ADVANCES IN PROTEIN CHEMISTRY Volume 67 Proteins in Eukaryotic Transcription
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ADVANCES IN PROTEIN CHEMISTRY EDITED BY FREDERIC M. RICHARDS
DAVID S. EISENDBERG
Department of Molecular Biophysics and Biochemistry, Yale University New Haven, Connecticut
Department of Chemistry and Biochemistry University of California, Los Angeles Los Angeles, California
JOHN KURIYAN Department of Molecular Biophysics Howard Hughes Medical Institute Rockefeller University New York, New York
VOLUME 67
Proteins in Eukaryotic Transcription EDITED BY Ronald C. Conaway
Joan Weliky Conaway
Stowers Institute for Medical Research Kansas City, Missouri
Stowers Institute for Medical Research Kansas City, Missouri
Elsevier Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK
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CONTENTS PREFACE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
ix
Structure and Function of RNA Polymerase II Patrick Cramer
I. II. III. IV. V.
Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Structure of RNA Polymerase II . . . . . . . . . . . . . . . . . . . . . . . . . . Function of RNA Polymerase II . . . . . . . . . . . . . . . . . . . . . . . . . . Comparison with Other Polymerases . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1 3 13 27 30 31
The Mediator Complex Stefan Bjo € rklund and Claes M. Gustafsson
I. II. III. IV. V.
Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Saccharomyces cerevisiae Mediator . . . . . . . . . . . . . . . . . . . . . . . . . . Mediator Complexes in Higher Eukaryotes . . . . . . . . . . . . . . . . Mechanism of Transcriptional Activation. . . . . . . . . . . . . . . . . . Concluding Remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
43 43 52 55 62 62
Structure and Function of the TFIID Complex Oranart Matangkasombut, Roy Auty, and Stephen Buratowski
I. II. III. IV. V. VI. VII.
TFIID and Transcription Initiation . . . . . . . . . . . . . . . . . . . . . . . TFIID Components and Structure. . . . . . . . . . . . . . . . . . . . . . . . TFIID Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The Requirement for TFIID In Vivo . . . . . . . . . . . . . . . . . . . . . . Regulation of TFIID . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . The TFIID-Chromatin Connection . . . . . . . . . . . . . . . . . . . . . . . Future Questions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v
67 68 76 82 84 85 87 87
vi
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Tetratricopeptide Repeats of TFC4 and a Limiting Step in the Assembly of the Initiation Factor TFIIIB Robyn D. Moir and Ian M. Willis
I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. Brf1-TFIIIC: A Limiting Interaction in Preinitiation Complex Assembly . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . III. Brf1-Tfc4 Interactions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Bdp1-Tfc4 Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Ligand Binding by TPR Arrays in Tfc4 . . . . . . . . . . . . . . . . . . . . VI. Ordered Binding of TFIIIB Subunits to TFIIIC-DNA and Dynamic Interactions with Tfc4 . . . . . . . . . . . . . . . . . . . . . . . . . . VII. Tfc4 and Other Pol III Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
93 95 97 105 107 111 114 115 116
Mechanism of RNA Polymerase I Transcription Lucio Comai
I. II. III. IV. V. VI. VII. VIII. IX.
Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ribosomal DNA Gene Structure . . . . . . . . . . . . . . . . . . . . . . . . . . Species-Specificity of Ribosomal RNA Transcription. . . . . . . . . Factors Involved in Ribosomal RNA Transcription . . . . . . . . . . Assembly of an RNA Polymerase I Initiation Complex in Vertebrates. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Factors Required for RNA Polymerase I Transcription in Yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulatory Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chromatin and RNA Polymerase I Transcription . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
123 124 127 128 136 138 140 147 149 149 149
Functional Properties of ATP-Dependent Chromatin Remodeling Enzymes Anthony N. Imbalzano and Hengyi Xiao
I. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. History of Nucleosome-Remodeling Complexes . . . . . . . . . . . .
157 158
CONTENTS
III. IV. V. VI.
Requirements for Nucleosome-Remodeling Enzymes . . . . . . . Mechanisms of ATP-Dependent Chromatin Remodeling . . . . Initiation of ATP-Dependent Chromatin Remodeling . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
vii 163 168 174 176 177 177
Histone Acetyltransferase Proteins Contribute to Transcriptional Processes at Multiple Levels Michael S. Torok and Patrick A. Grant
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . II. HATs and HAT Complexes: New Insights into HAT Regulation and Effects on Transcription . . . . . . . . . . . . . . . . . . III. HAT Complexes Functionally Interact with ChromatinRemodeling Complexes and Influence Transcription . . . . . . . IV. HAT Proteins Function in Regulating Transcriptional Elongation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. The Histone Code: Insights into Epigenetic Regulation of Transcription . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI. HATs: New Insight into Transcription and DNA Repair . . . . . VII. Conclusion and Perspectives. . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
181 182 187 192 194 195 196 196 197
Posttranslational Modifications of Histones by Methylation Adam Wood and Ali Shilatifard
I. II. III. IV.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lysine Methyltransferases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Arginine Methyltransferases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Epilogue . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
201 207 215 218 219
AUTHOR INDEX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
223 245
SUBJECT INDEX. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
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PREFACE Transcription is the first and often most critically regulated step in gene expression. In eukaryotes, multisubunit RNA polymerases transcribe genes, whose DNA is packaged together with histones and other nonhistone chromosomal proteins into chromatin, which is notoriously intractable and not easily transcribed by those polymerases in vitro. A central question driving research on eukaryotic transcription has therefore been, ‘‘How is timely and efficient transcription of chromatin-embedded genes achieved in eukaryotic cells?’’ Over the last 30 years, biochemical and genetic studies have revealed that eukaryotes have evolved remarkably sophisticated and unexpectedly complex means of regulating transcription of their genes. Studies teasing apart the components of the eukaryotic transcriptional apparatus through biochemical fractionation and genetic screens have led to the identification of nearly 100 proteins that appear to participate as ‘‘general’’ players in eukaryotic transcription. In addition, studies dissecting the transcriptional regulatory mechanisms operating on eukaryotic genes have revealed that these transcriptional proteins fall into multiple functional classes, including not only the multisubunit RNA polymerases that transcribe eukaryotic genes but also accessory transcription factors that support those polymerases and a diverse collection of enzymes that manipulate chromatin to make it more readily transcribed. These studies began with the discovery that eukaryotes express three distinct nuclear RNA polymerases that transcribe at least three distinct classes of genes. These polymerases are designated Pols I, II, and III and transcribe large ribosomal RNAs (rRNAs), messenger RNAs (mRNAs), and transfer and small ribosomal RNAs (tRNAs and 5S RNA), respectively. The development of methods for preparing crude cell extracts that recapitulate promoter-specific transcription in vitro by Pols I, II, and III led to the discovery that all three polymerases require multiple accessory transcription factors to initiate transcription on even the simplest, nonchromatin DNA templates. The ultimate purification of most of these ‘‘general’’ initiation factors for Pols I, II, and III has provided well-defined enzyme systems for dissection of eukaryotic transcriptional regulatory mechanisms and revealed further insights into the striking complexity of the eukaryotic transcriptional apparatus. Promoter-specific transcription by Pol II, for example, requires at minumun five general initiation factors designated TFIIB, TFIID, TFIIE, TFIIF, and TFIIH; with the exception of TFIIB, all are multisubunit ix
x
PREFACE
complexes ranging in size from two to more than 10 subunits. Pol II is also unique among eukaryotic RNA polymerases because of its requirement for a special coactivator, referred to as the Mediator, which appears to be indispensible for activation of Pol II transcription by DNA binding transcription factors. The Mediator is conserved from yeast to humans. Elegant biochemical studies have demonstrated that the Mediator is composed of more than 20 proteins and interacts with both DNA binding transcription factors and Pol II to transduce a variety of signals into specific transcriptional events in the nuclei of eukaryotic cells. In lines of research paralleling those exploring the structure and function of eukaryotic RNA polymerases and their associated transcription factors, investigations of the nature and regulation of chromatin structure have led to the discovery of multiple classes of novel enzymes, whose aggregate task is to remodel chromatin in anticipation of RNA polymerase and in ways that ensure timely and efficient transcription. These enzymes include multisubunit histone acetyltransferases that covalently modify nucleosomal histones to regulate their affinities for DNA; multisubunit ATP-dependent nucleosome remodeling complexes that reposition or entirely remove nucleosomes from DNA to provide RNA polymerase and its transcription factors unimpeded access to their DNA templates; and histone methyltransferases that have roles in transcription elongation and silencing. Finally, in addition to significant progress in biochemical and genetic studies of the mechanism and regulation of eukaryotic transcription, groundbreaking advances in biophysical studies of the eukaryotic transcriptional machinery are providing, for the first time, insights into the workings of the eukaryotic RNA polymerases and their associated transcription factors at atomic resolution. The recent reports of the highresolution x-ray structures of free and transcribing yeast Pol II and of yeast Pol II in association with one of its many transcription factors, TFIIS, most likely mark just the beginning of a structural revolution that promises to provide hitherto undreamed of insights into eukaryotic transcriptional regulation. This Advances in Protein Chemistry volume on Proteins in Eukaryotic Transcription seeks to provide an up-to-date account of the proteins and mechanisms of eukaryotic transcription and to illuminate the intimate cross-talk among eukaryotic RNA polymerases, their transcription factors, and the enzymes that expedite their journeys through chromatin. The first three chapters are devoted to Pol II and its associated transcription factors. Chapter 1 by Cramer describes the structure and function of Pol II, with special emphasis on the recent landmark high-resolution crystal resolutions of the enzyme. Chapter 2 provides an account by Bjorklund and
PREFACE
xi
Gustafsson of the structure and function of the multisubunit Mediator complex and its roles in basal and activated Pol II transcription. Chapter 3 by Matangkasombut, Auty, and Buratowski focuses on the structure and function of the multisubunit TFIID complex and the roles of its individual subunits in targeting Pol II to promoters. In Chapters 4 and 5, Moir and Willis describe recent developments in studies on the mechanism and regulation of transcription by Pol III, and Comai describes the Pol I transcriptional machinery, respectively. Chapters 6 through 8 are devoted to regulation of chromatin structure. In Chapter 6, Imbalzano and Xiao describe the structure and function of members of the family of ATPdependent chromatin remodeling enzymes. In Chapter 7, Torok and Grant provide an account of the growing family of histone acetyltransferases and their diverse roles in transcription and such other DNA transactions as recombination and repair. Finally, in Chapter 8, Wood and Shilatifard report on the burgeoning study of the roles of histone methyltransferases in eukaryotic transcription. Ronald C. Conaway Joan Weliky Conaway
STRUCTURE AND FUNCTION OF RNA POLYMERASE II By PATRICK CRAMER Institute of Biochemistry and Gene Center, University of Munich, 81377 Munich, Germany
I. Perspective . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . II. Structure of RNA Polymerase II . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . A. Overview of Structure Determinations. . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . B. Ten-Subunit Core Polymerase.. . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . C. Rpb4/7 Complex.. . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . D. Complete 12-Subunit Polymerase. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . E. Polymerase-TFIIS Complex . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . III. Function of RNA Polymerase II. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . A. Overview of the Transcription Cycle . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . B. Initiation Complex Assembly and Promoter DNA Loading . . . . . . . . . . . .. . . . . . . . C. Initiation-Elongation Transition . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . D. Elongation, Processivity, and Transcription Bubble Maintenance. . . . .. . . . . . . . E. Catalysis, Fidelity, Specificity, and Translocation. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . F. Backtracking, Pausing, Arrest, and Proofreading . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . G. Coupling to RNA Processing and Other Nuclear Events . . . . . . . . . . . . . . .. . . . . . . . H. Termination, Polymerase Recycling, Reinitiation, and Regulation. . . .. . . . . . . . IV. Comparison with Other Polymerases . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . A. Eukaryotic RNA Polymerases I and III. . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . B. Bacterial and Archaeal RNA Polymerases . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . C. Single-Subunit DNA and RNA Polymerases. . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . V. Conclusions . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . .
1 3 3 4 9 9 11 13 13 14 15 18 21 23 26 26 27 27 28 29 30 31
I. Perspective RNA polymerase II (Pol II) is the central enzyme that catalyses DNAdirected mRNA synthesis during the transcription of protein-coding genes. Pol II consists of a 10-subunit catalytic core, which alone is capable of elongating the RNA transcript, and a complex of two subunits, Rpb4/7, that is required for transcription initiation. Structures of individual Pol II subunits and subunit domains have been determined by nuclear magnetic resonance and X-ray analysis (Table I), and various forms and complexes of Pol II have been studied by electron microscopy (Asturias et al., 1997; Darst et al., 1991a; Jensen et al., 1998; Leuther et al., 1996). Here, however, I will concentrate on high-resolution structures of the 10-subunit Pol II core (Bushnell et al., 2002; Cramer et al., 2000; Cramer et al., 2001; Gnatt et al., 2001), an archaeal counterpart of Rpb4/7 (Todone et al., 2001), and x-ray crystallographic backbone models of the complete 12-subunit Pol II 1 ADVANCES IN PROTEIN CHEMISTRY, Vol. 67
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2
CRAMER
Table I High-Resolution Structural Studies of RNA Polymerase II Structure Rpb5
Organism
Method
Reference
1.9
1dzf
NMR*
—
1qk1
S. cerevisiae
NMR
—
1ald
Thermococcus celer
NMR
—
1qyp
Methanobacterium NMR thermoautotrophicum S. cerevisiae X-ray
—
1ef4
3.1
1i3q
S. cerevisiae
X-ray
2.8
1i50
S. cerevisiae
X-ray
3.3
1i6h
S. cerevisiae
X-ray
2.8
1k83
Bushnell et al., 2002
X-ray
1.75
1go3
Pol II
Methanococcus jannaschii S. cerevisiae
X-ray
4.2
1nt9
Pol II
S. cerevisiae
X-ray
4.1
1nik
Pol II-TFIIS complex
S. cerevisiae
X-ray
3.8
1pqv
Todone et al., 2001 Armache et al., 2003 Bushnell and Kornberg, 2003 Kettenberger et al., 2003
Rpb6
Saccharomyces cerevisiae Human
Rpb8 Rpb9 C-terminal domain Rpb 10 homolog Pol II core1 form 1 Pol II core1 form 2 Pol II core1 tailed-template elongation complex Pol II core1 -amanitin complex Rpb4/7 complex
X-ray
Resolution PDB* [A˚] code
Todone et al., 2000 del Rio-Portilla et al., 1999 Krapp et al., 1998 Wang et al., 1998 Mackereth et al., 2000 Cramer et al., 2000 Cramer et al., 2001 Gnatt et al., 2001
1 Pol II core comprises 10 subunits, Rpb1, Rpb2, Rpb3, Rpb4, Rpb5, Rpb6, Rpb7, Rpb8, Rpb9, Rpb10, Rpb11, Rpb12 and lacks the Rpb4/7 complex. *PDB: protein data bank; NMR: nuclear magnetic resonance.
(Armache et al., 2003; Bushnell and Kornberg, 2003), and of Pol II in complex with the elongation factor TFIIS (Kettenberger, et al., 2003) (Table I). These structures were published over the last 3 years and will be described in Chapter 2. Interpretation of the structures alongside biochemical and genetic data has provided valuable insights into many aspects of the transcription mechanism and will be discussed in Chapter 3. In Chapter 4, the conservation of the Pol II structure throughout species, its use as a model
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
3
for RNA polymerases I and III, and the consequences for understanding other polymerases are described.
II. Structure of RNA Polymerase II A. Overview of Structure Determinations Pol II is an asymmetric and large multiprotein complex with a total molecular weight of 0.5 MDa. High-resolution structural studies of Pol II by x-ray crystallography required large amounts of pure protein that cannot be obtained by overexpression because of the complexity of the enzyme. These difficulties have so far limited crystallographic studies of Pol II to the endogenous enzyme from Saccharomyces cerevisiae, which can be purified in milligram quantitites from yeast culture. Yeast Pol II preparations, however, contain substoichiometric amounts of the Rpb4/7 complex, giving rise to heterogeneity that impedes crystallization. This problem was overcome with the use of a rpb4 deletion strain of yeast. Purification from this strain yields the Pol II core, lacking both Rpb4 and Rpb7 (Darst et al., 1991b; Edwards et al., 1990). Initial studies of Pol II by electron microscopy (Asturias et al., 1997; Darst et al., 1991a; Jensen et al., 1998; Leuther et al., 1996) laid the ground for structural studies at high resolution, but several experimental difficulties had to be overcome first. Three-dimensional crystals were obtained (Fu et al., 1998, 1999) and were improved by induced crystal shrinkage (Cramer et al., 2000). Phase determination relied on heavy atom clusters (Cramer et al., 2000; Fu et al., 1999) and nonstandard heavy-metal compounds (Cramer et al., 2000). Interpretation of the experimental electron density maps was facilitated by placement of subunit structures that had been determined previously (Table I). Map interpretation also relied on phase combination and on the use of sequence markers (Cramer et al., 2000), including partially incorporated selenomethionine (Bushnell et al., 2001). These efforts first resulted in a backbone model of the Pol II core, which revealed the subunit architecture of the enzyme and functional elements (Cramer et al., 2000). Nucleic acids could also be placed on the Pol II backbone model (Cramer et al., 2000) with the use of electron microscopy data, which had earlier revealed the location of downstream DNA (Poglitsch et al., 1999). One year later, refined atomic structures of the Pol II core were reported in two crystal forms at 2.8- and 3.1-A˚ resolution (Cramer et al., 2001). The atomic core structures then enabled structure determination by molecular replacement of a minimal elongation complex of the yeast Pol II core (Gnatt et al., 1997; Gnatt et al.,
4
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2001) and a complex of the Pol II core with the mushroom toxin -amanitin (Bushnell et al., 2002). Lacking from the Pol II core structures was the Rpb4/7 complex. The structure of an archaeal counterpart of an isolated Rpb4/7 complex was, however, determined (Todone et al., 2001), and the location of the Rpb4/7 complex on the Pol II core surface was revealed by cryo-electron microscopy (Craighead et al., 2002). Recently, backbone models of the complete 12-subunit Pol II were derived by two groups independently, with the use of x-ray crystallographic data to around 4-A˚ resolution (Armache et al., 2003; Bushnell and Kornberg, 2003), and a model for the complex of the complete Pol II with the elongation factor TFIIS at 3.8-A˚ resolution was also reported (Kettenberger et al., 2003). The described structural studies of yeast Pol II are directly relevant for the Pol II enzymes in higher organisms, since the Pol II subunits are very well conserved in sequence and function. Approximately half of the amino acid residues in the twelve Pol II subunits are identical between yeast and human sequences. Furthermore, most yeast subunits can functionally replace their human counterparts (Woychik, 1998). The human Rpb4/7 complex can also functionally replace its yeast counterpart (Khazak et al., 1995), indicating that the core-Rpb4/7 interface is conserved.
B. Ten-Subunit Core Polymerase Five Pol II subunits, Rpb1, Rpb2, Rpb3, Rpb6, and Rpb11, show sequence and structural similarity in all cellular RNA polymerases and are referred to as the ‘‘core’’ subunits (Table II). One of the core subunits, Rpb6, and four other subunits, Rpb5, Rpb8, Rpb10, and Rpb12, are shared between the three eukaryotic RNA polymerases I, II, and III, and are referred to as the ‘‘common’’ subunits. The 10-subunit Pol II core comprises the core and common subunits and in addition, subunit Rpb9. The Pol II core structures show that the two large subunits, Rpb1 and Rpb2, form the central mass of the enzyme and opposite sides of a positively charged ‘‘cleft’’ that contains the active center (Fig. 1). The two large subunits are bridged on one side by a module of subunits Rpb3, Rpb10, Rpb11, and Rpb12. Around the periphery of the enzyme, Rpb5, Rpb6, and Rpb8 assemble with Rpb1, and Rpb9 binds to both Rpb1 and Rpb2. Subunits can be divided into domain-like regions, to aid interpretation of genetic and biochemical data and to facilitate the design of mutagenesis experiments (Cramer et al., 2001). Subunits Rpb1 and Rpb9 each bind two zinc ions, and subunits Rpb2, Rpb3, Rpb10, and Rpb12 each bind one zinc ion. All eight zinc ions are near the Pol II surface, apparently stabilizing the enzyme.
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STRUCTURE AND FUNCTION OF RNA POLYMERASE II
Table II RNA Polymerase Subunits Eukaryotes Bacteria
Class1
A0 + A00 B (B0 + B00 ) D L K
0 !
H — N P X F E þ1 other
— — — — — — — —
Core Core Core Core Core and common Common Common Common Common Unclear Rpb4/7 Rpb4/7 Specific Specific Specific
Pol I
Pol II
Pol III
Archaea
A190 A135 AC40 AC19 Rpb6
Rpb1 Rpb2 Rpb3 Rpb11 Rpb6
C160 C128 AC40 AC19 Rpb6
Rpb5 Rpb8 Rpb10 Rpb12 A12.2 A14 A43 A34.5 A49
Rpb5 Rpb8 Rpb10 Rpb12 Rpb9 Rpb4 Rpb7 —
Rpb5 Rpb8 Rpb10 Rpb12 C11 C17 C25 C82 C34 C31
1
Core: Sequence partially homologous in all RNA polymerases. Common: shared by all eukaryotic RNA polymerases, Rpb4/7: Rpb4/7 heterodimer and its structural counterparts. Unclear: It is unclear if A12.2 and C11 are true Rpb9 homologs. It appears that the C-terminal domain of the Pol II subunit C11 is functionally and structurally homologous to the Pol II transcript cleavage factor TFIIS.
Structural elements of Pol II have been given generic names if they appeared to be functionally relevant (Table III). The Rpb1 side of the cleft is formed by a mobile ‘‘clamp,’’ whereas the Rpb2 side consists of two domains, termed ‘‘lobe’’ and ‘‘protrusion.’’ The entrance to the cleft is formed between the ‘‘upper jaw’’ and the ‘‘lower jaw’’ of Pol II, which include subunits Rpb9 and Rpb5, respectively. The end of the cleft is blocked by a protein ‘‘wall.’’ The active center is formed by the floor of the cleft at its end and is located between the protrusion, the wall, and the clamp. Before the active center and opposite of the wall, a long ‘‘bridge’’ helix spans the cleft. The bridge partially lines a ‘‘pore’’ in the active center, which widens toward the other side of the enzyme, creating an inverted ‘‘funnel.’’ The rim of the pore also includes the highly conserved ‘‘aspartate loop’’ of Rpb1 that forms part of the active site. This loop comprises three invariant aspartate residues that stably bind a Mg2þ ion, termed ‘‘metal A.’’ The aspartate loop was identified as part of the active site by site-specific hydroxy radical cleavage (Zaychikov et al., 1996).
