 activity. The mechanism by which this interaction occurs is unknown, but does not appear to be facilitated by additional effector proteins as these are not present in an in vitro transcription system. While CARM1 was previously shown to methylate H3R2 as a minor substrate, recent evidence suggests that PRMT6 serves as the primary enzyme modifying this site.31,86,87 Methylation of H3R2 is mutually exclusive with H3K4 methylation by a mixed lineage leukemia (MLL) complex in vivo. Increased expression of PRMT6 and concomitant H3R2 methylation inhibits gene expression activated by MLL methylation of H3K4. Thus, in contrast to PRMT1 and CARM1, asymmetric dimethylation by PRMT6 can act as a transcriptional repressor. The mechanism of inhibition is due to steric hindrance of enzyme binding, as in vitro methylation studies indicate that PRMT6 is capable of methylating trimethylated H3K4 but to a lesser extent than mono or dimethylated H3K4, and methylation of H3R2 alters the binding surface recognized by MLL.87 2. HISTONE MODIFICATION CATALYZED BY TYPE II PRMTs PRMT5‐mediated symmetric dimethylation is usually a repressive mark on histones. Initially demonstrated on the Cyclin E1 promoter through association with the cyclin E1 repressor complex (CERC), PRMT5‐mediated histone methylation at H4R3 decreases gene expression and negatively regulates cell cycle progression.39,40,88 This phenomenon is further supported by the PRMT5‐mediated repression of the C‐myc target gene CAD and the tumor repressors ST7 and NM23 via methylation at H3R8 and H4R3.44,45 That methylation of H4R3 by PRMT1 and PRMT5 gives rise to different transcriptional outcomes indicates that one modification of one histone residue can signal either transcriptional activation or repression. The differential regulation

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that occurs through methylation of the same site further indicates that histone modifications are highly dependent on their cellular context, likely influenced by surrounding modifications, effector proteins, and local chromatin structure. Type I and Type II methylation do not appear to be inherently activating or inhibiting signals. Type I methylation results in both transcriptional activation and repression as discussed above. Additionally, even a specific PRMT enzyme may not be committed to be an activating or repressing entity, as PRMT5 can also activate gene expression. Methylation of H3R8 by PRMT5 corresponds with activation at the myogenin promoter during muscle differentiation, and PRMT5 expression is necessary for this activation.89 However, it is unclear whether PRMT5‐mediated histone methylation accounts for gene activation, as methylation of other coregulators by PRMT5 may also play a role.90,91 Another histone substrate for PRMT5 is H2AR3. This residue is dimethylated in concert with H4R3 during the regulation of primordial germ cell (PGC) differentiation, and appears to play a repressive role with H4R3 in this context.92 PRMT1 is also able to methylate H2A,22,64 although the function and importance of H2A methylation by PRMT1 is still unclear. It has been proposed that this also occurs at H2AR3 as the primary sequence surrounding R3 on H4 and H2A is similar and both are recognized by a H4R3 methyl specific antibody.64 Mutation of the H4R3 to glutamine (Q) also results in a significant increase in H2A methylation by PRMT1.85 PRMT1‐mediated activation of in vitro transcription is not significantly reduced using this H4R3Q mutant, suggesting that H2A methylation may have some redundant function with H4R3 methylation. PRMT7 also catalyzes symmetric dimethylation on histones.70 PRMT7 methylates H2A and H4 alone in vitro, and the addition of CTCFL significantly increases the methylation of both substrates. An H4R3Me2 antibody recognized H4 incubated with PRMT7, further expanding the number of PRMTs recognizing this arginine residue. FBXO11/PRMT9 (2p16) was recently identified as a novel and structurally distinct type II methyltransferase that contains an F‐box and a Zn finger motif.50 In vitro methylation studies show that PRMT9 methylates H2A and H4 but the specific residues and role of these modifications has yet to be assessed. 3. HISTONE ARGININE DEMETHYLATION Until recently protein methylation was thought to be a permanent mark, and loss of methylation was only due to protein degradation and turnover. Peptidyl deiminase 4 (PAD4) was the first enzyme identified to remove methylarginine residues from histones, resulting in a physiologically relevant citrulline residue.93,94 Deimination by PAD4 prevents arginine methylation by CARM1 and inhibits ER target gene pS2 expression by removing the activating histone marks. This dynamic regulation is not limited to histone targets, as

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PAD4 can also deiminate p300 on residues methylated by CARM1.95 While PAD4 activity does not constitute a completely reversible mechanism, deimination suggested that arginine methylation is not a permanent mark. Jumonji (JmjC) domain containing proteins that demethylate lysine residues established a highly reversible and dynamic model for epigenetic regulation. While the identification of the JmjC domain presented researchers with a seemingly good list of sequence handles with which to search for the arginine demethylase, an entire family of lysine demethylases was identified before the long‐awaited JMJD6 was identified.96 In analogy with the lysine demethylases, JMJD6 only demethylates designated arginine residues: JMJD6 demethylates H3R2 and H4R3 but not the prominent CARM1 methylation sites, H3R17 and H3R26.96 JMJD6 prefers to demethylate dimethylated arginines over those that are monomethylated. JMJD6 is able to demethylate either symmetric or asymmetric dimethylation sites, although symmetric demethylation gave a higher catalytic rate. The discovery of JMJD6 demonstrates arginine methylation is dynamically regulated and completely reversible, opening new avenues for epigenetic regulation. The net effect of combinatorial histone modification is now just emerging. As novel technologies arise, more combinatory histone marks and modification enzymes are emerged. For example, novel mass spectrometry approaches can detect over 150 different combinations of modifications on histone H3.2 alone.97 This enormous complexity indicates that future models of transcriptional regulation will have to consider the combinatory ‘‘histone code’’ (i.e., the dependence of one modification on another is the central theme of histone‐ mediated transcriptional regulation).

B. PRMTs Involved in Nuclear Receptor Regulation 1. CARM1 AS A MODEL COACTIVATOR FOR NUCLEAR RECEPTOR TRANSCRIPTION Initially identified as a GRIP1 interacting protein, CARM1 has been shown to play an important role in the transcriptional activation of multiple nuclear receptors including ERa, androgen receptor (AR), thyroid hormone receptor (TR),28 and retinoid acid receptor (RAR),98 and this activation is dependent on CARM1’s methyltransferase activity. CARM1 association with these nuclear receptors required the presence of the C‐terminal domain of GRIP1 as a binding intermediate between CARM1 and the nuclear receptor. PRMT1 was then shown to have a similar transactivation activity for these nuclear receptors and could synergistically enhance coactivation in cooperation with CARM1.99 This synergistic coactivation was also extended to the orphan

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nuclear receptors ERRa, ERRb, and ERRg.99 These initial studies defined PRMTs as secondary coactivators for nuclear receptors and set the stage for the significant discoveries to follow. The synergy among PRMTs and other nuclear receptor coactivators are best defined by studies of CARM1 promoting expression of hormone response element (HRE) containing reporter constructs. The histone acetyltransferase (HAT) p300 and CBP bind cooperatively to the activation domain 1 (AD1) of GRIP1 and further contribute to CARM1‐mediated transcriptional coactivation.100 While direct interaction with GRIP1 has not been examined, the HAT enzyme PCAF synergistically activates transcription with CARM1 in a GRIP1 dependent manner on the MMP‐9 promoter.101 The SAM binding domain is not the only region important for CARM1‐ mediated transcriptional activation. CARM1 deletion mutants have shown that the C‐terminal domain is also essential for transcriptional coactivation of an ER reporter, even though this region is dispensable for CARM1’s methyltransferase activity and specificity.102 This suggests that the C‐terminus of CARM1 facilitates protein–protein interactions in the coactivator complex assembly, but few C‐terminal binding partners have been identified. 2. COACTIVATORS METHYLATED BY CARM1 In addition to modifying histones, CARM1 has also been shown to methylate CBP directly in its KIX domain, blocking CBP binding to CREB and repressing CREB‐mediated transcription.71 CARM1 also methylates R714, R742, and R768 residues on CBP.103 Mutation of these three sites creates a dominant negative CBP, which does not coactivate GRIP1‐mediated transcription but can inhibit coactivation of an ER reporter. Finally, CARM1 was shown to methylate two residues (R2088 and R2142) in the GRIP1 binding domain of p300, inhibiting the interaction between p300 and GRIP1.95 The highly similar interface between CBP and a p160 protein ACTR exhibits synergistic folding, including the formation of a salt bridge between ACTR and CBP residue R2105 (R2088 in p300).104 Methylation of this site would block formation of the salt bridge, reducing CBP/ p300 binding with p160 proteins. Thus, in addition to the synergistic coactivation of transcription by CBP/p300 and CARM1 through histone modifications, these results provide evidence for a coregulator code where the posttranscriptional status of these coregulators affects their function. The role of CARM1’s methyltransferase activity in its interaction with the p160s is complex. Although CARM1 binding to GRIP1 is not significantly altered by the methyltransferase deficient E267Q mutation, cell‐based reporter assays indicate the CARM1 methyltransferase defective mutant exhibits a reduction in nuclear receptor gene activation when expressed with either b‐catenin or GRIP1.100,105 More recently it has been determined that SRC3/ ACTR is methylated by CARM1 proximal to the CARM1 binding domain,

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AD2, at residues conserved in p160 proteins.106,107 Methylation appears to decrease the binding affinity between CARM1 and SRC3, thus possibly serving to facilitate complex disassembly.106 SRC3 methylation also increases SRC3 degradation.107 Both of these are consistent with the observed increase in transcriptional coactivation by hypomethylated mutant SRC3. The increase in p300 binding to hypomethylated SRC3 mutants over wild-type SRC3106 and the increase in CBP binding to SRC3 in the absence of CARM1107 indicate that CARM1 methylation of SRC3 promotes coactivator complex disassembly. Thus, while the methyltransferase activity has been shown to be important for CARM1‐mediated gene expression, the intrinsic mechanisms by which CARM1 modulates the transcription are complex. 3. THE NONSUBSTRATE CARM1 INTERACTING PROTEINS In addition to CARM1’s methyltransferase activity, coregulator interactions also facilitate transcriptional activation. CARM1 binds to a nucleosomal methylation activator complex (NUMAC) made up of components of the Swi/Snf complex through a direct interaction with Brg1.98,108 This interaction alters the substrate specificity of CARM1 such that CARM1 in NUMAC preferentially methylates nucleosomes over free histones. The presence of CARM1 also increases Brg1’s ATPase activity in this complex, facilitating ER target gene expression.108 Another CARM1 interacting coregulator is the secondary coactivator TIF1a. Coexpression of CARM1 and TIF1a induce synergistic activation of reporter constructs for AR, ER, TR, and GR in a GRIP1 dependent fashion.109 This work again highlights the importance of the C‐terminal domain of CARM1 in transcriptional coactivation, as this region was sufficient for activation in a Gal4 reporter system. While CBP/p300, PRMT1, and CARM1 all bind to the C‐terminal domain of GRIP1, TIF1a bind to the N‐terminus of GRIP1. Additionally, TIF1a is the first coactivator that may interact with the transcriptionally essential C‐terminus of CARM1. Flightless‐I (Fli‐1), identified as a CARM1 interacting protein, is a nuclear receptor coactivator for TR and ER.110 Promoter binding by Fli‐1 occurs in concert with ERa, the p160 ACTR, CARM1, and p300. Like TIF1a, Fli‐1 binds to the N‐terminal domain of GRIP1 and synergistically coactivates a TR reporter with CARM1 and GRIP1. The lack of evidence of synergy between Fli‐1 or TIF1a and p300 suggests that while these proteins may reside in the same coactivator complex, they do not interact directly. The binding of Fli‐1 to the actin‐like Brm/Brg‐associated factor 53 (BAF53) is important for Fli‐1‐ mediated ER target gene expression.110 This observation, together with the finding that CARM1 associates with Swi/Snf components,108 supports the putative activation role of actin or actin‐related molecules in other chromatin remodeling complexes.111

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GRIP1 N‐terminal binding protein GAC63 has also been identified as nuclear receptor coactivator that significantly enhances CARM1‐mediated transcription.112 GAC63 is recruited to the pS2 promoter with ERa and GRIP1 in a ligand‐dependent manner. The N‐terminus of GAC63 containing a zinc finger domain and a central region with two leucine zippers both independently bind to bHLH‐PAS domain at N‐terminus of GRIP1, while the C‐terminal region of GAC63 is necessary for transcriptional activation in a GAL4 reporter system. Consistent with other GRIP1 N‐terminal binding proteins, GAC63 exhibits selective synergistic activation with CARM1 and GRIP1, but not with p300. Although a number of secondary coactivators bind to the N‐terminus of GRIP1 to enhance CARM1‐mediated gene expression, the mechanism of activation has not been elucidated. Coiled coil coactivator (CoCoA), another secondary nuclear receptor coactivator mediated by GRIP1, acts synergistically with CARM1 to facilitate transcription.113 CoCoA’s central coiled coil domain containing three leucine zippers interacts with the GRIP1 bHLH‐PAS. In contrast to the other GRIP1 N‐terminal binding proteins, there is evidence of functional interaction between p300 and CoCoA in the coactivator complex.114 CoCoA binds directly to the KIX and CH3 regions of p300 through its C‐terminal AD independent of GRIP1. Absence of the CoCoA AD abrogated CoCoA‐mediated reporter gene activation in the presence of CARM1 and p300. Methylation of H3K9 by G9a has been associated with recruitment of effector protein heterochromatin binding protein 1 (HP‐1) to repress gene transcription. G9a can also act as a transcriptional coactivator, acting synergistically with the GRIP1–CARM1–p300 complex to activate nuclear receptors including ER and AR.115 Synergistic activity is specific to CARM1, as PRMT1, ‐2, ‐3, and RMT1 do not cooperate with G9a. Additionally, siRNA targeting G9a in the prostate cancer cell line LNCaP reduces AR‐mediated expression of prostate specific antigen (PSA) mRNA and protein.115 In its conventional role as a transcriptional repressor, G9a opposes CARM1‐mediated activation of the TR target gene type 1 deiodinase (D1).116 CARM1 and the Swi/Snf complex component 5 (SNF5) are both necessary for T3‐mediated activation of the D1 promoter. Loss of CARM1 or SNF5 induces an increase in G9a binding to the repressed promoter, a reversed event of T3 treatment. Thus, the interaction between CARM1 and G9a in transcriptional coregulation is context specific. 4. OTHER PRMTS–NUCLEAR RECEPTOR INTERACTIONS PRMT1 serves as a coactivator for orphan nuclear receptor hepatocyte nuclear factor 4 (HNF4). HNF4 is expressed in liver, intestine, kidney, and pancreas, and plays a role in development, differentiation, and metabolism.

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Activation of HNF4 by PRMT1 depends on both conventional and novel mechanisms. In a conventional way, PRMT1 acts synergistically with the coactivators SRC‐1 and p300 to mediate HNF4 expression.117 The novel mechanism utilized by PRMT1 is via methylation of HNF4 to affect its DNA binding affinity. This methylation of the DNA binding domain increases the affinity between HNF4 and chromatinized DNA, thus acting as an additional activation signal from PRMT1. PRMT1 and CARM1 are also transcription coactivators for an orphan nuclear receptor farnesoid X receptor (FXR).118,119 Increased methylation of H3R17 and H4R3 and acetylation of H3K9 occurs in a methyltransferase dependant manner. CARM1‐mediated activation of an FXR reporter is dependent on the presence of SRC‐1, while the mode of recruitment of PRMT1 on FXR‐target genes remains to be determined. Recently, a nongenomic interaction between ERa and PRMT1 was identified. In addition to a p160‐mediated interaction between PRMT1 and ERa in the nucleus, PRMT1 directly binds to ERa and methylates R260 site.120 This occurred exclusively in cytoplasm, in a rapid, transient manner in response to 17b‐estradiol treatment. Loss of ERa methylation inhibits the formation of the ERa/Src/FAK/PI3K complex and blocks PI3K‐mediated phosphorylation of AKT, ultimately results in a change in cell cycle progression. PRMT2 interacts with a number of nuclear receptors including ERa, ERb, TRb, PR, RARa, PPARg, and RXRa in a ligand independent manner whereas it is a ligand dependent coactivator for ERa.53 While PRMT2 does not appear to have methyltransferase activity in vitro, the mutation of the SAM binding site reduces its coactivation of an ERa reporter. The PRMT2 C‐terminal domain also interacts directly with AR in a ligand dependent manner.54 Similar to ERa coactivation, the PRMT2 SAM binding domain is necessary for transcriptional activation of an AR reporter. The apparently unique ability of PRMT2 to bind directly to multiple nuclear receptors and its lack of methyltransferase activity in vitro suggests that more studies are needed to delineate the intricacies of PRMTs mediate transcriptional regulation.

C. PRMTs Regulate Transcription Mediated by Other Transcription Factors 1. PRMT1 PRMTs can regulate gene expression through the direct modification of transcription factors. FOXO1 is a transcription factor whose stability is determined by subcellular localization. PRMT1 is the primary enzyme catalyzing FOXO1 methylation at two arginine residues in its nuclear localization sequence.121 Depletion of PRMT1 results in cytoplasmic localization and degradation of FOXO1, indicating that PRMT1 methylation stabilizes FOXO1.

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Consistent with these results, the activity of FOXO1 is decreased after PRMT1 depletion. Thus, in addition to indirect regulation of transcription through histone and coregulator methylation, arginine methylation can regulate the activity of transcription factors directly through modulation of their posttranslational status. 2. CARM1 Like its binding partner p160 and p300/CBP, CARM1 is widely utilized by a variety of transcription factors. The recent identification of a number of CARM1 regulated transcription factors and the cooperation between CARM1 and PRMT1 suggests that PRMT family members have multifaceted functionality. The transcription factor myocyte enhancer factor‐2 (MEF‐2) is regulated by p160s and p300 as well as CARM1. Initially surprising, the observation that CARM1 binds directly to MEF‐2 instead of indirectly through a p160 protein is supported by later studies.122 GRIP1 and CARM1 bind to the N‐ and C‐terminus of MEF‐2, respectively. In a Gal4 reporter system, the activation by CARM1 was shown to be GRIP1 dependent, indicating that although CARM1 binds MEF‐2 independently of GRIP1, CARM1 still relies on GRIP1 for coactivation. This extends the dependence of CARM1 on p160 proteins in transcription activation beyond the known bridging effect. The transcription factor complex AP‐1 is a heterodimer of basic region leucine zipper (bZIP) proteins. Members of the Fos protein family are often found in AP‐1 complexes and regulate expression of matrix metalloproteinase (MMP) genes. SiRNA‐mediated depletion of endogenous c‐fos and p160s in CARM1 expressing MEF cells reduces mRNA levels of MMP‐1b, MMP‐3, and MMP‐13.123 Depletion of either c‐fos or p160s in CARM1
/
MEFs further inhibits the mRNA levels of these three MMP family members. While CARM1 appears to act synergistically with the p160 proteins in a Gal4‐c‐fos reporter system, ectopic expression of any p160 was not necessary for CARM1‐ mediated coactivation or coimmunoprecipitation with c‐Fos. This is analogous to the additive but independent relationship between GRIP1 and CARM1 on MEF2C‐mediated transcription.122 3. PRMT5 PRMT5 was initially identified to be important for the assembly of splicing factors and to possibly play a role in the regulation of cell signaling pathways, both of which involve cytoplasmic localization. However, more recently PRMT5 has been identified as an important transcriptional regulator. In contrast to PRMT1 and CARM1, PRMT5 interacts with a different set of binding partners and usually facilitates repression of gene expression.

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PRMT5 recruits BRG1‐ and BRM‐associated chromatin remodeling factors to target genes.44 In contrast to the CARM1‐associated Swi/Snf activator complex, the PRMT5‐associated remodeling complex contains mSin3A and histone deacetylase 2 (HDAC2), which generally facilitate transcriptional repression.108 The PRMT5 complex exhibits a higher methylation activity on deacetylated histones, suggesting that the histone modifiers work in concert to repress transcription. A Smad2, 3, 4, HDAC, and PRMT5 interaction was also observed in Smad7 repression in the TGF‐b signaling pathway.124 MEP50 is another binding protein for PRMT5 that appears to enhance its methyltransferase activity.13 MEP50 may also facilitate the association between PRMT5 and specific substrates. The C‐terminal domain of MEP50 is able to bind specifically to histone H2A, a known target for PRMT5.125 Furthermore, transcriptional repression of a Gal4 reporter construct by PRMT5 is dependent on the presence of MEP50. PRMT5 and MEP50 were also shown to be involved in SNAIL‐mediated repression of E‐cadherin.126,127 Cooperator of PRMT5 (COPR5) is a PRMT5 binding partner that mediates interactions between PRMT5 and histone H4.128 Exogenous PRMT5 precipitated from U2OS cells exhibit a substrate preference for H4R3 over H3R8, while PRMT5 precipitated from cells with siRNA‐disrupted COPR5 show a shift toward preference for H3R8. COPR5 knockdown also reduces PRMT5 binding to the cyclin E1 promoter and induces an increase in cyclin E1 expression. This result suggests COPR5, and perhaps other PRMT5 binding partners, can regulate its substrate specificity and expression of a subset of its target genes.

D. Interplay of PRMTs During Transcriptional Regulation The posttranslational modification of histones by individual PRMTs has been discussed above. There is also evidence that multiple PRMTs act on the same transcription factors, either collaborating with or opposing each other. For example, the combination of CARM1 and PRMT1 synergistically activate a number of nuclear receptors.99 This section highlights the increasingly complex and significant role of PRMTs in gene expression. 1. p53 PRMT1 and CARM1 are both p53 coactivators in in vitro transcription assays.85 In contrast to their role as secondary coactivators for nuclear receptors, PRMT1 and CARM1 directly bind to p53. PRMT1 methylation of H2A can facilitate partial coactivation activity by PRMT1 in the presence of an unmethylatable H4 mutant.85 This result suggests that transcription activation couples with methylation of H3 and H4/(possibly H2A) by CARM1 and

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PRMT1, respectively. This synergistic activation of p53 mediated transcription is dependent on the presence of p300, and occurs in an ordered manner where addition of PRMT1 and p300 prior to CARM1 yields the highest transcription yield than other sequential combinations. Demonstrating the multifunctional character of the PRMT family, p53 function is also regulated through methylation of its C‐terminal end by PRMT5.129 This region is important for p53 oligomerization and cellular localization. A p53 methylation defective mutant exhibits changes in localization and altered expression of p53 target genes. Increased PRMT5 enhanced p53‐mediated cell cycle arrest, while loss of PRMT5 enhanced p53‐mediated apoptosis. Thus, methylation functions can decouple the cellular programs controlled by p53. 2. NF‐kB CARM1 and PRMT1 bind directly to the p65 subunit of NF‐kB and synergistically activate NF‐kB‐dependent transcription.130,131 However, while p300 significantly increases NF‐kB‐mediated transcription in concert with CARM1 and PRMT1, it was not necessary for PRMT synergy as observed for p53. Moreover, PRMT2 and PRMT5 inhibit NF‐kB‐mediated transcription. The mechanism of PRMT5 repression appears to be partially dependent on its methyltransferase activity, whereas PRMT1 and CARM1 synergy rely on their methyltransferase activity exclusively. PRMT2 appears to inhibit NF‐kB function through a nonenzymatic mechanism. PRMT2 directly binds to IkB‐a and thus retain it in nucleus; the increased levels of nuclear IkB‐a could inhibit NF‐kB signaling.132 3. E2Fs PRMT5 is often regarded as a repressor of transcription. A Cyclin E1 repressor complex contains Rbp48, E2F4, and PRMT5. PRMT5 is recruited to the cyclin E1 transcription start site and overexpression of PRMT5 represses the endogenous cyclin E1 levels.88 Transient transfection of PRMT5 leads to a reduction in cellular proliferation, an effect dependent on the methyltransferase activity of PRMT5.88 PRMT2 also exhibits a repressive role in E2F‐mediated gene expression. PRMT2 formed a ternary complex with E2F1 in the presence of retinoblastoma (Rb).133 PRMT2 is able to repress Gal4‐DBD‐E2F1‐mediated activation of a Gal4 reporter in a dose dependent manner. Repression was not observed in Rb null Saos2 cells while exogenous Rb can restore it. Cell cycle progression in Saos2 was arrested in the presence of PRMT2 and Rb further supporting their interdependence. PRMT2
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MEFs exhibit increased E2F activity and early S‐phase entry, suggesting that Rb function is suppressed in these cells. This represents another example that E2F activity is regulated by PRMTs.

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PRMTs also play a role in the activation of the cyclin E1 gene through E2F. While PRMT5 is found on the CCNE1 promoter in cells at G0 phase, PRMT1 was observed in cells at G1/S phase coinciding with a small decrease in H4R3 methylation.83 CARM1 binding and dimethylation at H3R17 and H3R26 was also observed on cyclin E1 gene in late G1/S. CARM1 is a potent activator of E2F target genes including cyclin E1, DHFR, and cdc6, while it does not regulate cyclin D1. Again, CARM1 recruitment at the cyclin E1 gene requires E2Fs and ACTR, a member of p160 proteins. A recent study showed that expression of E2F1 itself is regulated by CARM1.134 Collectively, multiple PRMTs are able to directly interact with the E2F transcription factors.

E. PRMTs in Transcription Elongation SPT4 and SPT5 form a heterodimeric transcription factor that mediates transcriptional elongation, which can be enhanced by Tat, a viral transcription activator. PTEF‐b, made up of CDK9 and cyclin T1, is also essential components of the elongation complex.135 SPT5 was found to bind PRMT5, pICln, and enhancer of rudimentary homolog (ERH). SPT5 is methylated by PRMT1 and PRMT5 and this methylation reduces its interaction with RNA pol II.46 SPT5 methylation defective mutant exhibits enhanced binding to CDK9 and cyclin T1, increased basal transcriptional elongation, and enhanced Tat activation of elongation. Concomitantly, PRMT1 and PRMT5 are able to reduce HIV‐1 gene expression in a methyltransferase dependent manner. These results suggest arginine methylation of SPT5 downregulates transcriptional elongation in response to cellular and viral factors. Another RNA pol II elongation factor that binds to PRMT5 and ERH is FCP1. FCP1 is methylated by PRMT5 and may also play a role in the regulation of elongation.136

IV. PRMTs in Posttranscriptional Regulation Prior to the discovery of PRMTs, a number of arginine methylation sites were identified through biochemical analysis and mass spectrometry. The asymmetric and symmetric dimethylated arginines were used to trace PRMT activity through classical biochemical methods. Much of this work identified RNA binding proteins as major targets for arginine methylation. For example, the methylation of heterogeneous nuclear ribonucleoproteins (hnRNPs) accounts for over half of the total nuclear arginine dimethylation.137 This early work facilitated the identification of common PRMT recognition motifs, including the RGG box or RG repeat domains.16,138 These common methylation motifs are often present within RNA and protein binding domains. PRMT1 methylation regulates these interactions, affecting both localization and function of methylated targets.139–141 Uniquely, PRMT3 regulates

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ribosomal assembly through methylation of ribosomal proteins.25,142 While CARM1 does not have the same specificity for RG rich sequences, it too can methylate a number of RNA binding proteins to alter both mRNA stability and splicing.123,143,144 PRMT5 methylation of RNA binding proteins regulates the assembly of the pre‐mRNA splicing complex.43,145 Thus, PRMT activity also acts downstream of transcriptional coactivation. The regulation of both transcription and posttranscriptional processes by PRMTs suggests that there may be a significant link between these two systems. However, the functional aspects of such a link have not been investigated.

V. Structural Analysis of PRMTs Crystallography has significantly increased our comprehension of PRMT enzymatic function. PRMT1, PRMT3, CARM1, and the yeast RMT HMT1 have been successfully crystallized.61,146–151 Crystallization reveals a number of common PRMT characteristics. First, the catalytic module is folded into two domains connected by a conserved cis‐proline residue. The structure‐based sequence alignment for PRMTs with known structures is shown in Fig. 3A. A topology diagram of the secondary structure elements for CARM1 is illustrated as an example (Fig. 3B). The first domain at the N‐terminal end is where S-adenosyl-homocysteine (AdoHcy) binds contains two terminal helices aX and aY and a typical Rossmann fold; the second domain is b‐barrel like. The target arginine is situated in a cone‐shaped pocket between the AdoMet binding domain and the b‐barrel domain. Second, PRMTs are predominantly found to form homo‐oligomers, usually in the form of dimers, which are stabilized by a ‘‘dimerization arm.’’ The dimer interface is formed between the arm and the outer surface of the AdoMet binding site. Deletion of the dimerization arm abrogates AdoMet binding and methyltransferase activity of PRMT1. Thus, dimerization is necessary for AdoMet binding. Another putative function of dimerization is to allow processive production of asymmetric dimethylarginine. The ring‐like dimer might allow the intermediate monomethylarginine to enter the active site of the second PRMT molecule in the dimer, without releasing the substrate from the ring.61,149 Recently, two groups have solved crystal structures of truncated forms of mouse CARM1.61,147 The first 25 amino acids and the last 120 amino acids are predicted to be disordered from sequence analysis and thus the full‐length protein fails to crystallize. Interestingly, crystallization of CARM128–140 reveals an unexpected pleckstrin homology domain (PH) domain‐like fold, which in other proteins facilitates protein–protein interactions.147 CARM128–140 does not have sequence similarity with any known PH domains. However, it folds as

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FIG. 3. (A) Structure‐based sequence alignment for rat PRMT1, rat PRMT3, yeast RMT1/Hmt1, and mCARM1. The helices and strands are displayed for mCARM1 and corresponding residue numbering is shown below the sequences. The four motifs characteristic of the PRMT domain are marked above sequence. Amino acids highlighted are either invariant (gray) or similar (black) as defined by the following grouping: F, Y, and W; I, L, M, and V; R and K; D and E; G and A; S, T, N, and Q. (B) Topology diagram of the secondary structure elements. The four motifs characteristics of PRMT domain (I, II, III, and IV) are marked. N‐terminal helices aX and aY and the b‐barrel are in gray, the dimerization arm is labeled with E, F, G0 , and G, the last C‐terminal strand (b‐16) is in dark gray. (Adapted from Ref. 147 with permission from the author and publisher.)