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Table III Structural Elements of RNA Polymerase II Pol II element and subelement
Subunit
Homology region
Rpb1 Rpb1 Rpb2 Rpb1
D D F F
Clamp
Rpb1 Rpb2 Rpb1 Rpb2 Rpb2, Rpb1 Rpb1 Rpb1
A, B, C, H
Rpb1
A
Rpb1 Rpb9
G
Switch 1, 2 Switch 3 Switch 4,5 Rudder Lid Zipper Jaws Rpb1/9 jaw (upper jaw)
H, C I I, H B
Catalysis (Catalysis) (NTP binding) Positioning of nascent base pair, stabilization of twist between bases in the template strand, maintenance of downstream end of the bubble, -amanitin-binding, (translocation) Processivity, template strand binding, hybrid retention, bubble maintenance, (initiation factor binding) Template strand binding, clamp mobility, processivity Template strand binding, processivity Clamp mobility Stabilization of the elongation complex (maintenance of upstream end of hybrid, creation of RNA exit tunnel) (maintenance of upstream edge of bubble) TFIIS binding, (interaction with downstream DNA during initiation and elongation)
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Active center Metal A site Metal B site Bridge
Function (proposed if in parenthesis)
Mobile modules Jaw-lobe Shelf Trigger loop Pocket Tip
Rpb5 Rpb2 Rpb2 Rpb1 Rpb2 Rpb1, Rpb6, Rpb11 Rpb1, Rpb8 Rpb1 Rpb1 Rpb2 Rpb2 Rpb1 Rpb2 Rpb1
Rpb1, Rpb2, Rpb9 Rpb1, Rpb5 Rpb1 Rpb1, Rpb2, Rpb6 Rpb7
Interaction with downstream DNA during elongation G, H
Hybrid binding, (maintenance of upstream end of the bubble) (initiation factor interaction, RNA exit tunnel formation), (RNA exit) (RNA exit)
I (Alternative RNA exit routes beyond the saddle) F, G F
C H
G Rpb1 H, Rpb2 I
-amanitin-binding, TFIIS binding, (NTP entry) TFIIS binding, (NTP entry, RNA exit during backtracking and arrest) TFIIS binding, crevice opening triggers conformational changes
(Maintenance of downstream end of bubble) (Initiation factor interactions) (Interaction with downstream DNA) CTD flexibility, Rpb7 binding modulation of Pol II activity throughout the transcription cycle, binding of Mediator and RNA processing factors
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
Rpb5 jaw (lower jaw) Wall (flap) Flap loop Saddle between wall and clamp RNA exit grooves 1, 2 Funnel Pore Crevice Fork Fork loop 1 Fork loop 2 Dock domain Lobe Linker CTD
TFIIS binding, (translocation) Binds the Rpb7 tip (allosteric regulation of clamp) Binds into the pocket below the clamp of the Pol II core
7
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A second metal ion, ‘‘metal B,’’ is weakly bound further in the pore, between the Rpb1 aspartate loop and one or two conserved acidic residues in Rpb2 (Cramer et al., 2001). Low occupancy of this second metal binding site indicates that the metal may be exchangeable. Both metal ions are accessible from one side. Two adjacent metal binding sites were also observed in a high-resolution structure of a bacterial RNA polymerase holoenzyme (Vassylyev et al., 2002). The clamp is a mobile domain that was suggested to retain nucleic acids in the cleft (Cramer et al., 2000; Fu et al., 1999). The clamp is trapped in two different open states in the free core structures (Cramer et al., 2001) but is rotated and closed in the structure of the core elongation complex (Gnatt et al., 2001). In the elongation complex structure, the clamp binds the DNA template strand before and within the DNA-RNA hybrid (Fig. 2A). Template strand binding involves three out of five ‘‘switch’’ regions. The switch regions form the base of the clamp that connects the clamp to the remainder of Pol II. On clamp closure, the switches change conformation or undergo folding transitions. The closed conformation of the clamp is also observed in electron microscopic images of the 12-subunit Pol II (Craighead et al., 2002). The clamp is formed by two regions in Rpb1, located at the N terminus and near the C terminus, and the C-terminal region of Rpb2. Three zinc ions stabilize the unique clamp fold. The C-terminal region of Rpb1 protrudes from the base of the clamp on the outside of Pol II and gets disordered after a few residues. These last ordered residues of Rpb1 constitute the beginning of a ‘‘linker’’ that connects to the C-terminal repeat domain (CTD) of Rpb1. The linker comprises about 100 and 150 residues in yeast and human, respectively, and is not conserved. The CTD is a unique feature of Pol II and consists of repeats of a heptapeptide with the consensus sequence Tyr-Ser-ProThr-Ser-Pro-Ser. A total of 26 and 52 CTD repeats are found in yeast and human Rpb1, respectively. The CTD and most of the linker are not ordered in the Pol II crystal structures. Nuclear magnetic resonance and circular dichroism studies of CTD peptides in solution revealed little residual structure (Cagas and Corden, 1995). If the linker and CTD would adopt a fully extended conformation, the C-terminus of Rpb1 could extend almost 1000 A˚ from the Pol II surface, about seven times the diameter of Pol II. Thus, the CTD could in principle reach anywhere on the Pol II surface. However, it is likely that the unphosphorylated CTD adopts a compacted state near the beginning of the linker on the Pol II surface (Cramer et al., 2001). A compacted weak protein density was detected near the Pol II core by electron microscopy (Meredith et al., 1996).
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
9
C. Rpb4/7 Complex The core structures lacked subunits Rpb4 and Rpb7, which form a stable heterodimer that can dissociate from the yeast Pol II core under mild denaturing conditions and on ion exchange chromatography (Edwards et al., 1991). Whereas Rpb7 is well conserved in sequence, Rpb4 shows only weak sequence conservation. Rpb7 is an essential protein (McKune et al., 1993). Rpb4 is not essential in S. cerevisiae (Woychik and Young, 1989), whereas it is required for viability of the fission yeast Saccharomyces pombe (Sakurai et al., 1999). Counterparts of the Rpb4/7 complex exist in the eukaryotic RNA polymerases Pol I and Pol III (Hu et al., 2002; Peyroche et al., 2002; Sadhale and Woychik, 1994; Shematorova and Shpakovski, 1999; Siaut et al., 2003) and in the archaeal enzymes (Werner et al., 2000). The structure of an archaeal Rpb4/7 counterpart revealed that Rpb7 spans an elongated complex and is organized in two domains, an N-terminal ribonucleoprotein (RNP)-like domain and a C-terminal domain that includes an oligonucleotide/oligosaccharide-binding fold (Todone et al., 2001). The Rpb4 homolog binds at the connection between the two Rpb7 domains and forms a conserved hydrophobic interface with the Rpb7 homolog. Conservation of the interface is demonstrated by the formation of chimeric heterodimers with Rpb4 and Rpb7 from various species (Guilfoyle and Larkin, 1998; Sakurai et al., 1999; Werner et al., 2000). Mutagenesis and surface conservation indicate a potential nucleic acid binding face of the Rpb4/7 complex that could account for binding of single-stranded nucleic acids in vitro (Orlicky et al., 2001; Todone et al., 2001). Cryo-electron microscopy of the 12-subunit yeast Pol II revealed an additional density on the outside of the core that was interpreted as the Rpb4/7 complex (Craighead et al., 2002). This density coincides with a stalk of protein protruding from the core of Pol I in electron microscopy images (Bischler et al., 2002). With the use of immunolabeling, the stalk in Pol I was shown to contain counterparts of Rpb4 and Rpb7 (Bischler et al., 2002). In the electron microscopic reconstructions, most of the Rpb4/7 surface appears to be exposed and easily accessible for interactions with other proteins or nucleic acids. Electron microscopy in solution further revealed that the clamp adopts a closed state in the 12-subunit Pol II that includes the Rpb4/7 complex (Craighead et al., 2002).
D. Complete 12-Subunit Polymerase The above findings and proposals about the location and function of the Rpb4/7 complex were generally confirmed and extended by recent crystallographic backbone models of the complete Pol II that includes the
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Rpb4/7 complex. These models were derived independently by two groups (Armache et al., 2003; Bushnell and Kornberg, 2003) and show that Rpb4/7 protrudes from the polymerase surface near the base of the clamp (Fig. 1). The Rpb4/7 complex interacts with the Pol II core through Rpb7, which binds to regions of Rpb1, Rpb2, and Rpb6. Most of the Rpb4/7 surface is exposed and accessible for interactions with proteins or nucleic
Fig. 1. Two views of the complete yeast Pol II (Armache et al., 2003). The 12 protein subunits are shown as ribbon diagrams in different colors, as indicated in the schematic diagram. The active site metal ion A is depicted as a pink sphere. Zinc ions are shown as cyan spheres. A highly similar model was reported by Bushnell and Kornberg, 2003. CTD, C-terminal repeat domain. (See Color Insert.)
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
11
acids. Rpb4/7 binds to the Pol II core with the N-terminal RNP-like domain of Rpb7: termed the tip. Consistent with the core-Rpb7 interaction, Rpb7 alone can bind to core (Sheffer et al., 1999), and Rpb7 is essential for yeast growth (McKune et al., 1993), whereas Rpb4 is not (Woychik and Young, 1989). Deletion of the rpb4 gene in yeast facilitates dissociation of Rpb7 from core (Edwards et al., 1991). The models indicate that loss of the Rpb4-Rpb7 interface on Rpb4 deletion destabilizes Rpb7 and facilitates Rpb7 dissociation. The Rpb4/7 complex forms a wedge between the clamp and the linker, apparently restricting the clamp to a closed position. In particular, the Rpb7 tip partially fills a surface ‘‘pocket’’ formed between the clamp, the linker, and the core subunit Rpb6. The pocket is lined by five protein regions: three in Rpb1 and one each in Rpb2 and Rpb6. Rpb4/7 binding to the pocket thus holds together three subunits and may stabilize the Pol II subunit assembly.
E. Polymerase-TFIIS Complex Very recently, a backbone model for the complex of the complete Pol II with the elongation factor TFIIS (or SII) was reported at 3.8-A˚ resolution (Kettenberger et al., 2003). To obtain this structure, recombinant TFIIS comprising domains II and III of the three-domain factor was soaked into harvested crystals of the complete Pol II. Successful protein soaking was enabled by the very large solvent channels of the crystals and the fact that the TFIIS-binding site on Pol II is not obstructed by crystal contacts. The resulting 13-polypeptide asymmetric complex has a molecular weight of 536 kDa. The crystal lattice accommodated extensive structural changes induced by TFIIS around the active site of Pol II and in the periphery of the enzyme. The structure shows that TFIIS extends along the Pol II surface, spanning a distance of 100 A˚ (Fig. 2B). TFIIS domain II docks to the exposed Rpb1 jaw domain of Pol II. The TFIIS interdomain linker extends from domain II along the Pol II surface into the funnel. Domain III inserts into the Pol II pore, and approaches the polymerase active site from the bottom face of the enzyme as predicted (Cramer et al., 2000). TFIIS domain III reaches the Pol II active site with the highly conserved loop of the protruding hairpin. The domain II hairpin complements the polymerase active site with acidic groups that are essential for TFIIS function. Two invariant acidic residues in this loop, D290 and E291, are in close proximity of the Pol II catalytic metal ion A and are essential for TFIIS activity ( Jeon et al., 1994).
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Fig. 2. Structure of the Pol II core elongation complex and the Pol II-TFIIS complex. (A) Schematic cut-away view of the tailed-template yeast Pol II core complex (Gnatt et al., 2001). The view is related to the one on the bottom of Figure 1 by a 90-degrees rotation around a vertical axis. The DNA template and nontemplate strands are shown in blue and green, respectively, and the RNA in red. Four bases in the template strand are highlighted as sticks protruding from the DNA backbone. The yellow oval indicates the presumed location of the binding site for the incoming NTP. During polymerization, Pol II moves to the right. (B) Backbone model of the complete 12-subunit Pol II (grey) in complex with the elongation factor TFIIS (orange, Kettenberger et al., 2003). Parts of Pol II are omitted for clarity. DNA and RNA have been modeled according to the structure in (A). During backtracking, Pol II moves to the left. (See Color Insert.)
In addition to the active site complementation, TFIIS induces structural changes in the Pol II active center. Binding of TFIIS domain III induces folding of the Rpb1 ‘‘trigger loop’’ (Vassylyev et al., 2002) and shifts the bridge helix. These changes probably result in a repositioning of nucleic
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
13
acids in the active center. TFIIS further induces a coordinated repositioning of about one-third of the polymerase mass, which includes the jaws, the clamp, and the Rpb1 cleft and foot domains and corresponds essentially to three mobile polymerase modules (Cramer et al., 2001). The repositioning seems to be caused by insertion of TFIIS into the Pol II funnel and pore, where it opens an additional crevice.
III. Function of RNA Polymerase II A. Overview of the Transcription Cycle The transcription cycle may be divided in three major phases: initiation, elongation, and termination. Steps during transcription initiation include promoter DNA binding, DNA melting, and initial synthesis of short RNA transcripts. The transition from initiation to elongation, referred to as ‘‘promoter escape,’’ also occurs in a stepwise fashion. Promoter escape leads to a stable elongation complex that is characterized by an open DNA region, the ‘‘transcription bubble.’’ The incoming and exiting DNA duplex, located before and after the bubble, respectively, is referred to as downstream and upstream DNA. The bubble contains the DNA-RNA hybrid, a heteroduplex of eight to nine base pairs. At one end of the hybrid, the growing RNA 30 -end is engaged with the active site. At the other end of the hybrid, the DNA and RNA strands are separated. After successful RNA chain elongation, transcription terminates and Pol II dissocitates from the template. Some of the steps during the transcription cycle can be carried out by Pol II alone. Pol II can maintain an open transcription bubble, translocate along the template DNA, synthesize RNA from the template, and proofread the nascent RNA. For all other steps during the transcription cycle, however, Pol II requires additional proteins. Several steps of the transcription cycle are accompanied by phosphorylation or dephosphorylation of the Pol II CTD (Dahmus, 1996; O’Brien et al., 1994). During initiation, the CTD gets phosphorylated and the CTD phosphorylation pattern changes during elongation. CTD phosphorylation patterns govern specific interaction with RNA processing factors, thereby coupling transcription to RNA maturation events. Recycling of Pol II after termination involves CTD dephosphorylation, as initiation requires unphosphorylated Pol II. Proteins involved in phosphorylation and dephosphorylation of Pol II and other regulatory proteins influence the transcription cycle at various steps. For each step in the transcription cycle, insights coming from the Pol II structures are discussed below. At several points, supporting biochemical data are included that were
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obtained with bacterial RNA polymerase, for which a large amount of mechanistic information has accumulated (von Hippel, 1998). As discussed in Chapter 4, bacterial and eukaryotic RNA polymerases show a conserved core and share many functional features.
B. Initiation Complex Assembly and Promoter DNA Loading To bind and melt promoter DNA, Pol II requires the general transcription factors TFIIB, TFIID, TFIIE, TFIIF, and TFIIH (Buratowski, 1994; Buratowski et al., 1989; Kornberg, 1999), which in yeast consist of one, 14, two, three, and nine polypeptides, respectively. The general transcription factors assemble with Pol II on promoter DNA and are involved in sequence-specific promoter recognition (TFIIB, TFIID), prevention of nonspecific DNA binding (TFIIF), DNA melting (TFIIE, TFIIH), and phosphorylation of the CTD (TFIIE, TFIIH). Many Pol II promoters contain a TATA box about 25–30 base pairs upstream of the transcription start site. TFIID binds to the TATA box via its TATA box–binding protein (TBP) subunit. According to order-of-addition experiments, stepwise assembly of the initiation complex starts with the formation of a TFIID/TBP-DNA complex, followed by binding of TFIIB to TBP and to a promoter element adjacent to the TATA box, the TFIIB response element BRE (Buratowski et al., 1989; Lagrange et al., 1998). Assembly of TBP and TFIIB on TATA box DNA has been studied biochemically and structurally (Cox et al., 1997; Kim et al., 1993; Kim et al., 1993; Kosa et al., 1997; Littlefield et al., 1999; Nikolov et al., 1995; O’Brien et al., 1998; Sigler and Tsai, 2000). In addition to TBP and TFIIB, loading of promoter DNA onto Pol II minimally requires TFIIF (Killeen et al., 1992), which forms a stable complex with Pol II. An additional factor, TFIIA, can stabilize the TFIID-DNA complex (Pugh, 2000). Other core promoter elements are known, including the initiator element (Smale et al., 1998) and the downstream promoter element (Burke et al., 1998). Depending on the specific promoter structure, there are apparently various routes to the initiation complex. As an alternative to the stepwise assembly of the initiation complex, it has been suggested that a large Pol II ‘‘holoenzyme’’ can be recruited to a promoter in a single step. Such holoenzymes were purified from yeast (Koleske and Young, 1994) and mammalian cells (Ossipow et al., 1995) and comprise Pol II, general transcription factors, and various other proteins (Greenblatt, 1997; Myer and Young, 1998). The position of the general transcription factors with respect to promoter DNA in the initiation complex can be inferred from site-specific protein–DNA crosslinking (Ebright, 1998). The crosslinking data, taken
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
15
together with topological considerations and with structural data, predict that TBP, TFIIB, and TFIIF interact with the ‘‘upstream face’’ of Pol II. The upstream face of the enzyme includes parts of the Rpb4/7 complex, parts of the clamp, the outside of the wall, the ‘‘saddle’’ between the clamp and the wall, and the ‘‘dock’’ domain of the largest Pol II subunit. Biochemical data indicate that Rpb4/7 stabilizes a minimal initiation complex ( Jensen et al., 1998), suggesting that Rpb4/7 interacts with one or more general transcription factors. There is evidence that TFIIB binds adjacent to the Rpb4/7 complex, because Rpb4/7 binds near Rpb6 (Bischler et al., 2002; Craighead et al., 2002) and the archaeal homolog of TFIIB binds the archaeal Rpb6 homolog (Magill et al., 2001). Initiation factors interact with counterparts of the Rpb4/7 complex in the two other eukaryotic RNA polymerases, Pol I and Pol III. In Pol III, the Rpb4 homolog binds to a region corresponding to the linker in Pol II (Siaut et al., 2003) and to the TFIIB-related initiation factor Brf1 (Ferri et al., 2000). The Rpb7 homolog of Pol I also binds an initiation factor, called Rrn3/TIF-IA (Peyroche et al., 2000; Yuan et al., 2002). Thus Rpb4/7 and its counterparts seem to bridge the polymerase core with initiation factors. Differences between Rpb4/7 and its counterparts in other polymerases may contribute to promoter specificity. One function of TBP, TFIIB, and TFIIF is apparently to bring the promoter DNA duplex to a location on the Pol II surface that is appropriate for DNA melting and initiation of RNA synthesis at the transcription start site. There are two prominent possible locations of the initially loaded promoter DNA duplex. The promoter duplex may initially bind above the cleft on the enzyme surface. Alternatively, promoter DNA may be bound inside the Pol II cleft, closer to the active site. The structure of the free Pol II core showed that dramatic opening of the clamp can create sufficient space to allow for loading of duplex DNA into the Pol II cleft (Cramer et al., 2001). However, the Rpb4/7 complex acts as a wedge that prevents entry of the promoter DNA duplex into the active center cleft (Armache et al., 2003; Bushnell and Kornberg, 2003). Because the Rpb4/7 complex is apparently not dissociating rapidly in all species, it is likely that the promoter DNA duplex initially binds outside the cleft far above the active center.