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two nearly orthogonal b‐sheets made up of seven antiparallel b‐strains followed by a C‐terminal amphipathic a‐helix, characteristics of PH domains. In contrast to other PH domains, CARM128–140 forms a dimer and the surface buried at the interface of the dimer hides the ligand‐binding site found in all PH domains. No ligand has been reported to bind to CARM1 PH domain. CARM1 is the first PRMT to be crystallized in both unbound (apo) form as well as bound to the cofactor product AdoHcy or SAH. Comparison of Apo‐ CARM1 with SAH bound CARM1 reveals induced folding of the N‐terminal helices over SAH, which creates a ridge over one side of the substrate binding pocket. This structural evidence explains the ordered mechanism in which SAM binding is a prerequisite for substrate binding. Unique to CARM1 is a b‐strand (b16) located at the C‐terminal end of the PRMT core that folds back into the central cavity of the CARM1 dimer and constitute CARM1 active site (Fig. 4A). Thus, the active site of CARM1 is delineated on one side by helices aX and aY, on the top by helix aZ and by the loop of motif IV, and on the other side by b16 strand (Fig. 4A). The unique C‐extension in CARM1 constitutes a lower ridge of the substrate binding groove, where neutral molecular surface charge is maintained. The equivalent position in PRMT1 is enriched with acidic residues. These features may explain that CARM1 does not methylate substrates containing highly basic GAR domains, while those are the preferred substrate structures for PRMT1/3. Functional significance of the N‐ and C‐extension in CARM1 was manifested in the in vitro methylation assay using truncated CARM1 and histone H3 substrate. The C‐extension is essential for CARM1 methylating histone H3 at all sites including R2, R17, and R26, whereas that N‐terminal PH domain deletion is required for methylation of R26, but not R17 on H3 tail.61 The H3 R17 and R26 are differentially methylated by the full‐length protein and by N‐terminal truncated CARM1, implicating that PH‐like domain may provide additional docking site to make R26 a suitable substrate, independent of the methylation status of R17. CARM128–507 reveals large structural changes in the catalytic site of PRMTs as compared with CARM1140–480. Three major structural changes are observed. One is on peptide 147–179, in which part of helix aX has been changed to a b‐strand and helices aY and aZ have been merged together and the kink does not exist anymore. The other major changes relates to structural rearrangement of all motifs of the core PRMT catalytic domain (Fig. 4B and C), leading to formation of a nonproductive active site. Third, the electron density map of PH domain is not seen in the larger CARM128–507 structure, suggesting that the N‐terminal PH fold behaves as a wobbly domain that moves independently of the catalytic core. However, the structural changes at the N‐terminal end of CARM1 catalytic domain observed in SAH‐CARM1140–480 and

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FIG. 4. The catalytic site of CARM1 in apo‐CARM1140–480 crystal structure. (A) Overview of the SAH‐binding site. Backbone conformations of the loops of motifs I–IV are labeled. The ordered water molecules (small spheres) occupy the binding site of the arginine substrate. (B) The residues 144–154 of helix aX of SAH‐CARM1140–480 are disordered in monomer structure. (C) Structure of CARM128–507 monomer. Part of helix aX has been changed in a strand b0 and the kink between helices aY and aZ has disappeared, helices aY and aZ have been merged together. Strand b0 forms a two‐stranded antiparallel b‐sheet with strand b16. Motifs II–IV also adopt new conformations. (Adapted from Troffer‐Charlier et al.,147,148 with permission from the author and publisher.)

CARM128–507 presumably lead to significantly different position of the PH domain.147 The biological significance of those alternative positions remains to be proven. In summary, the structural studies provide mechanistic insights that the central domain of PRMTs and dimerization are both critical for their catalytic activities. The N‐ and C‐terminal extension could modulate the enzymatic activity or specificity of PRMTs, moreover, they might be involved in the formation of large protein complex. These structures suggest that while the enzymatic activity of PRMTs is important for their roles as transcriptional coregulators, the nonenzymatic terminal extensions may also play a role in transcriptional regulation.

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VI. Small Molecule Inhibitors for PRMTs Inhibitors of the PRMTs used in many literatures are not specific to PRMTs. These inhibitors can be divided into two categories, both of which are competitive inhibitors of AdoMet. S‐adenosyl‐homocysteine (AdoHcy), the product of methyltransferase reaction, can compete with AdoMet for binding to the active site of the enzyme, thus inhibiting a wide array of AdoMet‐ dependent methylases. The second class contains analogues of AdoMet such as sinefungin, a natural streptomycin antibiotic. Sinefungin resembles the structure of AdoMet and forms a futile complex with a variety of AdoMet‐ dependent methyltransferases (reviewed in Ref. 13). Both types of inhibitors are nonspecific, which can inhibit methylation on DNA, RNA, and protein indiscriminately. Since PRMTs share conserved catalytic central domain and sometimes could methylate the common substrate, developing specific inhibitors for individual PRMT are essential for exploring their cellular functions and identifying the in vivo substrates. In recent years, small molecule inhibitors for PRMT have emerged and this development can be divided into multiple phases. Gray and coworkers took ‘‘bump‐and‐hole’’ approach in which a mutation is created in the active site of the enzyme that renders the enzyme susceptible to selective inhibition by a small molecule.152 The small molecule, however, cannot inhibit wild‐type enzyme. The mutant PRMT Rmt1/Hmt1 is genetically engineered in S. cerevisiae to replace the wild‐type enzyme, rendering selection of the ‘‘bumped’’ SAH analogues as HMT1‐specific inhibitors. This approach proved the concept that an orthogonal methyltransferase/inhibitor pair can be designed through synergistic protein engineering and ligand design; however, this approach cannot be easily adapted in the mammalian system. In the second phase, Bedford and coworkers took high throughput chemical screening approach to identify nine arginine methyltransferase inhibitors (AMIs) and two arginine methyltransferase activators (AMAs)153 that are specific for R but not K methyltransferases. The IC50 of the inhibitors to PRMT1 and Hmt1p are mostly in the mM range. Interestingly, some compounds are ineffective for a PRMT towards one substrate but are effective inhibitors for the same PRMT towards another substrate. The substrate‐dependent nature of enzyme/inhibitor pairs suggests different lodging sites of substrate in relation to the active site of the same PRMT. AMI‐1, which is a symmetrical sulfonated urea, was identified as the lead compound. AMI‐1 is cell‐permeable, inhibits all PRMTs in in vitro methylation assays and inhibits gene activation mediated by ER and AR.153 AMI‐1 is not AdoMet analogue and does not compete with AdoMet for binding to PRMTs, rather it shares some similarity to peptidyl arginine. Although there is no evidence to support that AMI‐1 is competitive

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with peptidyl arginine binding, this study opens new avenues for designing of bisubstrate inhibitors and improving specificity for PRMT inhibitors via molecular remodeling. Homology modeling and virtual screening approaches were undertaken in the design of novel PRMT inhibitors.154–156 Several PRMT1 specific inhibitors based on AMI‐1 structure were identified and shown to inhibit histone hypomethylation and ER activation in cancer cells.154 Analogues of AMI‐5 (eosin), a recently reported inhibitor of both protein arginine and histone lysine methyltransferase, have been developed bearing two ortho‐bromo‐ and ortho,ortho‐ dibromophenol moieties. These compounds exhibit differential effects on histone/protein methyltransferases, acetyltransferases, and Class III deacetylase (Sirtuin), some compounds induce apoptosis and differentiation of human leukemia U937 cells.156 One inhibitor with two ortho‐bromo moieties appears to be specific for CARM1. However, the IC50 is still in mM range and cellular H3R17Me2 level is not altered when the compound reaches 50 mM. The later development of PRMT specific inhibitors focuses on improving selectivity and cell permeability. A peptide‐based bisubstrate analogue is developed for PRMT1 which will be useful for ‘‘chemical genetics’’ studies of PRMT1 function;59 however, it is unlikely to possess drug‐like properties. The other bisubstrate analogue is cell permeable, but the IC50 for PRMT1 is 55 mM, and 150 mM is needed to inhibit cellular histone hypomethylation in HepG2 cells.157 These studies suggest that bisubstrate inhibitor docking both AdoMet binding pocket and R binding site could potentially provide new opportunities in structure‐based drug design. Thus far, the most selective, potent inhibitor for PRMTs is a pyrazole inhibitor for CARM1.158 A compound shown in Fig. 5 exhibits 0.08 mM IC50 for CARM1, while it is ineffective for inhibiting PRMT1 and PRMT3 (IC50 > 25 mM). Although this compound has not been characterized for inhibiting AdoMet and peptidyl arginine binding, it has characteristics of bisubstrate inhibitor. Cocrystal of this compound with CARM1 would provide structural insights into its inhibition mechanism. In summary, the existing PRMT inhibitors and the crystal structures of PRMTs provide a basis for homology modeling, molecular docking, and targeted‐based approaches for future development of more selective, potent inhibitors for PRMTs.

VII. Biological Functions of PRMTs While studies of PRMTs’ functions in human disease are increasing rapidly, we will focus this chapter on differentiation and development, cancer, viral infection, metabolism, DNA methylation, and repair.

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F3C

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HN NH2 O FIG. 5. Chemical structure of a CARM1 specific pyrazole inhibitor.158

A. PRMTs in Differentiation and Development 1. PRMTs IN MUSCLE DIFFERENTIATION CARM1 and PRMT5 have been implicated in muscle differentiation. MEF2 family members are responsible for the expression of genes essential for skeletal muscle differentiation. In addition to activation of MEF2C‐ mediated genes as discussed above (see Section C), CARM1 directly regulates expression of MEF2 and myogenin‐1 in the myogenic C2C12 cell line.122 Moreover, CARM1 expression at embryonic day 8.25 coincides with MEF2‐ mediated gene expression in muscle precursor cells and localization of CARM1 becomes increasingly nuclear during differentiation in C2C12 cells. PRMT5 regulates myogenic differentiation through coactivation of MyoD. MyoD is a transcription factor essential for myogenic determination, which induces critical genes such as desmin, myogenin, and skeletal actin for muscle differentiation.89 PRMT5 forms a tertiary complex with MyoD and the Swi/Snf ATPase BRG1. Reduction of PRMT5 abrogates BRG1 and MyoD binding to the myogenin reporter, where both chromatin remodeling and H3R8 dimethylation catalyzed by PRMT5 appear to be important for myogenin activation.89 Thus, the unusual activating role of PRMT5 mediates muscle differentiation.

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2. PRMTs IN PRECURSOR MAINTENANCE The maintenance and reprogramming of precursor cells is of significant interest to the field of stem cell research, as these cells present a viable tool for therapeutics in treating human disease. Differential expression of CARM1 and H3R26 methylation in mouse blastomeres correlates with an altered cell fate. Blastomeres with higher levels of CARM1 contributed to the inner cell mass (ICM) and exhibit a more pluripotent phenotype.159 Injection of CARM mRNA into blastomeres also induces expression of Nanog and Sox2 and formation of ICM. This effect was dependent on the methyltransferase activity of CARM1, suggesting that CARM1 may maintain cells in pluripotent state through transcriptional activation. Pluripotent germ cell fate in mice also appears to be controlled by PRMT5mediated regulation of transcription. Blimp1 is a SET‐domain transcription factor. Blimp1 forms a repressor complex with PRMT5 during PGC migration to the genital ridges.92 Nuclear localization of Blimp1‐PRMT5 corresponds with symmetric dimethylation of H2AR3 and H4R3. FGF‐2 dependent downregulation of Blimp1 in PGCs correlates with upregulation of Blimp1 target genes expressed in embryonic germ cells.160 This indicates that the Blimp1‐ PRMT5 repressor complex is necessary for PGC specific gene regulation, whose loss may trigger reprogramming of PGC cells into embryonic germ cells. 3. PRMTs IN DEVELOPMENT—WHAT WE HAVE LEARNED FROM MOUSE MODELS The creation of PRMT knockout mice has been useful tool in the characterization of PRMT function, primarily as a source of null mouse embryonic fibroblast (MEF) cells. A subset of PRMTs is essential for mouse development. PRMT1
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embryos die shortly after implantation but before the onset of gastrulation (E4.5–E6.5). These embryos are smaller in size and have reduced organization when compared to wild‐type embryos, and also show signs of resorption.161 Early embryonic death usually reflects mutations disrupting basic cellular processes, so the phenotype of PRMT1 hypomorph is consistent with the importance of PRMT1 in regulation of RNA metabolism. PRMT1 expression is highest along the midline of the neural plate (E7.5–E8.5) and in the developing central nervous system (E8.5–E13.5). CARM1
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mice are grossly normal at E12.5, but show some lethality around E18.5 with smaller size.162 No CARM1
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pups survive birth, and caesarian‐section of neonatal (E19.5) embryos reveals reduced size of alveolar air spaces and an absence of lung inflation. PRMT3
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mice are not embryonic lethal. E13.5 embryos are 25% smaller and E18.5 are 18% smaller than wild‐type embryos, but there is no difference in size at weaning.26 It is worthy to note that PRMT3
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ES cells generated through insertional mutagenesis have 5–7% of wild‐type PRMT3

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protein. The mild phenotype of PRMT3 knockout may be attributed to the hypomorphic allele or potential functional redundancy of different PRMTs. PRMT2 mice lack any morphological or observable developmental defects. While cell cycle regulation is altered in PRMT2
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MEFs, this does not appear to result in a significant phenotype or malignancy in PRMT2
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mice.133 Thus, PRMT1 and CARM1 are essential for mouse development while PRMT2 and PRMT3 are not. It is likely that PRMT1 and CARM1 are essential due to their pervasive roles in the regulation of gene expression, which is supported by the role of CARM1 in precursor maintenance. The importance of PRMT5 in germ cell differentiation suggests that it too may be essential in development. Deletion mutants of PRMT5 and other PRMTs, as well as conditional and tissue specific knockouts will help to delineate the role of PRMTs in development and differentiation. These initial studies indicate that future research may identify PRMTs as important players in stem cell growth and regulation.

B. PRMTs in Cancer PRMT1 has recently been implicated to have contradictory roles in different types of leukemia. The transcription factor RUNX1 is important for hematopoiesis. However, the RUNX1–ETO fusion protein disregulates hematopoiesis and contributes to the development of acute myeloid leukemia (AML). PRMT1 methylates RUNX1 in fusion protein and blocks the recruitment of SIN3A, a repressor that contributes to AML development.19. Moreover, PRMT1 activates RUNX1 target gene expression and enhances cellular differentiation. In another study, PRMT1 was found to induce hematopoietic self renewal, supporting transformation of T cells.163 MLL is a histone lysine methyltransferase that frequently forms oncogenic fusion proteins. The specific ability of the fusion protein MLL–EEN but not MLL alone to transform primary hematopoietic cells is dependent on association with PRMT1 and Sam68.163 In fact, an MLL–PRMT1 fusion was also able to transform these primary cells, supporting the role of PRMT1 in MLL‐mediated transformation. CARM1 also regulates T cell differentiation, at least one mechanism is via methylating the thymocyte cyclic AMP‐regulated phosphoprotein (TARPP). TARPP is expressed in early T cell progenitors and loss of CARM1‐mediated methylation of TARPP results in a decrease in total thymocytes number.164 These studies highlight the role of PRMTs in T cell differentiation, self‐ renewal, and possibly survival. PRMT5 downregulates a number of cell cycle regulators including CDK4, Cyclin B2, Cyclin E1, Cyclin E2, and CDC20 as well as tumor suppressors ST7, NM23, GAS1, GAS2, LOX1, and members of the Rb family.45,88 Higher expression of PRMT5 increases proliferation and cell cycle turnover and facilitates anchorage independent colony formation. Consistent with its growth stimulatory role, PRMT5 protein levels were found elevated in various

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lymphoma and leukemia cancer cells over that of normal B cells. Increased expression of PRMT5 in these cancers associates with H3R8 and H4R3 methylation, decreased tumor suppressor expression, and either an increase in PRMT5 mRNA stability or an increase in PRMT5 translation efficiency, possibly regulated by miRNAs.63,165 Thus, PRMT5 may play an oncogenic role in hematopoietic cancers. PRMTs appear to be involved in solid tumor growth as well. PRMT3 binds to a tumor suppressor DAL‐1/4/4.1B, which reduces cancer cell growth and migration. 4.1B binding decreases methyltransferase activity of PRMT3 in vitro27 and inhibits its methylation of Sm proteins.166 In contrast to PRMT3, 4.1B regulates PRMT5 activity by either inhibiting (Sm proteins) or enhancing (myelin basic protein) protein methylation. 4.1B‐mediated apoptosis is dependent on cellular arginine methylation in MCF7 breast cancer cells, although the mechanism is not yet defined.167 The role of CARM1 in solid tumors is still poorly defined, but a number of observations suggest CARM1 may have function in prostate and breast cancer by regulating AR and ER,28 two prominent therapeutic cancer targets. Additionally, levels of cellular CARM1 are altered in both prostate and breast cancer. Comparison of normal prostate tissues, prostate hyperplasia, and aggressive androgen‐independent prostate cancer show elevated nuclear CARM1 levels in late stage tumors.30,168 Moreover, cytoplasmic CARM1 was observed in late stage prostate cancer. In contrast to CARM1 in prostate cancer, CARM1 level is inversely correlated with grades of ER‐positive breast tumors, where more aggressive, high‐grade cancers show a significantly lower level of CARM1 proteins (Leigh Murphy, personal communication). Another study showed that high‐grade breast cancers carry elevated CARM1 mRNA.83 The reason for this discrepancy is unclear.

C. PRMTs in Viral Infection and Immune Response CARM1 appears to enhance viral gene expression by regulating transcription by Tax, a viral transcription factor of human T cell lymphotropic virus type I (HTLV1).169 CARM1 is found on the same promoter with p300 and CREB in a Tax‐dependent manner and its methyltransferase activity is indispensable for Tax‐mediated transactivation. Thus, the concurrent interaction between CARM1 and p300 during transcriptional activation is preserved in viral gene regulation. Human immunodeficiency virus type 1 (HIV‐1) expresses an early transactivator protein, Tat, which enhances RNA polymerase II transcription of viral genes. PRMT6 is a negative regulator of Tat; it methylates a central arginine rich motif on Tat.170 Elevated PRMT6 interrupts the formation of a Tat‐cyclin T1‐RNA complex, thus preventing RNA polymerase II elongation.171 Loss of PRMT6 results in increased HIV‐1 replication probably due to increased Tat

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transactivation. Thus, PRMT6 serves as a transcriptional repressor of viral gene expression and suppresses viral replication. PRMT6 also methylates an N‐terminal arginine rich motif in the multifunctional HIV‐1 protein, Rev.33 This methylation appears to have two effects: one is to reduce Rev‐mediated transcriptional activation and the other is to inhibit Rev‐mediated RNA export. PRMT6 is also able to methylate the HIV‐1 nucleocapsid (NC) protein, one of the Gag polyprotein cleavage products172 to inhibit reverse transcription. In aggregate, these studies indicate a consistent role for PRMT6 in the inhibition of HIV‐1 infection. Not only do PRMTs interact directly with viral factors but they also play a role in cytokine responses to pathogenic infections. PRMT1 binds directly to the intracytoplasmic domain of the interferon (IFN) receptor 1 (IFNAR‐1),173 and appears to facilitate IFN‐mediated antiviral and antiproliferative effects. CARM1 also plays a role in IFN signaling. IFNg activates major histocompatibility (MHC‐II) expression through the Class II transactivator (CIITA). CARM1 associates with CIITA and CBP on the MHCII promoter and methylates H3R17 to activate MHC‐II gene expression.174 Thus, CARM1 plays a role in IFN‐g‐induced transcription and potentially facilitates development of specific immune responses.

D. PRMTs in Metabolism 1. PRMTs IN GLUCOSE METABOLISM Improper regulation of circulating glucose levels is a key determinant of metabolic diseases such as diabetes. Cyclic adenosine monophosphate (cAMP)/protein kinase A (PKA) dependant signaling pathways induce PEPCK and glucose‐6‐phophatase (G6Pase) expression, which are essential for gluconeogenesis. Elevated gluconeogenesis can result in peripheral insulin resistance. CARM1 coactivates both PEPCK and G6Pase reporter constructs in hepatocyte cell lines in a PKA dependent manner.175 A CREB response element was identified on PEPCK promoter, supporting the idea that CARM1 interact with CREB to facilitate PEPCK expression. Thus, CARM1 appears to play a role in gluconeogenesis, whether arginine methylation is involved in metabolic regulation remains to be determined. 2. PRMTs IN ADIPOGENESIS PPARg is a master regulator for fatty acid metabolism, whose function is regulated by a plethora of cofactors.176 Adipogenesis was recently shown to be regulated by CARM1. CARM1 null embryos and mouse embryonic fibroblasts (MEFs) were utilized to explore CARM1’s role in PPARg signaling. CARM null embryos exhibit a 40% reduction in lipid accumulation in brown adipose tissue.177 This loss of lipid development is likely due to altered differentiation

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of preadipocytes, a process regulated by PPARg. CARM1 activates PPARg transcription in transient transfection assays. Moreover, CARM1 and RNA Pol II are bound to the endogenous aP2 promoter concomitant with H3R17 methylation. These observations are consistent with our findings that aP2‐ directed adipose‐specific knockout of CARM1 in mice exhibits less lipid storage in brown fat, whereas no obvious differences were observed in white fat (Xu and Evans, unpublished data). To our knowledge, PRMT2 is the only other PRMT that has been shown to interact with PPARg and stimulate its transcriptional activity.53 Receptor interacting protein 140 (RIP140) is a corepressor for nuclear receptors. RIP140 null mice exhibit multisystem energy metabolic derangements.178 RIP140 is methylated by PRMT1 at three distal arginine residues in vitro.179 Methylation at R240 appears to inhibit the interaction between RIP140 and HDAC3179 and also induces the nuclear export of RIP140. The export of RIP140 is associated with release of transcriptional repression and an increase in adipogenesis in MEF cells. However, the specific transcription factors and promoters targeted by RIP140 in adipogenesis remain to be defined.

E. PRMTs and DNA Methylation DNA methylation at CpG islands is inhibitory to transcription. Methyl DNA binding domain (MBD) proteins are major effector proteins for methyl‐CpG sequences. The NuRD complexes that associate with MBDs contain known transcriptional repressors including HDACs, RbAp48, and RbAp46. PRMT5 is present in MBD2 containing NuRD complex, where it methylates MBD2 in RG repeat region.180 Inhibition of methyltransferase activity decreases association of PRMT5 and MBD2 on methyl‐CG containing promoters. MBD2 is also a target of PRMT1.181 Loss of either PRMT increases NuRD complex stability, while methylation of MBD2 decreases its methyl‐ CpG affinity. The decreased stability of the MBD2–HDAC interaction in the presence of PRMT1 or PRMT5 suggests that PRMTs may be transiently present in this repressor complex. Further studies indicate other methylation events may affects the MBD–NuRD complex, thus specific PRMT inhibitors would be desirable to elucidate function of individual PRMT.

F. PRMTs and DNA Repair MRE11 is part of the MRN complex, a key player in the repair of DNA double strand breaks. PRMT1 methylates MRE11 at a GAR motif in the DNA binding domain prior to MRE11–RAD50–NBS1 (MSN) complex assembly.182,183 Methylation of MRE11 appears to be important for the proper localization of MRE11 to double strand DNA breaks. Deletion of the GAR

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motif inhibits MRE11 binding to DNA. Together this indicates the PRMT1 regulates MRE11 localization and DNA binding activity through methylation of GAR motif. In addition to the regulation of double strand DNA repair, PRMTs are also involved in excision repair and single strand breaks. DNA polymerase b fills in short single‐stranded DNA gaps at the sites of DNA damage. PRMT6 methylates two residues in the lyase domain of DNA polymerase b.184 Polymerase b methylation defective mutant exhibits reduced ability to respond to DNA damage, resulting in delayed cell cycle progression and DNA fragmentation repair. In this scenario, methylation of DNA binding proteins appears to facilitate protein–DNA interactions. Consistent with this observation, PRMT1, PRMT3, and PRMT6 also methylate the three AT hooks of high mobility group A1a (HMGA1a), regions necessary for protein–DNA interactions.32,185,186 While the functional role of this methylation has not been assessed, like RNA binding domains, DNA binding domains appear to be major targets for arginine methylation.

G. PRMTs and Chromatin Domains Recently it has been shown that PRMT1‐mediated H4R3 dimethylation may not be limited to temporal gene regulation, but may also play a role in the maintenance of euchromatin structure. Insulator regions can act as a barrier for the encroachment of condensed heterochromatin into transcriptionally active euchromatin; however the mechanism for establishment of euchromatin domains remains to be defined. The transcription factor USF1 is known to bind to insulator sequences and facilitates the maintenance of euchromatin. PRMT1 associates with USF1 in concert with HAT PCAF and SRC‐1 within the HS4 insulator region; this is accompanied by H4R3 methylation and histone acetylation.187 Knocking down PRMT1 leads to a loss of activating acetylation marks with a specific increase in repressive marks on the folate receptor promoter and the adjacent HS4 insulator.187 Thus, PRMT1 can methylate H4R3 in gene promoters to activate transcription, but the same mark can facilitate maintenance of chromatin structure in insulator regions as well. CTCFL, a testes paralog of the insulator protein CTCF, binds to insulator regions and mediates imprinting through DNA methylation. The interaction of CTCFL with PRMT7 facilitates PRMT7‐mediated methylation of H4R3 and H2A, which directs DNA methylation at the H19 imprinting control region.70 This suggests a role for PRMT7 in the developmental control of germ cells in the testes. The interaction between PRMT1 and PRMT7 with insulator proteins involved in DNA methylation suggests a common mechanism where DNA and arginine methylation concurrently control gene expression.

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VIII. Concluding Remarks Protein arginine methylation catalyzed by PRMTs is an important posttranslational modification whose influence on cellular processes has been well‐acknowledged in recent years. Regulation of gene expression by PRMTs utilizes diverse mechanisms and targets transcription, posttranscriptional processes, and even translation. Many PRMT substrates and transcriptional targets are important in human disease; however our understanding of the role of PRMTs at both the molecular and organismal level is still quite limited. Regulation of PRMT activity (both transcriptional and enzymatic) will likely be a major target for future research, and will require characterization of posttranslational modifications, identification of novel PRMT binding proteins, and further structural characterization. At the organismal level, development of animal models and design of specific inhibitors will facilitate the delineation of the net effects of these multifunctional proteins and the development of therapeutics targeting PRMT enzymes. Acknowledgments We thank Jean Cavarelli, Steven Clarke, Laurence Vandel, and Mariam Al‐Dhaheri for critical reading of the manuscript, Mark Bedford for helpful discussion, and Shaun Hernandez for help with graphics. This work is supported by R01CA125387 to W.X., a grant from Susan Komen Foundation and Shaw Scientist Award (to W.X.) by Greater Milwaukee Foundation.

References 1. Martin JL, McMillan FM. SAM (dependent) I AM: the S‐adenosylmethionine‐dependent methyltransferase fold. Curr Opin Struct Biol 2002;12:783–93. 2. Bedford MT. Arginine methylation at a glance. J Cell Sci 2007;120:4243–6. 3. Katz JE, Dlakic M, Clarke S. Automated identification of putative methyltransferases from genomic open reading frames. Mol Cell Proteomics 2003;2:525–40. 4. Schubert HL, Blumenthal RM, Cheng X. Many paths to methyltransfer: a chronicle of convergence. Trends Biochem Sci 2003;28:329–35. 5. Kagan RM, Clarke S. Widespread occurrence of three sequence motifs in diverse S‐adenosylmethionine‐dependent methyltransferases suggests a common structure for these enzymes. Arch Biochem Biophys 1994;310:417–27. 6. Gary JD, Clarke S. RNA and protein interactions modulated by protein arginine methylation. Prog Nucleic Acid Res Mol Biol 1998;61:65–131. 7. Paik WK, Kim S. Enzymatic methylation of protein fractions from calf thymus nuclei. Biochem Biophys Res Commun 1967;29:14–20. 8. Lin WJ, Gary JD, Yang MC, Clarke S, Herschman HR. The mammalian immediate‐early TIS21 protein and the leukemia‐associated BTG1 protein interact with a protein‐arginine N‐methyltransferase. J Biol Chem 1996;271:15034–44.

BIOLOGICAL FUNCTIONS OF PROTEIN ARGININE METHYLTRANSFERASES

333

9. Bachand F. Protein arginine methyltransferases: from unicellular eukaryotes to humans. Eukaryot Cell 2007;6:889–98. 10. Niewmierzycka A, Clarke S. S‐Adenosylmethionine‐dependent methylation in Saccharomyces cerevisiae Identification of a novel protein arginine methyltransferase. J Biol Chem 1999;274:814–24. 11. Scebba F, De Bastiani M, Bernacchia G, Andreucci A, Galli A, Pitto L. PRMT11: a new Arabidopsis MBD7 protein partner with arginine methyltransferase activity. Plant J 2007;52:210–22. 12. Boulanger MC, Miranda TB, Clarke S, Di Fruscio M, Suter B, Lasko P, et al. Characterization of the Drosophila protein arginine methyltransferases DART1 and DART4. Biochem J 2004;379:283–9. 13. Krause CD, Yang ZH, Kim YS, Lee JH, Cook JR, Pestka S. Protein arginine methyltransferases: evolution and assessment of their pharmacological and therapeutic potential. Pharmacol Ther 2007;113:50–87. 14. Frankel A, Yadav N, Lee J, Branscombe TL, Clarke S, Bedford MT. The novel human protein arginine N‐methyltransferase PRMT6 is a nuclear enzyme displaying unique substrate specificity. J Biol Chem 2002;277:3537–43. 15. Najbauer J, Johnson BA, Young AL, Aswad DW. Peptides with sequences similar to glycine, arginine‐rich motifs in proteins interacting with RNA are efficiently recognized by methyltransferase(s) modifying arginine in numerous proteins. J Biol Chem 1993;268:10501–9. 16. Kiledjian M, Dreyfuss G. Primary structure and binding activity of the hnRNP U protein: binding RNA through RGG box. EMBO J 1992;11:2655–64. 17. Wooderchak WL, Zang T, Zhou ZS, Acuna M, Tahara SM, Hevel JM. Substrate profiling of PRMT1 reveals amino acid sequences that extend beyond the ‘‘RGG’’ paradigm. Biochemistry 2008;47:9456–66. 18. Smith JJ, Rucknagel KP, Schierhorn A, Tang J, Nemeth A, Linder M, et al. Unusual sites of arginine methylation in Poly(A)‐binding protein II and in vitro methylation by protein arginine methyltransferases PRMT1 and PRMT3. J Biol Chem 1999;274:13229–34. 19. Zhao X, Jankovic V, Gural A, Huang G, Pardanani A, Menendez S, et al. Methylation of RUNX1 by PRMT1 abrogates SIN3A binding and potentiates its transcriptional activity. Genes Dev 2008;22:640–53. 20. Mowen KA, Tang J, Zhu W, Schurter BT, Shuai K, Herschman HR, et al. Arginine methylation of STAT1 modulates IFNalpha/beta‐induced transcription. Cell 2001;104:731–41. 21. Teyssier C, Ma H, Emter R, Kralli A, Stallcup MR. Activation of nuclear receptor coactivator PGC‐1alpha by arginine methylation. Genes Dev 2005;19:1466–73. 22. Tang J, Frankel A, Cook RJ, Kim S, Paik WK, Williams KR, et al. PRMT1 is the predominant type I protein arginine methyltransferase in mammalian cells. J Biol Chem 2000;275:7723–30. 23. Tang J, Gary JD, Clarke S, Herschman HR. PRMT 3, a type I protein arginine N‐ methyltransferase that differs from PRMT1 in its oligomerization, subcellular localization, substrate specificity, and regulation. J Biol Chem 1998;273:16935–45. 24. Frankel A, Clarke S. PRMT3 is a distinct member of the protein arginine N‐methyltransferase family. Conferral of substrate specificity by a zinc‐finger domain. J Biol Chem 2000;275:32974–82. 25. Bachand F, Silver PA. PRMT3 is a ribosomal protein methyltransferase that affects the cellular levels of ribosomal subunits. EMBO J 2004;23:2641–50. 26. Swiercz R, Cheng D, Kim D, Bedford MT. Ribosomal protein rpS2 is hypomethylated in PRMT3‐deficient mice. J Biol Chem 2007;282:16917–23. 27. Singh V, Miranda TB, Jiang W, Frankel A, Roemer ME, Robb VA, et al. DAL‐1/4.1B tumor suppressor interacts with protein arginine N‐methyltransferase 3 (PRMT3) and inhibits its ability to methylate substrates in vitro and in vivo. Oncogene 2004;23:7761–71.

334

KUHN AND XU

28. Chen D, Ma H, Hong H, Koh SS, Huang SM, Schurter BT, et al. Regulation of transcription by a protein methyltransferase. Science 1999;284:2174–7. 29. Schurter BT, Koh SS, Chen D, Bunick GJ, Harp JM, Hanson BL, et al. Methylation of histone H3 by coactivator‐associated arginine methyltransferase 1. Biochemistry 2001;40:5747–56. 30. Hong H, Kao C, Jeng MH, Eble JN, Koch MO, Gardner TA, et al. Aberrant expression of CARM1, a transcriptional coactivator of androgen receptor, in the development of prostate carcinoma and androgen‐independent status. Cancer 2004;101:83–9. 31. Guccione E, Bassi C, Casadio F, Martinato F, Cesaroni M, Schuchlautz H, et al. Methylation of histone H3R2 by PRMT6 and H3K4 by an MLL complex are mutually exclusive. Nature 2007;449:933–7. 32. Sgarra R, Lee J, Tessari MA, Altamura S, Spolaore B, Giancotti V, et al. The AT‐hook of the chromatin architectural transcription factor high mobility group A1a is arginine‐methylated by protein arginine methyltransferase 6. J Biol Chem 2006;281:3764–72. 33. Invernizzi CF, Xie B, Richard S, Wainberg MA. PRMT6 diminishes HIV‐1 Rev binding to and export of viral RNA. Retrovirology 2006;3:93. 34. Aubert J, Stavridis MP, Tweedie S, O’Reilly M, Vierlinger K, Li M, et al. Screening for mammalian neural genes via fluorescence‐activated cell sorter purification of neural precursors from Sox1‐gfp knock‐in mice. Proc Natl Acad Sci USA 2003;100(Suppl 1):11836–41. 35. Lee J, Sayegh J, Daniel J, Clarke S, Bedford MT. PRMT8, a new membrane‐bound tissue‐ specific member of the protein arginine methyltransferase family. J Biol Chem 2005;280:32890–6. 36. Taneda T, Miyata S, Kousaka A, Inoue K, Koyama Y, Mori Y, et al. Specific regional distribution of protein arginine methyltransferase 8 (PRMT8) in the mouse brain. Brain Res 2007;1155:1–9. 37. Sayegh J, Webb K, Cheng D, Bedford MT, Clarke SG. Regulation of protein arginine methyltransferase 8 (PRMT8) activity by its N‐terminal domain. J Biol Chem 2007;282:36444–53. 38. Pahlich S, Zakaryan RP, Gehring H. Identification of proteins interacting with protein arginine methyltransferase 8: the Ewing sarcoma (EWS) protein binds independent of its methylation state. Proteins 2008;72:1125–37. 39. Pollack BP, Kotenko SV, He W, Izotova LS, Barnoski BL, Pestka S. The human homologue of the yeast proteins Skb1 and Hsl7p interacts with Jak kinases and contains protein methyltransferase activity. J Biol Chem 1999;274:31531–42. 40. Branscombe TL, Frankel A, Lee JH, Cook JR, Yang Z, Pestka S, et al. PRMT5 (Janus kinase‐ binding protein 1) catalyzes the formation of symmetric dimethylarginine residues in proteins. J Biol Chem 2001;276:32971–6. 41. Teng Y, Girvan AC, Casson LK, Pierce Jr WM, Qian M, Thomas SD, et al. AS1411 alters the localization of a complex containing protein arginine methyltransferase 5 and nucleolin. Cancer Res 2007;67:10491–500. 42. Liang JJ, Wang Z, Chiriboga L, Greco MA, Shapiro E, Huang H, et al. The expression and function of androgen receptor coactivator p44 and protein arginine methyltransferase 5 in the developing testis and testicular tumors. J Urol 2007;177:1918–22. 43. Friesen WJ, Wyce A, Paushkin S, Abel L, Rappsilber J, Mann M, et al. A novel WD repeat protein component of the methylosome binds Sm proteins. J Biol Chem 2002;277:8243–7. 44. Pal S, Yun R, Datta A, Lacomis L, Erdjument‐Bromage H, Kumar J, et al. mSin3A/histone deacetylase 2‐ and PRMT5‐containing Brg1 complex is involved in transcriptional repression of the Myc target gene cad. Mol Cell Biol 2003;23:7475–87. 45. Pal S, Vishwanath SN, Erdjument‐Bromage H, Tempst P, Sif S. Human SWI/SNF‐associated PRMT5 methylates histone H3 arginine 8 and negatively regulates expression of ST7 and NM23 tumor suppressor genes. Mol Cell Biol 2004;24:9630–45.

BIOLOGICAL FUNCTIONS OF PROTEIN ARGININE METHYLTRANSFERASES

335

46. Kwak YT, Guo J, Prajapati S, Park KJ, Surabhi RM, Miller B, et al. Methylation of SPT5 regulates its interaction with RNA polymerase II and transcriptional elongation properties. Mol Cell 2003;11:1055–66. 47. Miranda TB, Miranda M, Frankel A, Clarke S. PRMT7 is a member of the protein arginine methyltransferase family with a distinct substrate specificity. J Biol Chem 2004;279:22902–7. 48. Lee JH, Cook JR, Yang ZH, Mirochnitchenko O, Gunderson SI, Felix AM, et al. PRMT7, a new protein arginine methyltransferase that synthesizes symmetric dimethylarginine. J Biol Chem 2005;280:3656–64. 49. Sayegh J, Clarke SG. Hsl7 is a substrate‐specific type II protein arginine methyltransferase in yeast. Biochem Biophys Res Commun 2008;372:811–5. 50. Cook JR, Lee JH, Yang ZH, Krause CD, Herth N, Hoffmann R, et al. FBXO11/PRMT9, a new protein arginine methyltransferase, symmetrically dimethylates arginine residues. Biochem Biophys Res Commun 2006;342:472–81. 51. Katsanis N, Yaspo ML, Fisher EM. Identification and mapping of a novel human gene, HRMT1L1, homologous to the rat protein arginine N‐methyltransferase 1 (PRMT1) gene. Mamm Genome 1997;8:526–9. 52. Scott HS, Antonarakis SE, Lalioti MD, Rossier C, Silver PA, Henry MF. Identification and characterization of two putative human arginine methyltransferases (HRMT1L1 and HRMT1L2). Genomics 1998;48:330–40. 53. Qi C, Chang J, Zhu Y, Yeldandi AV, Rao SM, Zhu YJ. Identification of protein arginine methyltransferase 2 as a coactivator for estrogen receptor alpha. J Biol Chem 2002;277:28624–30. 54. Meyer R, Wolf SS, Obendorf M. PRMT2, a member of the protein arginine methyltransferase family, is a coactivator of the androgen receptor. J Steroid Biochem Mol Biol 2007;107:1–14. 55. Bedford MT. The family of protein arginine methyltransferases. Amsterdam: Academic Press; 2006. 56. Osborne TC, Obianyo O, Zhang X, Cheng X, Thompson PR. Protein arginine methyltransferase 1: positively charged residues in substrate peptides distal to the site of methylation are important for substrate binding and catalysis. Biochemistry 2007;46:13370–81. 57. Obianyo O, Osborne TC, Thompson PR. Kinetic mechanism of protein arginine methyltransferase 1. Biochemistry 2008;47:10420–7. 58. Lakowski TM, Frankel A. A kinetic study of human protein arginine N‐methyltransferase 6 reveals a distributive mechanism. J Biol Chem 2008;283:10015–25. 59. Osborne T, Roska RL, Rajski SR, Thompson PR. In situ generation of a bisubstrate analogue for protein arginine methyltransferase 1. J Am Chem Soc 2008;130:4574–5. 60. Lee DY, Ianculescu I, Purcell D, Zhang X, Cheng X, Stallcup MR. Surface‐scanning mutational analysis of protein arginine methyltransferase 1: roles of specific amino acids in methyltransferase substrate specificity, oligomerization, and coactivator function. Mol Endocrinol 2007;21:1381–93. 61. Yue WW, Hassler M, Roe SM, Thompson‐Vale V, Pearl LH. Insights into histone code syntax from structural and biochemical studies of CARM1 methyltransferase. EMBO J 2007;26:4402–12. 62. Rho J, Choi S, Seong YR, Cho WK, Kim SH, Im DS. Prmt5, which forms distinct homo‐ oligomers, is a member of the protein‐arginine methyltransferase family. J Biol Chem 2001;276:11393–401. 63. Pal S, Baiocchi RA, Byrd JC, Grever MR, Jacob ST, Sif S. Low levels of miR‐92b/96 induce PRMT5 translation and H3R8/H4R3 methylation in mantle cell lymphoma. EMBO J 2007;26:3558–69. 64. Wang H, Huang ZQ, Xia L, Feng Q, Erdjument‐Bromage H, Strahl BD, et al. Methylation of histone H4 at arginine 3 facilitating transcriptional activation by nuclear hormone receptor. Science 2001;293:853–7.

336

KUHN AND XU

65. Pahlich S, Bschir K, Chiavi C, Belyanskaya L, Gehring H. Different methylation characteristics of protein arginine methyltransferase 1 and 3 toward the Ewing Sarcoma protein and a peptide. Proteins 2005;61:164–75. 66. Fronz K, Otto S, Kolbel K, Kuhn U, Friedrich H, Schierhorn A, et al. Promiscuous modification of the nuclear poly(A)‐binding protein by multiple protein‐arginine methyltransferases does not affect the aggregation behavior. J Biol Chem 2008;283:20408–20. 67. Robin‐Lespinasse Y, Sentis S, Kolytcheff C, Rostan MC, Corbo L, Le Romancer M. hCAF1, a new regulator of PRMT1‐dependent arginine methylation. J Cell Sci 2007;120:638–47. 68. Miyata S, Mori Y, Tohyama M. PRMT1 and Btg2 regulates neurite outgrowth of Neuro2a cells. Neurosci Lett 2008;445:162–5. 69. Choi S, Jung CR, Kim JY, Im DS. PRMT3 inhibits ubiquitination of ribosomal protein S2 and together forms an active enzyme complex. Biochim Biophys Acta 2008;1780:1062–9. 70. Jelinic P, Stehle JC, Shaw P. The testis‐specific factor CTCFL cooperates with the protein methyltransferase PRMT7 in H19 imprinting control region methylation. PLoS Biol 2006;4: e355. 71. Xu W, Chen H, Du K, Asahara H, Tini M, Emerson BM, et al. A transcriptional switch mediated by cofactor methylation. Science 2001;294:2507–11. 72. Higashimoto K, Kuhn P, Desai D, Cheng X, Xu W. Phosphorylation‐mediated inactivation of coactivator‐associated arginine methyltransferase 1. Proc Natl Acad Sci USA 2007;104:12318–23. 73. Allfrey VG, Faulkner R, Mirsky AE. Acetylation and methylation of histones and their possible role in the regulation of RNA synthesis. Proc Natl Acad Sci USA 1964;51:786–94. 74. Berger SL. The complex language of chromatin regulation during transcription. Nature 2007;447:407–12. 75. Sims III RJ, Reinberg D. Is there a code embedded in proteins that is based on post‐ translational modifications?. Nat Rev Mol Cell Biol 2008;9:815–20. 76. Stallcup MR. Role of protein methylation in chromatin remodeling and transcriptional regulation. Oncogene 2001;20:3014–20. 77. Wysocka J, Allis CD, Coonrod S. Histone arginine methylation and its dynamic regulation. Front Biosci 2006;11:344–55. 78. Strahl BD, Briggs SD, Brame CJ, Caldwell JA, Koh SS, Ma H, et al. Methylation of histone H4 at arginine 3 occurs in vivo and is mediated by the nuclear receptor coactivator PRMT1. Curr Biol 2001;11:996–1000. 79. Ma H, Baumann CT, Li H, Strahl BD, Rice R, Jelinek MA, et al. Hormone‐dependent, CARM1‐directed, arginine‐specific methylation of histone H3 on a steroid‐regulated promoter. Curr Biol 2001;11:1981–5. 80. Huang S, Litt M, Felsenfeld G. Methylation of histone H4 by arginine methyltransferase PRMT1 is essential in vivo for many subsequent histone modifications. Genes Dev 2005;19:1885–93. 81. Wagner S, Weber S, Kleinschmidt MA, Nagata K, Bauer UM. SET‐mediated promoter hypoacetylation is a prerequisite for coactivation of the estrogen‐responsive pS2 gene by PRMT1. J Biol Chem 2006;281:27242–50. 82. Miao F, Li S, Chavez V, Lanting L, Natarajan R. Coactivator‐associated arginine methyltransferase‐1 enhances nuclear factor‐kappaB‐mediated gene transcription through methylation of histone H3 at arginine 17. Mol Endocrinol 2006;20:1562–73. 83. El Messaoudi S, Fabbrizio E, Rodriguez C, Chuchana P, Fauquier L, Cheng D, et al. Coactivator‐associated arginine methyltransferase 1 (CARM1) is a positive regulator of the Cyclin E1 gene. Proc Natl Acad Sci USA 2006;103:13351–6. 84. Daujat S, Bauer UM, Shah V, Turner B, Berger S, Kouzarides T. Crosstalk between CARM1 methylation and CBP acetylation on histone H3. Curr Biol 2002;12:2090–7.

BIOLOGICAL FUNCTIONS OF PROTEIN ARGININE METHYLTRANSFERASES

337

85. An W, Kim J, Roeder RG. Ordered cooperative functions of PRMT1, p300, and CARM1 in transcriptional activation by p53. Cell 2004;117:735–48. 86. Hyllus D, Stein C, Schnabel K, Schiltz E, Imhof A, Dou Y, et al. PRMT6‐mediated methylation of R2 in histone H3 antagonizes H3 K4 trimethylation. Genes Dev 2007;21:3369–80. 87. Iberg AN, Espejo A, Cheng D, Kim D, Michaud‐Levesque J, Richard S, et al. Arginine methylation of the histone H3 tail impedes effector binding. J Biol Chem 2008;283:3006–10. 88. Fabbrizio E, El Messaoudi S, Polanowska J, Paul C, Cook JR, Lee JH, et al. Negative regulation of transcription by the type II arginine methyltransferase PRMT5. EMBO Rep 2002;3:641–5. 89. Dacwag CS, Ohkawa Y, Pal S, Sif S, Imbalzano AN. The protein arginine methyltransferase Prmt5 is required for myogenesis because it facilitates ATP‐dependent chromatin remodeling. Mol Cell Biol 2007;27:384–94. 90. Hosohata K, Li P, Hosohata Y, Qin J, Roeder RG, Wang Z. Purification and identification of a novel complex which is involved in androgen receptor‐dependent transcription. Mol Cell Biol 2003;23:7019–29. 91. Richard S, Morel M, Cleroux P. Arginine methylation regulates IL‐2 gene expression: a role for protein arginine methyltransferase 5 (PRMT5). Biochem J 2005;388:379–86. 92. Ancelin K, Lange UC, Hajkova P, Schneider R, Bannister AJ, Kouzarides T, et al. Blimp1 associates with Prmt5 and directs histone arginine methylation in mouse germ cells. Nat Cell Biol 2006;8:623–30. 93. Wang Y, Wysocka J, Sayegh J, Lee YH, Perlin JR, Leonelli L, et al. Human PAD4 regulates histone arginine methylation levels via demethylimination. Science 2004;306:279–83. 94. Cuthbert GL, Daujat S, Snowden AW, Erdjument‐Bromage H, Hagiwara T, Yamada M, et al. Histone deimination antagonizes arginine methylation. Cell 2004;118:545–53. 95. Lee YH, Coonrod SA, Kraus WL, Jelinek MA, Stallcup MR. Regulation of coactivator complex assembly and function by protein arginine methylation and demethylimination. Proc Natl Acad Sci USA 2005;102:3611–6. 96. Chang B, Chen Y, Zhao Y, Bruick RK. JMJD6 is a histone arginine demethylase. Science 2007;318:444–7. 97. Garcia BA, Pesavento JJ, Mizzen CA, Kelleher NL. Pervasive combinatorial modification of histone H3 in human cells. Nat Methods 2007;4:487–9. 98. Xu W, Cho H, Evans RM. Acetylation and methylation in nuclear receptor gene activation. Methods Enzymol 2003;364:205–23. 99. Koh SS, Chen D, Lee YH, Stallcup MR. Synergistic enhancement of nuclear receptor function by p160 coactivators and two coactivators with protein methyltransferase activities. J Biol Chem 2001;276:1089–98. 100. Lee YH, Koh SS, Zhang X, Cheng X, Stallcup MR. Synergy among nuclear receptor coactivators: selective requirement for protein methyltransferase and acetyltransferase activities. Mol Cell Biol 2002;22:3621–32. 101. Zhao X, Benveniste EN. Transcriptional activation of human matrix metalloproteinase‐9 gene expression by multiple co‐activators. J Mol Biol 2008;383:945–56. 102. Teyssier C, Chen D, Stallcup MR. Requirement for multiple domains of the protein arginine methyltransferase CARM1 in its transcriptional coactivator function. J Biol Chem 2002;277:46066–72. 103. Chevillard‐Briet M, Trouche D, Vandel L. Control of CBP co‐activating activity by arginine methylation. EMBO J 2002;21:5457–66. 104. Demarest SJ, Martinez‐Yamout M, Chung J, Chen H, Xu W, Dyson HJ, et al. Mutual synergistic folding in recruitment of CBP/p300 by p160 nuclear receptor coactivators. Nature 2002;415:549–53.

338

KUHN AND XU

105. Koh SS, Li H, Lee YH, Widelitz RB, Chuong CM, Stallcup MR. Synergistic coactivator function by coactivator‐associated arginine methyltransferase (CARM) 1 and beta‐catenin with two different classes of DNA‐binding transcriptional activators. J Biol Chem 2002;277:26031–5. 106. Feng Q, Yi P, Wong J, O’Malley BW. Signaling within a coactivator complex: methylation of SRC‐3/AIB1 is a molecular switch for complex disassembly. Mol Cell Biol 2006;26:7846–57. 107. Naeem H, Cheng D, Zhao Q, Underhill C, Tini M, Bedford MT, et al. The activity and stability of the transcriptional coactivator p/CIP/SRC‐3 are regulated by CARM1‐dependent methylation. Mol Cell Biol 2007;27:120–34. 108. Xu W, Cho H, Kadam S, Banayo EM, Anderson S, Yates III JR, et al. A methylation‐mediator complex in hormone signaling. Genes Dev 2004;18:144–56. 109. Teyssier C, Ou CY, Khetchoumian K, Losson R, Stallcup MR. Transcriptional intermediary factor 1alpha mediates physical interaction and functional synergy between the coactivator‐ associated arginine methyltransferase 1 and glucocorticoid receptor‐interacting protein 1 nuclear receptor coactivators. Mol Endocrinol 2006;20:1276–86. 110. Lee YH, Campbell HD, Stallcup MR. Developmentally essential protein flightless I is a nuclear receptor coactivator with actin binding activity. Mol Cell Biol 2004;24:2103–17. 111. Thompson M. Polybromo‐1: The chromatin targeting subunit of the PBAF complex. Biochimie 2008;31:309–19. 112. Chen YH, Kim JH, Stallcup MR. GAC63, a GRIP1‐dependent nuclear receptor coactivator. Mol Cell Biol 2005;25:5965–72. 113. Kim JH, Li H, Stallcup MR. CoCoA, a nuclear receptor coactivator which acts through an N‐terminal activation domain of p160 coactivators. Mol Cell 2003;12:1537–49. 114. Kim JH, Yang CK, Stallcup MR. Downstream signaling mechanism of the C‐terminal activation domain of transcriptional coactivator CoCoA. Nucleic Acids Res 2006;34:2736–50. 115. Lee DY, Northrop JP, Kuo MH, Stallcup MR. Histone H3 lysine 9 methyltransferase G9a is a transcriptional coactivator for nuclear receptors. J Biol Chem 2006;281:8476–85. 116. Choi HK, Choi KC, Oh SY, Kang HB, Lee YH, Haam S, et al. The functional role of the CARM1‐SNF5 complex and its associated HMT activity in transcriptional activation by thyroid hormone receptor. Exp Mol Med 2007;39:544–55. 117. Barrero MJ, Malik S. Two functional modes of a nuclear receptor‐recruited arginine methyltransferase in transcriptional activation. Mol Cell 2006;24:233–43. 118. Rizzo G, Renga B, Antonelli E, Passeri D, Pellicciari R, Fiorucci S. The methyl transferase PRMT1 functions as co‐activator of farnesoid X receptor (FXR)/9‐cis retinoid X receptor and regulates transcription of FXR responsive genes. Mol Pharmacol 2005;68:551–8. 119. Ananthanarayanan M, Li S, Balasubramaniyan N, Suchy FJ, Walsh MJ. Ligand‐dependent activation of the farnesoid X‐receptor directs arginine methylation of histone H3 by CARM1. J Biol Chem 2004;279:54348–57. 120. Le Romancer M, Treilleux I, Leconte N, Robin‐Lespinasse Y, Sentis S, Bouchekioua‐Bouzaghou K, et al. Regulation of estrogen rapid signaling through arginine methylation by PRMT1. Mol Cell 2008;31:212–21. 121. Yamagata K, Daitoku H, Takahashi Y, Namiki K, Hisatake K, Kako K, et al. Arginine methylation of FOXO transcription factors inhibits their phosphorylation by Akt. Mol Cell 2008;32:221–31. 122. Chen SL, Loffler KA, Chen D, Stallcup MR, Muscat GE. The coactivator‐associated arginine methyltransferase is necessary for muscle differentiation: CARM1 coactivates myocyte enhancer factor‐2. J Biol Chem 2002;277:4324–33. 123. Fauquier L, Duboe C, Jore C, Trouche D, Vandel L. Dual role of the arginine methyltransferase CARM1 in the regulation of c‐Fos target genes. FASEB J 2008;22:3337–47.

BIOLOGICAL FUNCTIONS OF PROTEIN ARGININE METHYLTRANSFERASES

339

124. Tabata T, Kokura K, Ten Dijke P, Ishii S. Ski co‐repressor complexes maintain the basal repressed state of the TGF‐beta target gene, SMAD7, via HDAC3 and PRMT5. Genes Cells 2008. 125. Furuno K, Masatsugu T, Sonoda M, Sasazuki T, Yamamoto K. Association of Polycomb group SUZ12 with WD‐repeat protein MEP50 that binds to histone H2A selectively in vitro. Biochem Biophys Res Commun 2006;345:1051–8. 126. Hou Z, Peng H, Ayyanathan K, Yan KP, Langer EM, Longmore GD, et al. The LIM protein AJUBA recruits protein arginine methyltransferase 5 to mediate SNAIL‐dependent transcriptional repression. Mol Cell Biol 2008;28:3198–207. 127. Langer EM, Feng Y, Zhaoyuan H, Rauscher III FJ, Kroll KL, Longmore GD. Ajuba LIM proteins are snail/slug corepressors required for neural crest development in Xenopus. Dev Cell 2008;14:424–36. 128. Lacroix M, Messaoudi SE, Rodier G, Le Cam A, Sardet C, Fabbrizio E. The histone‐binding protein COPR5 is required for nuclear functions of the protein arginine methyltransferase PRMT5. EMBO Rep 2008;9:452–8. 129. Jansson M, Durant ST, Cho EC, Sheahan S, Edelmann M, Kessler B, et al. Arginine methylation regulates the p53 response. Nat Cell Biol 2008;10:1431–9. 130. Covic M, Hassa PO, Saccani S, Buerki C, Meier NI, Lombardi C, et al. Arginine methyltransferase CARM1 is a promoter‐specific regulator of NF‐kappaB‐dependent gene expression. EMBO J 2005;24:85–96. 131. Hassa PO, Covic M, Bedford MT, Hottiger MO. Protein arginine methyltransferase 1 coactivates NF‐kappaB‐dependent gene expression synergistically with CARM1 and PARP1. J Mol Biol 2008;377:668–78. 132. Ganesh L, Yoshimoto T, Moorthy NC, Akahata W, Boehm M, Nabel EG, et al. Protein methyltransferase 2 inhibits NF‐kappaB function and promotes apoptosis. Mol Cell Biol 2006;26:3864–74. 133. Yoshimoto T, Boehm M, Olive M, Crook MF, San H, Langenickel T, et al. The arginine methyltransferase PRMT2 binds RB and regulates E2F function. Exp Cell Res 2006;312:2040–53. 134. Frietze S, Lupien M, Silver PA, Brown M. CARM1 regulates estrogen‐stimulated breast cancer growth through up‐regulation of E2F1. Cancer Res 2008;68:301–6. 135. Bres V, Yoh SM, Jones KA. The multi‐tasking P‐TEFb complex. Curr Opin Cell Biol 2008;20:334–40. 136. Amente S, Napolitano G, Licciardo P, Monti M, Pucci P, Lania L, et al. Identification of proteins interacting with the RNAPII FCP1 phosphatase: FCP1 forms a complex with arginine methyltransferase PRMT5 and it is a substrate for PRMT5‐mediated methylation. FEBS Lett 2005;579:683–9. 137. Boffa LC, Karn J, Vidali G, Allfrey VG. Distribution of NG, NG,‐dimethylarginine in nuclear protein fractions. Biochem Biophys Res Commun 1977;74:969–76. 138. Liu Q, Dreyfuss G. In vivo and in vitro arginine methylation of RNA‐binding proteins. Mol Cell Biol 1995;15:2800–8. 139. Rho J, Choi S, Jung CR, Im DS. Arginine methylation of Sam68 and SLM proteins negatively regulates their poly(U) RNA binding activity. Arch Biochem Biophys 2007;466:49–57. 140. Cote J, Boisvert FM, Boulanger MC, Bedford MT, Richard S. Sam68 RNA binding protein is an in vivo substrate for protein arginine N‐methyltransferase 1. Mol Biol Cell 2003;14:274–87. 141. Iwasaki H. Involvement of PRMT1 in hnRNPQ activation and internalization of insulin receptor. Biochem Biophys Res Commun 2008;372:314–9. 142. Swiercz R, Person MD, Bedford MT. Ribosomal protein S2 is a substrate for mammalian PRMT3 (protein arginine methyltransferase 3). Biochem J 2005;386:85–91. 143. Cheng D, Cote J, Shaaban S, Bedford MT. The arginine methyltransferase CARM1 regulates the coupling of transcription and mRNA processing. Mol Cell 2007;25:71–83.