C. Initiation-Elongation Transition After loading of promoter DNA onto Pol II, duplex DNA is melted upstream of the transcription start site (Holstege et al., 1997; Pan and Greenblatt, 1994; Wang et al., 1992). DNA melting requires TFIIH, which
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comprises two ATP-dependent helicase activities that unwind DNA. Two alternative models for DNA opening have been proposed (Fiedler and Marc Timmers, 2000). Crosslinking data indicate that TFIIH interacts with the downstream DNA and acts from a distance (Kim et al., 2000). TFIIE bridges between Pol II and TFIIH and stimulates TFIIH activity (Maxon et al., 1994; Ohkuma, 1997). In Pol I, the initial melted DNA region is about nine base pairs long, and the mature transcription bubble extends over approximately 19 base pairs (Kahl et al., 2000). After RNA synthesis has initiated within the bubble, the bubble size remains flexible during early transcription (Fiedler and Timmers, 2001). If DNA would be melted in the cleft, the DNA nontemplate strand must be expelled from the cleft before the clamp can close. It is, however, possible that DNA is loaded on top of the cleft and remains above the cleft for melting. In this case, the template strand could pass the clamp after DNA melting; it could slip into the cleft and bind to the site formed by switch regions 1–3, as observed in the core elongation complex (Gnatt et al., 2001). Until the nascent transcript is about 15 nucleotides long, the early transcribing complex is functionally unstable. In some cases the transcript can even slip upstream along the DNA template by several bases and can be reextended (Luse and Pal, 2002). Early transcribing Pol II complexes have to undergo three transitions (Dvir, 2002). In the beginning, short RNAs are frequently released and Pol II has to restart transcription (‘‘abortive cycling’’). There is a decline in the level of abortive transcription when the RNA reaches a length of about four nucleotides, and this transition is termed ‘‘escape commitment’’ (Goodrich and Kugel, 2000, 2002). A second barrier has to be overcome when the RNA reaches a length of about 10 nucleotides. A third transition is reflected in the continued requirement for the ATP cofactor and TFIIH until the RNA is about 15 nucleotides long. Successful passage of early Pol II elongation complexes through all three transitions has been referred to as ‘‘promoter clearance.’’ The early initiation-elongation transitions limit the rate of Pol II transcription and can be enhanced by TFIIE, TFIIH, and ATP (Goodrich and Kugel, 1998). Transitions that underlie promoter clearance may be rationalized with the Pol II structures. At the very beginning of transcription, contacts of Pol II with nascent RNA are crucial. To allow for the synthesis of the first phosphodiester bond, nucleoside triphosphates must be held by the protein. The resulting dinucleotide RNA must still be held by protein– RNA contacts, as observed in the core elongation complex structure (Gnatt et al., 2001), as the energy of base-pairing alone is insufficient for its retention. Equally, RNA is still bound by Pol II at the position of the third nucleotide. Despite the observed RNA-Pol II contacts, short RNA
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
17
dinucleotides and trinucleotides are often lost, RNA synthesis must restart, and repetitive RNA loss and transcription initiation results in abortive cycling. RNA that has grown to a length of at least four nucleotides is generally not contacted by Pol II any more and is apparently held in the elongation complex solely by base pairing with the DNA template strand. This change in RNA interactions reflects the first transition in stability of the early transcribing complex that occurs at a transcript length of four residues, beyond which the RNA is generally retained. Maybe the Pol II–RNA contacts are limited to the crucial contacts of the first few nucleotides, to facilitate RNA mobility and translocation of nucleic acids. In the bacterial RNA polymerase, a portion of the initiation factor apparently interferes with the path of the early transcript, inducing abortive cycling (Murakami et al., 2002). It is possible that in the Pol II system, one of the general transcription factors is located similarly and plays a similar role. The Pol II structures also provide an explanation for the second transition in stability of the transcribing complex that occurs at an RNA length of around 10 nucleotides. The 50 -end of a 10-residue RNA would be located just beyond the DNA-RNA hybrid, after its removal from the DNA template strand. At this point, the RNA is apparently redirected to the Pol II ‘‘saddle’’ and an exit tunnel (compare section D). Threading of RNA into the exit tunnel and binding of RNA to its exit groove may underlie the second transition in elongation complex stability. The third transition, which occurs when the RNA is about 15 nucleotides long, may reflect successful positioning of all bubble-maintaining structural elements of Pol II with respect to the bubble and detachment of RNA from the Pol II surface. Two possible RNA exit grooves have been suggested beyond the saddle, and binding of RNA to the saddle and to one of the exit grooves could account for an additional gain in stability of the elongation complex. In addition, RNA may bind to a nearby potential nucleic-acid binding face of the Rpb4/7 complex. The described transitions may involve reshaping of protein–nucleic acid contacts and may require slight changes in the clamp position. Because downstream DNA contributes to the stability of early transcribing complexes (Wang et al., 2003), it is likely that the downstream DNA contacts Pol II during initiation and promoter escape. A candidate subunit for such interaction is Rpb9, as it is located at an appropriate position and as mutations in Rpb9 lead to changes in the position of the transcription start site (Furter-Graves et al., 1994; Hull et al., 1995). Indeed, a domain in the bacterial enzyme at an approximately corresponding location contacts downstream DNA (Ederth et al., 2002). After successful promoter clearance, the early elongation complex can pause in a promoter proximal position (Albert et al., 1997; Li et al., 1996; Raschke et al., 1999). This promoter-proximal pausing of polymerase
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provides a means of rapid response to stimulatory signals. Exonuclease III footprinting indicates another transition from initiation to elongation that occurs around 25 bases downstream of the transcription start site (Luse and Samkurashvili, 1998). In other studies it was found that the transition to full elongation competence is dependent on the synthesis of even longer RNAs of a length of 50 nucleotides (Ujvari et al., 2002). This late transition is reversible by shortening the nascent RNA. The structural basis of these transitions is unclear. The production of a fully competent elongation complex is referred to as ‘‘promoter escape.’’ In addition to the structural changes described, the initiationelongation transition involves phosphorylation of the Pol II CTD. Elongationally competent polymerases show a phosphorylated CTD (Cadena and Dahmus, 1987; O’Brien et al., 1994) that adopts a far more extended structure than the unphosphorylated CTD (Corden and Zhang, 1991). There is a temporal relationship between CTD phosphorylation and the progression of Pol II through the transcription cycle (Dahmus, 1996). Both initiation and elongation are regulated by phosphorylation/dephosphorylation events (Greenblatt and Kobor, 2002). Several kinases (Prelich, 2002) and at least one phosphatase, Fcp1 (Kobor et al., 1999), control the phosphorylation state of the Pol II CTD. In addition, several general transcription factors and elongation factors are phosphoproteins (Greenblatt and Kobor, 2002). Five out of the seven amino acids in the CTD consensus repeat may in principle be phosphorylated. During initiation, the CTD is phosphorylated mainly at serine 5, a reaction catalyzed by the kinase Cdk7/Kin28 within TFIIH. Serine 5 phosphorylation is detected primarily at promoter regions and serine 2 is phosphorylated in coding regions (Komarnitsky et al., 2000), indicating that a change in the phosphorylation pattern accompanies the transition from initiation to elongation. This change apparently plays a role in the first RNA processing event, 50 -RNA capping (cf. section III.G). Substitution of serines 2 or 5 to alanine is lethal in yeast (Corden and West, 1995). In addition, changing tyrosine 1 to phenylalanine is lethal (Corden and West, 1995), indicating that tyrosine 1 is also a target for phosphorylation. Indeed, tyrosine 1 is phosphorylated in mammalian cells by the Abl kinase (Baskaran et al., 1997). The significance of this CTD modification is, however, unclear. The CTD is also a target for the modulation by peptidyl prolyl isomerases that catalyzes isomerization of prolines (Hunter, 1998; Shaw, 2002).
D. Elongation, Processivity, and Transcription Bubble Maintenance A functional model of the elongation complex was derived for bacterial RNA polymerase from biochemical data (Korzheva et al., 1998; Nudler, 1999). X-ray crystallographic data and site-specific protein–nucleic acid
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crosslinking provided a three-dimensional model of the bacterial RNA polymerase elongation complex (Korzheva et al., 2000). The location of nucleic acids in the Pol II elongation complex was modeled on the basis of electron crystallographic analysis (Cramer et al., 2000; Poglitsch et al., 1999) and site-specific polymerase–nucleic acid crosslinking data (Burgess and Wooddell, 2000). The later X-ray structure of a Pol II elongation complex allowed direct observation of the course of the nucleic acid strands in the DNA-RNA hybrid and of the DNA template strand just before the hybrid (Gnatt et al., 2001). The crystallized complex was formed by transcription of a DNA with a single-strand extension, a ‘‘tailed template,’’ in the presence of only three nucleoside triphosphates, leading to pausing at a discrete site (Gnatt, 2002; Gnatt et al., 1997; Gnatt et al., 2001). From these studies has emerged the following view of the elongation complex: Downstream DNA enters Pol II near two mobile ‘‘jaws’’ and extends through the cleft toward the active site. Beyond the active site, the DNA-RNA hybrid extends upward, toward the wall. The axis of the downstream DNA duplex and the DNA-RNA hybrid heteroduplex enclose an angle of almost 90 degrees. The growing RNA 30 -end is located above the pore, which allows entry of nucleoside triphosphates from below during RNA synthesis. In the crystal structure of the Pol II elongation complex, the incoming DNA duplex is mobile and badly ordered. However, three nucleotides before the active site, the DNA template strand becomes well ordered by binding to the bridge helix and to two ‘‘switch’’ regions at the base of the clamp, switches 1 and 2. A 90-degree twist between subsequent nucleotides orients a DNA base toward the active site for base pairing with an incoming RNA nucleotide. This base pair is the first of nine base pairs of the DNA-RNA hybrid that emanate from the active site. The hybrid length agrees with the length observed biochemically (Kireeva et al., 2000; Nudler et al., 1997). The DNA template strand within the hybrid is partly bound by switch region 3. The DNA nontemplate strand is disordered in the Pol II core elongation complex structure, maybe because the complex lacks the upstream DNA duplex and a complete bubble. The location of the nontemplate strand and the upstream DNA duplex during Pol II elongation is still unclear and may change during transcription. The property of the polymerase to stay attached to the template, even during transcription of long genes, is often referred to as processivity. The major cause of processivity is believed to be the high stability of the Pol II elongation complex. Elongation complex stability is caused by tight binding of the DNA-RNA hybrid to RNA polymerase (Kireeva et al., 2000; Sidorenkov et al., 1998). This stability can be accounted for by a highly complementary hybrid-binding site, in which the hybrid is imbedded. Enclosure of the hybrid results in protection of the RNA from digestion
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by RNAses (Komissarova and Kashlev, 1998). The complementary hybridbinding site is partially created upon clamp closure and folding of switches 1–3, which interact with the DNA template strand. Overextended hybrids have a negative effect on elongation complex stability (Kireeva et al., 2000). Interaction of the hybrid with its binding site ensures that the stability of the elongation complex, and thus processive transcription, is coupled with the presence of RNA. Because the Pol II core alone is sufficient to maintain the transcription bubble and the DNA-RNA hybrid during RNA chain elongation, there must be exposed elements on the enzyme surface that keep the nucleic acid strands apart. Protein elements are needed to separate the DNA strands downstream of the active site and to separate the RNA from the DNA template strand at the upstream end of the hybrid. On the basis of their location with respect to nucleic acids, several Pol II structural elements are predicted to maintain the bubble and the hybrid. These proposals are currently tested by site-directed mutagenesis. Separation of the DNA strands at the downstream edge of the bubble may be attributed to binding of the DNA template strand by switch regions 1 and 2 and to blocking of the path of the nontemplate strand by ‘‘fork loop 2.’’ In the Pol II-TFIIS complex structure, fork loop 2 is ordered and restricts the cleft to a diameter of 15 A˚, consistent with the proposal that this loop removes the DNA nontemplate strand from the template strand before the active site. Maintenance of the upstream end of the hybrid and the bubble may involve three loops protruding from the edge of the clamp into the cleft. The two lower loops, called ‘‘rudder’’ and ‘‘lid,’’ are close to the upstream end of the hybrid. Mutagenesis of the rudder in bacterial RNA polymerase showed that this element stabilizes the elongation complex but that it is not involved in maintaining the hybrid length (Kuznedelov et al., 2002). The lid may be involved in separating RNA from DNA at the upstream end of the hybrid. The upper loop, called the ‘‘zipper,’’ could help maintain the upstream end of the transcription bubble. All three loops show some mobility and are present in all cellular RNA polymerases. The lid in bacterial polymerase interacts with the factor (Murakami et al., 2002; Vassylyev, 2002), indicating that the lid in Pol II could contact a general transcription factor. The lid approaches another loop that protrudes from the opposite side, from the top of the wall (‘‘flap loop’’). The saddle, lip, and flap loop create a putative RNA exit tunnel. The flap loop in bacterial RNA polymerase binds to nascent RNA hairpins that pause or terminate transcription (Landick, 2001; Toulokhonov et al., 2001).
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E. Catalysis, Fidelity, Specificity, and Translocation The location of metal ions A and B is generally consistent with the geometry of substrate binding observed in the Pol II core elongation complex structure and in x-ray structures of nucleic acid complexes of single-subunit DNA polymerases (Doublie et al., 1998; Franklin et al., 2001; Pelletier et al., 1994; Sawaya et al., 1997), although the metal ions in the free enzyme may not be observed at the exact same location where they would be found during catalysis. On this basis, a working model for the nucleotide addition cycle during RNA chain elongation by Pol II was suggested. According to this model, the cycle starts with entry of the nucleoside triphosphate (NTP) substrate together with metal B, and its binding between the bridge helix and the end of the hybrid, to form a base pair with the ‘‘coding’’ DNA base. The NTP binding site of Pol II has not been defined, but it can be inferred from the site observed in structures of single-subunit DNA polymerases. Correct orientation of the substrates and metal ions would lead to synthesis of a new phosphodiester bond and to release of pyrophosphate, maybe together with metal ion B. The resulting complex adopts the pretranslocation state, which was apparently trapped in the core Pol II elongation complex structure, with the RNA 30 -terminal nucleotide occupying the NTP binding site (Gnatt et al., 2001). Subsequent translocation of nucleic acids would align the new RNA 30 end with metal A and would free the NTP binding site, preparing Pol II for another cycle of nucleotide addition. Fidelity of transcription may be defined as the property of Pol II that generally ensures incorporation of the correct nucleotide complementary to the base in the template strand. Fidelity must rely on correct positioning of the incoming NTP to optimize Watson–Crick base pairing between the NTP and the coding base in the DNA template strand, which together form the nascent base pair. Understanding the mechanistic basis for Pol II fidelity would require a structure of Pol II with bound DNA, RNA, and incoming NTP, which is currently not available. However, it is likely that fidelity relies in part on binding and positioning of the nascent base pair from the minor groove side, as observed in single-subunit polymerases (Chapter IV,C). Another important property of Pol II is its specificity for RNA synthesis rather than DNA synthesis. Specificity for synthesizing RNA may be achieved by at least three mechanisms. First, the discriminating 20 -OH group of the incoming NTP may be hydrogen-bonded by a conserved Pol II residue (Cramer et al., 2001; Gnatt et al., 2001). Second, 20 -OH groups of the last few nucleotides that were incorporated into the growing RNA are
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directly hydrogen-bonded by Pol II residues, such that accidentally incorporated deoxyribonucleotides would destabilize the elongation complex, resulting in a proofreading reaction (see following). Finally, the active center of Pol II is complementary to the resulting DNA-RNA hybrid duplex that adopts a specific conformation intermediary between canonical A-forms and B-forms. DNA synthesis would lead to canonical B-form DNA that would not fit into the hybrid binding site. Pol II apparently binds DNA and RNA tightly to create a stable and processive elongation complex. At the same time, Pol II allows for precise translocation of nucleic acids over its surface and moves along the DNA template with a considerable speed of several hundred nucleotides per minute. The question of how rapid translocation and tight nucleic acid binding can be achieved at the same time is a central mystery of the Pol II mechanism and of the mechanism of other nucleic acid metabolizing enzymes. Hints for understanding translocation are provided by the Pol II structures. First, nucleic acids are only contacted via their backbones, and base interactions that would impede translocation are not observed. Second, there are many positively charged protein groups that form a ‘‘second shell’’ around the nucleic acids, at a distance of up to 8 A˚ from the nucleic acid backbones. Such long-range electrostatic interactions may enable tight binding of nucleic acids without restricting their movement. Finally, translocation may be accompanied by conformational changes in Pol II regions around the nucleic acids. Such conformational changes could maintain some of the protein–nucleic acid contacts, resulting in a lowering of the energy barrier between pretranslation and posttranslocation states. One such conformational change may be bending of the bridge helix, as observed indirectly by a comparison of structures of Pol II and the bacterial RNA polymerase (Cramer et al., 2001; Darst, 2001; Gnatt et al., 2001). A corresponding ‘‘O-helix’’ in singlesubunit polymerases also stacks against the template-product nucleic acid duplex and can also change its conformation (Li et al., 1998). A highresolution structure of a bacterial RNA polymerase holoenzyme revealed a ‘‘trigger’’ loop that may cooperate with the bridge helix (Vassylyev et al., 2002). Indeed, a corresponding trigger loop in Pol II is mobile but becomes ordered on TFIIS binding (Kettenberger et al., 2003). In addition to the bridge helix, conformational changes in other Pol II structural elements may accompany translocation of nucleic acids, such as relative movements of mobile modules that surround incoming DNA (Cramer et al., 2001). Pol II elongation may be inhibited by binding of the cyclic octapeptide -amanitin, the toxin of the ‘‘death cap’’ mushroom. -amanitin does not greatly influence NTP binding (Chafin et al., 1995), and a phosphodiester
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bond can still be formed when the toxin is added to an elongation complex (Gu et al., 1993). However, the rate of transcription is dramatically reduced, such that only several nucleotides are incorporated per minute. These biochemical observations are consistent with the structure of a Pol II core--amanitin complex. In this structure, -amanitin is seen binding to the bridge helix from below. Thus, -amanitin cannot interfere with access of nucleic acids to the cleft or with entry of NTP substrates through the pore. Instead, -amanitin could possibly restrain the bridge helix movement and thereby block conformational changes that are important for translocation. However, given the speculative nature of the conformational change of the bridge helix during translocation, understanding the exact mechanism of Pol II inhibition by -amanitin requires further study. The binding sites for -amanitin and domain III of TFIIS overlap, explaining why the toxin interferes with TFIIS activity (Izban and Luse, 1992; Weilbaecher et al., 2003).
F. Backtracking, Pausing, Arrest, and Proofreading Pol II does not move along the DNA template in a unidirectional manner. The polymerase, rather, oscillates between forward and backward movements. Reverse movement of Pol II along DNA and RNA is referred to as ‘‘backtracking.’’ As a result of backtracking, RNA polymerase elongation complexes can adopt different conformational states (Erie, 2002). Oscillation back and forth along DNA and RNA was demonstrated for bacterial RNA polymerase (Kashlev and Komissarova, 1997a). Oscillatory movement of the polymerase can explain DNA and RNA footprints that are irregular in length. Shorter and longer footprints are apparently reflections of mixed populations of the elongation complex in productive and backtracked states. Before the concept of oscillatory movement, the irregular footprints were interpreted as an ‘‘inchworming’’ motion of the polymerase, with the enzyme contracting and expanding along the template. Inchworming requires independent movement of two flexibly linked parts of Pol II and is, thus, inconsistant with the Pol II structure. During backtracking, a Pol II structural element must keep the two DNA strands at the upstream end of the bubble separated, but it is unclear which Pol II element this is. The bridge helix apparently removes the RNA 30 -end from the DNA template strand during backtracking. The backtracked RNA is apparently extruded through the pore into the funnel. Backtracking of Pol II during the elongation phase can lead to transcriptional pausing and arrest. Pausing and arrest are blocks to transcription that
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can be signaled intrinsically, by certain DNA sequences, or extrinsically, through additional protein factors (Uptain et al., 1997). Pausing is defined as a temporary block to elongation, from which Pol II can escape by itself, without the need for accessory factors. Pol II that has paused at certain DNA sites has generally backtracked by several nucleotides. The DNA-RNA hybrid normally prevents backtracking of Pol II and maintains the register of transcription (Nudler et al., 1997). However, destabilization of the hybrid at specific DNA sites leads to backtracking (Nudler et al., 1997) and appears to be the primary determinant for pausing (Landick and Palangat, 2001). Mutations that affect pausing are found in homology block F of the largest Pol II subunit (Thuillier et al., 1996), a region that lines the funnel into which RNA is extruded during backtracking. A single-molecule study showed that pausing is a reversible intermediary state between arrest and normal elongation (Davenport et al., 2000). Single-molecule analysis further revealed uniform elongation kinetics, but differences in the frequency and duration of pausing (Adelman et al., 2002). In addition to pausing that involves backtracking, another type of pausing has been observed for the bacterial polymerase, in which the RNA 30 -end disengages from the active site by hypertranslocation of the enzyme (Artsimovitch and Landick, 2000). During transcriptional arrest, RNA polymerase also translocates backwards, leaving the RNA 30 -end intact and extruded (Kashlev and Komissarova, 1997b). RNA polymerase cannot be rescued from arrest by mechanical force (Forde et al., 2002). The need for backtracking and extrusion of the RNA explains why blocking translocation is not sufficient to cause arrest (Samkurashvili and Luse, 1996). Arrest goes along with an increased accomodation of RNA in Pol II (Gu et al., 1996). During normal elongation, about 18 nucleotides of RNA are protected from ribonuclease cleavage, whereas at an arrest site up to 27 nucleotides are protected. The major difference between transcriptional pausing and arrest is that arrested Pol II, in contrast to the paused enzyme, cannot escape without transcript cleavage and the help of an extrinsic protein, the transcript cleavage factor TFIIS. Pol II has a weak intrinsic 30 ! 50 nuclease activity that is greatly stimulated by TFIIS. Bacterial RNA polymerase has been shown to have an intrinsic transcript cleavage activity (Orlova et al., 1995). In the presence of TFIIS, Pol II can cleave the RNA from its 30 -end primarily in dinucleotide increments, although mononucleotides and longer oligonucleotides are also observed (Gu and Reines, 1995; Izban and Luse, 1992, 1993a, 1993b; Hawley and Wang, 1993). Dinucleotides and 7–9-mer oligonucleotides are released from paused and arrested complexes, respectively (Gu and Reines, 1995), showing that arrest involves more extensive backtracking than pausing. TFIIS contacts the 30 -end of the RNA in the Pol II elongation complex (Powell et al., 1996).
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The recent Pol II-TFIIS structure provides a detailed picture of how TFIIS gains access to the Pol II active center via the funnel and pore from below and how TFIIS complements the active site with two functionally essential acidic residues (Kettenberger et al., 2003). TFIIS, however, does not fill the entire pore, it only restricts it. There is enough space for simultaneous binding of backtracked RNA and TFIIS in the pore, as required during TFIIS-mediated rescue of an arrested Pol II elongation complex. Structural and biochemical data indicate that the mechanism of TFIIS-induced RNA cleavage by Pol II involves positioning and activation of a nucleophilic water molecule with the help of a metal ion, to allow for an in-line attack of the phosphodiester bond to be cleaved (Sosunov et al., 2003; Kettenberger et al., 2003). In vitro, TFIIS can also stimulate ‘‘proofreading’’ of the nascent transcript (Agarwal and Jeon, 1996; Thomas et al., 1998), an activity that removes incorrectly incorporated nucleotides from the growing RNA. The following view of the mechanism of mRNA proofreading has emerged from many biochemical observations and the available Pol II structures. Incorporation of the correct nucleotide drives rapid forward translocation (Nedialkov et al., 2003). However, misincorporation of a nucleotide leads to slow forward translocation (Thomas et al., 1998), opening a time window for hydrolytic RNA cleavage and removal of the misincorporated nucleotide. Because a misincorporated nucleotide and the resulting mismatch base pair destabilizes the DNA-RNA hybrid and the elongation complex, misincorporation can also trigger backtracking (Nudler et al., 1997). Backtracking by one nucleotide would lead to cleavage of an RNA dinucleotide. Cleavage of mononucleotides (from the pretranslocation state) and of dinucleotides (from a backtracked state) would both result in a new RNA 30 -end at the active site, from which polymerization can continue. The Pol II-TFIIS complex structure provides evidence that Pol II contains a single tunable active site for both RNA polymerization and cleavage/proofreading, instead of two catalytic sites with distinct locations (Kettenberger et al., 2003). It had been suggested previously that the active sites for RNA polymerization and cleavage are close together or even identical (Powell et al., 1996; Rudd et al., 1994; Hawley and Wang, 1993). In addition to TFIIS, several other proteins influence Pol II elongation, pausing, and arrest, and some of these factors are involved in disease (Conaway and Conaway, 1999; Shilatifard, 1998a, 1998b). Deregulation of Pol II elongation can lead to certain types of cancer (Groudine and Krumm, 1995).
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G. Coupling to RNA Processing and to Other Nuclear Events In addition to the Pol II machinery, expression of protein-coding genes requires multicomponent machines in the nucleus that carry out various steps of mRNA processing, RNA export, and RNA surveillance. Over the last years, a large number of experimental observations showed that there is extensive coupling between these nuclear gene expression machines (Bentley, 2002; Hirose and Manley, 2000; Maniatis and Reed, 2002; Orphanides and Reinberg, 2002). The physical basis for coupling between transcription and mRNA processing appears to be the interaction of RNA processing factors with the phosphorylated Pol II CTD (Hirose and Manley, 2000; Proudfoot, 2000; Steinmetz, 1997). The CTD is flexibly linked to a region beyond the saddle, from which RNA exits, consistent with its role in coupling transcription to mRNA processing (Cramer et al., 2001). There is a tight coupling between transcription and the first RNA processing event, 50 -RNA capping. Capping occurs already when the nascent RNA has reached a length of 25–30 nucleotides and, thus, must take place near the Pol II surface. There is accumulating evidence for the existence of an early elongation checkpoint that ensures that the nascent RNA has received its 50 -cap structure that protects it from degradation (Orphanides and Reinberg, 2002). Other RNA processing events, splicing and 30 -end formation, also occur in a transcription-coupled manner. Pol II transcription elongation is further coupled to events of chromatin remodeling and modification (Orphanides and Reinberg, 2000). Recently it was also found that Pol II elongation is coupled to the export of mRNA out of the nucleus (Hammell et al., 2002; Strasser et al., 2002). Taken together, it now appears that Pol II stands at the heart of a giant mRNA factory that comprises several coupled multicomponent machines (Cook, 1999; Sawadogo and Szentirmay, 2000). Details of these coupling phenomena are beyond the scope of this review.
H. Termination, Polymerase Recycling, Reinitiation, and Regulation Transcription termination occurs in a reaction coupled to RNA 30 -end processing. Most eukaryotic mRNA precursors are cleaved in a site-specific manner in the 30 -untranslated region followed by polyadenylation of the upstream cleavage product. A large number of proteins is involved in these reactions, which are beyond the scope of this review (Barabino and Keller, 1999; Proudfoot, et al., 2002; Manley and Shatkin, 2000). The exact mechanism of coupling between 30 -end processing and transcription termination remains unclear. Termination goes along with dephosphorylation of the Pol II CTD, but the exact time point of Pol II dephosphorylation is also unclear. Dephosphorylation is required for
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the reinitiation of transcription, as Pol II can only join an initiation complex in its unphosphorylated form. The CTD phosphatase Fcp1 plays a key role in Pol II dephosphorylation and recycling (Cho et al., 1999; Kobor et al., 1999). Fcp 1 binds to Pol II via Rpb4 (Kimura et al., 2002). Rpb4 apparently recruits Fcp 1 to the vicinity of the CTD, as the Rpb4/7 complex binds to the beginning of the linker that connects the Pol II core to the CTD (Armache et al., 2003; Bushnell and Kornberg, 2003; Craighead et al., 2002). Reinitiation of Pol II transcription apparently occurs by a mechanism different from initiation (Hahn, 1998). After initiation, a subset of the transcription machinery remains at the promoter, forming a scaffold for assembly of a new initiation complex. This scaffold comprises TFIIA, TFIID, TFIIE, TFIIH, and the multisubunit Mediator complex and can be stabilized by transcriptional activators (Yudkovsky et al., 2000). Reinitiation as well as initiation are important targets for Pol II regulation. High levels of transcription may rely on rapid initiation and on reinititation of polymerases that have terminated. The transcription elongation phase is also subject to regulation, and Pol II elongation can be stimulated by transcriptional activators (Yankulov et al., 1994). The many levels of Pol II regulation befit the central role of Pol II as the end point of signal transduction pathways. In higher eukaryotes, hundreds of transcription factors use Pol II as a regulatory target to induce changes in gene expression. These regulatory proteins generally affect Pol II indirectly, via so-called coactivator complexes, which include the generally required and conserved Mediator complex. The Mediator complex can physically bridge between Pol II and transcriptional activator and repressor proteins. A recent electron microscopic reconstruction of the Pol II–Mediator complex revealed that the interface between Mediator and Pol II includes the polymerase subunit Rpb3 (Davis et al., 2002). Interestingly, two amino acid substitutions on the Rpb3 surface cause a defect in activated transcription (Tan et al., 2000a). Bacterial RNA polymerase contains a target for transcription activation at a similar location on the enzyme surface (Ebright, 2000; Tan et al., 2000b). Mediator is the subject of another review in this volume.
IV. Comparison with Other Polymerases A. Eukaryotic RNA Polymerases I and III Pol II belongs to the family of multisubunit RNA polymerases, which also comprises the two other eukaryotic RNA polymerases, Pol I and Pol III. Pol I and Pol III are mainly responsible for synthesis of ribosomal RNA
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and transfer RNA, respectively. All three eukaryotic RNA polymerases share the five common subunits Rpb5, Rpb6, Rpb8, Rpb10, and Rpb12 (Table II). Four core subunits of Pol II, Rpb1, Rpb2, Rpb3, and Rpb11, all have close homologues in Pol I and Pol III (Table II). The largest subunits of Pol I and Pol III, however, lack a C-terminal repeat domain. Recent studies show that the Rpb4/7 complex of Pol II also has structural and functional counterparts in Pol I and Pol III (Hu et al., 2002; Peyroche et al., 2002; Sadhale and Woychik, 1994; Shematorova and Shpakovski, 1999) and in the archaeal RNA polymerase (Werner et al., 2000). Indeed, the Pol II core-Rpb7 interaction is apparently conserved in all eukaryotic and archaeal RNA polymerases, but not in the bacterial enzyme (Kettenberger et al., 2003). In conclusion, the 12 subunits of Pol II are either identical or homologous in all three eukaryotic enzymes, and Pol II is thus a good model for all eukaryotic RNA polymerases. There are, however, minor differences on the enzymes’ surfaces caused by amino acid insertions and deletions. These differences are most likely responsible for conferring specificity toward the interaction with factors specific for Pol I, II, and III. In addition to the 12 subunits that are either identical or homologous, Pol I contains two specific subunits, A34.5 and A49, and Pol III contains a subcomplex of three specific subunits, called C82, C34, and C31, in yeast. The location of the two Pol I–specific subunits has been determined by electron microscopy and immunolabeling (Bischler et al., 2002). The Pol I subunit A49 binds to the top of the clamp, and subunit A34.5 is located near the jaws. The location of the specific C82/C34/C31 complex of Pol III can be inferred from subunit– subunit interaction studies (Ferri et al., 2000; Flores et al., 1999). These studies indicate that the specific subcomplex is located between the largest polymerase subunit and the Rpb4/7 complex counterpart C17/C25. The C11 subunit of Pol III contains a C-terminal domain that apparently corresponds structurally and functionally to domain III of TFIIS (Chedin et al., 1998; Kettenberger et al., 2003), which inserts into the polymerase pore. Thus, in Pol III, the RNA cleavage stimulatory activity is incorporated into a polymerase subunit, in contrast to Pol II, where it is provided by the additional factor TFIIS.
B. Bacterial and Archaeal RNA Polymerases Bacteria and archaea contain a single multisubunit RNA polymerase. X-ray crystallographic structures were determined of a bacterial RNA polymerase from Thermus aquaticus at 3.3-A˚ resolution (Darst, 2001; Zhang et al., 1999). Comparison of this bacterial RNA polymerase structure
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with the structure of yeast Pol II revealed that five ‘‘core’’ subunits underlie a general RNA polymerase architecture with an active center cleft (Cramer, 2002b). The core subunits show a total of 22 regions of sequence homology ( Jokerst et al., 1989; Minakhin et al., 2001; Sweetser et al., 1987; Darst and Zhang, 1998). These homology regions cluster around the active site and generally adopt the same structure in the bacterial and yeast RNA polymerases. Many additional regions share the same structure, although they differ in sequence between the bacterial and yeast polymerases. Thus structure is conserved better than sequence. The structurally conserved core includes the functional elements of the active center, indicating that all multisubunit RNA polymerases share common mechanistic features. Bacteria do not have a homolog of TFIIS, but the transcript cleavage factors GreA and GreB appear to function essentially like TFIIS, as revealed in an electron microscopic study recently (Opalka et al., 2003). A coiled coil of GreB binds in the secondary channel of bacterial polymerase, which corresponds to the Pol II pore, and reaches the active site with an acidic tip. These findings demonstrate in a powerful way the conserved strategies employed for RNA cleavage stimulation by the structurally unrelated bacterial and eukaryotic RNA polymerase cleavage factors. Bacterial RNA polymerase consists of the five core subunits only. In eukaryotic RNA polymerases, up to 10 additional subunits are found around the periphery of the enzymes (Table II). Archeael RNA polymerases comprise between five and seven subunits in addition to the five core subunits (Darcy et al., 1999; Langer et al., 1995). For all Pol II subunits except Rpb8, homologues have been reported in archaeal RNA polymerases. Thus, the overall structure of archaeal RNA polymerases must be very similar to the yeast Pol II structure. Although the archaeal enzymes lack some external domains, they are apparently closely related to the eukaryotic Pol II. The similarity between archaeal RNA polymerases and Pol II extends to the initiation complex. Archaea contain homologues of three Pol II general transcription factors, TBP, TFIIB, and TFIIE (Bell and Jackson, 2001). The archaeal RNA polymerase machinery is thus more closely related to the eukaryotic machinery than to the bacterial system. Indeed, an archaeal TFIIS homologue, TFS, has also been described (Hausner et al., 2000).
C. Single-Subunit DNA and RNA Polymerases Structures of multisubunit RNA polymerases are strikingly different from structures of polymerases of other families, such as the many single-subunit DNA and RNA polymerases. X-ray crystallography of
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single-subunit DNA and RNA polymerases revealed a great structural diversity (Beard and Wilson, 2001; Brautigam and Steitz, 1998; Doublie et al., 1999; Ellenberger and Silvian, 2001; Jager and Pata, 1999; Loeb and Patel, 2001; Steitz, 1999). Nevertheless, most single-subunit polymerases show a similar overall architecture and considerable structural conservation of the active center. Representative structures of the diverse singlesubunit polymerases were compared to Pol II by overlaying corresponding nucleic acids in functional complexes (Cramer, 2002a). In functional complexes of these diverse enzymes, nucleic acids take a similar course through the active center. In all cases studied, the entering DNA duplex encloses an angle of almost 90 degrees with the exiting template-product duplex. At the location of the bend, subsequent DNA template bases are twisted. This twist aligns the ‘‘coding’’ base with the binding site for the incoming nucleoside triphosphate substrate. The nucleoside triphosphate enters through an opening that is found in all polymerases. The nucleotide substrate often binds between an -helix and two catalytic metal ions. The exiting template-product duplex is bound from the minor groove side in all polymerases. Conformational changes on nucleic acid binding have been detected for several different polymerases, but the nature of this ‘‘induced fit’’ differs. Recent structures of elongation complexes of RNA polymerase from bacteriophage T7 have revealed dramatic changes in the conformation of the N-terminal domain on transition from initiation to elongation (Tahirov et al., 2002; Yin and Steitz, 2002). In the Pol II system, corresponding changes remain to be discovered but may be predominantly found in the general transcription factors rather than the polymerase itself. Structural and functional analysis of Pol II supports the idea that DNA and RNA polymerases follow different strategies for nucleic acid cleavage and proofreading. In the Klenow DNA polymerase, the growing DNA shuttles between widely separated active sites for DNA synthesis and cleavage, whereas in Pol II the growing RNA appears to remain at a single tunable active site that switches between RNA synthesis and cleavage modes, with the latter being dramatically enhanced by TFIIS. Despite this difference in strategy, both classes of polymerases may use the same general two–metal ion mechanism for both polymerization and cleavage of nucleic acids.
VI. Conclusions Detailed structures are now available for the Pol II core enzyme in free form, in the form of a minimal elongation complex with bound nucleic acids, and in an inhibited form with bound -amanitin. In addition,
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backbone models of the complete initiation-competent Pol II, including the Rpb4/7 complex, and of the complete Pol II with bound elongation factor TFIIS have recently been described. The structures together with many functional studies have given many insights into the mechanism of mRNA transcription. Structural and functional studies of bacterial RNA polymerase allow for interesting comparisons and evolutionary considerations. The Pol II structures now guide mutagenesis experiments aimed at a dissection of the transcription mechanism. In the future, further structures of Pol II complexes with transcription factors will provide more mechanistic details of the mRNA transcription cycle.
References Adelman, K., La Porta, A., Santangelo, T. J., Lis, J. T., Roberts, J. W., and Wang, M. D. (2002). Single molecule analysis of RNA polymerase elongation reveals uniform kinetic behavior. Proc. Natl. Acad. Sci. USA 99(21), 13538–13543. Albert, T., Mautner, J., Funk, J. O., Hortnagel, K., Pullner, A., and Eick, D. (1997). Nucleosomal structures of e-myc promoters with transcriptionally engaged RNA polymerase II. Mol. Cell. Biol. 17(8), 4363–4371. Armache, K.-J., Kettenberger, H., and Cramer, P. (2003). Architecture of the initiationcompetent 12-subunit RNA polymerase II. Proc. Natl. Acad. Sci. USA 100(12), 6964–6968. Artsimovitch, I., and Landick, R. (2000). Pausing by bacterial RNA polymerase is mediated by mechanistically distinct classes of signals. Proc. Natl. Acad. Sci. USA 97(13), 7090–7095. Asturias, F. J., Meredith, G. D., Poglitsch, C. L., and Kornberg, R. D. (1997). Two conformations of RNA polymerase II revealed by electron crystallography. J. Mol. Biol. 272(4), 536–540. Barabino, S. M., and Keller, W. (1999). Last but not least: regulated poly(A) tail formation. Cell 99(1), 9–11. Baskaran, R., Chiang, G. G., Mysliwiec, T., Kruh, G. D., and Wang, J. Y. (1997). Tyrosine phosphorylation of RNA polymerase II carboxyl terminal domain by the Ab1 related gene product. J. Biol. Chem. 272(30), 18905–18909. Beard, W. A., and Wilson, S. H. (2001). DNA lesion by pass polymerases open up. Structure (Camb) 9(9), 759–764. Bell, S. D., and Jackson, S. P. (2001). Mechanism and regulation of transcription in archaea. Curr. Opin. Microbiol. 4(2), 208–213. Bentley, D. (2002). The mRNA assembly line: transcription and processing machines in the same factory. Curr. Opin. Cell. Biol. 14(3), 336–342. Bischler, N., Brino, L., Carles, C., Riva, M., Tschochner, H., Mallouh, V., and Schultz, P. (2002). Localization of the yeast RNA polymerase I-specific subunits. EMBO J. 21(15), 4136–4144. Brautigam, C. A., and Steitz, T. A. (1998). Structural and functional insights provided by crystal structures of DNA polymerases and their substrate complexes. Curr. Opin. Struct. Biol. 8(1), 54–63. Buratowski, S. (1994). The basics of basal transcription by RNA polymerase II. Cell 77(1), 1–3.
32
CRAMER
Buratowski, S., Hahn, S., Guarente, L., and Sharp, P. A. (1989). Five intermediate complexes in transcription initiation by RNA polymerase II. Cell 56, 549–561. Burke, T. W., Willy, P. J., Kutach, A. K., Butler, J. E., and Kadonaga, J. T. (1998). The DPE, a conserved downstream core promoter element that is functionally analogous to the TATA box. Cold Spring Harb. Symp. Quant. Biol. 63, 75–82. Bushnell, D. A., Cramer, P., and Kornberg, R. D. (2001). Selenomethionine incorporation in Saccharomyces cerevisiae RNA polymerase II. Struct. Fold. Des. 9(1), R3–R9. Bushnell, D. A., Cramer, P., and Kornberg, R. D. (2002). Structural basis of transcription: -amanitin RNA polymerase II cocrystal at 2.8 A˚ resolution. Proc. Natl. Acad. Sci. USA 99, 1218–1222. Bushnell, D. A., and Kornberg, R. D. (2003). Complete 12-subunit RNA polymerase II at 4.1 A˚ resolution: implications for the initiation of transcription. Proc. Natl. Acad. Sci. USA 100(12), 6969–6972. Cadena, D. L., and Dahmus, M. E. (1987). Messenger RNA synthesis in mammalian cells is catalyzed by the phosphorylated form of RNA polymerase II. J. Biol. Chem. 262(26), 12468–12474. Cagas, P. M., and Corden, J. L. (1995). Structural studies of a synthetic peptide derived from the carboxyl-terminal domain of RNA polymerase II. Proteins 21(2), 149–160. Chafin, D. R., Guo, H., and Price, D. H. (1995). Actions of -amanitin during pyrophosphoryolysis and elongation by RNA polymerase II. J. Biol. Chem. 270, 19114–19119. Chedin, S., Riva, M., Schultz, P., Sentenac, A., and Carles, C. (1998). The RNA cleavage activity of RNA polymerase III is mediated by an essential TFHS-like subunit and is important for transcription termination. Genes Dev. 12, 3857–3871. Cho, H., Kim, T. K., Mancebo, H., Lane, W. S., Flores, O., and Reinberg, D. (1999). A protein phosphatase functions to recycle RNA polymerase II. Genes Dev. 13(12), 1540–1552. Conaway, J. W., and Conaway, R. C. (1999). Transcription elongation and human disease. Annu. Rev. Biochem. 68, 301–319. Cook, P. R. (1999). The organization of replication and transcription. Science 284(5421), 1790–1795. Cox, J. M., Hayward, M. M., Sanchez, J. F., Gegnas, L. D., van der Zee, S., Dennis, J. H., Sigler, P. B., and Schepartz, A. (1997). Bidirectional binding of the TATA box binding protein to the TATA box. Proc. Natl. Acad. Sci. USA 94(25), 13475–13480. Craighead, J. L., Chang, W. H., and Asturias, F. J. (2002). Structure of yeast RNA polymerase II in solution: implications for enzyme regulation and interaction with promoter DNA. Struct. Fold. Des. 10(8), 1117–1125. Cramer, P. (2002a). Common structural features of nucleic acid polymerases. Bioessays 24, 724–729. Cramer, P. (2002b). Multisubunit RNA polymerases. Curr. Op. Struct. Biol. 12, 89–97. Cramer, P., Bushnell, D. A., Fu, J., Gnatt, A. L., Maier-Davis, B., Thompson, N. E., Burgess, R. R., Edwards, A. M., David, P. R., and Kornberg, R. D. (2000). Architecture of RNA polymerase II and implications for the transcription mechanism. Science 288(5466), 640–649. Cramer, P., Bushnell, D. A., and Kornberg, R. D. (2001). Structural basis of transcription: RNA polymerase II at 2.8 angstrom resolution. Science 292(5523), 1863–1876. Dahmus, M. E. (1996). Reversible phosphorylation of the C-terminal domain of RNA polymerase II. J. Biol. Chem. 271(32), 19009–19012. Darcy, T. J., Hausner, W., Awery, D. E., Edwards, A. M., Thomm, M., and Reeve, J. N. (1999). Methanobacterium thermoautotrophicum RNA polymerase and transcription in vitro. J. Bacteriol. 181(14), 4424–4429.
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
33
Darst, S. A. (2001). Bacterial RNA polymerase. Curr. Opin. Struct. Biol. 11(2), 155–162. Darst, S. A., Edwards, A. M., Kubalek, E. W., and Kornberg, R. D. (1991a). Threedimensional structure of yeast RNA polymerase II at 16 A˚ resolution. Cell 66, 121–128. Darst, S. A., Kubalek, E. W., Edwards, A. M., and Kornberg, R. D. (1991b). Twodimensional and epitaxial crystallization of a mutant form of yeast RNA polymerase II. J. Mol. Biol. 221, 347–357. Davenport, R. J., Wuite, G. J., Landick, R., and Bustamante, C. (2000). Single-molecule study of transcriptional pausing and arrest by E. coli RNA polymerase. Science 287(5462), 2497–2500. Davis, J. A., Takagi, Y., Kornberg, R. D., and Asturias, F. A. (2002). Structure of the yeast RNA polymerase II holoenzyme: Mediator conformation and polymerase interaction. Mol. Cell 10(2), 409–415. del Rio-Portilla, F., Gaskell, A., Gilbert, D., Ladias, J. A., and Wagner, G. (1999). Solution structure of the hRPABC14.4 subunit of human RNA polymerases. Nat. Struct. Biol. 6(11), 1039–1042. Doublie, S., Sawaya, M. R., and Ellenberger, T. (1999). An open and closed case for all polymerases. Struct. Fold. Des. 7(2), R31–R35. Doublie, S., Tabor, S., Long, A. M., Richardson, C. C., and Ellenberger, T. (1998). Crystal structure of a bacteriophage T7 DNA replication complex at 2.2 A˚ resolution. Nature 391, 251–258. Dvir, A. (2002). Promoter escape by RNA polymerase II. Biochim. Biophys. Acta. 1577(2), 208–223. Ebright, R. H. (1998). RNA polymerase-DNA interaction. Structures of intermediate, open, and elongation complexes. Cold Spring Harb. Symp. Quant. Biol. 63, 11–20. Ebright, R. H. (2000). RNA polymerase: structural similarities between bacterial RNA polymerase and eukaryotic RNA polymerase II. J. Mol. Biol. 304(5), 687–698. Ederth, J., Artsimovitch, I., Isaksson, L. A., and Landick, R. (2002). The downstream DNA jaw of bacterial RNA polymerase facilitates both transcriptional initiation and pausing. J. Biol. Chem. 277(40), 37456–37463. Edwards, A. M., Darst, S. A., Feaver, W. J., Thompson, N. E., Burgess, R. R., and Kornberg, R. D. (1990). Purification and lipid layer crystallization of yeast RNA polymerase II. Proc. Natl. Acad. Sci. USA 87, 2122–2126. Edwards, A. M., Kane, C. M., Young, R. A., and Kornberg, R. D. (1991). Two dissociable subunits of yeast RNA polymerase II stimulate the initiation of transcription at a promoter in vitro. J. Biol. Chem. 266, 71–75. Ellenberger, T., and Silvian, L. F. (2001). The anatomy of infidelity. Nat. Struct. Biol. 8(10), 827–828. Erie, D. A. (2002). The many conformational states of RNA polymerase elongation complexes and their roles in the regulation of transcription. Biochim. Biophys. Acta. 1577(2), 224–239. Ferri, M. L., Peyroche, G., Siaut, M., Lefebvre, O., Carles, C., Conesa, C., and Sentenac, A. (2000). A novel subunit of yeast RNA polymerase III interacts with the TFIIBrelated domain of TFIIIB70. Mol. Cell. Biol. 20(2), 488–495. Fiedler, U., and Marc Timmers, H. T. (2000). Peeling by binding or twisting by cranking: models for promoter opening and transcription initiation by RNA polymerase II. Bioessays 22(4), 316–326. Fiedler, U., and Timmers, H. T. (2001). Analysis of the open region of RNA polymerase II transcription complexes in the early phase of elongation. Nucleic Acids Res. 29(13), 2706–2714.