340

KUHN AND XU

144. Fujiwara T, Mori Y, Chu DL, Koyama Y, Miyata S, Tanaka H, et al. CARM1 regulates proliferation of PC12 cells by methylating HuD. Mol Cell Biol 2006;26:2273–85. 145. Meister G, Eggert C, Buhler D, Brahms H, Kambach C, Fischer U. Methylation of Sm proteins by a complex containing PRMT5 and the putative U snRNP assembly factor pICln. Curr Biol 2001;11:1990–4. 146. Weiss VH, McBride AE, Soriano MA, Filman DJ, Silver PA, Hogle JM. The structure and oligomerization of the yeast arginine methyltransferase, Hmt1. Nat Struct Biol 2000;7:1165–71. 147. Troffer‐Charlier N, Cura V, Hassenboehler P, Moras D, Cavarelli J. Functional insights from structures of coactivator‐associated arginine methyltransferase 1 domains. EMBO J 2007;26:4391–401. 148. Troffer‐Charlier N, Cura V, Hassenboehler P, Moras D, Cavarelli J. Expression, purification, crystallization and preliminary crystallographic study of isolated modules of the mouse coactivator‐associated arginine methyltransferase 1. Acta Crystallogr Sect F Struct Biol Cryst Commun 2007;63:330–3. 149. Zhang X, Cheng X. Structure of the predominant protein arginine methyltransferase PRMT1 and analysis of its binding to substrate peptides. Structure 2003;11:509–20. 150. Zhang X, Zhou L, Cheng X. Crystal structure of the conserved core of protein arginine methyltransferase PRMT3. EMBO J 2000;19:3509–19. 151. Min JR, Wu H, Zeng H, Loppnau P, Sundstrom M, Arrowsmith CH, et al. Human HMT1 hnRNP methyltransferase‐like 3 (S. cerevisiae) protein. In Structural Genomics Consortium PDB ID: 2FYT ed. 2006. 152. Lin Q, Jiang F, Schultz PG, Gray NS. Design of allele‐specific protein methyltransferase inhibitors. J Am Chem Soc 2001;123:11608–13. 153. Cheng D, Yadav N, King RW, Swanson MS, Weinstein EJ, Bedford MT. Small molecule regulators of protein arginine methyltransferases. J Biol Chem 2004;279:23892–9. 154. Spannhoff A, Heinke R, Bauer I, Trojer P, Metzger E, Gust R, et al. Target‐based approach to inhibitors of histone arginine methyltransferases. J Med Chem 2007;50:2319–25. 155. Ragno R, Simeoni S, Castellano S, Vicidomini C, Mai A, Caroli A, et al. Small molecule inhibitors of histone arginine methyltransferases: homology modeling, molecular docking, binding mode analysis, and biological evaluations. J Med Chem 2007;50:1241–53. 156. Mai A, Cheng D, Bedford MT, Valente S, Nebbioso A, Perrone A, et al. epigenetic multiple ligands: mixed histone/protein methyltransferase, acetyltransferase, and class III deacetylase (sirtuin) inhibitors. J Med Chem 2008;51:2279–90. 157. Spannhoff A, Machmur R, Heinke R, Trojer P, Bauer I, Brosch G, et al. A novel arginine methyltransferase inhibitor with cellular activity. Bioorg Med Chem Lett 2007;17:4150–3. 158. Purandare AV, Chen Z, Huynh T, Pang S, Geng J, Vaccaro W, et al. Pyrazole inhibitors of coactivator associated arginine methyltransferase 1 (CARM1). Bioorg Med Chem Lett 2008;18:4438–41. 159. Torres‐Padilla ME, Parfitt DE, Kouzarides T, Zernicka‐Goetz M. Histone arginine methylation regulates pluripotency in the early mouse embryo. Nature 2007;445:214–8. 160. Durcova‐Hills G, Tang F, Doody G, Tooze R, Surani MA. Reprogramming primordial germ cells into pluripotent stem cells. PLoS ONE 2008;3:e3531. 161. Pawlak MR, Scherer CA, Chen J, Roshon MJ, Ruley HE. Arginine N‐methyltransferase 1 is required for early postimplantation mouse development, but cells deficient in the enzyme are viable. Mol Cell Biol 2000;20:4859–69. 162. Yadav N, Lee J, Kim J, Shen J, Hu MC, Aldaz CM, et al. Specific protein methylation defects and gene expression perturbations in coactivator‐associated arginine methyltransferase 1‐deficient mice. Proc Natl Acad Sci USA 2003;100:6464–8.

BIOLOGICAL FUNCTIONS OF PROTEIN ARGININE METHYLTRANSFERASES

341

163. Cheung N, Chan LC, Thompson A, Cleary ML, So CW. Protein arginine‐methyltransferase‐ dependent oncogenesis. Nat Cell Biol 2007;9:1208–15. 164. Kim J, Lee J, Yadav N, Wu Q, Carter C, Richard S, et al. Loss of CARM1 results in hypomethylation of thymocyte cyclic AMP‐regulated phosphoprotein and deregulated early T cell development. J Biol Chem 2004;279:25339–44. 165. Wang L, Pal S, Sif S. Protein arginine methyltransferase 5 suppresses the transcription of the RB family of tumor suppressors in leukemia and lymphoma cells. Mol Cell Biol 2008;28:6262–77. 166. Jiang W, Roemer ME, Newsham IF. The tumor suppressor DAL‐1/4.1B modulates protein arginine N‐methyltransferase 5 activity in a substrate‐specific manner. Biochem Biophys Res Commun 2005;329:522–30. 167. Jiang W, Newsham IF. The tumor suppressor DAL‐1/4.1B and protein methylation cooperate in inducing apoptosis in MCF‐7 breast cancer cells. Mol Cancer 2006;5:4. 168. Majumder S, Liu Y, Ford III OH, Mohler JL, Whang YE. Involvement of arginine methyltransferase CARM1 in androgen receptor function and prostate cancer cell viability. Prostate 2006;66:1292–301. 169. Jeong SJ, Lu H, Cho WK, Park HU, Pise‐Masison C, Brady JN. Coactivator‐associated arginine methyltransferase 1 enhances transcriptional activity of the human T‐cell lymphotropic virus type 1 long terminal repeat through direct interaction with Tax. J Virol 2006;80:10036–44. 170. Boulanger MC, Liang C, Russell RS, Lin R, Bedford MT, Wainberg MA, et al. Methylation of Tat by PRMT6 regulates human immunodeficiency virus type 1 gene expression. J Virol 2005;79:124–31. 171. Xie B, Invernizzi CF, Richard S, Wainberg MA. Arginine methylation of the human immunodeficiency virus type 1 Tat protein by PRMT6 negatively affects Tat Interactions with both cyclin T1 and the Tat transactivation region. J Virol 2007;81:4226–34. 172. Invernizzi CF, Xie B, Frankel FA, Feldhammer M, Roy BB, Richard S, et al. Arginine methylation of the HIV‐1 nucleocapsid protein results in its diminished function. AIDS 2007;21:795–805. 173. Altschuler L, Wook JO, Gurari D, Chebath J, Revel M. Involvement of receptor‐bound protein methyltransferase PRMT1 in antiviral and antiproliferative effects of type I interferons. J Interferon Cytokine Res 1999;19:189–95. 174. Zika E, Fauquier L, Vandel L, Ting JP. Interplay among coactivator‐associated arginine methyltransferase 1, CBP, and CIITA in IFN‐gamma‐inducible MHC‐II gene expression. Proc Natl Acad Sci USA 2005;102:16321–6. 175. Krones‐Herzig A, Mesaros A, Metzger D, Ziegler A, Lemke U, Bruning JC, et al. Signal‐ dependent control of gluconeogenic key enzyme genes through coactivator‐associated arginine methyltransferase 1. J Biol Chem 2006;281:3025–9. 176. Powell E, Kuhn P, Xu W. Nuclear receptor cofactors in PPARgamma‐mediated adipogenesis and adipocyte energy metabolism. PPAR Res 2007;2007:53843. 177. Yadav N, Cheng D, Richard S, Morel M, Iyer VR, Aldaz CM, et al. CARM1 promotes adipocyte differentiation by coactivating PPARgamma. EMBO Rep 2008;9:193–8. 178. Christian M, Kiskinis E, Debevec D, Leonardsson G, White R, Parker MG. RIP140‐targeted repression of gene expression in adipocytes. Mol Cell Biol 2005;25:9383–91. 179. Mostaqul Huq MD, Gupta P, Tsai NP, White R, Parker MG, Wei LN. Suppression of receptor interacting protein 140 repressive activity by protein arginine methylation. EMBO J 2006;25:5094–104. 180. Le Guezennec X, Vermeulen M, Brinkman AB, Hoeijmakers WA, Cohen A, Lasonder E, et al. MBD2/NuRD and MBD3/NuRD, two distinct complexes with different biochemical and functional properties. Mol Cell Biol 2006;26:843–51.

342

KUHN AND XU

181. Tan CP, Nakielny S. Control of the DNA methylation system component MBD2 by protein arginine methylation. Mol Cell Biol 2006;26:7224–35. 182. Boisvert FM, Hendzel MJ, Masson JY, Richard S. Methylation of MRE11 regulates its nuclear compartmentalization. Cell Cycle 2005;4:981–9. 183. Dery U, Coulombe Y, Rodrigue A, Stasiak A, Richard S, Masson JY. A glycine‐arginine domain in control of the human MRE11 DNA repair protein. Mol Cell Biol 2008;28:3058–69. 184. El‐Andaloussi N, Valovka T, Toueille M, Steinacher R, Focke F, Gehrig P, et al. Arginine methylation regulates DNA polymerase beta. Mol Cell 2006;22:51–62. 185. Miranda TB, Webb KJ, Edberg DD, Reeves R, Clarke S. Protein arginine methyltransferase 6 specifically methylates the nonhistone chromatin protein HMGA1a. Biochem Biophys Res Commun 2005;336:831–5. 186. Zou Y, Webb K, Perna AD, Zhang Q, Clarke S, Wang Y. A mass spectrometric study on the in vitro methylation of HMGA1a and HMGA1b proteins by PRMTs: methylation specificity, the effect of binding to AT‐rich duplex DNA, and the effect of C‐terminal phosphorylation. Biochemistry 2007;46:7896–906. 187. Huang S, Li X, Yusufzai TM, Qiu Y, Felsenfeld G. USF1 recruits histone modification complexes and is critical for maintenance of a chromatin barrier. Mol Cell Biol 2007;27:7991–8002.

Roles of Histone H3‐Lysine 4 Methyltransferase Complexes in NR‐Mediated Gene Transcription Seunghee Lee,* Robert G. Roeder,{ and Jae W. Lee* *Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas 77030 {

Laboratory of Biochemistry and Molecular Biology, The Rockefeller University, New York 10021

I. Introduction ................................................................................ II. Activating Signal Cointegrator‐2 (ASC‐2)............................................ A. Expression of ASC‐2 and Its Isoforms ........................................... B. Autonomous Transactivation Domains of ASC‐2 .............................. C. Two NR Boxes in ASC‐2 ............................................................ D. Homodimerization Domain in ASC‐2............................................ III. Set1‐Like H3K4MT Complexes........................................................ A. Methylation of H3K4 ................................................................ B. Multiple Set1‐Like H3K4MT Complexes in Vertebrates .................... C. A Subcomplex of WDR5, RbBP5, and ASH2L in Set1‐Like Complexes ................................................................ D. H3K4 Methyl‐Binding Effectors .................................................. IV. ASCOM in NR‐Mediated Transactivation ........................................... A. ASCOM‐MLL3 and ASCOM‐MLL4 ............................................ B. ASCOMs as Crucial H3K4MT Complexes for a Subset of NRs ........... C. Recruitment of Set1‐Like H3K4MT Complexes............................... V. Cross Talk of ASCOMs with Other Coactivators ................................... A. CBP/p300............................................................................... B. RNA‐Binding Proteins............................................................... C. Swi/Snf .................................................................................. VI. Physiological Roles of Key Subunits of ASCOM ................................... A. ASC‐2 ................................................................................... B. Metabolic Phenotypes of MLL3................................................... C. ASCOM in Cancers .................................................................. VII. Future Challenges ........................................................................ References ..................................................................................

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Transcriptional regulation by nuclear hormone receptors (NRs) requires multiple coregulators that modulate chromatin structures by catalyzing a diverse array of posttranslational modifications of histones. Different combinations of these modifications yield dynamic functional outcomes, constituting an Progress in Molecular Biology and Translational Science, Vol. 87 DOI: 10.1016/S1877-1173(09)87010-5

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epigenetic histone code. This code is inscribed by histone‐modifying enzymes and decoded by effector proteins that recognize specific covalent marks. One important modification associated with active chromatin structures is methylation of histone H3‐lysine 4 (H3K4). Crucial roles for this modification in NR transactivation have been recently highlighted through our purification and subsequent characterization of a steady‐state complex associated with ASC‐2, a coactivator of NRs and other transcription factors. This complex, designated ASCOM for ASC‐2 complex, contains H3K4‐methyltransferase MLL3/HALR or its paralogue MLL4/ALR and represents the first Set1‐like H3K4‐methyltransferase complex to be reported in vertebrates. This review focuses on recent progress in our understanding of how ASCOM‐MLL3 and ASCOM‐ MLL4 influence NR‐mediated gene transcription and of their physiological function.

I. Introduction Nuclear hormone receptors (NRs) share a common modular structure comprised an N‐terminal variable domain, a central DNA‐binding domain (DBD), and a C‐terminal ligand‐binding domain (LBD).1 While the LBD harbors a ligand‐dependent activation function referred to as AF2, the N‐terminal domains of some NRs harbor AF1, a constitutive activation function. A variety of endocrine hormones, fatty acids, cholesterol, and lipid metabolites act as ligands for various NRs. During ligand‐dependent transcriptional regulation by NRs, multiple transcriptional coregulators have been demonstrated to operate in a ligand‐dependent, combinatorial manner, as dictated by the context of the individual target gene and cell type.2–4 Importantly, genes are normally compacted in closed chromatin structures in a transcriptionally inactive state.5 Thus, the establishment of open chromatin structures is an essential step toward successful transactivation. Consistent with this notion, NR transactivation depends on the ability of NR to recruit multiple coactivators whose primary function is to remodel or modify chromatin structures. In particular, histone‐modifying enzymes catalyze a diverse array of posttranslational modifications of core and linker histones within chromatin.6,7 Interestingly, different combinations of these modifications appear to yield dynamic functional outcomes, constituting an epigenetic histone code. While this code is inscribed by histone‐modifying enzymes catalyzing site‐selective modifications, it is proposed to be decoded by effector proteins that recognize specific covalent marks.6,7 Methylation of histone H3‐lysine 4 (H3K4) is one particular histone modification that has been recently highlighted for its roles in NR transactivation through our characterization of ASCOM, a novel NR coactivator complex

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associated with H3K4‐methyltransferase (H3K4MT) MLL3/HALR or its paralogue MLL4/ALR.8,9 ASCOM belongs to a family of Set1‐like H3K4MT complexes (Table I). It has been proposed that ASC‐2 may play crucial roles in reproduction and a variety of endocrine functions.10 Our recent results also implicate ASCOM as a Set1‐like complex specialized to regulate genes involved with metabolic homeostasis.11,12 Moreover, ASCOM contains another histone modifier, UTX, that removes trimethylated‐H3K27, a mark for inactive chromatin structures.13–17 Thus, ASCOM is associated with two histone modifiers linked to transactivation and should serve as an excellent model system to study the potential cross talk between H3K4 and H3K27‐methylation events. Interestingly, ASCOM appears to have a unique function as a platform for integrating the activities of several other coactivators,10 including the ATPase‐ dependent Swi/Snf chromatin remodeling complexes (see below), during NR transactivation. Here, we present an overview of current literature on the function of ASCOM‐MLL3 and ASCOM‐MLL4 in NR transactivation and further discuss future challenges in fully elucidating their physiological functions.

TABLE I A FAMILY OF SET1‐LIKE COMPLEXES ySet1_C

hSet1_C

MLL1_C

MLL2_C

ASCOM

Set1

hSet1a/b

MLL1

MLL2

MLL3/4

Bre2

ASH2L

ASH2L

ASH2L

ASH2L

Swd1

RbBP5

RbBP5

RbBP5

RbBP5

Swd3

WDR5

WDR5

WDR5

WDR5

Sdc1

hDPY‐30

hDPY‐30

hDPY‐30

hDPY‐30

Menin

Menin

ASC‐2

Swd2

hSwd2

Spp1

CXXC1 HCF1

HCF1/HCF2

PTIP PA1 UTX a/b‐Tubulins

The yeast complex (left) is aligned with six human complexes. Three common subunits, which assemble an independent subcomplex, are shaded. This subcomplex forms a functional core H3K4MT complex, along with each individual H3K4MT enzyme. Unique subunits in each complex are shown in bold. Notably, additional subunit proteins may remain unidentified in each complex.

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II. Activating Signal Cointegrator‐2 (ASC‐2) NRs bind to specific response elements in target genes and regulate transcriptional initiation in a ligand‐dependent manner.1 In the unliganded state, a subset of the NRs repress transcription by recruiting corepressors.2–4 Upon ligand binding, the conserved C‐terminal LBD of NRs undergoes a dramatic conformational change, which is recognized by an a‐helical LXXLL motif, named the NR box, which is often found associated with transcriptional coactivators.18,19 ASC‐2, also named NCOA6 (nuclear receptor coactivator‐6), TRBP (thyroid hormone receptor‐binding protein), RAP250 (nuclear receptor‐activating protein‐250), NRC (nuclear receptor coregulator), and PRIP (peroxisome proliferator‐activated receptor‐interacting protein), has been shown to function as a coactivator of many NRs.10 In addition, AIB3 (amplified in breast cancer‐3) has been identified as a human ASC‐2 isoform in which the N‐terminal 26 amino acids replace the first 88 amino acids of ASC‐2. Of note, ASC‐2 has two NR interaction boxes10 (Fig. 1A). While NR1 binds

A

DD (849−995)

1528

ARID

2063

AD2 AD3 (622−783) (940−1057) 970 1043

259 305

1 67

AD1 (1−218)

ASC-2

D/E Q-rich

LxxLL

Q/P-rich

S/T-rich

1

2000

B

5262

LxxLL

MLL4 PHD

PHD

HMG

SET W R

A

FIG. 1. Schematic representations of ASC‐2 and MLL4. Various domains that include the AD1–3, DD (dimerization domain), and ARID (AR‐interacting domain) in ASC‐2, as well as the PHD finger, HMG box, and SET domains in MLL4, are as indicated. The SET domain in MLL3/4 also serves as a docking site for a subcomplex consisting of WDR5 (W)‐RbBP5 (R)‐ASH2L (A).

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multiple NRs, NR2 interacts primarily with the liver X receptors (LXRs).20 The physiological importance of ASC‐2 and its two NR boxes in transactivation by ASC‐2‐interacting NRs has been documented by studies with various ASC‐ 2 mouse models.21–27 Notably, ASC‐2 also functions as an important coactivator for an array of other classes of transcription factors.10

A. Expression of ASC‐2 and Its Isoforms ASC‐2 is a 250 kDa protein of 2063 amino acids that is expressed from a
8–9 kb mRNA in different human tissues.28 ASC‐2 mRNAs of 6.8, 4.5, and 3.6 kb also have been described.28 While the 3.6 kb transcript is the predominant ASC‐2 mRNA species detected in heart and skeletal muscle, the 4.5 kb transcript is the major form in testes.28 The latter transcript has been identified in human testis as an alternatively spliced form of ASC‐2 mRNA that encodes an ASC‐2 isoform of 1070 amino acids.29 This isoform lacks the NR box 2 and a large internal region (amino acids 972–1964) toward the C‐terminus. Mouse ASC‐2 mRNA (
8–9 kb) encodes a protein of 2068 amino acids.28 A partial rat ASC‐2 cDNA encoding the NR box 1 and AD3 followed by a stop codon and a poly A tail has also been isolated from a rat pituitary GH4C1 cell cDNA library.28 The mouse ASC‐2 gene, containing 13 exons, is localized on chromosome 2,28 while rat ASC‐2 resides on chromosome 3. The organization of human ASC‐2 on chromosome 20q11 predicts a single gene, with 15 exons and 14 introns, that spans
111 kb.30 Thus, the various sized ASC‐2 mRNAs described above likely represent alternative splicing products of the ASC‐2 gene. The sequence of the ASC‐2 gene predicts a TATA‐less promoter with four binding sites for Sp1, as well as sites for C/EBP and Myc/Max.30 However, the functional significance of these binding sites in regulating ASC‐2 gene expression remains to be established. Human, mouse, and rat ASC‐2 proteins show over 90% sequence identity at the amino acid level. The distribution of the ASC‐2 protein in mice has been studied by immunochemistry using antibody against ASC‐2. These studies indicate relatively higher levels of ASC‐2 in cells of endocrine target tissues, including testicular sertoli cells, follicular granulosa cells, and epithelial cells of the prostate, uterus, mammary gland, kidney tubules and urothelia23,31 (PNAS 2009;106:8513-8, and our unpublished results). ASC‐2 protein expression has also been detected in thyroid and parathyroid cells and in the pancreatic islets of langerhans.23 Although medium to low expression is reported throughout a variety of tissues, the relatively higher levels of ASC‐2 observed in reproductive and many types of endocrine tissues suggest that ASC‐2 supports NR functions in reproduction and regulation of the endocrine system.23,31 ASC‐2 also is relatively abundant in a variety of metabolic tissues and cell types, including

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the liver, pancreas, adipose tissues, and hypothalamic metabolic neurons23,31 (our unpublished results). These results are consistent with our proposed roles for ASCOM in metabolic homeostasis (see below).

B. Autonomous Transactivation Domains of ASC‐2 Structure–function analysis of human ASC‐2 has revealed a number of functional domains. First, we have mapped three autonomous transactivation domains, AD1, AD2, and AD3, that in human ASC‐2 comprise residues 1–218, 622–849, and 849–1057, respectively.20 Mahajan and Samuels have also mapped two autonomous transactivation domains in ASC‐2 to residues 302– 783 and 940–1124.28 Thus, the boundaries of AD2 and AD3 are likely to be refined to ASC‐2 residues 622–783 and 940–1057, respectively (Fig. 1A). Interestingly, while AD2 is dispensable for NR transactivation, the AD3 domain appears to play crucial roles for NR signaling and AF2‐dependent enhancement of NR transactivation by ASC‐2.32 The
600 residue ASC‐2 C-terminal region that is rich in Ser, Thr, and Leu (Fig. 1A) inhibits the autonomous transactivation function of ASC‐2 in the context of full length ASC‐2, as its deletion enhances the intrinsic activation potential of AD1, AD2, and AD3 in ASC‐2.28 Notably, this region contains an array of putative phosphorylation sites for different protein kinases, raising the interesting possibility that the activity of ASC‐2 may potentially be modulated by kinases and phosphatases. These results also suggest that this C‐terminal region may function to control the transcriptional output of ASC‐2 through inter‐ and intramolecular interactions. One interesting possibility is that the activation properties of ASC‐2 could be modulated through exposure of the cryptic activation domain in response to liganded NRs, as previously exemplified by the finding that PGC‐1 undergoes a conformational change upon NR binding.33 In support of this notion, a LexA fusion protein containing ASC‐2 AD2 and AD3 domains is transcriptionally mild in yeast but shows a dramatically increased activity when coexpressed with RXR in the presence of its ligand. Similar results are also observed in mammalian cells. When an ASC‐2 region containing the NR1 and AD2 regions is expressed in mammalian cells, it is moderately active when expressed alone. However, its activity is markedly enhanced when coexpressed with the liganded TR.28 These results lead to a proposal that the NR‐coactivator function of ASC‐2 is masked until it associates with liganded NRs.28

C. Two NR Boxes in ASC‐2 ASC‐2 contains three distinct NR‐interacting surfaces. Two distinct NR boxes mediate direct interactions of ASC‐2 with a variety of NRs.10 In addition, we have shown that ASC‐2 contains a binding site for the retinoblastoma (Rb)

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tumor suppressor protein,34 which is known to interact with androgen receptor (AR).35 We have further shown that this region indeed functions as an indirect binding site for AR, as it recruits AR via Rb.34 1. NR1 Two functional NR boxes have been identified in ASC‐2 (Fig. 1A). The NR box 1 shows ligand‐dependent interactions with a wide variety of NRs that include RARs, RXRs, TRs, GR, ERs, VDR, and the PPARs.28,29,36 Accordingly, mutation of the leucine residues in NR1 abolishes the association of ASC‐2 with NRs. Through the use of combinatorial peptide libraries, the NR boxes have been characterized as Class 1, 2, or 3 depending on the nature of the amino acid residue at position 1 or 2 of the LXXLL motif.37 The Class 2 NR box with Pro at 2 has been shown to interact with various NRs. The NR box 1 region of ASC‐2 (LTSPLLVNLLQSDIS) resembles the Class 2 NR box and contains a Pro at the 2 position. Like other NR box‐dependent coactivators, helix 12 of the NR AF2 domain is pivotal in the formation of an interface that contacts the ASC‐2 NR1. Thus, either a point mutation (L398R) in helix 12 of chicken TRa or a deletion of helix 12 abolishes the association with ASC‐2.28 Deletion of helix 12 of PPARg also disrupts the interaction with ASC‐2.38 We have found that the interactions of TRb with the NR1 of ASC‐2 are abolished by specific mutations in the TRb helices 3, 5, and 6,28 which are known to form the binding pocket for SRC‐1.39,40 Overall, these results indicate that the ASC‐2:NR interactions are similar to those of other coactivators. The Ser at the 3 position (Ser 884) of the ASC‐2 NR1 has been demonstrated to be important for binding to LBDs of NRs, particularly for ERb, TR, and RXR.41 Although in vitro phosphorylation of Ser 884 by MAPK reduces its interaction with NRs, it remains to be determined whether such regulation by phosphorylation occurs in vivo.41 Interestingly, the ASC‐2 NR1 has been shown to associate more strongly with mouse ERb1 than with ERb2, consistent with the notion that mouse ERb1 is a more potent estradiol‐induced transcriptional activator than mouse ERb2.42 Studies with a
300 amino acid rat ASC‐2 region that contains NR1 and AD3 indicate that NR1 enhances both the ligand‐dependent activity and the intrinsic basal activity of AD3.32 Thus, the NR1 of ASC‐2 embedded in AD3 appears to influence not only the association of ASC‐2 with NRs, but also the activation potential of AD3. Correspondingly, the AD3 region has been shown to be necessary for transcriptional activation by NRs.32 2. NR2 ASC‐2 contains a second NR box (Fig. 1A) that does not contain a conserved hydrophobic amino acid residue at the 1 position (EAPTSLSQLLDNSGA).28 While NR2 strongly interacts with LXRs,20 it does not recognize most NRs other

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than ERa, which shows
10‐fold lower affinity toward NR2 compared with NR1.28 The ASC‐2 NR2 does not resemble any of the known NR boxes identified thus far in various NR coactivators or from combinatorial phage peptide libraries.37 RXR has been shown to enhance LXR transactivation, even in the absence of an LXR ligand, via a unique mechanism of allosteric regulation.43 Interestingly, we have found that LXR binding to the ASC‐2 NR2 is enhanced by RXR and even more substantially by liganded RXR.44 We also have identified specific NR2 region residues that are involved both in its interaction with LXR and in the ASC‐2‐mediated transactivation of LXR in mammalian cells. Using these mutants, we have demonstrated that the NR2–LXR interaction surface is not altered by the presence of RXR and RXR ligand and that Ser 1490 is the critical determinant for the LXR‐specific interaction of NR2.44 Notably, NR2, but not NR1, is essential for ASC‐2‐mediated transactivation of LXR in vivo and for the interaction between LXR–RXR and ASC‐2 in vitro. These results indicate that RXR does not interact directly with the NR1 of ASC‐2, but functions as an allosteric activator of LXR binding to the ASC‐2 NR2.44

D. Homodimerization Domain in ASC‐2 Most NRs activate transcription from target genes either as homodimers or as heterodimers with RXR.1 SRC family coactivators contain multiple NR boxes, such that a single SRC coactivator could interact simultaneously, through distinct NR boxes, with various NR dimmers.45 Similarly, the TRAP220/MED1 Mediator subunit has two NR boxes. One NR box interacts with RXR, while the other interacts preferentially with VDR or TR.46 Thus, a TRAP220/MED1 monomer has the potential to bind to NR–RXR heterodimers through two interfaces. However, ASC‐2 contains a single NR box that is involved in interactions with multiple NRs. Potentially resolving the issue of how ASC‐2 with a single functional NR box binds to and activates NR dimers, ASC‐2 has been found to homodimerize.28 The homodimerization region (DD) of ASC‐2 (Fig. 1A) has been mapped to a region of human ASC‐2 containing amino acid 849–995.28,32 Embedded in this region is the NR1 motif, which is involved in NR interactions, but not homodimerization of ASC‐2. Thus, a homodimer of ASC‐2 with two functional NR box motifs has the potential to bind NR homo‐ and heterodimers with high affinity.28,32 However, in vivo evidence for formation of an ASC‐2 homodimer has yet to be established.