34
CRAMER
Flores, A., Briand, J. F., Gadal, O., Andrau, J. C., Rubbi, L., Van Mullem, V., Boschiero, C., Goussot, M., Marck, C., Carles, C., Thuriaux, P., Sentenac, A., and Werner, M. (1999). A protein-protein interaction map of yeast RNA polymerase III. Proc. Natl. Acad. Sci. USA 96(14), 7815–7820. Forde, N. R., Izhaky, D., Woodcock, G. R., Wuite, G. J., and Bustamante, C. (2002). Using mechanical force to probe the mechanism of pausing and arrest during continuous elongation by Escherichia coli RNA polymerase. Proc. Natl. Acad. Sci. USA 99(18), 11682–11687. Franklin, M. C., Wang, J., and Steitz, T. A. (2001). Structure of the replicating complex of a pol alpha family DNA polymerase. Cell 105(5), 657–667. Fu, J., Gerstein, M., David, P. R., Gnatt, A. L., Bushnell, D. A., Edwards, A. M., and Kornberg, R. D. (1998). Repeated tertiary fold of RNA polymerase II and implications for DNA binding. J. Mol. Biol. 280, 317–322. Fu, J., Gnatt, A. L., Bushnell, D. A., Jensen, G. J., Thompson, N. E., Burgess, R. R., David, P. R., and Kornberg, R. D. (1999). Yeast RNA polymerase II at 5 A˚ resolution. Cell 98, 799–810. Furter-Graves, E. M., Hall, B. D., and Furter, R. (1994). Role of a small RNA pol II subunit in TATA to transcription start site spacing. Nucleic Acids Res. 22(23), 4932–4936. Gnatt, A. (2002). Elongation by RNA polymerase II: structure function relationship. Biochim. Biophys. Acta. 1577(2), 175–190. Gnatt, A., Fu, J., and Kornberg, R. D. (1997). Formation and crystallization of yeast RNA polymerase II elongation complexes. J. Biol. Chem. 272, 30799–30805. Gnatt, A. L., Cramer, P., Fu, J., Bushnell, D. A., and Kornberg, R. D. (2001). Structural basis of transcription: an RNA polymerase II elongation complex at 3.3 A˚ resolution. Science 292(5523), 1876–1882. Greenblatt, J. (1997). RNA polymerase II holoenzyme and transcriptional regulation. Curr. Opin. Cell. Biol. 9(3), 310–319. Gu, W., Powell, W., Mote, J., and Reines, D. (1993). Nascent RNA cleavage by arrested RNA polymerase II does not require upstream translocation of the elongation complex on DNA. J. Biol. Chem. 268, 25604–25616. Gu, W., and Reines, D. (1995). Variation in the size of nascent RNA cleavage products as a function of transcript length and elongation competence. J. Biol. Chem. 270, 30441–30447. Gu, W., Wind, M., and Reines, D. (1996). Increased accommodation of nascent RNA in a product site on RNA polymerase II during arrest. Proc. Natl. Acad. Sci. USA 93, 6935–6940. Hahn, S. (1998). Activation and the role of reinitiation in the control of transcription by RNA polymerase II. Cold Spring Harb. Symp. Quant. Biol. 63, 181–188. Hammell, C. M., Gross, S., Zenklusen, D., Health, C. V., Stutz, F., Moore, C., and Cole, C. N. (2002). Coupling of termination, 30 processing, and mRNA export. Mol. Cell. Biol. 22(18), 6441–6457. Hausner, W., Lange, U., and Musfeldt, M. (2000). Transcription factor S, a cleavage induction factor of the archaeal RNA polymerase. J. Biol. Chem. 275, 12393–12399. Hirose, Y., and Manley, J. L. (2000). RNA polymerase II and the integration of nuclear events. Genes Dev. 14(12), 1415–1429. Holstege, F. C. P., Fiedler, U., and Timmers, H. T. M. (1997). Three transitions in the RNA polymerase II transcription complex during initiation. EMBO J. 16, 7468–7480. Hu, P., Wu, S., Sun, Y., Yuan, C. C., Kobayashi, R., Myers, M. P., and Hernandez, N. (2002). Characterization of human RNA polymerase III identifies orthologues for
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
35
Saccharomyces cerevisiae RNA polymerase III subunits. Mol. Cell. Biol. 22(22), 8044–8055. Hull, W. W., McKune, K., and Woychik, N. A. (1995). RNA polymerase II subunit RPB9 is required for accurate start site selection. Genes Dev. 9, 481–490. Hunter, T. (1998). Prolyl isomerases and nuclear function. Cell 92(2), 141–143. Izban, M. G., and Luse, D. S. (1992). The RNA polymerase II ternary complex cleaves the nascent transcript in a 30 -50 direction in the presence of elongation factor SII. Genes Dev. 6, 1342–1356. Izban, M. G., and Luse, D. S. (1993a). The increment of SII facilitated transcript cleavage varies dramatically between elongation competent and incompetent RNA polymerase II ternary complexes. J. Biol. Chem. 268, 12874–12885. Izban, M. G., and Luse, D. S. (1993). SII-facilitated transcript cleavage in RNA polymerase II complexes stalled early after initiation occurs in primarily dinucleotide increments. J. Biol. Chem. 268, 12864–12873. Jager, J., and Pata, J. D. (1999). Getting a grip: polymerases and their substrate complexes. Curr. Opin. Struct. Biol. 9(1), 21–28. Jensen, G. J., Meredith, G., Bushnell, D. A., and Kornberg, R. D. (1998). Structure of wild type yeast RNA polymerase II and location of Rpb4 and Rpb7. EMBO J. 17(8), 2353–2358. Jeon, C-J., and Agarwal, K. (1996). Fidelity of RNA polymerase II transcription controlled by elongation factor TFIIS. Proc. Natl. Acad. Sci. USA 93, 13677–13682. Jeon, C., Yoon, H., and Agarwal, K. (1994). The transcription factor TFIIS zinc ribbon dipeptide Asp-Glu is critical for stimulation of elongation and RNA cleavage by RNA polymerase II. Proc. Natl. Acad. Sci. USA 91, 9106–9110. Jokerst, R. S., Weeks, J. R., Zehring, W. A., and Greenleaf, A. L. (1989). Analysis of the gene encoding the largest subunit of RNA polymerase II in Drosophila. Mol. Gene Genet. 215, 266–275. Kahl, B. F., Li, H., and Paule, M. R. (2000). DNA melting and promoter clearance by eukaryotic RNA polymerase I. J. Mol. Biol. 299(1), 75–89. Kettenberger, H., Armache, K.-J., and Cramer, P. (2003). Architecture of the RNA polymerase II-TFIIS complex and implications for mRNA cleavage. Cell 114(3), 347–357. Khazak, V., Sadhale, P. P., Woychik, N. A., Brent, R., and Golemis, E. A. (1995). Human RNA polymerase II subunit hsRPB7 functions in yeast and influences stress survival and cell morphology. Mol. Biol. Cell 6(7), 759–775. Killeen, M., Coulombe, B., and Greenblatt, J. (1992). Recombinant TBP, transcription factor IIB, and RAP30 are sufficient for promoter recognition by mammalian RNA polymerase II. J. Biol. Chem. 267(14), 9463–9466. Kim, J. L., Nikolov, D. B., and Burley, S. K. (1993). Co-crystal structure of TBP recognizing the minor groove of a TATA element. Nature 365, 520–527. Kim, T. K., Ebright, R. H., and Reinberg, D. (2000). Mechanism of ATP-dependent promoter melting by transcription factor IIH. Science 288(5470), 1418–1422. Kim, Y., Geiger, J. H., Hahn, S., and Sigler, P. B. (1993). Crystal structure of a yeast TBP/TATA-box complex. Nature 365, 512–519. Kimura, M., Suzuki, H., and Ishihama, A. (2002). Formation of a carboxy-terminal domain phosphatase (Fcp1)/TFIIF/RNA polymerase II (pol II) complex in Schizosaccharomyces pombe involves direct interaction between Fep1 and the Rpb4 subunit of pol II. Mol. Cell. Biol. 22(5), 1577–1588. Kireeva, M. L., Komissarova, N., Waugh, D. S., and Kashlev, M. (2000). The 8-nucleotide-long RNA:DNA hybrid is a primary stability determinant of the RNA polymerase II elongation complex. J. Biol. Chem. 275(9), 6530–6536.
36
CRAMER
Kobor, M. S., Archambault, J., Lester, W., Holstege, F. C., Gileadi, O., Jansma, D. B., Jennings, E. G., Kouyoumdjian, F., Davidson, A. R., Young, R. A., and Greenblatt, J. (1999). An unusual eukaryotic protein phosphatase required for transcription by RNA polymerase II and CTD dephosphorylation in S. cerevisiae. Mol. Cell 4(1), 55–62. Kobor, M. S., and Greenblatt, J. (2002). Regulation of transcription elongation by phosphorylation. Biochim. Biophys. Acta. 1577(2), 261–275. Komarnitsky, P., Cho, E. J., and Buratowski, S. (2000). Different phosphorylated forms of RNA polymerase II and associated mRNA processing factors during transcription. Genes Dev. 14(19), 2452–2460. Komissarova, N., and Kashlev, M. (1997a). RNA polymerase switches between inactivated and activated states by translocating back and forth along the DNA and the RNA. J. Biol. Chem. 272, 15329–15338. Komissarova, N., and Kashlev, M. (1997b). Transcription arrest: Escherichia coli RNA polymerase translocates backward, leaving the 30 end of the RNA intact and extruded. Proc. Natl. Acad. Sci. USA 94, 1755–1760. Komissarova, N., and Kashlev, M. (1998). Functional topography of nascent RNA in elongation intermediates of RNA polymerase. Proc. Natl. Acad. Sci. USA 95, 14699–14704. Kornberg, R. D. (1999). Eukaryotic transcriptional control. Trends Cell. Biol. 9(12), M46–M49. Korzheva, N., Mustaev, A., Kozlov, M., Malhotra, A., Nikiforov, V., Goldfarb, A., and Darst, S. A. (2000). A structural model of transcription elongation. Science 289(5479), 619–625. Korzheva, N., Mustaev, A., Nudler, E., Nikiforov, V., and Goldfarb, A. (1998). Mechanistic model of the elongation complex of Escherichia coli RNA polymerase. Cold Spring Harb. Symp. Quant. Biol. 63, 337–345. Kosa, P., Ghosh, G., DeDecker, B. S., and Sigler, P. B. (1997). The 2.1-A˚ crystal structure of an archaeal preinitiation complex: TATA box binding protein/transcription factor (II)B core/TATA box. Proc. Natl. Acad. Sci. USA 94, 6042–6047. Krapp, S., Kelly, G., Reischl, J., Weinzierl, R. O. J., and Matthews, S. (1998). Eukaryotic RNA polymerase subunit RPB8 is a new relative of the OB family. Nat. Struct. Biol. 5, 110–114. Krumm, A., and Groudine, M. (1995). Tumor suppression and transcription elongation: the dire consequences of changing partners. Science 269(5229), 1400–1401. Kugel, J. F., and Goodrich, J. A. (1998). Promoter escape limits the rate of RNA polymerase II transcription and is enhanced by TFIIE, TFIIH, and ATP on negatively supercoiled DNA. Proc. Natl. Acad. Sci. USA 95(16), 9232–9237. Kugel, J. F., and Goodrich, J. A. (2000). A kinetic model for the early steps of RNA synthesis by human RNA polymerase II. J. Biol. Chem. 275(51), 40483–40491. Kugel, J. F., and Goodrich, J. A. (2002). Translocation after synthesis of a four nucleotide RNA commits RNA polymerase II to promoter escape. Mol. Cell. Biol. 22(3), 762–773. Kuznedelov, K., Korzheva, N., Mustaev, A., and Severinov, K. (2002). Structure-based analysis of RNA polymerase function: the largest subunits rudder contributes critically to elongation complex stability and is not involved in the maintenance of RNA-DNA hybrid length. EMBO J. 21(6), 1369–1378. Lagrange, T., Kapanidis, A. N., Tang, H., Reinberg, D., and Ebright, R. H. (1998). New core promoter element in RNA polymerase II dependent transcription: sequence specific DNA binding by transcription factor IIB. Genes Dev. 12(1), 34–44.
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
37
Landick, R. (2001). RNA polymerase clamps down. Cell 105(5), 567–570. Langer, D., Hain, J., Thuriaux, P., and Zillig, W. (1995). Transcription in archaea: similarity to that in eukarya. Proc. Natl. Acad. Sci. 92, 5768–5772. Larkin, R. M., and Guilfoyle, T. J. (1998). Two small subunits in Arabidopsis RNA polymerase II are related to yeast RPB4 and RPB7 and interact with one another. J. Biol. Chem. 273(10), 5631–5637. Leuther, K. K., Bushnell, D. A., and Kornberg, R. D. (1996). Two-dimensional crystallography of transcription factor IIB- and IIE-RNA polymerase II complexes: implications for start site selection and initiation complex formation. Cell 85, 773–779. Li, B., Weber, J. A., Chen, Y., Greenleaf, A. L., and Gilmour, D. S. (1996). Analyses of promoter-proximal pausing by RNA polymerase II on the hsp 70 heat shock gene promoter in a Drosophila nuclear extract. Mol. Cell. Biol. 16(10), 5433–5443. Li, Y., Korolev, S., and Waksman, G. (1998). Crystal structures of open and closed forms of binary and ternary complexes of the large fragment of Thermus aquaticus DNA polymerase I: structural basis for nucleotide incorporation. EMBO J. 17(24), 7514–7525. Littlefield, O., Korkhin, Y., and Sigler, P. B. (1999). The structural basis for the oriented assembly of a TBP/TFB/promoter complex. Proc. Natl. Acad. Sci. USA 96(24), 13668–13673. Mackereth, C. D., Arrowsmith, C. H., Edwards, A. M., and McIntosh, L. P. (2000). Zinc-bundle structure of the essential RNA polymerase subunit RPB10 from Methanobacterium thermoautotrophicum. Proc. Natl. Acad. Sci. USA 97(12), 6316–6321. Magill, C. P., Jackson, S. P., and Bell, S. D. (2001). Identification of a conserved archaeal RNA polymerase subunit contacted by the basal transcription factor TFB. J. Biol. Chem. 276(50), 46693–46696. Maniatis, T., and Reed, R. (2002). An extensive network of coupling among gene expression machines. Nature 416(6880), 499–506. Maxon, M. E., Goodrich, J. A., and Tjian, R. (1994). Transcription factor IIE binds preferentially to RNA polymerase IIA and recruits TFIIH: a model for promoter clearance. Genes Dev. 8, 515–524. McKune, K., Richards, K. L., Edwards, A. M., Young, R. A., and Woychik, N. A. (1993). Rpb7, one of two dissociable subunits of yeast RNA polymerase II, is essential for cell viability. Yeast 9, 295–299. Meredith, G. D., Chang, W.-H., Li, Y., Bushnell, D. A., Darst, S. A., and Kornberg, R. D. (1996). The C-terminal domain in the structure of RNA polymerase II. J. Mol. Biol. 258, 413–419. Minakhin, L., Bhagat, S., Brunning, A., Campbell, E. A., Darst, S. A., Ebright, R. H., and Severinov, K. (2001). Bacterial RNA polymerase subunit omega and eukaryotic RNA polymerase subunit RPB6 are sequence, structural, and functional homologs and promote RNA polymerase assembly. Proc. Natl. Acad. Sci. USA 98(3), 892–897. Murakami, K. S., Masuda, S., Campbell, E. A., Muzzin, O., and Darst, S. A. (2002). Structural basis of transcription initiation: an RNA polymerase holoenzyme-DNA complex. Science 296(5571), 1285–1290. Myer, V. E., and Young, R. A. (1998). RNA polymerase II holoenzymes and subcomplexes. J. Biol. Chem. 273(43), 27757–27760. Nedialkov, Y. A., Gong, X. Q., Hovde, S. L., Yamaguchi, Y., Handa, H., Geiger, J. H., Yan, H., and Burton, Z. F. (2003). NTP-driven translocation by human RNA polymerase II. J. Biol. Chem. 278, 18303–18312. Nikolov, D. B., Chen, H., Halay, E. D., Usheva, A. A., Hisatake, K., Lee, D. K., Roeder, R. G., and Burley, S. K. (1995). Crystal structure of a TFIIB-TBP-TATA-element ternary complex. Nature 377, 119–128.
38
CRAMER
Nudler, E. (1999). Transcription elongation: structural basis and mechanisms. J. Mol. Biol. 288(1), 1–12. Nudler, E., Mustaev, A., Lukhtanov, E., and Goldfarb, A. (1997). The RNA-DNA hybrid maintains the register of transcription by preventing backtracking of RNA polymerase. Cell 89, 38–41. O’Brien, R., DeDecker, B., Fleming, K. G., Sigler, P. B., and Ladbury, J. E. (1998). The effects of salt on the TATA binding protein-DNA interaction from a hyperthermophilic archacon. J. Mol. Biol. 279(1), 117–125. O’Brien, T., Hardin, S., Greenleaf, A., and Lis, J. T. (1994). Phosphorylation of RNA polymerase II C-terminal-domain and transcriptional elongation. Nature 370, 75–77. Ohkuma, Y. (1997). Multiple functions of general transcription factors TFIIE and TFIIH in transcription. Possible points of regulation by trans acting factors. J. Biochem. 122, 481–489. Opalka, N., Chlenov, M., Chacon, P., Rice, W. J., Wriggers, W., and Darst, S. A. (2003). Structure and function of the transcription elongation factor GreB bound to bacterial RNA polymerase. Cell 114(3), 335–345. Orlicky, S. M., Tran, P. T., Sayre, M. H., and Edwards, A. M. (2001). Dissociable Rpb4, Rpb7 subassembly of rna polymerase II binds to single-strand nucleic acid and mediates a post recruitment step in transcription initiation. J. Biol. Chem. 276(13), 10097–10102. Orlova, M., Newlands, J., Das, A., Goldfarb, A., and Borukhov, S. (1995). Intrinsic transcript cleavage activity of RNA polymerase. Proc. Natl. Acad. Sci. USA 92(10), 4596–4600. Orphanides, G., and Reinberg, D. (2000). RNA polymerase II elongation through chromatin. Nature 407(6803), 471–475. Orphanides, G., and Reinberg, D. (2002). A unified theory of gene expression. Cell 108(4), 439–451. Ossipow, V., Iassan, J.-P., Nigg, E. A., and Schibler, U. (1995). A mammalian RNA polymerase II holoenzyme containing all components required for promoterspecific transcription initiation. Cell 83, 137–146. Pal, M., and Luse, D. S. (2002). Strong natural pausing by RNA polymerase II within 10 bases of transcription start may result in repeated slippage and reextension of the nascent RNA. Mol. Cell. Biol. 22(1), 30–40. Palangat, M., and Landick, R. (2001). Roles of RNA:DNA hybrid stability, RNA structure, and active site conformation in pausing by human RNA polymerase II. J. Mol. Biol. 311(2), 265–282. Pan, G., and Greenblatt, J. (1994). Initiation of transcription by RNA polymerase II is limited by melting of the promoter DNA in the region immediately upstream of the initiation site. J. Biol. Chem. 269(48), 30101–30104. Patel, P. H., and Loeb, L. A. (2001). Getting a grip on how DNA polymerases function. Nat. Struct. Biol. 8(8), 656–659. Peyroche, G., Levillain, E., Siaut, M., Callebaut, I., Schultz, P., Sentenac, A., Riva, M., and Carles, C. (2002). The A14-A43 heterodimer subunit in yeast RNA pol I and their relationship to Rpb4-Rpb7 pol II subunits. Proc. Natl. Acad. Sci. USA 99(23), 14670–14675. Peyroche, G., Milkereit, P., Bischler, N., Tschochner, H., Schultz, P., Sentenac, A., Carles, C., and Riva, M. (2000). The recruitment of RNA polymerase I on rDNA is mediated by the interaction of the A43 subunit with Rrn3. EMBO J. 19(20), 5473–5482.