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III. Set1‐Like H3K4MT Complexes A. Methylation of H3K4 H3K4‐methylation is an evolutionarily conserved mark that is linked to transcriptionally active chromatin and has been proposed to counter the generally repressive chromatin environment imposed by H3K9/H3K27‐methylation in higher eukaryotes.5 H3K4‐methylation has been demonstrated to be associated with transcriptional activation in a variety of eukaryotic species.47,48 Lysine residues can be mono‐, di‐, or trimethylated at the z‐amine in vivo. In particular, H3K4‐trimethylation is tightly associated with the 50 regions of transcriptionally active genes, as it shows a strong positive correlation with transcription rates, active polymerase II occupancy, and histone acetylation.49–54 Interestingly, the patterns of H3K4‐dimethylation differ significantly between yeast and vertebrates. In Saccharomyces cerevisiae, dimethylated H3K4 is found throughout genes that, transcriptionally, are either active or poised and peaks around the middle of the coding region, whereas monomethylation is most abundant at the 30 end of genes.49,50,54 In vertebrates, the majority of H3K4‐ dimethylation is coupled to H3K4‐trimethylation in discrete zones about 5–20 nucleosomes in length proximate to highly transcribed genes.51,52

B. Multiple Set1‐Like H3K4MT Complexes in Vertebrates The majority of histone lysine methyltransferases contain a SET domain, which catalyzes the addition of methyl groups to the specific lysine residues.48 The catalytic SET domains of H3K4MTs can be arbitrarily grouped into either Set1‐like H3K4MTs, which are related to the yeast Set1 and Drosophila Trx, or non‐Set1‐like H3K4MTs, which include ASH1, SET7/9, SMYD3, and Meisetz.55–59 The first H3K4MT to be identified is S. cerevisiae Set1, a single enzyme responsible for all H3K4‐methylation in yeast.60,61 Interestingly, this enzyme is found as a component of a large steady‐state Set1 complex62,63 (Table I). We subsequently described a similar mammalian complex containing mixed lineage leukemia 3 (MLL3, also named HALR) or MLL4 (also named MLL2/ALR),8,9 and others later described homologous vertebrate complexes containing MLL1, MLL2, SET1a, or SET1b64–68 (Table I). These evolutionarily conserved complexes are collectively named ‘‘Set1‐like H3K4MT complexes’’ (Table I). The multiplicity of Set1‐like H3K4MT complexes in vertebrate genomes suggests that these complexes are not redundant in their function. In strong support of this notion, MLL1, MLL2, and MLL3 mutant mice display distinguishable phenotypes.9,11,12,64,69 The potential functional

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specialization of these enzymes likely results from their differential expression patterns, recruitment to different target genes, and/or methylation of distinct nonhistone substrates.

C. A Subcomplex of WDR5, RbBP5, and ASH2L in Set1‐Like Complexes The Set1‐like H3K4MT complexes share at least three common subunits: WDR5, RbBP5, and ASH2L8,65–68,70 (Table I). Our recent biochemical reconstitution of a functional four‐component MLL1 core complex reveals that recombinant WDR5, RbBP5, ASH2L, and MLL1 are sufficient for recapitulating H3K4MT activity comparable to that of the MLL1 holocomplex purified from human cells.71 Importantly, the subcomplex of WDR5, RbBP5, and ASH2L associates with the MLL1 SET domain (Fig. 1B) but can exist independently of the catalytic subunit, thereby serving as a structural platform that can associate with the SET domains of different MLL‐family members8,71 (Fig. 2A). All three subcomplex components are required for H3K4‐methylation by MLL1 in vitro and in vivo,71,72 with different components affecting

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methyl states to different degrees.71,73,74 In particular, WDR5 mediates interactions of the MLL1 catalytic unit both with the common structural platform and with the histone substrate71 (Fig. 1B).

D. H3K4 Methyl‐Binding Effectors Although H3K4 methylation has been proposed to function through countering the generally repressive chromatin environment imposed by H3K9/ H3K27‐methylation in higher eukaryotes,5 its precise role in transcription remains poorly understood. Like other histone modifications, histone methylation is proposed to act through the recruitment of downstream effector proteins, which in turn carry out specific independent functions on the chromatin template. Indeed, recent work has uncovered a remarkably wide variety of domains that have the capacity to recognize methylated H3‐lysine residues.75 In particular, at least two distinct motifs have been shown to bind H3K4‐ trimethyl residues: the royal superfamily (including the chromodomains of CHD1 and tudor domains of JMJD2A) and the PHD‐finger superfamily (including the PHD fingers of BPTF and ING proteins).76,77 Interestingly, many of these H3K4‐methyl‐binding proteins reside within protein complexes associated with chromatin remodeling and modifications. For example, CHD1 and BPTF are involved in ATP‐dependent chromatin remodeling,78 ING proteins associate with and modulate the activity of histone acetyltransferase (HAT) and deacetylase (HDAC) complexes,79 and JMJD2A carries out histone demethylation.80 Thus, H3K4‐methylation can be directly coupled to further chromatin remodeling and modification mechanisms necessary to carry out specific biological functions. The recruitment of remodeling machinery, such as the BPTF‐containing NURF remodeling complex, may facilitate transcriptional activation by increasing the accessibility of the chromatin template to the transcriptional machinery.81 The association of H3K4‐methyl with ING3–5 and yeast Yng1 containing acetyltransferase complexes is also consistent with the role of H3K4‐methylation in transcriptional activation.82–84 Histone acetylation directly promotes more accessible chromatin structure and can further recruit bromodomain‐containing transcriptional regulators.85 Interplay between H3K4‐ methylation and acetylation is evidenced by a high correlation of hyperacetylation and H3K4‐methylation patterns in genomic location analyses.52–54,83 In further support of this notion, we have recently demonstrated that loss of the H3K4‐methyl mark on RAR‐target gene RAR‐b2 in ASC‐2‐null cells is accompanied by loss of H3/H4‐acetylation marks.9 Intriguingly, recent findings demonstrate that upon DNA damage ING2 recruits the Sin3/HDAC complex to silence transcription of cell proliferation genes, thus implicating H3K4‐ trimethylation in active gene repression as well.82 In support of this idea, H3K4‐methyl‐binding PHD fingers exist in well‐established HDAC complexes

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in organisms ranging from yeast to humans.79 Interestingly, human CHD1 recognition of trimethylated H3K4 has been shown to function through the recruitment of factors implicated in transcriptional elongation and pre‐mRNA processing.86 A PHD finger within RAG2, a protein involved in V(D)J recombination, has been shown to recognize trimethylated H3K4 and these interactions subsequently have been shown to play important roles in V(D)J recombination.87–89 The latter two sets of results expand the functional scope for H3K4‐methylation beyond its well‐defined role in transcriptional initiation.

IV. ASCOM in NR‐Mediated Transactivation A. ASCOM‐MLL3 and ASCOM‐MLL4 Our proteomic analysis of ASC‐2 in HeLa nuclei has led to the surprising finding that ASC‐2 does not exist as a single polypeptide but belongs to a large (
2 MDa) steady‐state H3K4MT complex that is highly homologous to the yeast Set1 complex.8 This complex, which we named ASCOM, was the first mammalian Set1‐like H3K4MT complex to be described.8,9 ASCOM, which contains either MLL3 or MLL4 as the H3K4MT, shares the RbBP5, ASH2L, and WDR5 subcomplex with other Set1‐like complexes (Table I). The components unique to ASCOM include PTIP, PTIP‐associated protein 1 (PA1), a‐ and b‐tubulins, UTX, and possibly other additional proteins8,90,91 (Table I). Like Set1 and MLL1/2, the C‐termini of MLL3 and MLL4 have a SET domain (Fig. 1B) that is associated with an intrinsic histone lysine‐specific methyltransferase activity.5 We and others have shown that ASCOM is indeed a genuine H3K4MT complex.8,90,91 Interestingly, UTX has been found to be a H3K27‐ specific demethylase.13–17 Because H3K27‐trimethylation is a repressive chromatin mark critical for maintaining embryonic stem (ES) cell pluripotency and plasticity in developing embryos, polycomb‐mediated gene silencing, and X chromosome inactivation,92 ASCOM has two distinct histone modifiers linked to active chromatin. Virtually identical complexes have also been purified as MLL491 and PTIP90 complexes from K562 and HeLa cells, respectively. Notably, in our original purification, we have failed to determine the identity of UTX, PTIP, PA1, and WDR5 due to technical problems.8 In addition, a similar complex enriched in MLL4 has also been reported as a coactivator complex of ERa from DU4475 cells.93 This complex has been claimed to be distinct from ASCOM based on the relatively unaltered amount of RbBP5, ASH2L, and MLL4 in lysates immunodepleted for ASC‐2.93 However, the reported purification includes ASC‐2, although it is less abundant than MLL4.93 Moreover, RbBP5 and ASH2L are associated with other Set1‐like complexes (Table I)94 and thus are not readily coimmunodepleted with ASC‐2.

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In addition, currently it is not clear whether MLL4 can also be found outside of ASCOM, either as a single polypeptide or as a subunit of other distinct complexes. If MLL4 also exists unassociated with ASC‐2, it would not be coimmunodepleted with ASC‐2. Thus, it needs to be further examined whether this MLL4 complex93 is indeed distinct from ASCOM. Considering all the relevant studies, we favor a model in which a shared subcomplex of RbBP5, ASH2L, and WDR5 forms a core complex with MLL3 or MLL4 that may also form a heterogeneous population of complexes with ASC‐2 and other proteins depending on cell types and/or target genes (Fig. 2B).

B. ASCOMs as Crucial H3K4MT Complexes for a Subset of NRs Transactivation by RXR, RAR, TR, and PPARg has been shown to be compromised in ASC‐2/ mouse embryo fibroblast (MEF) cells relative to wild‐type cells.24,25,27,95 Moreover, we have found that RAR transactivation is correlated with RA‐induced H3K4‐trimethylation, and that this modification is redundantly mediated by ASCOM‐MLL3 and ASCOM‐MLL4 but not by related menin‐containing MLL1/2‐complexes (Table I), primarily due to the ability of ASC‐2 to function as a specific linker for RARs to recruit ASCOM.9 ASC‐2/ MEF cells are refractory to PPARg‐mediated adipogenesis and fail to express adipogenic markers such as aP2,95 suggesting that ASC‐2 is required for the adipogenic program by PPARg. In addition, our recent results reveal that MLL3 also plays crucial roles in adipogenesis.12 First, MLL3D/D mice expressing an H3K4MT inactivated mutant of MLL3 have significantly less white fat. Second, MLL3D/D MEFs are mildly but consistently less responsive to inducers of adipogenesis than wild‐type MEFs. Third, ASC‐2, MLL3, and MLL4 are recruited to the PPARg‐activated aP2 gene during adipogenesis, and PPARg is shown to interact directly with the purified ASCOM. Moreover, while H3K4‐methylation of aP2 is readily induced in wild‐type MEFs, it is not induced in ASC‐2/ MEFs and only partially induced in MLL3D/D MEFs.12 These results suggest that ASCOM‐MLL3 and ASCOM‐MLL4 likely function as crucial but redundant H3K4MT complexes for PPARg‐dependent adipogenesis. ASC‐2 similarly functions as a key adaptor for LXR recruitment of MLL3 and MLL4.11 Moreover, H3K4‐trimethylation of a subset of metabolic target genes of LXRs, mediated redundantly by MLL3 and MLL4, is correlated with their expression.11 We also have found that ASCOM‐MLL3 and ASCOM‐ MLL4 play crucial roles in bile acid homeostasis as specific H3K4MT coactivator complexes for FXR (Mol Endo 2009, in press, and our unpublished results). Overall, our results expand the roles for H3K4‐trimethylation to

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transcriptional regulation of metabolic genes and suggest that ASCOM, among Set1‐like complexes, functions as a Set1‐like H3K4MT complex specialized for a subset of NRs that regulate metabolic genes. Interestingly, MLL4 has been proposed to play crucial roles in ERa transactivation via direct interactions with ERa.93 Moreover, MLL1/2‐complexes have also been shown to function with ERa via interactions between ERa and menin.96 These results highlight the important caveat that ASC‐2 is not a general adaptor for NRs and that other subunits of ASCOM may also function as adaptors for some NRs (e.g., MLL4 for ERa),93 and that ASCOM is not the only Set1‐like H3K4MTcomplex involved in NR functions.

C. Recruitment of Set1‐Like H3K4MT Complexes In S. cerevisiae, Set1 is thought to be recruited to chromatin by the phosphorylated form of RNA polymerase II carboxy‐terminal domain (CTD), the histone chaperone FACT, and the Paf1 elongation complex. Moreover, its H3K4‐trimethylation activity is dependent on H2B‐monoubiquitination.5,47 However, this model fails to explain several observations in higher eukaryotes. First, H3K4‐trimethylation is enriched in promoter regions. Second, interactions of methylated H3K4 and/or H3K4MTs with chromatin modifiers involved in transcription initiation, such as NURF81 and p300,97 occur at the promoter. Third, although direct interactions among FACT, the PAF complex, and the H2B‐ubiquitination machinery are well documented and although Paf1 interacts (directly or indirectly) with Set1 in yeast,98 direct interactions of these components with H3K4MTs are not yet established in mammalian cells. Finally, several reports demonstrate that genetic or siRNA‐mediated knockdown of components of the H2B ubiquitination machinery does not affect H3K4 mono‐ and dimethylation at target genes, suggesting intact recruitment of H3K4MTs.99,100 These results lead to our favored model in which the primary mechanisms of recruitment of at least some H3K4MTs to their target genes involve direct or indirect associations with transcription factors (Fig. 2C). In support of this notion, diverse site‐specific transcription factors have been shown to associate with MLL‐family complexes. For example, ERa associates with MLL2 and MLL4,93,96 b‐catenin associates with MLL1 and MLL2,101 E2F6 associates with MLL1,102 and SET1a associates with the viral transcription factor VP16.66 In addition, apart from binding directly to p53 and effecting p53‐dependent H3K4 methylation of chromatin templates independently of H2B ubiquitylation, the MLL1 complex stimulates p53‐dependent transcription from a chromatin template in a reconstituted in vitro transcription system.102 We have also found that RARs and LXRs recruit MLL3 and MLL4 to the target genes of

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RARs and LXRs via interactions of these NRs with ASC‐2, an integral subunit of MLL3/4 complexes.9,11 Importantly, associations with transcription factors likely function to determine the target specificity of Set1‐like H3K4MT complexes. For instance, ASCOM, but not the related Set1‐like MLL1/2 complexes, is targeted by RAR since ASCOM is capable of interacting with RAR via ASC‐ 2 while MLL1/2 complexes show no interactions with RAR.9 The mechanisms of recruitment of Set1‐like complexes to their target genes may also involve at least two additional types of interactions. First, interactions with the basal machinery have been proposed (Fig. 2D). Similar to yeast, Set1, MLL1, and MLL2 associate with the phosphorylated CTD of Pol II and components of the basal machinery,102,103 albeit not necessarily directly, and colocalize with the Pol II binding sites.104 This association with the basal machinery appears to be also conserved with Drosophila Trx.105 Second, interactions with modified histones may serve to further stabilize the association of Set1‐like complexes with chromatin, as suggested by the discovery that WDR5 presents the H3K4 side chain for further methylation by MLL171,73 (Fig. 2E). Because H3K4MTs often contain a series of distinct modified histone binding domains (e.g., PHD fingers in MLL4 in Fig. 1B), future studies may uncover roles for additional interactions with modified histones in recruiting Set1‐like complexes to their targets. Taken together, we propose that recruitment of mammalian Set1‐like H3K4MT complexes to their target genes may generally involve a two‐step mechanism. In this model, the primary recruitment event is proposed to be carried out through interactions with specific transcription factors (Fig. 2C), followed by further stabilization of the complex on chromatin through interactions with basal transcription machinery and modified histones (Fig. 2D and E). The secondary stabilization might allow for efficient association of the complex with chromatin over a larger domain than dictated by the presence of the transcription factor on its DNA regulatory element. For example, both NURF and ING2 complexes could be recruited by the site‐specific transcription factors and also be stabilized on chromatin by the specific recognition of H3 trimethylated at K4.82,106

V. Cross Talk of ASCOMs with Other Coactivators Although NR transactivation involves multiple coactivators, it is unclear how these factors are functionally integrated. Interestingly, ASC‐2 has been reported to bind both to HATs CBP and p30020,28,36 and, although direct interactions and functions were not established, to the TRAP/DRIP/ARC Mediator complex that links NRs to the basal transcription machinery.36 ASC‐2 has also been linked to other coregulators that function in

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transcriptional initiation and subsequent mRNA processing steps (see below). Our recent results further indicate that the NR coactivator function of ASCOM requires a novel interplay with Swi/Snf that is likely mediated through specific interactions of the SET domain of MLL3/4 with INI1, a core subunit of Swi/ Snf.107 These results raise the interesting possibility that ASCOM may serve as a novel platform for NR transactivation by integrating the functions of multiple coactivators through mutual interactions. Moreover, the two histone‐modifying activities in ASCOM (i.e., MLL3/4 and UTX) may directly and/or indirectly affect these integration processes.

A. CBP/p300 ASC‐2 has been shown to form a high‐affinity complex with CBP.20,28,36 Thus, expression of ASC‐2 in 293 cells followed by extraction, affinity adsorption, and immunoblotting have indicated that a significant amount of CBP in the cell is associated with ASC‐2. This association is likely an indirect event, as no direct interaction between full length ASC‐2 and full length CBP was detected in yeast.108 Other in vitro binding studies have demonstrated that the C‐terminal region of p300 (amino acids 1661–2414) can associate with the C‐terminal region of ASC‐2.36 Whether association of ASC‐2 with CBP/p300 plays essential roles for ligand‐dependent activation by NRs or other transcription factors activated by ASC‐2 has not been directly addressed using CBP//p300/ cells. However, ligand‐dependent activation of NRs by ASC‐2 is completely blocked by expression of E1A, which is known to inactivate CBP/ p300.20,28 We also have found that not only H3K4‐trimethylation, but also H3/ H4‐acetylation of the RAR‐target gene RAR‐b2, is ablated in ASC‐2‐null cells.9 These results implicate ASC‐2 in recruitment of a complex containing CBP/ p300 and associated factors to ligand‐bound NRs on gene promoters.20,28 However, because CBP/p300 appears to associate with primary coactivators, such as SRC‐1, that directly interact with NRs, the recruitment of CBP by NRs may not necessarily require ASCOM. Moreover, recruitment of CBP/p300 and ASCOM to NR‐target genes could be facilitated not only by mutual interactions but also by the possible presence of binding modules that recognize methylated H3K4 (by MLL3/4), demethylated H3K27 (by UTX), or acetylated H3/H4 (by CBP/p300). CBP/p300 complexes may also modify components of ASCOM and vice versa, resulting in enhanced function for these complexes. More studies are needed to distinguish between these possibilities. Nonetheless, these results are consistent with a possibility that ASCOM and CBP/p300 are functionally integrated during NR transactivation through interactions between these two complexes (Fig. 3A).

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B. RNA‐Binding Proteins Increasing evidence indicates that transcription and pre‐mRNA processing are functionally coupled to modulate gene expression. Interestingly, three proteins containing RNA recognition motifs (RRMs) have been identified as ASC‐2‐interacting proteins through yeast two‐hybrid screenings; CAPER (for coactivator protein for AP‐1 and ER receptor), PIMT (PRIP‐interacting methyltransferase), and CoAA (for coactivator activator). The recent results demonstrate that CAPER and CoAA are clearly involved with both transcription and splicing, suggesting that ASCOM may play important roles in functionally linking these two processes (Fig. 3B). We have isolated CAPER from a mouse liver cDNA library using the C‐terminal region (amino acids 1172–1729) of ASC‐2 as bait.109 This protein has been subsequently renamed CAPERa upon identification of its paralogue CAPERb.110 CAPERa is identical to the nuclear autoantigens HCC1.3 and HCC1.4 reported in hepatocarcinoma.111 HCC1.3 and HCC1.4 are identical except for six additional amino acids in HCC1.4. Mouse CAPERa (HCC1.3) enhances transactivation by ERa and c‐Jun.109 CAPERa contains three RRMs and associates with c‐Jun and the liganded ERa via RRM3, while the C‐terminus of CAPERa binds to ASC‐2.109 Interestingly, CAPERa does not appear to enhance activation of other NRs such as RARs, RXRs, TR, and GR. The basis for ERa specificity is unclear, but this specificity implies that CAPERa may not enhance ERa function through association with ASC‐2,

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which interacts with and enhances activation of many other NRs. Both CAPERa and CAPERb coactivate the progesterone receptor (PR) in luciferase transcription reporter assays and alter alternative splicing of a calcitonin/calcitonin gene‐related peptide minigene in a hormone‐dependent manner.110 The importance of CAPER coactivators in the regulation of alternative RNA splicing of an endogenous cellular gene (VEGF) has been substantiated by siRNA knockdown of CAPERa.110 Mutational analysis of CAPERb indicates that the transcriptional and splicing functions are located in distinct and separable domains of the protein.110 Further implicating CAPERs as potential splicing factors, CAPERa has been found in a purified spliceosome complex.112 Overall, these results indicate that NR‐regulated transcription and pre‐mRNA splicing can be directly linked via ASCOM and dual function coactivator molecules such as CAPERa and CAPERb (Fig. 3B). CoAA is another RRM‐containing factor that was isolated from a GC cell cDNA library in a yeast two‐hybrid screen using the C‐terminal region (amino acids 1641–2063) of ASC‐2 as bait.113 The 669 amino acid CoAA contains two RRMs near its N‐terminus and the C‐terminal auxiliary domain that interacts with ASC‐2. Expression of CoAA enhances activation by a number of transcription factors that include NF‐kB, CREB, AP‐1, PR, TR, ER, and GR.113,114 Using transcriptional and splicing reporter genes driven by different promoters, CoAA has been shown to mediate transcriptional and splicing effects in a promoter‐preferential manner.114 CoAA also associates with DNA–PK regulatory subunit Ku86 and poly (ADP‐ribose) polymerase (PARP) from GH3 cells, raising the interesting possibility that it may act to functionally link ASCOM to DNA repair machinery in vivo.113 PIMT has been isolated in a yeast two‐hybrid screen from a human liver cDNA library using a large C‐terminal region (amino acids 773–2067) of ASC‐2 as bait.115 PIMT is a ubiquitously expressed putative RNA methyltransferase that contains an invariant GXXGXXI motif near its N‐terminus found in K‐homology motifs of many RNA‐binding proteins. Expression of PIMT enhances PPARg and RXR activity, and this activity is further enhanced by expression of ASC‐2. The putative methyltransferase activity of PIMT does not appear to be involved in its role as an activator since deletion of the methyltransferase domain does not affect its activating function.115 Like ASC‐2, PIMT homodimerizes in vitro and may also form homo‐oligomers.115 PIMT does not appear to methylate histones, but associates with CBP/p300 and PBP/TRAP220/MED1.116 However, whether PIMT is also involved in splicing remains to be tested.

C. Swi/Snf Three lines of our recent results suggest the presence of a novel cross talk between ASCOM and the ATPase‐dependent chromatin remodeling complex Swi/Snf during NR‐mediated transactivation.107 First, these two complexes are

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colocalized in the nuclear matrix.117 Although transcriptionally active DNA has been suggested to be tightly associated with the nuclear matrix,118 the relevance of the nuclear matrix colocalization of ASCOM and Swi/Snf to their function remains to be further determined. Second, Swi/Snf promotes binding of ASC‐ 2 to NR‐target genes, and ASCOM assists binding of Swi/Snf to NR‐target genes.107 Finally, we have discovered that the C‐terminal SET domains of MLL3 and MLL4 directly interact with INI1, an integral subunit of Swi/Snf, and that these interactions play important roles in the mutually facilitated recruitment of ASCOM and Swi/Snf to NR‐target genes107 (Fig. 3C). The involvement of these interactions in the cross talk between ASCOM and Swi/Snf is clearly demonstrated by our identification of a specific INI1 point mutant that affects the MLL3/4‐SET interactions, but not the known hBRM and c‐Myc interactions, of INI1.107 Both ASCOM and Swi/Snf are known to be recruited to NR‐target genes through well‐defined interaction interfaces; for example, ligand‐dependent interactions of NRs and ASC‐2 in recruiting ASCOM to NRs10 and interactions between various subunits of Swi/Snf and NRs in tethering Swi/Snf to NR‐target genes.119 Thus, our results suggest a model in which the MLL3/4‐SET:INI1 interactions provide ‘‘additional’’ interactions that can lead to mutual facilitation of ASCOM and Swi/Snf in recruitment to NR‐target genes. Importantly, this study provides the molecular basis for a novel integration of two enzymatic complexes in NR transactivation, H3K4MT in ASCOM and the ATPase‐dependent chromatin remodeler Swi/Snf. These results, along with the possible interplay of ASCOM with CBP/ p30020,28,36 and the TRAP/DRIP/ARC Mediator complex36 (Fig. 3D), further support the possibility that ASCOM has a distinct platform function in facilitating the recruitment of multiple coactivators to NRs. As these include the chromatin regulators Swi/Snf and CBP/p300, and as ASCOM itself is a H3K4MT complex, ASCOM is expected to play crucial roles in establishing transcriptionally active chromatin on NR‐target genes. In addition, these chromatin remodelers and modifiers may ‘‘indirectly’’ affect recruitment of other coactivators. For instance, ASCOM‐mediated H3K4‐methylation may create a direct docking site for other chromatin remodeling/modifying complexes that contribute to the generation of a proper environment for transcription. Interestingly, despite the conserved nature of the SET domains throughout evolution, the SET–INI1 interactions, which were originally described in MLL1,120 are not shared with the SET domains of Ash1, E(Z), and SU(VAR)3–9.121 Given that MLL1 and MLL3/4 belong to the Set1‐like family of complexes, these results raise the interesting possibility that Set1a/b and MLL2, the remaining members of the Set1‐like complexes, may also communicate with Swi/Snf through INI1 interactions. This possibility is supported by the results that ASH2L and RbBP5, two common subcomplex subunits of all the Set1‐like complexes (Table I), are also found associated

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with the nuclear matrix107 and that SNR1, a Drosophila homologue of INI1 that interacts with the C‐terminal SET domain of Trx (a Drosophila orthologue of MLL1), has been shown to colocalize with approximately one‐half of the TRX binding sites on larval salivary gland polytene chromosomes.120

VI. Physiological Roles of Key Subunits of ASCOM A. ASC‐2 1. DN1 AND DN2 We have shown that an 80 amino acid ASC‐2 peptide (DN1) containing NR1 blocks transactivation by NRs when expressed in transgenic mice.22 DN1 transgenic lines display pathological abnormalities in many different organs that include heart, pituitary, adrenal glands, brain, spleen, liver, and lung. In particular, we have observed eyes with microphthalmia and posterior lenticonus with cataract.22 We have also found that transgenic expression of another ASC‐2 region (DN2; amino acids 1431–1511) containing NR2 blocks the activity of LXRs in vivo.21 Livers from DN2 transgenic mice show changes similar to those seen in livers from LXRa/ mice, suggesting that ASC‐2 plays a role in cholesterol and lipid metabolism in the liver, presumably through regulation of LXRs.21 On a high fat cholesterol diet, DN2 transgenic lines display rapid accumulation of large amounts of cholesterol and downregulation of known lipid metabolizing target genes of LXR.21 Unexpectedly, DN1 and DN2 selectively block the activity of endogenous ASC‐2, but not that of other coactivators such as TRAP220/MED1 or SRC‐1.21,22 Since DN1 and DN2 would be expected to compete with the binding of other coactivators for liganded NRs, the mechanism underlying this specificity is unclear and requires further study. Importantly, recent studies of a conditional knockout of ASC‐2 in liver122 suggest that the phenotypes observed in DN1 transgenic mice123 may not be simply ASC‐2‐dependent. Thus, while these transgenic studies have provided the initial clues to the probable biological role of ASC‐2 in vivo, they should be interpreted with caution. 2. ASC‐2 MUTANT MICE The ASC‐2 gene has been knocked out in mice in a mixed C57BL/6–129S6 (C57/129) genetic background.24–27 These studies reveal that the ASC‐2‐null mutation is embryonic lethal and that mutant embryos die in utero between E8.5 and E12.5, while C57/129 ASC‐2þ/ mutant mice appear normal and grow similar to wild‐type mice (see below). The cause of lethality in ASC‐2/ embryos is attributed to placental dysfunction and a number of developmental defects involving the heart, liver, and brain.24,25,27 ASC‐2/ embryos are growth retarded and half the size of wild‐type or ASC‐2þ/ embryos.26 Consistent with these

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TABLE II PHYSIOLOGICAL FUNCTIONS FOR ASC‐2 AND MLL3 Defects in

Genotype

Comments

Wound heeling

ASC‐2þ/

Defects occur in some old mice26

þ/

Insulin secretion

ASC‐2

Apoptosis and decreased proliferation of islets23

Growth

ASC‐2þ/ (129S6)

Stunted growth26 Stunted growth9

D/D

MLL3 Fertility

þ/

ASC‐2

(129S6)

D/D

26

MLL3

9

Ductal branching of mammary glands

ASC‐2/ in mammary glands

No milk production during lactation124

WAT (PPARg‐signaling)

MLL3D/D

MEFs show poorer adipogenic potential than wild‐type MEFs in vitro,12 while ASC‐2/ MEFs do not undergo adipogenesis95

Hepatic lipid synthesis (LXR‐signaling)

MLL3D/D

Less fats deposited to hepatocytes even with a high fat diet11

Bile acid homeostasis (FXR‐signaling)

MLL3D/D

Enlarged gall bladder (Mol Endo 2009, in press, and our unpublished results)

Mammary tumor formation suppression

ASC‐2þ/

Polyoma middle‐Tantigen expressed in ASC‐2þ/ mammary glands133

Urothelia tumor formation suppression

MLL3D/D

At least in part due to defects in p53‐ signaling (PNAS 2009;106:8513–8, and our unpublished results)

References and additional remarks are included as comments.

results, ASC‐2/ MEFs derived from E12.5 ASC‐2/ embryos exhibit growth retardation in culture compared to wild‐type MEFs. ASC‐2/ MEFs also undergo apoptosis as they enter into the late log phase of growth,26 while wild‐ type MEFs are resistant to apoptosis under the same growth conditions. RNAi‐ mediated knockdown of ASC‐2 in wild‐type MEFs leads to a level of apoptosis similar to that found with ASC‐2/ MEFs.26 Apoptosis of the ASC‐2/ cells seems caspase‐mediated since zVAD‐fmk, a pan‐caspase inhibitor, blocks apoptosis in ASC‐2/ MEFs.26 Overall, these findings suggest that ASC‐2 likely regulates antiapoptotic and/or prosurvival genes required for cell growth and development. Interestingly, older ASC‐2þ/ C57/129 mice have been found to exhibit a wound healing phenotype26 (Table II). They spontaneously develop skin lesions or ulcers around the neck, ears, snout, and facial area. These regions

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correspond to the areas where mice groom and likely scratch themselves. This occurs in
25% of both male and female ASC‐2þ/ C57/129 mice. These spontaneous lesions develop as early as 4–6 months of age in the ASC‐2þ/ C57/129 mice.26 Histopathology of the affected and normal regions of the skin show thickening of the epidermis, increased sebaceous glands, and a reduction or total loss of hair follicles. In addition, there is no leading edge of keratinocyte migration, which is seen in normal wound healing. This lack of keratinocyte migration has been reproduced using ex vivo skin explant cultures from 2‐day‐old C57/129 ASC‐2þ/ mice.26 Wild‐type keratinocytes show robust migration when incubated with epidermal growth factor (EGF), whereas keratinocytes from the ASC‐2þ/ explants show no response to EGF. The molecular mechanism by which a reduction of ASC‐2 in the skin of ASC‐2þ/ mice leads to chronic skin wounds remains to be further defined. In the pancreas, we have found that ASC‐2 is expressed in the endocrine cells of islets of langerhans.23 In addition, our results reveal that overexpressed ASC‐2 increases glucose‐elicited insulin secretion, whereas insulin secretion is decreased in islets from ASC‐2þ/ mice. Moreover, primary rat islets ectopically expressing DN1 or DN2 exhibit decreased insulin secretion. The mass and number of islets also decrease in heterozygous mice, likely due to increased apoptosis and decreased proliferation of ASC‐2þ/ islets.23 3. 129S6 ISOGENIC ASC‐2þ/ MICE In contrast to ASC‐2þ/ mice in a C57/129 mixed genetic background, isogenic ASC‐2þ/ 129S6 mice exhibit a neonatal growth phenotype and newborn pups are
10–15% smaller than their wild‐type littermates.26 However, within 2 months after weaning, ASC‐2þ/ 129S6 mice become similar in size to their wild‐type littermates. Interestingly,
3% of the ASC‐2þ/ 129S6 newborn pups are extremely growth stunted, weighing 70% less than their wild‐type littermates. These mice often die before weaning. However, a small number of these mice survive and also become equivalent to wild‐type littermates in weight.26 Interestingly, this phenotype seems distinct from MLL3D/D newborn mice whose uniformly smaller size and weight deficit persist through adulthood.12 The molecular mechanisms underlying impaired growth in MLL3D/D and isogenic 129S6 ASC‐2þ/ mice have yet to be determined. In contrast to the absence of reproductive phenotypes in ASC‐2þ/ mice in C57/129 mixed genetic background, the fertility of both sexes of isogenic ASC‐2þ/ mice in 129S6 background is compromised.26 Both male and female mice are hypofertile and
20% of isogenic ASC‐2þ/ females are sterile. These infertile females appear to mate based on the formation of vaginal plugs and exhibit normal estrus cycles. These results suggest abnormalities in oogenesis, implantation of fertilized ova, or a defect in placental function, as no embryos

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are detected between E8.5 and E12.5. The number of newborn ASC‐2þ/ pups obtained from crosses between ASC‐2þ/ 129S6 hypofertile males and females is far less than expected based on Mendelian distribution.26 This appears to result from a significant number of ASC‐2þ/ embryos dying in utero. Those ASC‐2þ/ 129S6 female mice which are hypofertile exhibit a progressive decline in fertility as they age and their newborn pups have a high rate of neonatal mortality. The reproductive phenotypes in isogenic ASC‐2þ/ 129S6 mice are similar to those found for MLL3D/D mice.9 Future studies should be directed at elucidating the molecular mechanism underlying male and female hypofertility in MLL3D/D and isogenic 129S6 ASC‐2þ/ mice. 4. PHENOTYPES FROM CONDITIONAL KNOCKOUTS To overcome the early embryonic lethality of ASC‐2/ mice and to study the role of ASC‐2 in select tissues of adult mice, conditional knockout mice for ASC‐2 have been constructed. In ASC‐2‐deficient mammary glands,124 the elongation of ducts during puberty is not affected, but the number of ductal branches is decreased. Moreover, during pregnancy, the null mammary glands exhibit decreased alveolar density. Interestingly, the lactating ASC‐2‐deficient glands contain scant lobuloalveoli with many adipocytes, while the wild‐type glands lack adipocytes. The null mammary glands fail to produce enough milk to nurse all the pups during lactation. These results suggest that ASC‐2 contributes to efficient ductal branching of mammary glands in response to estrogen. A slight increase in apoptosis has been observed in the terminal buds from ASC‐2‐deficient glands, raising the possibility that abnormal apoptosis contributes to the impaired ductal branching.124 ASC‐2 liver conditional knockout mice have been generated to study the in vivo role of ASC‐2 as a coactivator for PPARa and CAR.122 We have reported that ASC‐2 interacts with CAR and enhances the transcriptional activation by CAR in transfection assays.123 However, the in vivo data obtained from conditional knockout studies show that ASC‐2 deficiency in the liver fails to alter CAR function, as both wild‐type and ASC‐2 liver conditional knockout mice are susceptible to acetaminophen‐induced liver damage mediated by CAR and upregulate expression of CAR‐target genes (e.g., CYP1A2, CYP2B10, CYP3A11, and CYP2E1) in response to CAR ligand. CAR mRNA and protein levels between these mice are similar, suggesting that ASC‐2 does not alter hepatic CAR expression.122 Notably, these in vivo findings are somewhat contradictory to our earlier studies with transgenic mice expressing DN1, a fragment of ASC‐2 containing the NR1, as these mice fail to show acetaminophen‐induced hepatic necrosis.123 Similarly, ASC‐2 has been shown to activate PPARs in vitro.29,38 However, in conditional knockout mice, the degree of PPARa ligand‐mediated peroxisome proliferation in liver cells has been found to be

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essentially similar to that seen in wild‐type liver cells, implying that, in the liver, ASC‐2 is not involved for PPARa‐mediated proliferation. Accordingly, the PPARa‐specific target genes encoding fatty acyl‐CoA oxidase, enoyl‐CoA hydratase/L‐3 hydoxyacyl‐CoA hydrogenase (L‐bifunctional enzyme; L‐PBE), peroxisomal thiolase (PTL), and CYP4A1 increase markedly in both wild‐type and ASC‐2 liver conditional knockout mice, following treatment with the PPARa ligand. PPARa mRNA levels remain essentially similar in control and PPARa ligand‐treated mice.122 Based on these results, it has been suggested that ASC‐2 is dispensable in the liver for activating PPARa‐ and CAR‐mediated gene expression.122 However, it needs to be further determined whether MLL3/4 complexes are also dispensable for the hepatic function of PPARa and CAR, as it is possible that MLL3/4 complexes still form in the absence of ASC‐2 and that another subunit of ASCOM may function to tether ASCOM to PPARa and CAR. Alternatively, other Set1‐like complex(es) may function redundantly with ASCOM, and inactivation of ASCOM alone may not be sufficient to alter PPARa and CAR functions in the liver. Further analyses of MLL3/4 mutant mice should clarify these issues.

B. Metabolic Phenotypes of MLL3 H3K4‐trimethylation is an evolutionarily conserved mark for transcriptionally active chromatin.47,48 Interestingly, whereas yeast has a single enzyme responsible for this modification (H3K4MT), higher eukaryotes carry a number of H3K4MTs.94 This suggests that individual H3K4MTs in higher eukaryotes may have distinct target genes. However, the physiological roles for higher eukaryotic H3K4MTs and their target genes remain poorly understood. We have generated MLL3D/D mutant mice that express an MLL3 with a deletion of the Set catalytic domain that is involved in H3K4‐methylation.9 The MLL3D/D mutation in a mixed C57/129 genetic background leads to partial embryonic lethality,9 while isogenic C57BL/6 MLL3D/D mice are completely embryonic lethal (our unpublished results). Interestingly, MLL3D/D mice share some similar phenotypes with isogenic 129S6 ASC‐2þ/ mice,26 providing genetic evidence for the assembly of MLL3 into ASCOM. Similar to isogenic 129S6 ASC‐2þ/ mice,26 MLL3D/D mice are stunted in their overall growth and weigh 30–40% less at birth. MLL3D/D females also exhibit a range of reproductive phenotypes from infertility to hypofertility, while males are generally hypofertile. In addition, like ASC‐2/ MEFs, MEFs generated from MLL3D/D mice divide at approximately half the rate of wild‐type MEFs.9,26 Our phenotypic analyses of MLL3D/D mice reveal that MLL3 has multiple functions in vivo, likely as a component of ASCOM. In particular, our results reveal that MLL3, and possibly MLL4, may be specialized for regulating genes involved in metabolic homeostasis, thus revealing key roles for specific H3K4MTs in metabolism.

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1. WAT PHENOTYPES OF MLL3D/D MICE ASC‐2‐null MEFs are refractory to PPARg‐stimulated adipogenesis and fail to express the PPARg‐responsive, adipogenic marker gene aP2.95 However, the specific roles for MLL3 and MLL4 in adipogenesis remain undefined. We recently have found that MLL3 plays crucial roles in adipogenesis.12 First, MLL3D/D mice have a significantly decreased amount of WAT with a favorable overall metabolic profile, including improved insulin sensitivity and increased energy expenditure. Second, MLL3D/D MEFs are mildly but consistently less responsive to inducers of adipogenesis than wild‐type MEFs. Third, ASC‐2, MLL3, and MLL4 are recruited to the PPARg‐activated aP2 gene during adipogenesis.12 Consistent with earlier demonstrations that ASC‐2 binds directly to PPARg and C/EBPa,10 two factors important for adipogenesis,125 ASCOM directly interacts with both free and promoter‐bound PPARgRXRa heterodimers.12 Thus, it is possible that ASC‐2 plays a similar role in facilitating ASCOM function through PPARg and C/EBPa during adipogenesis. Moreover, while H3K4‐methylation of aP2 is readily induced in wild‐type MEFs, it is not induced in ASC‐2/ MEFs and only partially induced in MLL3D/D MEFs.12 These results suggest that ASCOM‐MLL3 and ASCOM‐MLL4 likely function as crucial but redundant H3K4MT complexes for PPARg‐dependent adipogenesis, uncovering an interesting connection between H3K4‐trimethylation and adipogenesis. Interestingly, we have observed that PPARg exhibits both ligand‐dependent and ligand‐independent interactions with different components of the ASCOM complex,12 uncovering a previously unappreciated complexity in recruiting ASCOM to NRs. This could reflect, for example, the presence of a dynamic equilibrium distribution of ASC‐2‐containing and ASC‐2 free complexes and an associated ligand‐independent recruitment of the core complex, possibly through interactions with a PPARg domain other than AF2, followed by a ligand‐ and AF2‐dependent recruitment or stabilization of ASC‐2. Another possibility is the presence in our purified PPARg of an active AF2 conformation, due either to an endogenous ligand or to a natural equilibrium between active and inactive AF2 states,45 and an AF2‐dependent recruitment of ASCOM through LXXLL motifs in MLL3/411,93 or ASC‐2 with further stabilization of MLL3/4 or ASC‐2 by the potent ectopic ligands.

2. LIPID AND BILE ACID PHENOTYPES OF MLL3D/D MICE In further support of the selective involvement of ASCOM in metabolism, we have discovered two additional lines of metabolic phenotypes, in MLL3D/D mice that are ascribed to the roles of ASCOM as a H3K4MT coactivator of LXRs and FXR.

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The second NR box of ASC‐2 has been shown to specifically recognize LXRs.20,44 However, no exact role for either ASC‐2 or MLL3/4 in LXR transactivation has been clearly defined. Our recent results reveal that the key function of ASC‐2 in LXR transactivation is to present MLL3 and MLL4 to LXRs.11 Thus, ASC‐2 is required for ligand‐induced recruitment of MLL3 and MLL4 to LXRs, and LXR ligand T1317 induces not only expression of LXR‐ target genes but also their H3K4‐trimethylation. Strikingly, both of these ligand effects are ablated in ASC‐2‐null cells but only partially suppressed in cells expressing an enzymatically inactivated mutant MLL3.11 Our results also show that LXR transactivation does not appear to require certain other Set1‐like complexes, because, while ASC‐2 is required for LXR transactivation, menin, an integral component of MLL1/2 Set1‐like complexes, is dispensable.11 In comparison, targeted deletion of menin or siRNA‐mediated reduction of MLL4 alone has been shown to significantly impair ERa transactivation.93,96 Taken together, these results suggest that ASCOM‐MLL3 and ASCOM‐MLL4 play redundant but essential roles in ligand‐dependent H3K4‐trimethylation and expression of LXR‐target genes and that ASC‐2 is likely a key determinant for LXR function through ASCOM (but not other Set1‐like complexes). We have proposed that a subunit of Set1‐like complexes determines the target specificity of each complex via direct protein–protein interactions with target transcription factors9 (Fig. 2C). The current study supports this model and validates the previously proposed specialized function for ASC‐2 as a key adaptor to present ASCOM to LXRs.11,20,21,44,126 In further support of the role of ASCOM as a coactivator of LXRs, our analysis of hepatic mRNAs shows that the hepatic lipogenic LXR‐target genes FAS and SREBP‐1c are suppressed in MLL3D/D mice.11 Intriguingly, expression of another direct LXR‐target gene Cyp7A1, that encodes a key enzyme in the conversion of cholesterol into bile acids is not downregulated.11 These results suggest that clearance of cholesterol via bile acid synthesis could be intact in MLL3D/D mice, which, along with the impaired expression of lipogenic genes, might have contributed to our failure to observe accumulation of oil‐red‐O positive lipid droplets in the livers of MLL3D/D mice even with a high fat diet.11 Thus, these results raise the interesting possibility that MLL3 may have some selectivity toward a subset of LXR‐target genes involved in hepatic de novo lipogenesis. Future studies should be directed toward elucidation of the molecular basis underlying this interesting selectivity. Our recent results reveal that expression of two FXR‐target genes, BSEP and SHP, is downregulated in MLL3D/D mice (Mol Endo 2009, in press, and our unpublished results). SHP encodes an atypical orphan nuclear receptor that lacks a DBD.127 SHP forms a novel regulatory loop with FXR, which functions to maintain bile acid homeostasis.127 SHP expression is induced in response to increased hepatic levels of bile acids that bind and activate FXR, and SHP in

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turn suppresses expression of Cyp7A1 by antagonizing the activity of another NR, ‘‘liver receptor homolog 1 (LRH‐1),’’ that functions as an inducer of Cyp7A1. This leads to a blockage in the further production of bile acids from cholesterol.127 Consistent with the decreased expression of SHP, we have found that Cyp7A1 is significantly upregulated in MLL3D/D mice (Mol Endo 2009, in press, and our unpublished results). In further support of these results, MLL3D/D mice exhibit a much higher level of plasma bile acid when fed a control diet supplemented with 0.5% (w/w) cholic acid (CA, a ligand for FXR) (Mol Endo 2009, in press, and our unpublished results), similar to FXR‐null mice.128 In further indication for impaired bile acid homeostasis, MLL3D/D mice fed a control diet supplemented with 0.5% (w/w) CA also have a significantly enlarged gallbladder filled with bile acids and thus the total pool of bile acids in MLL3D/D mice is much larger than that in wild‐type mice (Mol Endo 2009, in press, and our unpublished results). These results indicate that bile acid homeostasis is seriously disturbed in MLL3D/D mice. Interestingly, expression of BSEP and SHP is still induced and expression of Cyp7A1 is still suppressed in response to a control diet supplemented with 0.5% (w/w) CA in MLL3D/D mice, suggesting that another H3K4MT, likely MLL4, functions in the livers of MLL3D/D mice. Consistent with these results, both MLL3 and MLL4 are recruited to SHP via FXR, and suppression of SHP expression and SHP H3K4‐trimethylation is much greater with siRNA‐mediated downregulation of both MLL3 and MLL4 than with downregulation of MLL3 or MLL4 alone (Mol Endo 2009, in press, and our unpublished results). Overall, these results suggest that ASCOM‐MLL3 and ASCOM‐MLL4 act as redundant but crucial coactivators of FXR. Our newly established mice with a mutant MLL4 should help clarify whether MLL4 indeed functions redundantly with MLL3 in mediating FXR transactivation.

C. ASCOM in Cancers The ASC‐2 gene is amplified and overexpressed in human breast and other cancers.129–131 However, it has been difficult to determine whether ASCOM is involved in tumorigenesis or antitumorigenesis. This is mainly because ASC‐2 and MLL3/4 target a complex array of transcription factors that include both tumorigenic and tumor suppressive proteins.10 However, as these ASCOM target factors have been identified mostly through cell transfection and in vitro studies,10 the roles for ASCOM in their transactivation remain to be validated in more physiological settings. The results discussed below, along with the finding that MLL3 maps to 7q36, a chromosome region that is frequently deleted in myeloid disorders,132 are in favor of tumor suppressive roles for ASCOM (see below). Given the multifunctionality of ASCOM, however, it is possible that ASCOM may also function to trigger tumorigenesis in different cell types.

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1. AMPLIFICATION AND OVEREXPRESSION OF ASC‐2 IN HUMAN CANCERS Human ASC‐2 is localized on chromosome 20 (20q11). The ASC‐2 gene was initially mapped to the 20q11 segment of chromosome 20 during a search for amplified and overexpressed genes mapping to chromosome 20q in breast cancer.130 Subsequently, we have found, using FISH analysis, that the ASC‐2 copy number is increased to a moderate level (4–6 copies) in 14 out of 335 (4.2%) cases of breast cancer and to a high level (>6 copies) in 15 out of 335 (4.5%) cases of breast cancer.131 ASC‐2 mRNA is also detected in 11 different breast cancer cell lines, with the highest expression in BT‐474 cells.131 The ASC‐2 gene is also found to be amplified in lung and colon cancers.131 Notably, the high level of ASC‐2 expression in breast cancer cell lines does not correlate with the level of ER expression.131 2. MAMMARY TUMOR SUPPRESSIVE FUNCTION OF ASC‐2 To study the roles of ASC‐2 in mammary tumorigenesis, ASC‐2þ/ mice have been crossed with mice carrying polyoma middle‐T antigen (pyMT).133 This study has revealed that mammary tumor development in ASC‐2þ/ pyMT female and male mice is substantially accelerated compared with that in wild‐ type pyMT mice. Correspondingly, tumor formation in nude mice that receive premalignant ASC‐2þ/ pyMT mammary tissue is much faster than in nude mice that receive transplants from premalignant wild‐type pyMT mammary tissue.133 This tumor acceleration is reported to result from increased cell proliferation and ductal hyperplasia and mammary intraepithelial neoplasia. Although the mechanism of pyMT‐induced tumorigenesis is proposed to reflect a partial impairment of activated PPARg/RXR,133 it is important to note that ASC‐2 is a multifunctional coactivator and, thus, that the accelerated tumor phenotypes in ASC‐2þ/ pyMT mice may also involve ASC‐2 coactivator functions for other tumor suppressive transcription factors such as p53 (see below). 3. ASCOM AS A COACTIVATOR OF THE TUMOR SUPPRESSOR P53 D/D AND KIDNEY PHENOTYPES OF MLL3 MICE Further implicating ASCOM in tumor suppression pathways, we have found that ASCOM‐MLL3 and ASCOM‐MLL4 function as redundant but crucial coactivator complexes for the tumor suppressor p53 (PNAS 2009;106:8513–8, and our unpublished results). Our results reveal, first, that ASC‐2 acts as a coactivator of p53 in reporter assays and is required for H3K4‐trimethyation and expression of endogenous p53‐target genes in response to the DNA damaging agent doxorubicin and, second, that ureter epithelial tumors result from targeted inactivation of MLL3 H3K4‐methylation activity in the mouse (PNAS 2009;106:8513–8, and our unpublished results). Interestingly, this latter

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phenotype is exacerbated in a p53þ/ background and the tumorigenic cells are heavily immunostained for gH2AX (PNAS 2009;106:8513–8, and our unpublished results), indicating a contribution of MLL3 to the DNA damage response pathway through p53. In support of redundant functions for MLL3 and MLL4 in this process, siRNA‐mediated downregulation of both MLL3 and MLL4 is required to suppress doxorubicin‐inducible expression of p53‐target genes (PNAS 2009;106:8513–8, and our unpublished results). Importantly, this study identifies a physiologically relevant, specific H3K4‐trimethytransferase coactivator complex for p53. Notably, and related, independent siRNA‐based studies have indicated that p53 is regulated by ASC‐2.32 Interestingly, PTIP, which has been proposed to play important roles in cellular responses to DNA damage,134–136 has recently been identified as an additional component of ASCOM.90,91 The tumor suppressor p53 plays a key role countering the adverse effects of DNA damage,137,138 which otherwise can be lethal or lead to oncogenic transformation. DNA damage induces the transcriptional activity of p53 via damage sensors such as ATM. Interestingly, PTIP appears to be required for ATM‐mediated phosphorylation of p53 at Ser 15 and for DNA damage‐induced upregulation of the cyclin‐dependent kinase inhibitor p21.135 Correspondingly, the loss‐of‐function studies in mice indicate that PTIP is essential for the maintenance of genomic stability.134 One intriguing future challenge is to test whether this function of PTIP occurs in the context of ASCOM, particularly because ASC‐2‐null cells show increased phosphorylation of p53 at Ser 15 (PNAS 2009;106:8513–8, and our unpublished results). Regardless, as our results suggest that ASCOM acts as a tumor suppressive coactivator complex of p53, ASCOM, like PTIP, may also play crucial roles in the maintenance of genomic stability. Indeed, we show that targeted inactivation of MLL3 H3K4‐methylation activity in the mouse results in ureter epithelial tumors accompanied by an increased level of damaged DNA and that ASC‐2 appears to be important for DNA damage‐induced expression of p53‐target genes (PNAS 2009;106:8513–8, and our unpublished results). ASC‐2 appears to play a crucial role in effecting MLL3/4 function on p53‐target genes, as our results reveal that MLL3 and MLL4 are recruited to p53 and carry out H3K4‐trimehtylation of p53‐target genes in an ASC‐2‐ dependent manner (PNAS 2009;106:8513–8, and our unpublished results). The precise roles of ASC‐2 in p53 transactivation remain unclear, as ASC‐2 does not directly interact with p53 (PNAS 2009;106:8513–8, and our unpublished results). Our further search for the mechanistic basis underlying the associations between p53 and ASCOM reveal that 53BP1, a protein originally identified based on its interaction with p53,139,140 is recruited to p21‐p53REs. Moreover, the timing of this recruitment precisely overlaps that of ASC‐2 (PNAS 2009;106:8513–8, and our unpublished results), suggesting corecruitment of these two proteins. These results support the previously proposed role

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for 53BP1 as a coactivator of p53.140 Our proposal for an adaptor role for 53BP1 between p53 and ASCOM is further supported by our finding that 53BP1 and ASC‐2 are readily coimmunopurified and that 53BP1 appears to directly interact not only with p53139,140 but also with ASC‐2 (PNAS 2009;106:8513–8, and our unpublished results). However, because a significant level of ASC‐2/MLL3 recruitment to p21‐p53REs is still observed in 53BP1‐null cells, additional mechanisms are expected to exist (PNAS 2009;106:8513–8, and our unpublished results). Thus, the detailed molecular basis underlying the recruitment of ASCOM to p53 remains to be further delineated. Because the loss of p53 function is generally associated with most tumors, the seeming propensity of MLL3D/D mice to selectively develop urothelial tumors (PNAS 2009;106:8513–8, and our unpublished results) may appear paradoxical. On the one hand, this may reflect a redundancy between MLL3 and other MLLs in most tissues and, on the other hand, a more careful examination may yet uncover other types of tumors. In fact, ASCOM may guard against a broad range of epithelial tumors, and acceleration of pyMT‐induced mammary tumorigenesis by haploid inactivation of ASC‐2133 may similarly involve genomic instability caused by the impaired ability of ASCOM to support p53 transactivation. Nonetheless, it is possible that, along with a general redundancy between ASCOM‐MLL3 and ASCOM‐MLL4, an as‐yet‐to‐be‐characterized specific function of MLL3 in urothelium may selectively lead to urothelial tumors in MLL3D/D mice.