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
39
Poglitsch, C. L., Meredith, G. D., Gnatt, A. L., Jensen, G. J., Chang, W. H., Fu, J., and Kornberg, R. D. (1999). Electron crystal structure of an RNA polymerase II transcription elongation complex. Cell 98, 791–798. Powell, W., Bartholomew, B, and Reines, D. (1996). Elongation factor SII contacts the 30 end of RNA in the RNA polymerase II elongation complex. J. Biol. Chem. 271, 22301–22304. Proudfoot, N. (2000). Connecting transcription to messenger RNA processing. Trends Biochem. Sci. 25(6), 290–293. Proudfoot, N. J., Furger, A., and Dye, M. J. (2002). Integrating mRNA processing with transcription. Cell 108(4), 501–512. Pugh, B. F. (2000). Control of gene expression through regulation of the TATAbinding protein. Gene 255(1), 1–14. Raschke, E. E., Albert, T., and Eick, D. (1999). Transcriptional regulation of the Ig kappa gene by promoter-proximal pausing of RNA polymerase II. J. Immunol. 163(8), 4375–4382. Rudd, M. D., Izban, M. G., and Luse, D. S. (1994). The active site of RNA polymerase II participates in transcript cleavage within arrested ternary complexes. Proc. Natl. Acad. Sci. USA 91, 8057–8061. Sadhale, P. P., and Woychik, N. A. (1994). C25, an essential RNA polymerase III subunit related to the RNA polymerase II subunit RPB7. Mol. Cell. Biol. 14(9), 6164–6170. Sakurai, H., Mitsuzawa, H., Kimura, M., and Ishihama, A. (1999). The Rpb4 subunit of fission yeast Schizosaccharomyces pombe RNA polymerase II is essential for cell viability and similar in structure to the corresponding subunits of higher eukaryotes. Mol. Cell. Biol. 19(11), 7511–7518. Samkurashvili, I., and Luse, D. S. (1996). Translocation and transcriptional arrest during transcript elongation by RNA polymerase II. J. Biol. Chem. 271(38), 23495–23505. Samkurashvili, I., and Luse, D. S. (1998). Structural changes in the RNA polymerase II transcription complex during transition from initiation to elongation. Mol. Cell. Biol. 18(9), 5343–5354. Sawaya, M. R., Prasad, R., Wilson, S. H., Kraut, J., and Pelletier, H. (1997). Crystal structures of human DNA polymerase beta complexed with gapped and nicked DNA: evidence for an induced fit mechanism. Biochemistry 36(37), 11205–11215. Shatkin, A. J., and Manley, J. L. (2000). The ends of the affair: capping and polyadenylation. Nat. Struct. Biol. 7(10), 838–842. Shaw, P. E. (2002). Peptidyl-prolyl isomerases: a new twist to transcription. EMBO Rep. 3(6), 521–526. Sheffer, A., Varon, M., and Choder, M. (1999). Mol. Cell. Biol. 19, 2672–2680. Shilatifard, A. (1998a). Factors regulating the transcriptional elongation activity of RNA polymerase II. FASEB J. 12, 1437–1446. Shilatifard, A. (1998b). The RNA polymerase II general elongation complex. Biol. Chem. 379(1), 27–31. Shpakovski, G. V., and Shematorova, E. K. (1999). Characterization of the rpa43þ cDNA of Schizosacharomyces pombe: structural similarity of subunit Rpa43 of RNA polymerase I and subunit Rpe25 of RNA polymerase III. Russ. J. Bioorgan. Chem. 25(10), 700–704. Siaut, M., Zaros, C., Levivier, E., Ferri, M. L., Court, M., Werner, M., Callebaut, I., Thuriaux, P., Sentenac, A., and Conesa, C. (2003). An Rpb4/Rpb7-like complex in
40
CRAMER
yeast RNA polymerase III contains the orthologue of mammalian CGRP-RCP. Mol. Cell. Biol. 23(1), 195–205. Sidorenkov, I., Komissarova, N., and Kashlev, M. (1998). Crucial role of the RNA:DNA hybrid in the processivity of transcription. Mol. Cell 2, 55–64. Smale, S. T., Jain, A., Kaufmann, J., Emami, K. H., Lo, K., and Garraway, I. P. (1998). The initiator element: a paradigm for core promoter heterogeneity within metazoan protein-coding genes. Cold Spring Harb. Symp. Quant. Biol. 63, 21–31. Sosunov, V., Sosunova, E., Mustaev, A., Bass, I., Nikiforov, V., and Goldfarb, A. (2003). Unified two-metal mechanism of RNA synthesis and degradation by RNA polymerase. EMBO J. 22, 2234–2244. Steinmetz, E. J. (1997). Pre-mRNA processing and the CTD of RNA polymerase II: the tail that wags the dog? Cell 89(4), 491–494. Steitz, T. A. (1999). DNA polymerases: structural diversity and common mechanisms. J. Biol. Chem. 274(25), 17395–17398. Strasser, K., Masuda, S., Mason, P., Pfannstiel, J., Oppizzi, M., Rodriguez-Navarro, S., Rondon, A. G., Aguilera, A., Struhl, K., Reed, R., and Hurt, E. (2002). TREX is a conserved complex coupling transcription with messenger RNA export. Nature 417(6886), 304–308. Sweetser, D., Nonet, M., and Young, R. A. (1987). Prokaryotic and eukaryotic RNA polymerases have homologous core subunits. Proc. Natl. Acad. Sci. USA 84, 1192–1196. Szentirmay, M. N., and Sawadogo, M. (2000). Spatial organization of RNA polymerase II transcription in the nucleus. Nucleic Acids Res. 28(10), 2019–2025. Tahirov, T. H., Temiakov, D., Anikin, M., Patlan, V., McAllister, W. T., Vassylyev, D. G., and Yokoyama, S. (2002). Structure of a T7 RNA polymerase elongation complex at 2.9 resolution. Nature 420(6911), 43–50. Tan, Q., Linask, K. L., Ebright, R. H., and Woychik, N. A. (2000a). Activation mutants in yeast RNA polymerase II subunit RPB3 provide evidence for a structurally conserved surface required for activation in eukaryotes and bacteria. Genes Dev. 14(3), 339–348. Tan, Q., Linask, K. L., Ebright, R. H., and Woychik, N. A. (2000b). Activation mutants in yeast RNA polymerase II subunit RPB3 provide evidence for a structurally conserved surface required for activation in eukaryotes and bacteria. Genes Dev. 14(3), 339–348. Thomas, M. J., Platas, A. A., and Hawley, D. K. (1998). Transcriptional fidelity and proofreading by RNA polymerase II. Cell 93(4), 627–637. Thuillier, V., Brun, I., Sentenac, A., and Werner, M. (1996). Mutations in the alpha amanitin conserved domain of the largest subunit of yeast RNA polymerase III affect pausing, RNA cleavage and transcriptional transitions. EMBO J. 15, 618–629. Todone, F., Brick, P., Werner, F., Weinzierl, R. O., and Onesti, S. (2001). Structure of an archaeal homolog of the eukaryotic RNA polymerase II RPB4/RPB7 complex. Mol. Cell 8(5), 1137–1143. Todone, F., Weinzierl, R. O., Brick, P., and Onesti, S. (2000). Crystal structure of RPB5, a universal eukaryotic RNA polymerase subunit and transcription factor interaction target. Proc. Natl. Acad. Sci. USA 97(12), 6306–6310. Toulokhonov, I., Artsimovitch, I., and Landick, R. (2001). Allosteric control of RNA polymerase by a site that contacts nascent RNA hairpins. Science 292(5517), 730–733. Tsai, F. T., and Sigler, P. B. (2000). Structural basis of preinitiation complex assembly on human pol II promoters. EMBO J. 19(1), 25–36.
STRUCTURE AND FUNCTION OF RNA POLYMERASE II
41
Ujvari, A., Pal, M., and Luse, D. S. (2002). RNA polymerase II transcription complexes may become arrested if the nascent RNA is shortened to less than 50 nucleotides. J. Biol. Chem. 277(36), 32527–32537. Uptain, S. M., Kane, C. M., and Chamberlin, M. J. (1997). Basic mechanisms of transcript elongation and its regulation. Annu. Rev. Biochem. 66, 117–172. Vassylyev, D. G., Sekine, S., Laptenko, O., Lee, J., Vassylyeva, M. N., Borukhov, S., and Yokoyama, S. (2002). Crystal structure of a bacterial RNA polymerase holoenzyme at 2.6 A resolution. Nature 417(6890), 712–719. von Hippel, P. H. (1998). An integrated model of the transcription complex in elongation, termination, and editing. Science 281, 660–665. Wang, B., Jones, D. N., Kaine, B. P., and Weiss, M. A. (1998). High-resolution structure of an archaeal zinc ribbon defines a general architectural motif in eukaryotic RNA polymerases. Structure 6(5), 555–569. Wang, D., and Hawley, D. K. (1993). Identification of a 30 –50 exonucldase activity associated with human RNA polymerase II. Proc. Natl. Acad. Sci. USA 90, 843–847. Wang, W., Carey, M., and Gralla, J. D. (1992). Polymerase II promoter activation: closed complex formation and ATP-driven start site opening. Science 255, 450–452. Wang, X., Spangler, L., and Dvir, A. (2003). Promoter escape by RNA polymerase II: downstream promoter DNA is required during multiple steps of early transcription. J. Biol. Chem. 278, 10250–10256. Weilbaecher, R. G., Awrey, D. E., Edwards, A. M., and Kane, C. M. (2003). Intrinsic transcript cleavage in yeast RNA polymerase II elongation complexes. J. Biol. Chem. 278, 24189–24199. Werner, F., Eloranta, J. J., and Weinzierl, R. O. (2000). Archaeal RNA polymerase subunits F and P are bona fide homologs of eukaryotic RPB4 and RPB12. Nucleic Acids Res. 28(21), 4299–4305. West, M. L., and Corden, J. L. (1995). Construction and analysis of yeast RNA polymerase II CTD deletion and substitution mutations. Genetics 140(4), 1223–1233. Wooddell, C. I., and Burgess, R. R. (2000). Topology of yeast RNA polymerase II subunits in transcription elongation complexes studied by photoaffinity crosslinking [in process citation]. Biochemistry 39(44), 13405–13421. Woychik, N. A. (1998). Fractions to functions: RNA polymerase II thirty years later. Cold Spring Harb. Symp. Quant. Biol. 63, 311–317. Woychik, N. A., and Young, R. A. (1989). RNA polymerase II subunit RPB4 is essential for high- and low-temperature yeast cell growth. Mol. Cell. Biol. 9, 2854–2859. Yankulov, K., Blau, J., Purton, T., Roberts, S., and Bentley, D. L. (1994). Transcriptional elongation by RNA polymerase II is stimulated by transactivators. Cell 77(5), 749–759. Yin, Y. W., and Steitz, T. A. (2002). Structural basis for the transition from initiation to elongation transcription in T7 RNA polymerase. Science 298(5597), 1387–1395. Yuan, X., Zhao, J., Zentgraf, H., Hoffmann-Rohrer, U., and Grummt, I. (2002). Multiple interactions between RNA polymerase I, TIF-1A and TAFI subunits regulate preinitiation complex assembly at the ribosomal gene promoter. EMBO Rep. 3(11), 1082–1087. Yudkovsky, N., Ranish, J. A., and Hahn, S. (2000). A transcription reinitiation intermediate that is stabilized by activator. Nature 408(6809), 225–229. Zaychikov, E., Martin, E., Denissova, L., Kozlov, M., Markovtsov, V., Kashlev, M., Heumann, H., Nikiforov, V., Goldfarb, A., and Mustaev, A. (1996). Mapping of catalytic residues in the RNA polymerase active center. Science 273, 107–109.
42
CRAMER
Zhang, G., Campbell, E. A., Minakhin, L., Richter, C., Severinov, K., and Darst, S. A. (1999). Crystal structure of Thermus aquaticus core RNA polymerase at 3.3 A˚ resolution [see comments]. Cell 98(6), 811–824. Zhang, G., and Darst, S. A. (1998). Structure of the Escherichia coli RNA polymerase alpha subunit amino-terminal domain. Science 281(5374), 262–266. Zhang, J., and Corden, J. L. (1991). Phosphorylation causes a conformational change in the carboxyl-terminal domain of the mouse RNA polymerase II largest subunit. J. Biol. Chem. 266, 2297–2302.
THE MEDIATOR COMPLEX ¨ RKLUND* AND CLAES M. GUSTAFSSONÀ By STEFAN BJO
À
*Department of Medical Biochemistry, Umea˚ University, S-901 87 Umea˚, Sweden; Department of Medical Nutrition, Karolinska Institute, Novum, S-141 86 Huddinge, Sweden
I. Summary. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . II. Saccharomyces cerevisiae Mediator. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . A. Identification of S. cerevisiae Mediator . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . B. Interactions with RNA Polymerase . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . C. Subunit Composition of Yeast Mediator. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . D. Global Gene Regulation. . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . III. Mediator Complexes in Higher Eukaryotes. . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . A. Identification of Mammalian Mediator . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . B. Functional Studies of Metazoan Subunits . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . IV. Mechanism of Transcriptional Activation . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . A. Role of the RNA Polymerase II C-Terminal Domain . . . . . . . . . . . . . . . . . . . .. . . . . . . B. Structure–Function. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . V. Concluding Remarks . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . .
43 43 44 44 46 51 52 53 54 55 58 58 62 62
I. Summary The Mediator complex acts as a bridge, conveying regulatory information from enhancers and other control elements to the general transcription machinery. The Mediator was originally identified in Saccharomyces cerevisiae and is required for the basal and regulated expression of nearly all RNA Pol II–dependent genes. Mediator complexes were recently identified also in metazoans, confirming a role for Mediator in transcription regulation in higher eukaryotes as well. In spite of its general significance for transcription control, the exact mechanisms of Mediator function remain unclear. We here review our understanding of the structure and possible models for the function of Mediator in yeast and metazoan cells.
II. Saccharomyces cerevisiae Mediator RNA polymerase II (RNA Pol II)–dependent transcription initiation supposedly proceeds in two stages. First there is a relief of repression by remodeling of chromatin structure at the promoter. This step is dependent on the activity of chromatin modifying or remodeling complexes (Urnov and Wolffe, 2001), which are recruited to specific promoters by regulatory proteins. Second, after remodeling of the promoter, a preinitiation complex containing RNA Pol II and the general transcription 43 ADVANCES IN PROTEIN CHEMISTRY, Vol. 67
Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00
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factors (GTFs) TFIIB, TFIID, TFIIE, TFIIF, and TFIIH is formed. At this stage, activators recruit GTFs and stimulate the assembly of the preinitiation complex on to the promoter. However, direct interactions between different activators and general transcription factors do not seem to be sufficient for transcription activation, as activators fail to stimulate transcription in systems reconstituted from pure RNA Pol II, basal factors, and purified template DNA.
A. Identification of S. cerevisiae Mediator In a search for a factor that could enable a response to transcriptional activators in a pure in vitro transcription system, R. D. Kornberg and colleagues isolated an activity from S. cerevisiae that was termed Mediator (Flanagan et al., 1991; Kelleher et al., 1990). The assay used was based on naked DNA templates and thus reflects the second stage of the transcription initiation process described above. The Mediator activity was purified to homogeneity and shown to be a holoenzyme form of RNA Pol II, made up of core 12-subunit RNA Pol II and a Mediator complex (Kim et al., 1994). Mediator was later also isolated as a discrete entity and identified as a multiprotein complex of 20 individual polypeptides (Table I; Myers et al., 1998). The functional activities identified for the Mediator were stimulation of basal transcription, support of activated transcription, and enhancement of phosphorylation of RNA Pol II by TFIIH kinase (Kim et al., 1994; Myers et al., 1998). Later studies also identified a histone acetyltransferase activity in the S. cerevisiae Mediator (Lorch et al., 2000).
B. Interactions with RNA Polymerase The C-terminal domain (CTD) of the largest subunit in RNA Pol II plays an important role in the function of Mediator (Myers and Kornberg, 2000). The domain, which consists of multiple heptapeptide repeats of the sequence Tyr-Ser-Pro-Thr-Ser-Pro-Ser, is conserved in all eukaryotes studied to date. In S. cerevisiae, CTD truncations cause defects in transcriptional activation both in vivo and in vitro (Scafe et al., 1990). Two distinct forms of RNA pol II have been identified in S. cerevisiae. Most RNA Pol II molecules have an unphosphorylated CTD, but a portion of RNA Pol II molecules is highly phosphorylated. The unphosphorylated RNA Pol II associates with the promoter-bound initiation complex, whereas the phosphorylated form is responsible for active elongation (Cadena and Dahmus, 1987). The principal protein kinase involved in the phosphorylation of CTD has been identified as Kin28, a cyclin dependent kinase (cdk) and subunit of
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Table I Mediator Subunits in S. cerevisiae and Their Homologues in Saccharomyces pombe and Human Cells Saccharomyces cerevisiae Gene deletion phenotype
Protein mass (kD)
Nut1
Conditional
129
Gal11 Rgr1 Sin4 Srb4 Med1 Med2 Pgd1/Hrs1 Med4 Med6 Srb5 Med7 Med8 Rox3 Srb2 Nut2 Cse2 Srb7 Srb6 Med11 Srb8 Srb9 Srb10
Conditional Inviable Conditional Inviable Conditional Conditional Conditional Inviable Inviable Conditional Inviable Inviable Inviable Conditional Inviable Conditional Inviable Inviable Inviable Conditional Conditional Conditional
120 123 111 78 64 48 47 32 32 34 32 25 25 23 18 17 16 14 15 167 160 63
Srb11
Conditional
38
Subunit
Activity
S. pombe subunit
Human subunita
Pmc1
Med150
spSrb4
Med78
spMed4 spMed6
Med36 Med33
spMed7 spMed8 spRox3
Med34 Arc32b
spNut2
Med10
spSrb7 spSrb6
Med17
spSrb8
Med230
spSrb10
Cdk8
spSrb11
CyclinC
Histone acetyltransferase
Cyclin-dependent protein kinase Cyclin
a
We here use the nomenclature proposed by Rachez and Freeman (2001). The Arc32 protein has so far only been identified in the ARC complex (Naar et al., 1999). b
the general transcription factor TFIIH (Feaver et al., 1994). It is generally believed that Kin28-dependent phosphorylation of the CTD leads to a breakdown of the preinitiation complex and the transition from transcription initiation to elongation (Svejstrup et al., 1997). One of the cardinal activities of Mediator is its ability to stimulate phosphorylation of CTD by TFIIH (Kim et al., 1994; Myers et al., 1998). The level of stimulation can be more than 40-fold and is specific for the
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Kin28 kinase. The molecular basis for Mediator’s ability to stimulate CTD phosphorylation is unknown. However, the observation that Saccharomyces pombe Mediator is unable to stimulate TFIIH derived from S. cerevisiae indicates that specific interactions are formed between Mediator and TFIIH (Spahr et al., 2000).
C. Subunit Composition of Yeast Mediator The majority of genes encoding the yeast Mediator subunits had previously been identified in genetic screens for mutations affecting activation and repression of transcription. The presence of these well-characterized gene products in one single complex connected Mediator with a quartercentury of genetic analysis in yeast and at once established the relevance of Mediator function in vivo.
1. Srb Proteins As mentioned previously, CTD truncations cause defects in transcriptional activation both in vivo and in vitro (Scafe et al., 1990). The nine SRB genes were originally identified by R. A. Young and colleagues in a genetic screen for suppressors of RNA polymerase IIB, a version of the polymerase carrying only 11 instead of the normal 26–27 heptapeptide repeats in the CTD of the largest RNA Pol II subunit (Nonet and Young, 1989). The Srb proteins were later isolated in a complex with RNA Pol II, giving the first indication of the existence of an RNA Pol II holoenzyme (Koleske and Young, 1994; Thompson et al., 1993). Five of the SRB genes, SRB2, SRB4, SRB5, SRB6, and SRB7, encode core Mediator subunits, which are present in all Mediator preparations. Proteins Srb2, Srb4, Srb5, and Srb6 have been shown to interact in a subcomplex of Mediator together with Med6 and Rox3 (Lee and Kim, 1998). The Srb4, Srb6, and Srb7 proteins are all encoded by essential genes, and a temperature-sensitive(ts) mutation in the SRB4 gene shuts down nearly all RNA Pol II–dependent transcription at the nonpermissive temperature (Thompson and Young, 1995). SRB2 and SRB5 are nonessential genes with a slow growth phenotype (Nonet and Young, 1989; Thompson et al., 1993). SRB5 is needed for expression of genes involved in the pheromone response pathway, which is reflected in a defect in mating efficiency for srb5 cells (Holstege et al., 1998). A subgroup of Srb proteins (Srb8, Srb9, Srb10, and Srb11) forms a specific module that is present in holoenzyme preparations from cells growing exponentially in rich glucose medium, but is absent in stationary-phase
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cells (Hengartner et al., 1998). The SRB11 and SRB10 encode cyclin C and the cyclin C-dependent kinase, respectively (Liao et al., 1995). Genetic analysis indicates that the Srb8-11 module is involved in the negative regulation of a small subset of genes (Holstege et al., 1998). Srb8 is required for stable association of Srb10 and Srb11 with the holoenzyme inasmuch as holoenzyme preparations from Srb8 deletion strains lack Srb10 and Srb11 (Myer and Young, 1998). Homologues to Srb10 and Srb11 are found in some human Mediator preparations. Alleles of SRB8, SRB9, SRB10, and SRB11 have been identified as ssn (suppressors of snf1) mutations (Song et al., 1996). Two other genes encoding mediator subunits, ROX3 and SIN4, were also identified in the same genetic screen. The Snf1 kinase is a homologue of the mammalian AMP-activated protein kinase and is inactive in the presence of glucose (Woods et al., 1994). Snf1 functions by inactivating the Mig1 repressor, which binds to promoters of many glucose-repressed genes and recruits the Ssn6-Tup1 corepressor complex (Treitel and Carlson, 1995). Direct interactions have been demonstrated between Tup1 and the N-terminal domain of the Mediator subunit Srb7 (Gromoller and Lehming, 2000). Interestingly, Tup1 interaction with Srb7 precludes interaction between Srb7 and another Mediator subunit, Med6. The Srb7–Med6 contacts are believed to be part of a pathway that relays positive signals within Mediator, and the inhibition of this pathway could, therefore, explain the repressive activity of Tup1’s repressive activity. Cells lacking Snf1 cannot grow on any carbon source except glucose. Cells lacking both Snf1 and Mig1 can also grow on galactose and sucrose, but are still unable to grow on gluconeogenic carbon sources. Thus, some genes that are required for gluconeogenic growth are repressed by Mig1-independent mechanisms that operate downstream of Snf1. The SRB genes appear to also be involved in the Mig1-independent repression, as spontaneous mutations that allow snf1/mig1 cells to grow on gluconeogenic carbon sources have been identified in SRB8, SRB10, and SRB11 (Balciunas and Ronne, 1995). Even if the Srb8-11 module has mostly been implicated in negative regulation of transcription, it also appears to have a positive effect on some genes. When yeast cells enter stationary phase in response to certain types of nutrient limitations, there is a down-regulation of most RNA Pol II transcribed genes. However, the expression of some genes, such as YGP1, is induced under these conditions. In a genetic screen for mutants that are defective in the regulation of YGP1 expression (rye), Herman and colleagues showed that three of the RYE genes encode Srb9, Srb10, and Srb11 (Chang et al., 2001).
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2. Med Proteins The MED genes were grouped together because they were encoded by previously uncharacterized genes and their products were identified through peptide sequencing (Kim et al., 1994; Myers et al., 1998). Some of the Med proteins appear to be specific for S. cerevisiae, such as Med1, Med2, and Med11, whereas others like Med4, Med6, Med7, and Med8 are conserved in all Mediator-like complexes isolated to date (Table I). Deletion of MED1 causes a complex phenotype similar to mutations in SRB10 and SRB11, including suppression of snf1 (Balciunas et al., 1999). A MED1 deletion strain displays a partial loss of glucose repression and a slightly impaired induction of galactose-regulated genes. In contrast to many other Mediator subunits, the Med1 protein fused to the DNA-binding domain of LexA does not function as an activator in wild-type cells. However, LexA-Med1 fusion is a strong activator (400-fold) when expressed in an srb8, srb10, or srb11 deletion strain (Balciunas et al., 2003). It thus appears as if the Srb10-Srb11 cyclin kinase complex negatively regulates the function of Med1. Med2 forms a stable submodule with Hrs1/Pgd1 and Gal11 (Lee et al., 1999; Myers et al., 1999). A deletion of one of the genes encoding for these proteins will lead to a concomitant loss of all three proteins from the Mediator complex. Whole-genome analysis of MED2-dependent transcription indicates that the expression of about 200 genes is significantly decreased in med2 cells and, specifically, induction of several GAL genes was found to be defective in the med2 strain (Myers et al., 1999). However, it seems as it this Gal phenotype is caused by a delay in induction rather than a reduction in galactokinase levels (Balciunas et al., 1999). Med4 is an essential gene with weak sequence homology to the human Mediator subunit Trap36 (Myers et al., 1998; Spahr et al., 2001). So far, no genetic studies involving Med4 have been presented. Med6 is an essential subunit with highly conserved homologues in all Mediator complexes studied to date (Lee et al., 1997). A ts mutation in S. cerevisiae MED6 showed defects in activation of several inducible promoters, but no effect on uninduced or constitutively expressed genes. The same pattern has also been observed for the Caenorhabditis elegans homologue, which is required for developmental stage-specific transcriptional regulation but dispensable for the expression of two constitutively expressed genes tested (Kwon et al., 1999). The effect on inducible yeast promoters is specific and coupled with certain classes of transcriptional activators. No effect was observed for GCN4-regulated genes, whereas both GAL4- and MAT1-regulated genes require Med6 for activation. However, deletion of MED6 does not affect the interaction between activators and
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Mediator, and thus points to a function of Med6 at a stage subsequent to recruitment of Mediator to promoters (Lee et al., 1999). S. cerevisiae cells lacking MED8 are inviable, but the function of Med8 is ambiguous (Myers et al., 1998). It was recently reported that Med8 binds directly to control elements in the invertase (SUC2) and hexokinase 2 (HXK2) promoters (Chaves et al., 1999). However, it is not clear whether this interaction involves the entire Mediator complex or merely the free Med8 protein. In S. pombe, Med8 has been identified as SEP15 in a genetic screen for mutants defective in cell separation (Zilahi et al., 2000). It is still possible that this effect is indirect, as the penetrance of the SEP15 mutation is incomplete in cell separation. Med11 is an essential gene required for MF1 transcription (Han et al., 1999).
3. Nut1 and Nut2 Proteins HO transcription is dependent on Swi4p and Swi6p for relief of repression by the URS2 region upstream of the HO promoter. NUT1 and NUT2 were, together with SIN4, ROX3, SRB8, SRB9, SRB10, and SRB11, originally isolated in a screen for mutants that would suppress the Swi4p/Swi6p dependence of a synthetic reporter gene containing part of URS2 (Tabtiang and Herskowitz, 1998). Nut1 appears to be specific to S. cerevisiae, whereas homologs to the essential Nut2 protein have been identified in all eukaryotic Mediator complexes isolated to date (Table I). Nut1 has been demonstrated to have histone acetyltransferase (HAT) activity, and purified Mediator can interact directly with free nucleosomes (Lorch et al., 2000). The exact role for the Nut1 HAT activity in Mediator function remains to be established.
4. Rox3 Protein The ROX3 gene, which is essential, was found in a search for mutants leading to overexpression of the heme-regulated CYC7 gene and was later also identified as SSN7 (Rosenblum-Vos et al., 1991; Song et al., 1996). ROX3 is also synonymous with RMR1, whose mutation can relieve glucose repression of the CYB2 gene (Brown et al., 1995). ROX3 does not only play a role in repression, as it is needed for full induction of the GAL1 gene in the presence of galactose (Brown et al., 1995).
5. Gal11, Sin4, and Rgr1 Proteins Gal11 was first described as an auxiliary transcription activator for genes encoding galactose-metabolizing enzymes (Suzuki et al., 1988). It has also been implicated in enhancement of basal transcription (Sakurai et al., 1993), in negative regulation of the activity of the MCM1 transcription
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factor in Ty1 elements (Yu and Fassler, 1993), as an SNF gene (Vallier and Carlson, 1991), and as being involved in regulation of the structure and the position effect of telomeres (Suzuki and Nishizawa, 1994). However, the identification of Gal11 as a Mediator subunit has now led to a model in which Gal11 is part of a subcomplex of the Mediator that also includes Med2, Sin4, Pgd1, and possibly Nut1. This so-called Sin4 (or Gal11) module has been shown to be essential for the response to acidic transcriptional activator proteins such as Gal4VP16 (Lee et al., 1999; Myers et al., 1999). Gal11 interacts directly with the general transcription factors TFIIE and TFIIH (Sakurai et al., 1996), and a deletion of GAL11, MED2, or PGD1 causes synthetic lethality in combination with mutations in the large subunit of TFIIE. In addition, a ts mutation in KIN28, which encodes the kinase subunit of TFIIH, is lethal in a gal11 background (Sakurai and Fukasawa, 2000). RGR1 was isolated as a negative regulator of SUC2 (Sakai et al., 1990) but has also been identified as a negative regulator of the HO gene (Stillman et al., 1994). RGR1 is an essential gene, and an rgr1 strain shows pleiotropic effects such as resistance to glucose repression, ts lethality, sporulation deficiency in homozygous diploid cells, and abnormal cell morphology. SIN4, however, was identified as a negative regulator of GAL1 gene transcription, and it was also suggested that Sin4 alters chromatin structure in a way that affects transcriptional regulation ( Jiang and Stillman, 1992). However, several lines of evidence indicate that Rgr1 and Sin4 participate in the same regulatory pathways. RGR1 and SIN4 are negative transcriptional regulators of HO and IME1, and sin4 or rgr1 mutations have phenotypes similar to those caused by histone mutations, thus indicating that they act together in vivo to organize chromatin structure and to regulate transcription (Covitz et al., 1994; Jiang and Stillman, 1995; Stillman et al., 1994). These genetic interactions were confirmed biochemically in experiments in which the N-terminal domain of Rgr1 was shown to be important for the interaction between the Sin4 module and the rest of Mediator (Li et al., 1995).
6. Cse2 Protein Mutations in CSE1 and CSE2 lead to defects in chromosome segregation (CSE); (Xiao et al., 1993). CSE1 is an essential gene, whereas disruption of CSE2 causes chromosome missegregation, conditional lethality, and slow growth. Both Cse1 and Cse2 have been shown to interact physically with components of Mediator, using a high-throughput yeast two-hybrid system. Cse1 was shown to interact directly with Sin4, and Cse2 was identified as interacting directly with Med4 and indirectly, via Med4, to
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Srb7 and Med7. However, only Cse2 has been identified biochemically as a Mediator subunit (Gustafsson et al., 1998; Han et al., 1999). CSE2 encodes a 17-kDa protein that contains a putative leucine zipper motif, indicating that it may possess a DNA binding activity. CSE2 is required for Bas1/Bas2mediated basal transcription of amino acid biosynthetic genes, and holopolymerase isolated from cells lacking CSE2 display a 50% reduction in basal, nonregulated transcription (Han et al., 1999). On the basis of these results, it seems likely that the connection between CSE2 and chromosome segregation is indirect and involves transcription.
7. Soh1 Protein A human homologue of the yeast Soh1 protein has been identified as a subunit of the human mediator-like complexes TRAP, SMCC, and NC2 (Gu et al., 1999; Malik and Roeder, 2000), but Soh1 has not been demonstrated as a yeast Mediator subunit. The soh1, soh2, and soh4 mutants were isolated as suppressors of the temperature-dependent growth of the hyperrecombination mutant hpr1 (Fan et al., 1996). However, cloning of the corresponding genes indicates an involvement in RNA Pol II transcription. Soh2 is identical to the second-largest subunit of RNA Pol II, and Soh4 was identified as TFIIB. SOH1 encodes a novel 14-kD protein with limited sequence similarity to RNA polymerases. Mutations in SOH1 are synthetically lethal with mutations in RNA Pol II subunits and mutations in SUA7, which encodes yeast TFIIB.
D. Global Gene Regulation It is evident from the genetic characterization of Mediator that the general requirement for individual Mediator subunits in gene regulation will differ significantly. DNA microarray analysis of global gene expression supports this notion. Some Mediator components are needed for the regulated expression of nearly all genes, whereas others are only needed for a certain subset of genes (Holstege et al., 1998). The Srb4 ts strain demonstrates a decrease in the expression of 93% of all S. cerevisiae genes at the nonpermissive temperature. This value corresponds closely to that observed with a ts mutant in the largest subunit of RNA Pol II. In addition, the set of genes whose mRNAs are not significantly reduced in the RNA Pol II ts mutant exhibit the same behavior in the Srb4 ts experiment. The results indicate that genome-wide expression is as dependent on Srb4 as it is on core RNA Pol II, and that the Srb4-containing RNA Pol II holoenzyme is generally required for transcription. However, there are exceptions to the rule, and a small number of genes can indeed be expressed
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independently of Srb4, for example, CUP1 and SSN2 (Lee and Lis, 1998; McNeil et al., 1998). Interestingly, expression of the same subset of genes is also unaffected in cells lacking Kin28. This observation supports the notion that the function of Mediator is dependent of Kin28 and that regulation of TFIIH kinase activity is an essential part of Mediator’s ability to govern transcription in vivo. For most Mediator components, the effects on global gene expression are far less dramatic. Med6 is needed for expression of approximately 10% of all genes in the yeast genome, whereas about 16% of all genes are dependent on Srb5 function (Holstege et al., 1998). It should be noted that it is often difficult to distinguish between results that are a direct consequence of the loss of a specific Mediator subunit and those that are the result of a secondary effect. For example, global genome analysis has demonstrated a role for Med2 in the regulation of galactose inducible genes (Myers et al., 1999). However, as discussed above, Mediator purified from a med2 strain also lacks the Hrs1/Pgd1 and Gal11 proteins. Provided that the Hrs1/Pgd1 and Gal11 are also absent from Mediator in vivo, it would be impossible to distinguish the effects of a MED2 on global gene expression from the effects caused by deletion of HRS1/PGD1 and GAL11.
III. Mediator Complexes in Higher Eukaryotes Initially, it was unclear whether Mediator was specific for yeast or whether it had a counterpart in metazoan cells. The general view was that activators contacted TBP-associated factors, (TAFs), which in turn recruited TBP (TATA-binding protein) and subsequently other GTFs to specific promoters. This view was challenged by genetic studies in yeast, showing that TAFs are not required for transcriptional activation but, rather, contribute to the specificity of TBP-promoter interaction (Shen and Green, 1997). The first experimental indications of a metazoan Mediator complex came in 1996, when R. G. Roeder and coworkers isolated the multisubunit thyroid hormone receptor coactivator complex (TRAP), later identified as human Mediator (see Section III,A); (Fondell et al., 1996). The same year, R. A. Young and colleagues identified a human homologue to the Srb7 protein as a part of a larger RNA Pol II containing complex (Chao et al., 1996). Later, mammalian multiprotein complexes containing homologues of yeast Mediator proteins were identified in six laboratories (Boyer et al., 1998; Gu et al., 1999; Jiang et al., 1998; Naar et al., 1999; Rachez et al., 1999; Ryu et al., 1999; Sun et al., 1998). Similar to previous findings in yeast, human Mediator was shown to support activation in a fully reconstituted transcription system in the absence of TAFs (Oelgeschlager et al., 1998). These findings established Mediator as a major conduit of regulatory
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information from regulatory DNA elements to promoters across the whole range of eukaryotes, from yeast to man.
A. Identification of Mammalian Mediator The general importance of Mediator for both activated and repressed transcription in mammalian cells is also reflected in the multitude of ways the human Mediator complex was identified. The TRAP defined on p. 14 was purified by Roeder and coworkers as a complex associated with human thyroid hormone receptor alpha purified from HeLa cells grown in the presence of thyroid hormone (T3; Fondell et al., 1996). TRAP could also support activation of transcription in vitro from a promoter template containing T3-response elements. In parallel, the same laboratory used HeLa-derived cell lines expressing epitope-tagged hSrb7, hSrb10, or hSrb11 to identify a similar complex called SRB/MED-containing cofactor complex (SMCC; Gu et al., 1999). TRAP and SMCC have been shown to be identical (Ito et al., 1999). The DRIP (Vitamin D3 receptor [VDR] interacting proteins) complex was purified using a VDR ligand-binding domain affinity matrix (Rachez et al., 1998). DRIP is needed for full transcriptional activity of VDR on naked DNA templates in vitro. Another complex, ARC (activator-recruited cofactor), was identified as a complex that enhances transcription activation by SREBP-la, VP16, and the p65 subunit of NF-kappaB using chromatin-assembled DNA templates (Naar et al., 1999). Characterization of the subunits of DRIP and ARC showed that the two complexes are highly related—if not identical—to each other and also to the TRAP/SMCC complexes (Rachez et al., 1999). A role for Mediator in the transcription activation program that is initiated by viral infections of mammalian cells was revealed when Berk and colleagues identified a human homologue to the C. elegans Sur-2 protein as an in vivo target for the adenovirus E1A protein (Boyer et al., 1999). Further purification identified Sur-2 as a member of a mulitprotein complex containing human homologues to yeast Mediator proteins. This human Mediator could also support activation by Gal4-E1A as well as Gal4VP16 in a defined in vitro transcription system. TRAP/SMCC, DRIP, ARC, and human Mediator are virtually identical in their subunit composition (Malik and Roeder, 2000). Another set of Mediator-like complexes has also been isolated that appear to correspond to a submodule of the larger Mediator. This has led to speculations that the Mediator might exist in two separate forms. One of these smaller complexes is PC2, a component of the coactivator fraction USA (Malik et al.,
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2000; Meisterernst et al., 1991). PC2 can support activated transcription in vitro, but only in the presence of two other cofactors, PC3/topoisomerase I and PC4. Two other small Mediator complexes, CRSP (cofactor required for Sp1) and mouse Mediator, were both identified using a biochemical fractionation ( Jiang et al., 1998; Ryu et al., 1999). Interestingly, CRSP and PC2 could only support activation in the presence of TAFs (Malik et al., 2000; Ryu et al., 1999), This could indicate a yet-to-be-defined functional relationship between TAFs and the smaller form of Mediator. Another small Mediator complex is NAT (negative regulator of activated transcription). In contrast to the other Mediator complexes, NAT displayed an inhibitory effect on transcription in vitro (Sun et al., 1998). It is unclear whether the complexes described above represent distinct functional entities or whether the differences that exist between them are consequences of different purification methods. This was recently studied in experiments in which HeLa cell nuclear extracts were resolved directly by gel filtration (Wang et al., 2001). Only one peak of human Mediator, stoichiometric to the levels of GTFs in HeLa cells, was detected and revealed that the human Mediator had a molecular weight of about 2 MDa. This indicates that the smaller-sized complexes (CRSP, mouse mediator, PC2, and NAT) are either subcomplexes of the larger complexes (TRAP/SMCC, NAT, DRIP, ARC, and human Mediator) formed by dissociation of the larger complexes during fractionation or are much less abundant. It was also found that the 2-MDa human mediator is present in two forms of indistinguishable size, one containing and one lacking the Srb10/Srb11 CDK-cyclin pair. Recently, a smaller form of the Mediator was also identified in S. cerevisiae nuclear extracts and termed Medc (Liu et al., 2000). This complex contains all the subunits of Mediator with the exception of Rgr1, Rox3, Nut1, and the Sin4 module. Medc is less abundant than Mediator and is also less active in transcription. The functional role of Medc remains to be established, but it could lend biochemical support for the existence of two forms of the Mediator complex not only in higher eukaryotes but also in yeast.
B. Functional Studies of Metazoan Subunits The function of individual metazoan Mediator subunits has been studied by different methods; that is, by RNA interference (RNAi) and chemical mutagenesis in C. elegans, P insertions in Drosophila, homologous recombination in mouse, and studying spontaneous mutations in human cells (Ito et al., 2000; Kwon et al., 1999; Nilsson et al., 2000; Philibert et al., 1998; Singh and Han, 1995; Spradling et al., 1999; Tudor et al., 1999; Zhu et al., 2000).
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RNAi experiments in C. elegans showed that Med6, Med7, and Nut2 are essential for embryogenesis (Kwon et al., 1999). Interestingly, Med6, Med7, and Nut2 were found to be required for expression of two developmentally regulated genes but dispensable for expression of two ubiquitously expressed genes. RNAi experiments also show that the C. elegans homolog of Rgr1 is required early in embryogenesis (S. Tuck, personal communication). In contrast, worms homozygous for putative null mutations in the C. elegans Med130/sur-2 are viable and fertile (Singh and Han, 1995), as are worms with reducedMed230/sop-1 activity (Zhang and Emmons, 2000). sop-1(RNAi) worms are also viable, but it is not known whether such worms completely lack sop-1 activity. The sur-2 gene product appears to have a role in multiple developmental stages, operating downstream of Ras and MAP Kinase (Nilsson et al., 2000; Singh and Han, 1995). Animals with reduced sop-1 activity can bypass the requirement for PAL-1 (a homeobox protein) for neurogenesis in the male tail (Zhang and Emmons, 2000). In wild-type animals, sop-1 is thought to be a repressor of Wnt signaling. Two Mediator subunits, Med220/TRAP220 and Srb7, have been studied via inactivation of the corresponding genes in mice. In the Srb7 study, heterozygous ES cells and animals showed no phenotype (Tudor et al., 1999). However, no homozygous ES cells could be obtained, and homozygous embryos were only found up to the blastocyst stage, thus indicating that the Srb7 gene product is essential for both embryonic development and cell viability. Trap220 þ/ mice were fertile and phenotypically normal except for being slightly smaller compared with their Trap220 þ/þ littermates as a result of pituitary hypothyroidism (Ito et al., 2000; Zhu et al., 2000). However, Trap220 / embryos died at embryonic day 11 with defects in development of the central nervous system, cardiac and large vessel enlargement, and defects in placental vasculature. Finally, in Drosophila melanogaster, a mutation generated by P insertion demonstrated an essential function also for the Med78 subunit (Spradling et al., 1999).
IV. Mechanism of Transcriptional Activation The molecular mechanism for Mediator-dependent transcriptional activation is still not completely understood. Specific interactions have been demonstrated between various activators and Mediator subunits in both S. cerevisiae and mammalian Mediator. In yeast, the VP16, Gal4, and Gcn4 proteins all interact directly with Gal11, and in the case of Gcn4, additional contacts are made with Hrs1/Pgd1 (Park et al., 2000). Specific interactions have also been reported between the Gal4 activation domain
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and Srb4 (Koh et al., 1998). Direct interactions with mammalian Mediator have been demonstrated for a number of nuclear hormone receptors. These include TR, VDR, retinoic acid receptor (RAR), retinoic X receptor , peroxisome proliferator-activated receptor , proliferatoractivated receptor , estrogen receptor , and glucocorticoid receptor (Hittelman et al., 1999; Yuan et al., 1998; Zhu et al., 1997). Many of the receptors seem to interact with two closely located nuclear receptor (NR) interaction boxes (LXXLL motifs) in the Med220/TRAP220 subunit of mammalian Mediator. A number of other interacting partners (i.e., p160, p300CBP, pCAF/SAGA, and SWI/SNF) have also been identified for the NR family (Aalfs and Kingston, 2000; Lemon and Freedman, 1999). All these coactivator complexes possess chromatin modifying or remodelling activities, whereas Mediator is supposed to operate mainly on the basal transcription machinery. A model has been proposed in which unliganded NRs initially function by binding to their target sites in chromatin in complex with different corepressor complexes. Binding of ligand to promoter-bound NRs leads to an exchange of NR-bound factors from corepressors to chromatin-remodeling coactivators. The remodeling of the promoter sequence surrounding the NR binding site then leads to recruitment of Mediator and subsequent or concomitant formation of a functional preinitiation complex. Given the specific interactions described between activators and Mediator, as well as the gene-specific effects observed for individual Mediator subunits, it seems likely that recruitment of RNA Pol II to the preinitiation complex plays an important role for Mediator function. This notion is supported by the finding in yeast that LexA-fusions to many individual Mediator subunits strongly activate transcription from a reporter containing LexA-binding sites 50 to the promoter (Balciunas et al., 1999; Song et al., 1996). However, the recruitment model does not take into account the specific genetic, functional, and physical interactions demonstrated between Mediator and CTD. A model to explain these interactions has been proposed (Svejstrup et al., 1997). It is based on the observation that RNA Pol II engaged in active transcription lacks associated Mediator. Formation of the preinitiation complex is dependent on the holoenzyme form of RNA Pol II, but Mediator is then released at the end of initiation or early in RNA chain elongation, as shown by its absence from the transcribing polymerase. During the initiation of transcription, Mediator stimulates CTD phosphorylation by TFIIH. Because Mediator is unable to bind to the hyperphosphorylated form of RNA Pol II, this eventually leads to dissociation of RNA Pol II from the preinitiation complex as transcriptional elongation begins. After completing a round of transcription, the CTD is dephosphorylated by
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a CTD-phosphatase. The unphosphorylated RNA Pol II can finally enter a new cycle of transcription by interacting with Mediator. This model recently gained support from results obtained by Hahn and collaborators (Yudkovsky et al., 2000). Using an immobilized template assay, they demonstrated that Mediator remains at the promoter after transcriptional initiation (Yudkovsky et al., 2000). Mediator forms a scaffold together with TFIID, TFIIA, TFIIH, and TFIIE that facilitates reinitiation of transcription from the promoter. Interestingly, the scaffold is stabilized in the presence of certain activators, for example, Gal4-VP16, immediately suggesting one possible mechanism for Mediator-dependent transcriptional activation. The question of how RNA Pol II is recruited to a promoter on activation was recently addressed in two independent systems (Cosma et al., 2001; Park et al., 2001). In the first paper, the ordered binding of factors to the HO promoter was studied using chromatin immunoprecipitation. Activation of HO is initiated in late mitosis by inactivation of the Cdk1 kinase via anaphase-promoting complex–mediated proteolysis of its B-type cyclin partners. This leads to translocation of the Swi5 transcription factor from the cytoplasm to the nucleus where it recruits the Swi/Snf chromatinremodeling complex to the HO promoter. The promoter-bound Swi/Snf then recruits the SAGA HAT complex to the promoter. Remodeling of the HO promoter permits binding of the transcriptional activator SBF, which is essential for activation of HO. Although these initial steps were well described previously, the function of SBF in the final steps of HO-activation has been unclear. However, the chromatin immunoprecipitation experiments clearly show that SBF functions in two steps. First, SBF recruits Mediator, but not RNA Pol II or GTFs, to the promoter by a mechanism that is independent of Cdk1. Activation of HO by recruitment of RNA Pol II and GTFs does not occur until the G1 phase of the cell cycle, when Cdk1 is activated by binding to the G1 cyclins. The target for Cdk1 in this process in so far unknown. A similar stepwise mechanism for transcriptional activation has also been proposed on the basis of studies of recruitment of the transcriptional activator HSF, Mediator, and RNA Pol II to the heat shock promoter of Drosophila polythene chromosomes (Park et al., 2001). Using different techniques, it was observed that on heat shock, both HSF and Mediator are rapidly recruited to the hsp70 promoter in a manner that is independent of the presence of a core promoter. This recruitment was not accompanied with a corresponding increase in RNA Pol II or GTFs and was also independent of the presence of the RNA Pol II inhibitor -amanitine. The results above are in line with the results in yeast discussed earlier, demonstrating that recruitment of holoenzyme is not needed for each round of transcription (Yudkovsky et al., 2000). Rather, the
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Mediator–RNA Pol II interaction is dynamic, and both Mediator and several GTFs remain at the promoter after release of the polymerase and can function as a scaffold for reinitiation by polymerase devoid of Mediator and GTFs.