VII. Future Challenges The studies discussed in this review suggest that ASCOM likely serves as an essential H3K4‐methylation complex of multiple NRs and other transcription factors and accordingly regulates a diverse array of physiological processes. Most striking is the possibility that ASCOM may have a specialized function for a selective set of NRs involved in metabolism. However, more surprises for the physiological function of ASCOM are likely to be encountered as we continue to dissect the phenotypes of ASC‐2/MLL3/MLL4 mutant mice. To fully decipher the physiological function of ASCOM, many outstanding questions remain to be answered, and a few immediate challenges are summarized below. ASCOM is a unique Set1‐like complex, because it also contains UTX, an H3K27‐demethylase. H3K27‐methylation is a posttranslational modification that is highly correlated with genomic silencing. Thus, ASCOM may serve as an excellent model system to study the potential interplay between H3K4‐ and H3K27‐methylation events. Indeed, during retinoic acid signaling events, the recruitment of the UTX complex to HOX genes has been shown to result in

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H3K27‐demethylation and a concomitant methylation of H3K4.16 Our preliminary results also indicate that these two modifications are inversely coregulated in RAR‐target gene, RAR‐b2, through MLL3/4 and UTX (our unpublished results). These results suggest a concerted mechanism for transcriptional activation in which cycles of H3K4‐methylation by ASCOM are linked with the demethylation of H3K27 through UTX. It will be interesting to investigate whether these two distinct enzymes residing in the same complex modulate each other’s activity ‘‘directly.’’ Importantly, further studies of ASCOM may open new possibilities for pharmacological interventions of various disorders and diseases, because UTX and MLL3/4 should be chemically modulatable. In our studies, MLL3 and MLL4 appear to function redundantly in regulating target genes for RAR, LXRs, FXR, and PPARg9,11,12 (our unpublished results), likely due to the central adaptor role of ASC‐2 in recruiting ASCOM‐ MLL3 and ASCOM‐MLL4 to these NRs. Accordingly, the phenotypes of MLL3D/D mice elicited by these NRs may also be observed in MLL4 mutant animals. One important future challenge is to determine whether MLL3 and MLL4 also have their own unique sets of target genes. There are at least two related scenarios which would lead us to observe distinct functions between MLL3 and MLL4. First, if any component of ASCOM, which functions as an adaptor for specific NRs and/or transcription factors, is found only in ASCOM‐MLL3 or ASCOM‐MLL4, we would observe differences between MLL3 and MLL4 in regulating genes targeted by these NRs and transcription factors. Secondly, it is noted that MLL3 and MLL4 are gigantic in size but their overall homology is merely 30%.8,132 Thus, it is possible that these proteins may contain distinct interaction surfaces for some NRs and/or transcription factors and thereby act as specific adaptors for selective ASCOM‐ MLL3 or ASCOM‐MLL4 recruitment. Given our recent success in establishing MLL4 mutant mice (our unpublished results), we should be able to investigate these and other possibilities. Also, we should carefully examine whether any subunit of ASCOM is found selectively associated with MLL3 or MLL4. Our unpublished results indicate that multiple subunits of ASCOM undergo specific proteolysis during adipogenesis. We also have found that MEKK1 relieves the cryptic autonomous activation function of full‐length ASC‐2 (our unpublished results), suggesting that ASC‐2 is subjected to MAPK regulation through direct or indirect phosphorylation. Overall, ASC‐2 is likely to be subject to diverse posttranslational modifications, which may serve as a focal point to communicate and integrate regulatory events mediated by NRs or other transcription factors and the signaling cascades generated by cell surface receptors.

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Our finding of a coactivator role for ASCOM in p53 transactivation is consistent with some recent studies suggesting that ASC‐2 could be involved in DNA repair. The ASC‐2 C‐terminal region has been shown to interact with components of the DNA‐dependent protein kinase complex, including the catalytic subunit (DNA–PKc), regulatory subunits (Ku70 and Ku80), PARP, NuMA (p200), and DNA topoisomerase I36 (Fig. 2E). Although the role of DNA–PK in recombination and DNA repair has been well characterized,141 the involvement of DNA–PK in transcription is less clear. However, studies of ASC‐2 activity in cells deficient in DNA–PK reveal a marked reduction in ASC‐ 2‐mediated enhancement of GR transactivation.141 Moreover, DNA–PK phosphorylates an ASC‐2 fragment containing amino acids 714–999 in a DNA‐ dependent manner and a fragment containing amino acids 1237–2063 in a DNA‐independent manner.142 In addition, ASC‐2 stimulates the phosphorylation activity of DNA–PK in vitro, generating a phosphorylation pattern that differs from that of its DNA‐mediated activation form. Interestingly, PTIP, a component of ASCOM,90,91 and 53BP1, a putative adaptor protein between p53 and ASCOM (PNAS 2009;106:8513–8, and our unpublished results), have also been shown to modulate DNA repair,134–136,143 although it remains to be determined whether PTIP functions through ASCOM in DNA repair. ASC‐ 2 has also been shown to localize to a pS2 promoter nucleosome (NucE) that contains an ER binding site and TopoIIb‐mediated double strand breaks.144 Finally, with regard to the tumor suppressive function of ASCOM and its interplay with Swi/Snf, it is noted that at least two subunits of Swi/Snf, BRG1, and INI1, have been linked to tumor suppression pathways and that mice heterozygous for mutations at SNF5 and BRG1 are cancer‐prone.145–147 Thus, one interesting future challenge is to investigate whether the MLL3/4: INI1 interactions that play crucial roles in the cross talk between ASCOM and Swi/Snf107 play any roles in integrating the two tumor suppressive pathways directed by ASCOM and Swi/Snf, respectively. Acknowledgments Work from authors’ laboratories was supported by NIH/NIDDK grants to J.W.L. (DK064678) and to R.G.R. (DK071900).

References 1. Aranda A, Pascual A. Nuclear hormone receptors and gene expression. Physiol Rev 2001;81:1269–304. 2. Lonard DM, Lanz RB, O’Malley BW. Nuclear receptor coregulators and human disease. Endocr Rev 2007;28:575–87.

ASCOM AS A KEY H3K4MT COMPLEX OF NRS

375

3. Nettles KW, Greene GL. Ligand control of coregulator recruitment to nuclear receptors. Annu Rev Physiol 2005;67:309–33. 4. Rosenfeld MG, Glass CK. Coregulator codes of transcriptional regulation by nuclear receptors. J Biol Chem 2001;276:36865–8. 5. Shilatifard A. Chromatin modifications by methylation and ubiquitination: implications in the regulation of gene expression. Annu Rev Biochem 2006;75:243–69. 6. Jenuwein T, Allis CD. Translating the histone code. Science 2001;293:1074–80. 7. Wang Y, Fischle W, Cheung W, Jacobs S, Khorasanizadeh S, Allis CD. Beyond the double helix: writing and reading the histone code. Novartis Found Symp 2004;259:3–17; discussion 17–21, 163–9. 8. Goo YH, Sohn YC, Kim DH, Kim SW, Kang MJ, Jung DJ, et al. Activating signal cointegrator 2 belongs to a novel steady‐state complex that contains a subset of trithorax group proteins. Mol Cell Biol 2003;23:140–9. 9. Lee S, Lee DK, Dou Y, Lee J, Lee B, Kwak E, et al. Coactivator as a target gene specificity determinant for histone H3 lysine 4 methyltransferases. Proc Natl Acad Sci USA 2006;103:15392–7. 10. Mahajan MA, Samuels HH. Nuclear hormone receptor coregulator: role in hormone action, metabolism, growth, and development. Endocr Rev 2005;26:583–97. 11. Lee S, Lee J, Lee SK, Lee JW. Activating signal cointegrator‐2 is an essential adaptor to recruit histone H3 lysine 4 methyltransferases MLL3 and MLL4 to the liver X receptors. Mol Endocrinol 2008;22:1312–9. 12. Lee J, Saha PK, Yang QH, Lee S, Park JY, Suh Y, et al. Targeted inactivation of MLL3 histone H3‐Lys‐4 methyltransferase activity in the mouse reveals vital roles for MLL3 in adipogenesis. Proc Natl Acad Sci USA 2008;105:19229–34. 13. Agger K, Cloos PA, Christensen J, Pasini D, Rose S, Rappsilber J, et al. UTX and JMJD3 are histone H3K27 demethylases involved in HOX gene regulation and development. Nature 2007;449:731–4. 14. Hong S, Cho YW, Yu LR, Yu H, Veenstra TD, Ge K. Identification of JmjC domain‐containing UTX and JMJD3 as histone H3 lysine 27 demethylases. Proc Natl Acad Sci USA 2007;104:18439–44. 15. Lan F, Bayliss PE, Rinn JL, Whetstine JR, Wang JK, Chen S, et al. A histone H3 lysine 27 demethylase regulates animal posterior development. Nature 2007;449:689–94. 16. Lee MG, Villa R, Trojer P, Norman J, Yan KP, Reinberg D, et al. Demethylation of H3K27 regulates polycomb recruitment and H2A ubiquitination. Science 2007;318:447–50. 17. Smith ER, Lee MG, Winter B, Droz NM, Eissenberg JC, Shiekhattar R, et al. Drosophila UTX is a histone H3 Lys27 demethylase that colocalizes with the elongating form of RNA polymerase II. Mol Cell Biol 2008;28:1041–6. 18. Heery DM, Kalkhoven E, Hoare S, Parker MG. A signature motif in transcriptional co‐ activators mediates binding to nuclear receptors. Nature 1997;387:733–6. 19. Torchia J, Rose DW, Inostroza J, Kamei Y, Westin S, Glass CK, et al. The transcriptional co‐ activator p/CIP binds CBP and mediates nuclear‐receptor function. Nature 1997;387:677–84. 20. Lee SK, Jung SY, Kim YS, Na SY, Lee YC, Lee JW. Two distinct nuclear receptor‐interaction domains and CREB‐binding protein‐dependent transactivation function of activating signal cointegrator‐2. Mol Endocrinol 2001;15:241–54. 21. Kim SW, Park K, Kwak E, Choi E, Lee S, Ham J, et al. Activating signal cointegrator 2 required for liver lipid metabolism mediated by liver X receptors in mice. Mol Cell Biol 2003;23:3583–92. 22. Kim SW, Cheong C, Sohn YC, Goo YH, Oh WJ, Park JH, et al. Multiple developmental defects derived from impaired recruitment of ASC‐2 to nuclear receptors in mice: implication for posterior lenticonus with cataract. Mol Cell Biol 2002;22:8409–14.

376

LEE ET AL.

23. Yeom SY, Kim GH, Kim CH, Jung HD, Kim SY, Park JY, et al. Regulation of insulin secretion and beta‐cell mass by activating signal cointegrator 2. Mol Cell Biol 2006;26:4553–63. 24. Antonson P, Schuster GU, Wang L, Rozell B, Holter E, Flodby P, et al. Inactivation of the nuclear receptor coactivator RAP250 in mice results in placental vascular dysfunction. Mol Cell Biol 2003;23:1260–8. 25. Kuang SQ, Liao L, Zhang H, Pereira FA, Yuan Y, DeMayo FJ, et al. Deletion of the cancer‐ amplified coactivator AIB3 results in defective placentation and embryonic lethality. J Biol Chem 2002;277:45356–60. 26. Mahajan MA, Das S, Zhu H, Tomic‐Canic M, Samuels HH. The nuclear hormone receptor coactivator NRC is a pleiotropic modulator affecting growth, development, apoptosis, reproduction, and wound repair. Mol Cell Biol 2004;24:4994–5004. 27. Zhu YJ, Crawford SE, Stellmach V, Dwivedi RS, Rao MS, Gonzalez FJ, et al. Coactivator PRIP, the peroxisome proliferator‐activated receptor‐interacting protein, is a modulator of placental, cardiac, hepatic, and embryonic development. J Biol Chem 2003;278:1986–90. 28. Mahajan MA, Samuels HH. A new family of nuclear receptor coregulators that integrate nuclear receptor signaling through CREB‐binding protein. Mol Cell Biol 2000;20:5048–63. 29. Caira F, Antonson P, Pelto‐Huikko M, Treuter E, Gustafsson JA. Cloning and characterization of RAP250, a novel nuclear receptor coactivator. J Biol Chem 2000;275:5308–17. 30. Antonson P, Al‐Beidh F, Matthews J, Gustafsson JA. The human RAP250 gene: genomic structure and promoter analysis. Gene 2004;327:233–8. 31. Zhang H, Liao L, Kuang SQ, Xu J. Spatial distribution of the messenger ribonucleic acid and protein of the nuclear receptor coactivator, amplified in breast cancer‐3, in mice. Endocrinology 2003;144:1435–43. 32. Mahajan MA, Murray A, Levy D, Samuels HH. Nuclear receptor coregulator (NRC): mapping of the dimerization domain, activation of p53 and STAT‐2, and identification of the activation domain AD2 necessary for nuclear receptor signaling. Mol Endocrinol 2007;21:1822–34. 33. Puigserver P, Adelmant G, Wu Z, Fan M, Xu J, O’Malley B, et al. Activation of PPARgamma coactivator‐1 through transcription factor docking. Science 1999;286:1368–71. 34. Goo YH, Na SY, Zhang H, Xu J, Hong S, Cheong J, et al. Interactions between activating signal cointegrator‐2 and the tumor suppressor retinoblastoma in androgen receptor transactivation. J Biol Chem 2004;279:7131–5. 35. Lu J, Danielsen M. Differential regulation of androgen and glucocorticoid receptors by retinoblastoma protein. J Biol Chem 1998;273:31528–33. 36. Ko L, Cardona GR, Chin WW. Thyroid hormone receptor‐binding protein, an LXXLL motif‐containing protein, functions as a general coactivator. Proc Natl Acad Sci USA 2000;97:6212–7. 37. Chang C, Norris JD, Gron H, Paige LA, Hamilton PT, Kenan DJ, et al. Dissection of the LXXLL nuclear receptor‐coactivator interaction motif using combinatorial peptide libraries: discovery of peptide antagonists of estrogen receptors alpha and beta. Mol Cell Biol 1999;19:8226–39. 38. Zhu Y, Kan L, Qi C, Kanwar YS, Yeldandi AV, Rao MS, et al. Isolation and characterization of peroxisome proliferator‐activated receptor (PPAR) interacting protein (PRIP) as a coactivator for PPAR. J Biol Chem 2000;275:13510–6. 39. Hu X, Lazar MA. The CoRNR motif controls the recruitment of corepressors by nuclear hormone receptors. Nature 1999;402:93–6. 40. Perissi V, Staszewski LM, McInerney EM, Kurokawa R, Krones A, Rose DW, et al. Molecular determinants of nuclear receptor‐corepressor interaction. Genes Dev 1999;13:3198–208. 41. Ko L, Cardona GR, Iwasaki T, Bramlett KS, Burris TP, Chin WW. Ser‐884 adjacent to the LXXLL motif of coactivator TRBP defines selectivity for ERs and TRs. Mol Endocrinol 2002;16:128–40.

ASCOM AS A KEY H3K4MT COMPLEX OF NRS

377

42. Zhao C, Toresson G, Xu L, Koehler KF, Gustafsson JA, Dahlman‐Wright K. Mouse estrogen receptor beta isoforms exhibit differences in ligand selectivity and coactivator recruitment. Biochemistry 2005;44:7936–44. 43. Willy PJ, Mangelsdorf DJ. Unique requirements for retinoid‐dependent transcriptional activation by the orphan receptor LXR. Genes Dev 1997;11:289–98. 44. Son YL, Park OG, Kim GS, Lee JW, Lee YC. RXR heterodimerization allosterically activates LXR binding to the second NR box of activating signal co‐integrator‐2. Biochem J 2008;410:319–30. 45. Nolte RT, Wisely GB, Westin S, Cobb JE, Lambert MH, Kurokawa R, et al. Ligand binding and co‐activator assembly of the peroxisome proliferator‐activated receptor‐gamma. Nature 1998;395:137–43. 46. Ito M, Roeder RG. The TRAP/SMCC/Mediator complex and thyroid hormone receptor function. Trends Endocrinol Metab 2001;12:127–34. 47. Dehe PM, Geli V. The multiple faces of Set1. Biochem Cell Biol 2006;84:536–48. 48. Martin C, Zhang Y. The diverse functions of histone lysine methylation. Nat Rev Mol Cell Biol 2005;6:838–49. 49. Santos‐Rosa H, Schneider R, Bannister AJ, Sherriff J, Bernstein BE, Emre NC, et al. Active genes are tri‐methylated at K4 of histone H3. Nature 2002;419:407–11. 50. Ng HH, Robert F, Young RA, Struhl K. Targeted recruitment of Set1 histone methylase by elongating Pol II provides a localized mark and memory of recent transcriptional activity. Mol Cell 2003;11:709–19. 51. Schneider R, Bannister AJ, Myers FA, Thorne AW, Crane‐Robinson C, Kouzarides T. Histone H3 lysine 4 methylation patterns in higher eukaryotic genes. Nat Cell Biol 2004;6:73–7. 52. Bernstein BE, Kamal M, Lindblad‐Toh K, Bekiranov S, Bailey DK, Huebert DJ, et al. Genomic maps and comparative analysis of histone modifications in human and mouse. Cell 2005;120:169–81. 53. Schubeler D, MacAlpine DM, Scalzo D, Wirbelauer C, Kooperberg C, van Leeuwen F, et al. The histone modification pattern of active genes revealed through genome‐wide chromatin analysis of a higher eukaryote. Genes Dev 2004;18:1263–71. 54. Pokholok DK, Harbison CT, Levine S, Cole M, Hannett NM, Lee TI, et al. Genome‐wide map of nucleosome acetylation and methylation in yeast. Cell 2005;122:517–27. 55. Beisel C, Imhof A, Greene J, Kremmer E, Sauer F. Histone methylation by the Drosophila epigenetic transcriptional regulator Ash1. Nature 2002;419:857–62. 56. Nishioka K, Chuikov S, Sarma K, Erdjument‐Bromage H, Allis CD, Tempst P, et al. Set9, a novel histone H3 methyltransferase that facilitates transcription by precluding histone tail modifications required for heterochromatin formation. Genes Dev 2002;16:479–89. 57. Wang H, Cao R, Xia L, Erdjument‐Bromage H, Borchers C, Tempst P, et al. Purification and functional characterization of a histone H3‐lysine 4‐specific methyltransferase. Mol Cell 2001;8:1207–17. 58. Hamamoto R, Furukawa Y, Morita M, Iimura Y, Silva FP, Li M, et al. SMYD3 encodes a histone methyltransferase involved in the proliferation of cancer cells. Nat Cell Biol 2004;6:731–40. 59. Hayashi K, Yoshida K, Matsui Y. A histone H3 methyltransferase controls epigenetic events required for meiotic prophase. Nature 2005;438:374–8. 60. Briggs SD, Bryk M, Strahl BD, Cheung WL, Davie JK, Dent SY, et al. Histone H3 lysine 4 methylation is mediated by Set1 and required for cell growth and rDNA silencing in Saccharomyces cerevisiae. Genes Dev 2001;15:3286–95. 61. Roguev A, Schaft D, Shevchenko A, Pijnappel WW, Wilm M, Aasland R, et al. The Saccharomyces cerevisiae Set1 complex includes an Ash2 homologue and methylates histone 3 lysine 4. EMBO J 2001;20:7137–48.

378

LEE ET AL.

62. Miller T, Krogan NJ, Dover J, Erdjument‐Bromage H, Tempst P, Johnston M, et al. COMPASS: a complex of proteins associated with a trithorax‐related SET domain protein. Proc Natl Acad Sci USA 2001;98:12902–7. 63. Roguev A, Schaft D, Shevchenko A, Pim Pijnappel WWM, Wilm M, Aasland R, et al. The Saccharomyces cerevisiae Set1 complex includes an Ash2 homologue and methylates histone 3 lysine 4. EMBO J 2001;20:7137–48. 64. Glaser S, Schaft J, Lubitz S, Vintersten K, van der Hoeven F, Tufteland KR, et al. Multiple epigenetic maintenance factors implicated by the loss of Mll2 in mouse development. Development 2006;133:1423–32. 65. Milne TA, Briggs SD, Brock HW, Martin ME, Gibbs D, Allis CD, et al. MLL targets SET domain methyltransferase activity to Hox gene promoters. Mol Cell 2002;10:1107–17. 66. Wysocka J, Herr W. The herpes simplex virus VP16‐induced complex: the makings of a regulatory switch. Trends Biochem Sci 2003;28:294–304. 67. Hughes CM, Rozenblatt‐Rosen O, Milne TA, Copeland TD, Levine SS, Lee JC, et al. Menin associates with a trithorax family histone methyltransferase complex and with the hoxc8 locus. Mol Cell 2004;13:587–97. 68. Yokoyama A, Wang Z, Wysocka J, Sanyal M, Aufiero DJ, Kitabayashi I, et al. Leukemia proto‐ oncoprotein MLL forms a SET1‐like histone methyltransferase complex with menin to regulate Hox gene expression. Mol Cell Biol 2004;24:5639–49. 69. Yu BD, Hess JL, Horning SE, Brown GA, Korsmeyer SJ. Altered Hox expression and segmental identity in Mll‐mutant mice. Nature 1995;378:505–8. 70. Lee JH, Skalnik DG. CpG‐binding protein (CXXC finger protein 1) is a component of the mammalian Set1 histone H3‐Lys4 methyltransferase complex, the analogue of the yeast Set1/ COMPASS complex. J Biol Chem 2005;280:41725–31. 71. Dou Y, Milne TA, Ruthenburg AJ, Lee S, Lee JW, Verdine GL, et al. Regulation of MLL1 H3K4 methyltransferase activity by its core components. Nat Struct Mol Biol 2006;13:713–9. 72. Steward MM, Lee J, O’Donovan A, Wyatt M, Bernstein BE, Shilatifard A. Molecular regulation of H3K4 trimethylation by ASH2L, a shared subunit of MLL complexes. Nat Struct Mol Biol 2006;13:852–4. 73. Wysocka J, Swigut T, Milne TA, Dou Y, Zhang X, Burlingame AL, et al. WDR5 associates with histone H3 methylated at K4 and is essential for H3 K4 methylation and vertebrate development. Cell 2005;121:859–72. 74. Steward MM, Lee JS, O’Donovan A, Wyatt M, Bernstein BE, Shilatifard A. Molecular regulation of H3K4 trimethylation by ASH2L, a shared subunit of MLL complexes. Nat Struct Mol Biol 2006;13:852–4. 75. Sims III RJ, Reinberg D. Histone H3 Lys 4 methylation: caught in a bind? Genes Dev 2006;20:2779–86. 76. Maurer‐Stroh S, Dickens NJ, Hughes‐Davies L, Kouzarides T, Eisenhaber F, Ponting CP. The Tudor domain ‘‘Royal Family’’: tudor, plant Agenet, Chromo, PWWP and MBT domains. Trends Biochem Sci 2003;28:69–74. 77. Bienz M. The PHD finger, a nuclear protein‐interaction domain. Trends Biochem Sci 2006;31:35–40. 78. Neely KE, Workman JL. Histone acetylation and chromatin remodeling: which comes first? Mol Genet Metab 2002;76:1–5. 79. Shi X, Gozani O. The fellowships of the INGs. J Cell Biochem 2005;96:1127–36. 80. Shi Y, Whetstine JR. Dynamic regulation of histone lysine methylation by demethylases. Mol Cell 2007;25:1–14. 81. Mizuguchi G, Tsukiyama T, Wisniewski J, Wu C. Role of nucleosome remodeling factor NURF in transcriptional activation of chromatin. Mol Cell 1997;1:141–50.

ASCOM AS A KEY H3K4MT COMPLEX OF NRS

379

82. Shi X, Hong T, Walter KL, Ewalt M, Michishita E, Hung T, et al. ING2 PHD domain links histone H3 lysine 4 methylation to active gene repression. Nature 2006;442:96–9. 83. Taverna SD, Ilin S, Rogers RS, Tanny JC, Lavender H, Li H, et al. Yng1 PHD finger binding to H3 trimethylated at K4 promotes NuA3 HAT activity at K14 of H3 and transcription at a subset of targeted ORFs. Mol Cell 2006;24:785–96. 84. Martin DG, Baetz K, Shi X, Walter KL, MacDonald VE, Wlodarski MJ, et al. The Yng1p plant homeodomain finger is a methyl‐histone binding module that recognizes lysine 4‐methylated histone H3. Mol Cell Biol 2006;26:7871–9. 85. Yang XJ. Lysine acetylation and the bromodomain: a new partnership for signaling. Bioessays 2004;26:1076–87. 86. Sims III RJ, Millhouse S, Chen CF, Lewis BA, Erdjument‐Bromage H, Tempst P, et al. Recognition of trimethylated histone H3 lysine 4 facilitates the recruitment of transcription postinitiation factors and pre‐mRNA splicing. Mol Cell 2007;28:665–76. 87. Matthews AG, Kuo AJ, Ramon‐Maiques S, Han S, Champagne KS, Ivanov D, et al. RAG2 PHD finger couples histone H3 lysine 4 trimethylation with V(D)J recombination. Nature 2007;450:1106–10. 88. Liu Y, Subrahmanyam R, Chakraborty T, Sen R, Desiderio S. A plant homeodomain in RAG‐ 2 that binds Hypermethylated lysine 4 of histone H3 is necessary for efficient antigen‐ receptor‐gene rearrangement. Immunity 2007;27:561–71. 89. Ramon‐Maiques S, Kuo AJ, Carney D, Matthews AG, Oettinger MA, Gozani O, et al. The plant homeodomain finger of RAG2 recognizes histone H3 methylated at both lysine‐4 and arginine‐2. Proc Natl Acad Sci USA 2007;104:18993–8. 90. Cho YW, Hong T, Hong S, Guo H, Yu H, Kim D, et al. PTIP associates with MLL3‐ and MLL4‐containing histone H3 lysine 4 methyltransferase complex. J Biol Chem 2007;282:20395–406. 91. Issaeva I, Zonis Y, Rozovskaia T, Orlovsky K, Croce CM, Nakamura T, et al. Knockdown of ALR (MLL2) reveals ALR target genes and leads to alterations in cell adhesion and growth. Mol Cell Biol 2007;27:1889–903. 92. Simon JA, Lange CA. Roles of the EZH2 histone methyltransferase in cancer epigenetics. Mutat Res 2008;647:21–9. 93. Mo R, Rao SM, Zhu YJ. Identification of the MLL2 complex as a coactivator for estrogen receptor alpha. J Biol Chem 2006;281:15714–20. 94. Ruthenburg AJ, Allis CD, Wysocka J. Methylation of lysine 4 on histone H3: intricacy of writing and reading a single epigenetic mark. Mol Cell 2007;25:15–30. 95. Qi C, Surapureddi S, Zhu YJ, Yu S, Kashireddy P, Rao MS, et al. Transcriptional coactivator PRIP, the peroxisome proliferator‐activated receptor gamma (PPARgamma)‐interacting protein, is required for PPARgamma‐mediated adipogenesis. J Biol Chem 2003;278:25281–4. 96. Dreijerink KM, Mulder KW, Winkler GS, Hoppener JW, Lips CJ, Timmers HT. Menin links estrogen receptor activation to histone H3K4 trimethylation. Cancer Res 2006;66:4929–35. 97. Cho H, Orphanides G, Sun X, Yang XJ, Ogryzko V, Lees E, et al. A human RNA polymerase II complex containing factors that modify chromatin structure. Mol Cell Biol 1998;18:5355–63. 98. Krogan NJ, Dover J, Wood A, Schneider J, Heidt J, Boateng MA, et al. The Paf1 complex is required for histone H3 methylation by COMPASS and Dot1p: linking transcriptional elongation to histone methylation. Mol Cell 2003;11:721–9. 99. Pavri R, Zhu B, Li G, Trojer P, Mandal S, Shilatifard A, et al. Histone H2B monoubiquitination functions cooperatively with FACT to regulate elongation by RNA polymerase II. Cell 2006;125:703–17. 100. Shahbazian MD, Zhang K, Grunstein M. Histone H2B ubiquitylation controls processive methylation but not monomethylation by Dot1 and Set1. Mol Cell 2005;19:271–7.

380

LEE ET AL.

101. Sierra J, Yoshida T, Joazeiro CA, Jones KA. The APC tumor suppressor counteracts beta‐ catenin activation and H3K4 methylation at Wnt target genes. Genes Dev 2006;20:586–600. 102. Dou Y, Milne TA, Tackett AJ, Smith ER, Fukuda A, Wysocka J, et al. Physical association and coordinate function of the H3 K4 methyltransferase MLL1 and the H4 K16 acetyltransferase MOF. Cell 2005;121:873–85. 103. Milne TA, Dou Y, Martin ME, Brock HW, Roeder RG, Hess JL. MLL associates specifically with a subset of transcriptionally active target genes. Proc Natl Acad Sci USA 2005;102:14765–70. 104. Guenther MG, Jenner RG, Chevalier B, Nakamura T, Croce CM, Canaani E, et al. Global and Hox‐specific roles for the MLL1 methyltransferase. Proc Natl Acad Sci USA 2005;102:8603–8. 105. Smith ST, Petruk S, Sedkov Y, Cho E, Tillib S, Canaani E, et al. Modulation of heat shock gene expression by the TAC1 chromatin‐modifying complex. Nat Cell Biol 2004;6:162–7. 106. Wysocka J, Swigut T, Xiao H, Milne TA, Kwon SY, Landry J, et al. A PHD finger of NURF couples histone H3 lysine 4 trimethylation with chromatin remodelling. Nature 2006;442:86–90. 107. Lee S, Kim D, Goo Y, Lee YC, Lee SK, Lee JW. Crucial roles for interactions between MLL3/ 4 and INI1 in nuclear receptor transactivation. Mol Endocrinol 2009;23:610–19. 108. Tian H, Mahajan MA, Wong CT, Habeos I, Samuels HH. The N‐Terminal A/B domain of the thyroid hormone receptor‐beta2 isoform influences ligand‐dependent recruitment of coactivators to the ligand‐binding domain. Mol Endocrinol 2006;20:2036–51. 109. Jung DJ, Na SY, Na DS, Lee JW. Molecular cloning and characterization of CAPER, a novel coactivator of activating protein‐1 and estrogen receptors. J Biol Chem 2002;277:1229–34. 110. Dowhan DH, Hong EP, Auboeuf D, Dennis AP, Wilson MM, Berget SM, et al. Steroid hormone receptor coactivation and alternative RNA splicing by U2AF65‐related proteins CAPERalpha and CAPERbeta. Mol Cell 2005;17:429–39. 111. Imai H, Chan EK, Kiyosawa K, Fu XD, Tan EM. Novel nuclear autoantigen with splicing factor motifs identified with antibody from hepatocellular carcinoma. J Clin Invest 1993;92:2419–26. 112. Rappsilber J, Ryder U, Lamond AI, Mann M. Large‐scale proteomic analysis of the human spliceosome. Genome Res 2002;12:1231–45. 113. Iwasaki T, Chin WW, Ko L. Identification and characterization of RRM‐containing coactivator activator (CoAA) as TRBP‐interacting protein, and its splice variant as a coactivator modulator (CoAM). J Biol Chem 2001;276:33375–83. 114. Auboeuf D, Dowhan DH, Li X, Larkin K, Ko L, Berget SM, et al. CoAA, a nuclear receptor coactivator protein at the interface of transcriptional coactivation and RNA splicing. Mol Cell Biol 2004;24:442–53. 115. Zhu Y, Qi C, Cao WQ, Yeldandi AV, Rao MS, Reddy JK. Cloning and characterization of PIMT, a protein with a methyltransferase domain, which interacts with and enhances nuclear receptor coactivator PRIP function. Proc Natl Acad Sci USA 2001;98:10380–5. 116. Misra P, Qi C, Yu S, Shah SH, Cao WQ, Rao MS, et al. Interaction of PIMT with transcriptional coactivators CBP, p300, and PBP differential role in transcriptional regulation. J Biol Chem 2002;277:20011–9. 117. Reyes JC, Muchardt C, Yaniv M. Components of the human SWI/SNF complex are enriched in active chromatin and are associated with the nuclear matrix. J Cell Biol 1997;137:263–74. 118. Razin SV, Iarovaia OV, Sjakste N, Sjakste T, Bagdoniene L, Rynditch AV, et al. Chromatin domains and regulation of transcription. J Mol Biol 2007;369:597–607. 119. McKenna NJ, Xu J, Nawaz Z, Tsai SY, Tsai MJ, O’Malley BW. Nuclear receptor coactivators: multiple enzymes, multiple complexes, multiple functions. J Steroid Biochem Mol Biol 1999;69:3–12.

ASCOM AS A KEY H3K4MT COMPLEX OF NRS

381

120. Rozenblatt‐Rosen O, Rozovskaia T, Burakov D, Sedkov Y, Tillib S, Blechman J, et al. The C‐terminal SET domains of ALL‐1 and TRITHORAX interact with the INI1 and SNR1 proteins, components of the SWI/SNF complex. Proc Natl Acad Sci USA 1998;95:4152–7. 121. Rozovskaia T, Rozenblatt‐Rosen O, Sedkov Y, Burakov D, Yano T, Nakamura T, et al. Self‐ association of the SET domains of human ALL‐1 and of Drosophila TRITHORAX and ASH1 proteins. Oncogene 2000;19:351–7. 122. Sarkar J, Qi C, Guo D, Ahmed MR, Jia Y, Usuda N, et al. Transcription coactivator PRIP, the peroxisome proliferator‐activated receptor (PPAR)‐interacting protein, is redundant for the function of nuclear receptors PPARalpha and CAR, the constitutive androstane receptor, in mouse liver. Gene Exp 2007;13:255–69. 123. Choi E, Lee S, Yeom SY, Kim GH, Lee JW, Kim SW. Characterization of activating signal cointegrator‐2 as a novel transcriptional coactivator of the xenobiotic nuclear receptor constitutive androstane receptor. Mol Endocrinol 2005;19:1711–9. 124. Qi C, Kashireddy P, Zhu YT, Rao SM, Zhu YJ. Null mutation of peroxisome proliferator‐ activated receptor‐interacting protein in mammary glands causes defective mammopoiesis. J Biol Chem 2004;279:33696–701. 125. Tontonoz P, Spiegelman BM. Fat and beyond: the diverse biology of PPARgamma. Annu Rev Biochem 2008;77:289–312. 126. Li Q, Chu MJ, Xu J. Tissue‐ and nuclear receptor‐specific function of the C‐terminal LXXLL motif of coactivator NCoA6/AIB3 in mice. Mol Cell Biol 2007;27:8073–86. 127. Davis RA, Miyake JH, Hui TY, Spann NJ. Regulation of cholesterol‐7alpha‐hydroxylase: barely missing a SHP. J Lipid Res 2002;43:533–43. 128. Sinal CJ, Tohkin M, Miyata M, Ward JM, Lambert G, Gonzalez FJ. Targeted disruption of the nuclear receptor FXR/BAR impairs bile acid and lipid homeostasis. Cell 2000;102:731–44. 129. Tanner MM, Tirkkonen M, Kallioniemi A, Isola J, Kuukasjarvi T, Collins C, et al. Independent amplification and frequent co‐amplification of three nonsyntenic regions on the long arm of chromosome 20 in human breast cancer. Cancer Res 1996;56:3441–5. 130. Guan XY, Xu J, Anzick SL, Zhang H, Trent JM, Meltzer PS. Hybrid selection of transcribed sequences from microdissected DNA: isolation of genes within amplified region at 20q11‐ q13.2 in breast cancer. Cancer Res 1996;56:3446–50. 131. Lee SK, Anzick SL, Choi JE, Bubendorf L, Guan XY, Jung YK, et al. A nuclear factor, ASC‐2, as a cancer‐amplified transcriptional coactivator essential for ligand‐dependent transactivation by nuclear receptors in vivo. J Biol Chem 1999;274:34283–93. 132. Ruault M, Brun ME, Ventura M, Roize`s G, De Sario A. MLL3, a new human member of the TRX/MLL gene family, maps to 7q36, a chromosome region frequently deleted in myeloid leukaemia. Gene 2002;284:73–81. 133. Zhang H, Kuang SQ, Liao L, Zhou S, Xu J. Haploid inactivation of the amplified‐in‐breast cancer 3 coactivator reduces the inhibitory effect of peroxisome proliferator‐activated receptor gamma and retinoid X receptor on cell proliferation and accelerates polyoma middle‐T antigen‐induced mammary tumorigenesis in mice. Cancer Res 2004;64:7169–77. 134. Cho EA, Prindle MJ, Dressler GR. BRCT domain‐containing protein PTIP is essential for progression through mitosis. Mol Cell Biol 2003;23:1666–73. 135. Jowsey PA, Doherty AJ, Rouse J. Human PTIP facilitates ATM‐mediated activation of p53 and promotes cellular resistance to ionizing radiation. J Biol Chem 2004;279:55562–9. 136. Munoz M, Jowsey PA, Toth R, Rouse J. Phospho‐epitope binding by the BRCT domains of hPTIP controls multiple aspects of the cellular response to DNA damage. Nucleic Acids Res 2007;35:5312–22. 137. Bartek J, Bartkova J, Lukas J. DNA damage signalling guards against activated oncogenes and tumour progression. Oncogene 2007;26:7773–9.

382

LEE ET AL.

138. Riches LC, Lynch AM, Gooderham NJ. Early events in the mammalian response to DNA double‐strand breaks. Mutagenesis 2008;23:331–9. 139. Iwabuchi K, Bartel PL, Li B, Marraccino R, Fields S. Two cellular proteins that bind to wild‐type but not mutant p53. Proc Natl Acad Sci USA 1994;91:6098–102. 140. Iwabuchi K, Li B, Massa HF, Trask BJ, Date T, Fields S. Stimulation of p53‐mediated transcriptional activation by the p53‐binding proteins, 53BP1 and 53BP2. J Biol Chem 1998;273:26061–8. 141. Nishino T, Morikawa K. Structure and function of nucleases in DNA repair: shape, grip and blade of the DNA scissors. Oncogene 2002;21:9022–32. 142. Ko L, Chin WW. Nuclear receptor coactivator thyroid hormone receptor‐binding protein (TRBP) interacts with and stimulates its associated DNA‐dependent protein kinase. J Biol Chem 2003;278:11471–9. 143. Zgheib O, Huyen Y, DiTullio Jr RA, Snyder A, Venere M, Stavridi ES, et al. ATM signaling and 53BP1. Radiother Oncol 2005;76:119–22. 144. Ju BG, Rosenfeld MG. A breaking strategy for topoisomerase IIbeta/PARP‐1‐dependent regulated transcription. Cell Cycle 2006;5:2557–60. 145. Medina PP, Sanchez‐Cespedes M. Involvement of the chromatin‐remodeling factor BRG1/ SMARCA4 in human cancer. Epigenetics 2008;3:64–8. 146. Sansam CG, Roberts CW. Epigenetics and cancer: altered chromatin remodeling via Snf5 loss leads to aberrant cell cycle regulation. Cell Cycle 2006;5:621–4. 147. Roberts CW, Orkin SH. The SWI/SNF complex—chromatin and cancer. Nat Rev Cancer 2004;4:133–42.

Index

A Acf1 and ISWI complexes, NR regulation ACF complex and repressive role, 216–217 IGFBP3 and RANKL repression, 220 Isw2/Rpd3 complex, 220–221 NCoR/SMRT complex and repression, 218–219, 221 Snf2h repression activity, 218–219 TIP5 functions, 222 transactivation, 217–218 WINAC complex and repression, 221 Activating signal cointegrator-2 (ASC-2) expression and isoform, 347–348 homodimerization domain, 350 NR box 1, 349 NR box 2, 349–350 transactivation domains, 348 Acute promyelocytic leukemia (APL), 247 Adipose and intestine, ERR, 34–35 ANCCA coregulator role, cancer AIB1/ACTR target, 288–289 overexpression and potential role, 289–290 ASCOM, NR-mediated transactivation cancer activity amplification and overexpression, 370 MLL3/4 function, 371–372 p53 regulation, 370–371 PTIP and p53 phosphorylation, 371 suppressive function, 370 CBP/p300 cross talk, 358 ER-transactivation, 356 metabolic phenotypes FXR-target genes activation, 368–369 LXRs transactivation, 368 WAT, 367 MLL3 and MLL4, 354–355 physiological role cell growth and development, 362–364 DN1 and DN2, 362 fertility, 364–365

liver function, 365–366 mammary gland function, 365 PPAR -mediated adipogenesis, 355 recruitment mechanisms, 356–357 RNA-binding protein, cross talk, 359–360 Swi/Snf cross talk, 360–362 Assembly systems, chromatin analyzing methods, 154–156 biochemical fractionation, 153 cell extraction, 152–153 salt dilution/dialysis, 151–152 Atherosclerosis farnesoid X receptor (FXR), 23–24 liver X receptors (LXR), 15 peroxisome proliferator activated receptors (PPARs), 9–10 ATP-dependent chromatin remodeler Acf1 and ISWI complexes, regulation IGFBP3 and RANKL repression, 220 Isw2/Rpd3 complex, 220–221 NCoR/SMRT complex and repression, 218–219, 221 repressive role and ACF complex, 216–217 Snf2h repression activity, 218–219 TIP5 functions, 222 transactivation, 217–218 WINAC complex and repression, 221 SWItching/sucrose nonfermentors (SWI/SNF) complexes class I NRs coactivation, 213–214 VDR–WINAC connection, 214–216

B Bile metabolism control, FXR, 20–21 Biochemical methods chromatin analyzing methods, 154–156 assembling methods, 151–153

383

384

index

Biochemical methods (cont. ) DNA templates, 149 histones, 149–151 in vitro transcription, 156 nuclear receptor-dependent transcription coregulator proteins, purification, 148–149 DNA templates, 146–147 in vitro transcription systems, 145–146 transcription machinery, 147 transcripts detection, 147–148 Brahma-related gene 1 (BRG1), 201

C Cancer ANCCA coregulator role AIB1/ACTR target, 288–289 overexpression and potential role, 289–290 ASCOM amplification and overexpression, 370 MLL3/4 function, 371–372 p53 regulation, 370–371 PTIP and p53 phosphorylation, 371 suppressive function, 370 endometrial genes regulation, progestin, 63–64 glands and uterine leiomyomas, 61–62 morphological and biochemical evaluations, 62 progesterone action, 66 progestin therapy, 62–63 transcriptional activity, 65 ERR, 35–36 CAR. See Constitutive androstane receptor Chromatin acetylation, 204 analyzing methods imaging methods, 156 nuclease digestion, 154–155 physical properties, 155–156 binding kinetics, NR, 174 biochemical methods analyzing methods, 154–156 assembling methods, 151–153 DNA templates, 149 histones, 149–151 in vitro transcription, 156 histone code, 203–204

inhibitor of acetyltransferases (INHAT) role, 205–210 methylation, 205 NR function structural alterations, 144 Swi/Snf complexes, 145 nucleosomes, 202, 210–211 physiological template dynamic properties, 144 histones, 143 nucleosome anatomy, 144 Chromosomal alterations, p160/SRC gene amplification, ABI and SRC-1, 264–267 translocation and inversion, 267–270 Constitutive androstane receptor (CAR) activation mechanism, 107–108 CYP2B genes regulator, identification, 104–105 species differences, 108–109 transcriptional regulation CYP2C genes, 105–106 drug transporters, 106–107 UGT1A1 genes, 106 Coregulators, NR ATP-dependent CRCS, 163 mediator, 164–165 methyltransferases and kinases, 162–163 P300 AND CBP acetyltransferases, 161–162 P160/SRC proteins, 160–161

D DBD. See DNA-binding domain Diabetes estrogen related receptors (ERRs), 34 farnesoid X receptor (FXR), 24 liver X receptors (LXR), 17–18 DNA-binding domain (DBD), 57, 89, 91, 93 DNA methylation nuclear receptors (NRs), 247 protein arginine methyltransferases (PRMTs), 330 Domain structures, nuclear receptor DBD and AF-1 domain, 91 LBD and AF-2 domain, 89–90

E Endometrial cancer genes regulation, progestin, 63–64

385

index glands and uterine leiomyomas, 61–62 morphological and biochemical evaluations, 62 perspectives of progesterone action, 66 progestin therapy, 62–63 transcriptional activity, 65 Endometrium and myometrium, progesterone action activation function (AF) domains, 57–58 coregulators, 58–60 physiological response, 56–57 receptor expression, 60 Estrogen receptor related receptor (ERR) adipose and intestine, 34–35 and cancer, 35–36 and diabetes, 34 PGC-1 pathway, 32–34 therapeutic potential gene expression, 36 oxidative phosphorylation, 37 Extracellular matrix (ECM), 66, 68–70

F Farnesoid X receptor (FXR) and atherosclerosis, 23–24 bile metabolism control small heterodimeric partner (SHP), 20 synthetic pathway, 20–21 and diabetes, 24 gallstones, cholestasis, and bacterial growth, 21–22 and lipid metabolism, 22–23 liver regeneration and tumorigenesis, 24–25 therapeutic potential, 25–26

G Glucocorticoid resistance, 207 Glutathione S-transferase (GST) regulation, 96

H Histone H3-lysine 4-methyltransferase (H3K4MT) activating signal cointegrator-2 (ASC-2) expression and isoform, 347–348

homodimerization domain, 350 NR box 1, 349 NR box 2, 349–350 transactivation domains, 348 ASCOM, NR-mediated transactivation cancer activity, 370–372 CBP/p300 cross talk, 358 ER-transactivation, 356 MLL3 and MLL4, 354–355 PPAR -mediated adipogenesis, 355 RNA-binding protein, cross talk, 359–360 Set1-like complexes, recuitment, 356–357 Swi/Snf cross talk, 360–362 domains, ASC-2 and MLL4, 346 pharmacological possibilities, 372–373 physiological role cell growth and development, 362–364 DN1 and DN2, 362 fertility, 364–365 liver function, 365–366 mammary gland function, 365 metabolic phenotypes, 367–369 Set1-like complexes methylation, 351 methyl-binding effectors, 353–354 multiplicity, vertebrates, 351–352 subcomplexes, 352–353 working models, 352 Hormone response elements (HREs), 91 HREs. See Hormone response elements

I Inflammation liver X receptors (LXR), 15–17 peroxisome proliferator activated receptors (PPARs), 10–12 retinoid-related orphan receptors (RORs), 30–31 Inhibitor of acetyltransferases (INHAT) role bi-functional coregulator function, 207–208 glucocorticoid resistance, 207 and histone chaperones, 209–210 histone masking, 206 Jun dimerization protein 2 (JDP2), 208–209 multitasking role, 208 NR regulation, 206 repressor activity, 209

386

index J

Jumonji (JmjC) domain, 310 Jun dimerization protein 2 (JDP2), 208–209

L LBD. See Ligand-binding domain Leiomyoma ECM deposition, 68–70 growth factor regulation, 70–71 progesterone action and relevance, 67–68, 73–74 proliferation, apoptosis, 68–70 signaling pathways, 71–73 Ligand-binding domain (LBD), 57, 89–90, 93, 196 Ligands role, NR-mediated transcription agonist ligands, 157–158 coregulators interactions, 158–159 Liver X receptors (LXRs) and atherosclerosis, 15 and diabetes, 17–18 hepatic lipid metabolism regulation, 13 and inflammation, 15–17 reverse cholesterol transport regulation, 13–15 therapeutic potential, 18–19

M Metabolic phenotypes, ASCOM FXR-target genes activation, 368–369 LXRs transactivation, 368 WAT, 367 Mixed lineage leukemia (MLL) complex, 308, 327 MOZ gene structure–function, 269 TIF2 fusion, 267–268

N Nuclear hormone receptors estrogen receptor related receptor (ERR) adipose and intestine, 34–35 and cancer, 35–36 and diabetes, 34

PGC-1 pathway, 32–34 therapeutic potential, 36–37 farnesoid X receptor (FXR) and atherosclerosis, 23–24 bacterial growth, 21–22 bile metabolism control, 20–21 and diabetes, 24 gallstones, cholestasis, 21–22 and lipid metabolism, 22–23 liver regeneration and tumorigenesis, 24–25 therapeutic potential, 25–26 liver X receptors (LXRs) and atherosclerosis, 15 and diabetes, 17–18 hepatic lipid metabolism regulation, 13 and inflammation, 15–17 reverse cholesterol transport regulation, 13–15 therapeutic potential, 18–19 peroxisome proliferator activated receptors (PPARs) and atherosclerosis, 9–10 and inflammation, 10–12 PPAR , 5–6 PPAR, 7–9 PPAR, 4–5 retinoid-related orphan receptor (ROR) circadian rhythm control and metabolism, 28–30 and inflammation, 30–31 lipid metabolism regulation, 26–28 therapeutic potential, 31 Nuclear receptor corepressor (NCoR) ligand-dependent (LCoRs) hairless (Hr), 241 receptor-interacting protein (RIP) 140, 240–241 ligand-independent Alien protein, 239 NCoR/SMRT protein complex, 238–239 SIN3 complexes, 239 SUV39H1 methyltransferase, 240 Nuclear receptor-dependent transcription biochemical methods coregulator proteins, purification, 148–149 DNA templates, 146–147 in vitro transcription systems, 145–146 transcription machinery, 147 transcripts detection, 147–148

index chromatin assembly and transcription, role basal TFs, 159 biochemical methods, 149–156 chromatin structural alterations, 144 coactivators, 160–165 corepressors, 165–166 ligands, 157–159 NR function, 144–145 nucleosome anatomy, 144 nucleosome-binding proteins, 166–168 order and dynamics, 168–174 physiological template, 143–144 general structural and functional organization, 138 interactions, and functions biochemical activities, 139 coregulators, 139–141 RNA polymerase II transcription machinery, 139–140 transcriptional regulation, 142–143 ligand-dependent transcriptional regulation, 138–139 temporal aspects RNA Pol II binding, 168–169 SRC and p300/ CBP actions, 171–172 transcriptional activation genomic analyses, 176–177 single molecule biophysical analyses, 175–176 transcription repression, 175 transcription factors (TFs), 138–139 Nuclear receptor-mediated transcription phosphorylation pathways, 123 proteasome inhibition, 121–122 protein degradation, 120–121 ubiquitin proteasome system, 118–120 Nuclear receptors (NRs). See also Protein arginine methyltransferases (PRMTs) ATP-dependent chromatin remodeler, 223 (see also ATP-dependent chromatin remodeler) Acf1 and ISWI complexes, regulation, 216–222 SWI/SNF complexes, 212–216 chromatin acetylation, 204 histone code, 203–204 inhibitor of acetyltransferases (INHAT) role, 205–210

387 methylation, 205 nucleosomes, 202, 210–211 classification class I and II, 195 class III and IV, 195–196 corepressor (NCoR) ligand-dependent (LCoRs), 240–241 ligand-independent, 238–240 histone acetylation, 197 NCoR/SMRT complex, corepressor histone deacetylases (HDACs), 200–201 mammalian switch-independent 3 (mSin3), 200 structure, 199–200 SWI/SNF and Mi2/NuRD complexes, 201 physiological functions, repression metamorphosis regulation, 249 NCoR and SMRT activity, 250–251 PPAR regulator activity, 249–250 RIP140 activity, 251 unliganded thyroid hormone receptors (TR), 248–249 p160/steroid receptor coactivator (SRC), 198 repression types agonist-bound receptors, 243–244 antagonist-bound steroid receptors, 242–243 unliganded receptors, 241–242 structure conserved domains, 196 PPAR –RXR complex, 196–197 target gene regulation, 224 transcriptional repression, molecular mechanisms chromatin assembly/remodeling, 246–247 DNA methylation, 247 histone deacteylation, 244–245 histone methylation, 245–246 Nuclear receptor signaling SUMOylation, 128–129 ubiquitin proteasome system (UPS) coregulators, 123–125 nuclear receptor interactions, 119–123 protein degradation, 119 targeting system, 118–119 targets, 123–125 transcriptional activities, 118 Nuclear xenobiotic receptors constitutive androstane receptor (CAR) activation mechanism, 107–108

388

index

Nuclear xenobiotic receptors (cont. ) CYP2B genes regulator, identification, 104–105 regulation, other DMEs, 105–107 species differences, 108–109 domain structures DBD and AF-1 domain, 91 LBD and AF-2 domain, 89–90 ligands, 88–89 physiology and diseases, 92 pregnane X receptor (PXR) cloning and initial characterization, 93 drug transporter regulation, 96–97 endobiotic functions, 98–101 gene regulation, drug metabolism, 97–98 phase I CYP enzyme regulation, 94 phase II enzyme regulation, 94–96 species specificity, humanized mice, 101–104

P Peptidyl deiminase 4 (PAD4), 309–310 Peroxisome proliferator activated receptors (PPARs) and atherosclerosis, 9–10 and inflammation, 10–12 PPAR fatty acid regulation, 5 thiazolidinedione (TZD), 6 PPAR, 7–9 PPAR, 4–5 Platelet derived growth factor (PDGF), 71–72 Polyomavirus middle T antigen (PyMT) model, 287 Post-translational modification (PTM), histones acetylation, 204 histone code, 203–204 methylation, 205 PPAR-dependent endurance pathway, 8–9 PPARs. See Peroxisome proliferator activated receptors Pregnane X receptor (PXR) cloning and initial characterization, 93 drug transporter regulation, 96–97 endobiotic functions adrenal steroid homeostasis, 100 bile acid detoxification and cholestasis, 99

bilirubin detoxification and clearance, 99–100 drug–hormone interactions, 100 lipid metabolism, 100–101 gene regulation, drug metabolism, 97–98 phase I CYP enzyme regulation, 94 phase II enzyme regulation glutathione S-transferase (GST), 96 sulfotransferases (SULT), 95–96 UDP-glucuronosyltransferase (UGT), 94–95 species specificity, humanized mice creation and characterization, 102–103 drug–drug interactions, 104 drug metabolism models, 101 rodent AND hPXR, 101–102 Progesterone receptor endometrial cancer genes regulation, progestin, 63–64 glands and uterine leiomyomas, 61–62 morphological and biochemical evaluations, 62 progesterone action, 66 progestin therapy, 62–63 transcriptional activity, 65 endometrium and myometrium, progesterone action activation function (AF) domains, 57–58 coregulators, 58–60 physiological response, 56–57 receptor expression, 60 leiomyoma ECM deposition, 68–70 growth factor regulation, 70–71 mitogenic effect, progesterone, 67–68 progesterone action perspectives, 73–74 proliferation, apoptosis, 68–70 signaling pathways, 71–73 uterine leiomyoma, 66–67 uterus, 54–55 Protein arginine methyltransferases (PRMTs) biological functions chromatin domains, 331 developmental process, 326–327 DNA repair and methylation, 330–331 muscle differentiation, 325 precursor maintenance, 326 cancer and CARM1 levels, 328 MLL-mediated transformation, 327

389

index oncogenic role, 327–328 classification, 300 enzymatic activity asymmetric dimethylation, type I, 301–303 distributive vs. processive mechanism, 304–305 HRMT1L1 and PMRT10, 304 posttranslational regulation, 306 substrate specificity and regulation, 305–306 symmetric dimethylation, type II, 303–304 metabolic functions adipogenesis, 329–330 gluconeogenesis, 329 methylated arginine, 301 posttranscriptional regulation, 318–319 small molecule inhibitors arginine methyltransferase (AMI), 323–324 HMT1-specific, 323 pyrazole, 235, 324 Sinefungin, 323 Sirtuin, 324 structural analysis CARM1 and functional significance, 321 catalytic site, 321–322 dimerization, 319 pleckstrin homology (PH) domain, 319–321 sequence alignment, 320 transcriptional regulation CARM1 factor, 315 COPR5 factor, 316 E2F activation, 317–318 FOXO1 factor, 314–315 histone methyltransferases, 307–310 NF-B activation, 317 nuclear receptor regulation, 310–314 p53 activation, 316–317 transcription elongation, 318 viral infection and immune response HIV-1, 328–329 human T cell lymphotropic virus type I (HTLV1), 328 p160/SRC coactivator role, cancer aberrant expression overexpression and disease implication, 272–273 transcriptional deregulation, AIB1/ACTR

gene, 273–275 action mechanism cell proliferation, 275–277 IGF/AKT signaling and cell survival, 277–279 invasion and metastasis, 279–280 androgen independence, prostrate cancer binding affinity, 285 receptor (AR) activation, 284–285 chromosomal alterations AIB1 and SRC-1 amplification, 264–267 translocation and inversion, 267–270 functional domains, 263 genetic alteration, 265–266 polymorphism polyglutamine repeat number, 271–272 single nucleotide polymorphisms (SNPs), 270–271 SRC-3 identification, 263 tamoxifen resistance, breast cancer AIB1/ACTR and HER2 overexpression, 281–282 AIB1/ACTR isoform activity, 282–283 E2F-regulation and HER2 signaling, 282 SRC-1 interaction, 283 tumorigenesis function metastasis, mammary gland, 286–287 TRAMP model, 288 Pyrazole, PRMT inhibitor, 235, 324

R Retinoid-related orphan receptor (ROR) circadian rhythm control and metabolism, 28–30 and inflammation, 30–31 lipid metabolism regulation, 26–28 therapeutic potential, 31 RNA transcripts detection methods, 147–148 ROR. See Retinoid-related orphan receptor

S Set1-like complexes, H3K4MT methylation, 351 methyl-binding effectors, 353–354 multiplicity, vertebrates, 351–352 subcomplexes, 352–353 working models, 352

390

index

Sinefungin, PRMT inhibitor, 323 Sirtuin, PRMT inhibitor, 324 Steroid receptor coactivator (SRC), 198 Sulfotransferases (SULT) regulation, 95–96 SWItching/sucrose nonfermentors (SWI/SNF) complexes, NRs, 212–216

T Tamoxifen resistance, breast cancer AIB1/ACTR and HER2 overexpression, 281–282 AIB1/ACTR isoform activity, 282–283 E2F-regulation and HER2 signaling, 282 SRC-1 interaction, 283 Thyrotropin (TSH) repression, 243 Transcription initiation, reinitiation, 172–173 Transgenic adenocarcinoma, mouse prostate (TRAMP) model, 288

U Ubiquitin proteasome system (UPS)

coregulators, 123–125 nuclear hormone receptors, 118 nuclear receptor interactions nuclear receptor-mediated transcription, 119–123, 125 protein degradation, 120–121 nuclear receptor signal, 128–129 proteasome inhibition, 125 protein acetylation, 127 SRC-3 proteasome-mediated degradation, 125–126 UDP-glucuronosyltransferase (UGT) regulation, 94–95 UPS. See Ubiquitin proteasome system Uterine leiomyoma extracellular matrix (ECM), 61, 66, 68–70 gene expression, 73–74 growth factor regulation, 70–71 progesterone receptor action mitogenic effect, 68 pathogenesis, 67 proliferation, apoptosis, 68–70 signaling pathways activation, 71–73 Uterus, 54–55