A. Role of the RNA Polymerase II C-Terminal Domain The role of CTD for metazoan Mediator is still controversial. The S. cerevisiae Mediator interacts directly with CTD and needs CTD to stimulate basal transcription and support transcriptional activation (Myers et al., 1998). In contrast, in vitro studies of human Mediator have demonstrated activated transcription using a CTD-less polymerase (Gu et al., 1999). The molecular basis for the observed differences remains unclear. However, even if direct interactions have been demonstrated between CTD and Mediator, structural studies indicate that the most pronounced contacts are CTD independent (Asturias et al., 1999). It is therefore possible that Mediator may recruit RNA Pol II in a CTD independent fashion in vitro. RNA Pol II used by Mediator in the cell will undoubtedly contain an intact CTD. So what is then the molecular function of CTD? We favor a model in which the major role of CTD is to break the interaction between Mediator and RNA Pol II on CTD hyper-phosphorylation rather than being essential for formation of a Mediator-RNA Pol II complex. In support of this view, Reinberg and collaborators have demonstrated preferential binding of the human Mediator (NAT complex) to unphosphorylated CTD over phosphorylated CTD (Sun et al., 1998). In this respect it could be of interest to investigate the properties of Mediator isolated from Srb mutant strains, which suppress the mutant phenotypes of a truncated CTD in vivo. Perhaps the srb mutants weaken the CTD independent interactions formed between Mediator and RNA Pol II. This could facilitate dissociation of RNA Pol II and Mediator on CTD-hyperphosphorylation, and thus suppress the functional consequences of a truncated CTD. In support of this view, structural and biochemical studies indicate that the Srb2, Srb4, Srb5, and Srb6 proteins are located in the head domain of Mediator—the domain responsible for CTD-independent interactions with RNA Pol II (see following).
B. Structure–Function The subunit composition of the S. cerevisiae Mediator is clearly distinct from similar complexes found in other eukaryotic cells. Only eight out of 20 core Mediator subunits have a highly homologous counterpart in
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mouse and human Mediator (Table I). The low degree of conservation at the primary sequence level has evoked the idea that the metazoan Mediator is significantly different both in structure and function from yeast Mediator. Another possible interpretation is that Mediator functions as an interface between rapidly evolving gene-specific regulatory proteins and the highly conserved basal transcription machinery. According to this view, the conserved Mediator core of only eight proteins found in all eukaryotic cells is responsible for contacts with the basal transcription machinery. Subunits responsible for interactions with gene specific activators and repressors will be less conserved. In this view, the subunit composition may vary between eukaryotic cell types but the basic mechanisms of Mediator-dependent transcriptional regulation are the same. In support of this model, one can note that only essential gene products are conserved between the Mediator complex in yeast and higher eukaryotes (Table I). Nonessential gene products appear to be species specific. In fact, this is also true if one compares Mediator complex isolated from S. cerevisiae with the corresponding complex from fission yeast, S. pombe. The two species were separated early in evolution, and the 10 subunits conserved between them are all encoded by essential S. cerevisiae genes (Spahr et al., 2001). Support for the existence of a conserved Mediator core comes also from structural studies. Single-particle analysis by electron microscopy has demonstrated striking structural similarities between Mediator isolated from yeast, mouse, and human cells (Asturias et al., 1999; Dotson et al., 2000). In two-dimensional projections, the isolated Mediator purified from S. cerevisiae and mouse cells appears compact. When RNA Pol II is present, however, these Mediators adopt an extended conformation and embrace the globular Pol II. The extended structure reveals three distinct submodules of Mediator: a head, a middle, and a tail region. Direct contacts are formed between RNA Pol II and the head and middle region. The largest part of Mediator is made up of an elongated tail region, which does not appear to contact the RNA Pol II.
1. Tail Region ScMediator isolated from a sin4 strain lacks the Sin4 protein as well as Gal11, Med2, and Hrs1/Pgd1 (Myers et al., 1999). As mentioned previously, these proteins, the Sin4 module, are needed for the function of a wide variety of activators, including Gal4 and Gcn4. The module does not, however, appear to be important for other Mediator functions such as stimulation of basal transcription or CTD phosphorylation. The Sin4 module corresponds to the tail region, as image reconstruction of the sin4 Mediator lacks this part of Mediator (Dotson et al., 2000). In vivo
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and in vitro evidence thus indicates that this region plays an important role for activator and repressor interactions with Mediator. The Sin4 module is known to interact with the rest of the Mediator complex through the terminal domain of the Rgr1/TRAP170 subunit, because deletion of this domain causes the loss of the entire module (Li et al., 1995). These observations, together with the fact that head and tail domains do not interact in the extended conformation of Mediator (Asturias et al., 1999), indicate that Rgr1/TRAP170 constitutes the part of the middle domain that is located most proximal to the tail domain.
2. Middle Region Biochemical analysis has identified two stable subcomplexes within Mediator (Lee and Kim, 1998). One of these subcomplexes (the Rgr1 module) contains Rgr1 together with Med1, Med4, Med7, Med8, Srb7, and probably Nut2. Many of the subunits of the Rgr1 submodule have a conserved homolog in S. pombe and metazoan Mediator complexes (Table I). These proteins include Med7, Nut2, and Srb7, which have recently been shown experimentally to form a stable complex when coexpressed from recombinant baculoviruses in insect cells (Han et al., 2001). On the basis of the structure of the sin4 Mediator, it appears likely that the middle region corresponds to the Rgr1 module. The electron microscopy structures indicate a possible direct contact between RNA Pol II CTD and this middle region (Asturias et al., 1999).
3. Head Region By way of exclusion, the proteins not associated with the tail or middle region of the complex will probably correspond to the head region. These proteins correspond to second stable subcomplex of scMediator identified by biochemical analysis, the Srb4 module, which is composed of the Med6, Rox3, Srb2, Srb4, Srb5, and Srb6 proteins (Lee and Kim, 1998). Support for this module comes from the observation that the Med6, Srb2, Srb4, Srb5, and Srb6 proteins can form a stable complex on coexpression in insect cells (Lee et al., 1998). In fact, the Srb2 and Srb5 appear to form a subcomplex of the module, as Mediator purified from srb2 cells also lack the Srb5 subunit (S. Bjo¨rklund, unpublished observations). Genetics experiments also support the postulated module, as Med6 and Srb6 both have been identified as dominant suppressors of a ts mutation in SRB4 (Lee et al., 1998). EM structures show that the most pronounced contacts between Mediator and RNA Pol II take place in the head region, which seems to interact with a part of RNA Pol II different from the CTD (Asturias et al., 1999). In fact, Med6 copurify with the core RNA Pol II rather than with the rest of
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Fig. 1. The yeast RNA Polymerase II holoenzyme revealed by electron microscopy and image processing. (Asturias et al., 1999). (A) The extended Mediator contains three distinguishable regions; head (h), middle (m), and tail (t). The globular density embraced by Mediator is identified as RNA polymerase II. The outline of a projection of the previously determined polymerase three-dimensional structure is superimposed (dark line), with the point of attachment of the C-terminal domain (dark circle) and the location of the DNA-binding channel (c) indicated. (B) Tentative subunit organization for the holoenzyme. The model is based on available structural information and reported physical interactions. The surface of each subunit has been calculated by assuming a globular shape and drawn in scale. Subunits in red have reported homologs in Saccharomyces pombe and, with the exception of Rox3 and Srb6, also in mammalian Mediator. The yellow subunits are specific for Saccharomyces cerevisiae. (See Color Insert.)
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the Mediator in extracts from certain yeast strains (Balciunas et al., 1999). As expected, many of the proteins of the head region are also conserved in other species, including homologs to Med6 and Srb4 in mammalian Mediator complexes (Table I). In summary, we would like to propose an onion-like structure of Mediator (Fig. 1). It appears as if the head and middle regions of Mediator contain all the conserved and essential subunits of the S. cerevisiae complex. In concordance with the core Mediator model, these are also the polymerase-interacting regions, and it is likely that the conserved subunits of these domains are faced toward the RNA Pol II. In contrast, nonconserved subunits in the head and middle domains and all subunits in the tail region interact with activators, and repressors are facing outward to receive signals from regulatory proteins. In agreement with this, structural comparison between S. cerevisiae and human Mediator has demonstrated striking structural similarities in the head and middle regions (Asturias et al., 1999; Dotson et al., 2000). The tail region of the human complex is large and differs significantly in its structure from S. cerevisiae. One could speculate that this region of the human Mediator complex may contain a number of large and metazoan-specific subunits involved in activator and repressor interactions; for example, TRAP220, TRAP230, and TRAP240 (Malik and Roeder, 2000).
V. Concluding Remarks The discovery of Mediator has changed our view of transcriptional regulation. This multiprotein complex is now established as the main transducer of regulator information from enhancers and other control elements to the promoter. Mediator seems to form an interface between gene-specific regulatory proteins and the highly conserved basal transcription machinery. A conserved core of only eight proteins found in all eukaryotic cells is responsible for contacts formed with RNA Pol II and TFIIH. Other, species-specific subunits are mainly responsible for direct interactions with regulatory proteins. The subunit composition of Mediator may therefore vary between different eukaryotic cell types, but the mechanisms of Mediator-dependent transcriptional regulation are highly conserved.
References Aalfs, J. D., and Kingston, R. E. (2000). Trends Biochem. Sci. 25, 548–555. Asturias, F. J., Jiang, Y. W., Myers, L. C., Gustafsson, C. M., and Kornberg, R. D. (1999). Science 283, 985–987. Balciunas, D., Galman, C., Ronne, H., and Bjorklund, S. (1999). Proc. Natl. Acad. Sci. USA 96, 376–381.
THE MEDIATOR COMPLEX
63
Balciunas, D., Hallberg, M., Bjorklund, S., and Ronne, H. (2003). J. Biol. Chem. 278, 3831–3839. Balciunas, D., and Ronne, H. (1995). Nucleic Acids Res. 23, 4421–4425. Boyer, T. G., Martin, M. E., Lees, E., Ricciardi, R. P., and Berk, A. J. (1999). Nature 399, 276–279. Brown, T. A., Evangelista, C., and Trumpower, B. L. (1995). J. Bacteriol. 177, 6836–6843. Cadena, D. L., and Dahmus, M. E. (1987). J. Biol. Chem. 262, 12468–12474. Chang, Y. W., Howard, S. C., Budovskaya, Y. V., Rine, J., and Herman, P. K. (2001). Genetics 157, 17–26. Chao, D. M., Gadbois, E. L., Murray, P. J., Anderson, S. F., Sonu, M. S., Parvin, J. D., and Young, R. A. (1996). Nature 380, 82–85. Chaves, R. S., Herrero, P., and Moreno, F. (1999). Biochem. Biophys. Res. Commun. 254, 345–350. Cosma, M. P., Panizza, S., and Nasmyth, K. (2001). Mol. Cell 7, 1213–1220. Covitz, P. A., Song, W., and Mitchell, A. P. (1994). Genetics 138, 577–586. Dotson, M. R., Yuan, C. X., Roeder, R. G., Myers, L. C., Gustafsson, C. M., Jiang, Y. W., Li, Y., Kornberg, R. D., and Asturias, F. J. (2000). Proc. Natl. Acad. Sci. USA 97, 14307–14310. Fan, H. Y., Cheng, K. K., and Klein, H. L. (1996). Genetics 142, 749–759. Feaver, W. J., Svejstrup, J. Q., Henry, N. L., and Kornberg, R. D. (1994). Cell 79, 1103–1109. Flanagan, P. M., Kelleher, R. J. D., Sayre, M. H., Tschochner, H., and Kornberg, R. D. (1991). Nature 350, 436–438. Fondell, J. D., Ge, H., and Roeder, R. G. (1996). Proc. Natl. Acad. Sci. USA 93, 8329–8333. Gromoller, A., and Lehming, N. (2000). FEBS Lett. 484, 48–54. Gu, W., Malik, S., Ito, M., Yuan, C. X., Fondell, J. D., Zhang, X., Martinez, E., Qin, J., and Roeder, R. G. (1999). Mol. Cell 3, 97–108. Gustafsson, C. M., Myers, L. C., Beve, J., Spahr, H., Lui, M., Erdjument-Bromage, H., Tempst, P., and Kornberg, R. D. (1998). J. Biol. Chem. 273, 30851–30854. Han, S. J., Lee, Y. C., Gim, B. S., Ryu, G. H., Park, S. J., Lane, W. S., and Kim, Y. J. (1999). Mol. Cell. Biol. 19, 979–988. Han, S. J., Lee, J. S., Kang, J. S., and Kim, Y. J. (2001). J. Biol. Chem. 276, 37020–37026. Hengartner, C. J., Myer, V. E., Liao, S. M., Wilson, C. J., Koh, S. S., and Young, R. A. (1998). Mol. Cell 2, 43–53. Hittelman, A. B., Burakov, D., Iniguez-Lluhi, J. A., Freedman, L. P., and Garabedian, M. J. (1999). EMBO J. 18, 5380–5388. Holstege, F. C., Jennings, E. G., Wyrick, J. J., Lee, T. I., Hengartner, C. J., Green, M. R., Golub, T. R., Lander, E. S., and Young, R. A. (1998). Cell 95, 717–728. Ito, M., Yuan, C. X., Malik, S., Gu, W., Fondell, J. D., Yamamura, S., Fu, Z. Y., Zhang, X., Qin, J., and Roeder, R. G. (1999). Mol. Cell 3, 361–370. Ito, M., Yuan, C. X., Okano, H. J., Darnell, R. B., and Roeder, R. G. (2000). Mol. Cell 5, 683–693. Jiang, Y. W., and Stillman, D. J. (1992). Mol. Cell. Biol. 12, 4503–4514. Jiang, Y. W., and Stillman, D. J. (1995). Genetics 140, 103–114. Jiang, Y. W., Veschambre, P., Erdjument-Bromage, H., Tempst, P., Conaway, J. W., Conaway, R. C., and Kornberg, R. D. (1998). Proc. Natl. Acad. Sci. USA 95, 8538–8543. Kelleher, R. J., Flanagan, P. M., and Kornberg, R. D. (1990). Cell 61, 1209–1215. Kim, Y. J., Bjorklund, S., Li, Y., Sayre, M. H., and Kornberg, R. D. (1994). Cell 77, 599–608.
64
¨ RKLUND AND GUSTAFSSON BJO
Koh, S. S., Ansari, A. Z., Ptashne, M., and Young, R. A. (1998). Mol. Cell 1, 895–904. Koleske, A. J., and Young, R. A. (1994). Nature 368, 466–469. Kwon, J. Y., Park, J. M., Gim, B. S., Han, S. J., Lee, J., and Kim, Y. J. (1999). Proc. Natl. Acad. Sci. USA 96, 14990–14995. Lee, D., and Lis, J. T. (1998). Nature 393, 389–392. Lee, T. I., Wyrick, J. J., Koh, S. S., Jennings, E. G., Gadbois, E. L., and Young, R. A. (1998). Mol. Cell. Biol. 18, 4455–4462. Lee, Y. C., and Kim, Y. J. (1998). Mol. Cell. Biol. 18, 5364–5370. Lee, Y. C., Min, S., Gim, B. S., and Kim, Y. J. (1997). Mol. Cell. Biol. 17, 4622–4632. Lee, Y. C., Park, J. M., Min, S., Han, S. J., and Kim, Y. J. (1999). Mol. Cell. Biol. 19, 2967–2976. Lemon, B. D., and Freedman, L. P. (1999). Curr. Opin. Genet. Dev. 9, 499–504. Li, Y., Bjorklund, S., Jiang, Y. W., Kim, Y. J., Lane, W. S., Stillman, D. J., and Kornberg, R. D. (1995). Proc. Natl. Acad. Sci. USA 92, 10864–10868. Liao, S. M., Zhang, J., Jeffery, D. A., Koleske, A. J., Thompson, C. M., Chao, D. M., Viljoen, M., van Vuuren, H. J., and Young, R. A. (1995). Nature 374, 193–196. Liu, Y., Ranish, J. A., Aebersold, R., and Hahn, S. (2000). J. Biol. Chem. 11, 11. Lorch, Y., Beve, J., Gustafsson, C. M., Myers, L. C., and Kornberg, R. D. (2000). Mol. Cell 6, 197–201. Malik, S., Gu, W., Wu, W., Qin, J., and Roeder, R. G. (2000). Mol. Cell 5, 753–760. Malik, S., and Roeder, R. G. (2000). Trends Biochem. Sci. 25, 277–283. McNeil, J. B., Agah, H., and Bentley, D. (1998). Genes Dev. 12, 2510–2521. Meisterernst, M., Roy, A. L., Lieu, H. M., and Roeder, R. G. (1991). Cell 66, 981–993. Myer, V. E., and Young, R. A. (1998). J. Biol. Chem. 273, 27757–27760. Myers, L. C., Gustafsson, C. M., Bushnell, D. A., Lui, M., Erdjument-Bromage, H., Tempst, P., and Kornberg, R. D. (1998). Genes Dev. 12, 45–54. Myers, L. C., Gustafsson, C. M., Hayashibara, K. C., Brown, P. O., and Kornberg, R. D. (1999). Proc. Natl. Acad. Sci. USA 96, 67–72. Myers, L. C., and Kornberg, R. D. (2000). Annu. Rev. Biochem. 69, 729–749. Naar, A. M., Beaurang, P. A., Zhou, S., Abraham, S., Solomon, W., and Tjian, R. (1999). Nature 398, 828–832. Nilsson, L., Tiensuu, T., and Tuck, S. (2000). Genetics 156, 1083–1096. Nonet, M. L., and Young, R. A. (1989). Genetics 123, 715–724. Oelgeschlager, T., Tao, Y., Kang, Y. K., and Roeder, R. G. (1998). Mol. Cell 1, 925–931. Park, J. M., Kim, H. S., Han, S. J., Hwang, M. S., Lee, Y. C., and Kim, Y. J. (2000). Mol. Cell. Biol. 20, 8709–8719. Park, J. M., Werner, J., Kim, J. M., Lis, J. T., and Kim, Y. J. (2001). Mol. Cell 8, 9–19. Philibert, R. A., King, B. H., Winfield, S., Cook, E. H., Lee, Y. H., Stubblefield, B., Damschroder-Williams, P., Dea, C., Palotie, A., Tengstrom, C., Martin, B. M., and Ginns, E. I. (1998). Mol. Psychiatry 3, 303–309. Rachez, C., Lemon, B. D., Suldan, Z., Bromleigh, V., Gamble, M., Naar, A. M., Erdjument-Bromage, H., Tempst, P., and Freedman, L. P. (1999). Nature 398, 824–828. Rachez, C., Suldan, Z., Ward, J., Chang, C. P., Burakov, D., Erdjument-Bromage, H., Tempst, P., and Freedman, L. P. (1998). Genes. Dev. 12, 1787–1800. Rosenblum-Vos, L. S., Rhodes, L., Evangelista, C. C.Jr., Boayke, K. A., and Zitomer, R. S. (1991). Mol. Cell. Biol. 11, 5639–5647. Ryu, S., Zhou, S., Ladurner, A. G., and Tjian, R. (1999). Nature 397, 446–450. Sakai, A., Shimizu, Y., Kondou, S., Chibazakura, T., and Hishinuma, F. (1990). Mol. Cell. Biol. 10, 4130–4138. Sakurai, H., and Fukasawa, T. (2000). J. Biol. Chem. 275, 37251–37256.
THE MEDIATOR COMPLEX
65
Sakurai, H., Hiraoka, Y., and Fukasawa, T. (1993). Proc. Natl. Acad. Sci. USA 90, 8382–8386. Sakurai, H., Kim, Y. J., Ohishi, T., Kornberg, R. D., and Fukasawa, T. (1996). Proc. Natl. Acad. Sci. USA 93, 9488–9492. Scafe, C., Chao, D., Lopes, J., Hirsch, J. P., Henry, S., and Young, R. A. (1990). Nature 347, 491–494. Shen, W. C., and Green, M. R. (1997). Cell 90, 615–624. Singh, N., and Han, M. (1995). Genes Dev. 9, 2251–2265. Song, W., Treich, I., Qian, N., Kuchin, S., and Carlson, M. (1996). Mol. Cell. Biol. 16, 115–120. Spahr, H., Beve, J., Larsson, T., Bergstrom, J., Karlsson, K. A., and Gustafsson, C. M. (2000). J. Biol. Chem. 275, 1351–1356. Spahr, H., Samuelsen, C. O., Baraznenok, V., Ernest, I., Huylebroeck, D., Remacle, J. E., Samuelsson, T., Kieselbach, T., Holmberg, S., and Gustafsson, C. M. (2001). Proc. Natl. Acad. Sci. USA 98, 11985–11990. Spradling, A. C., Stern, D., Beaton, A., Rhem, E. J., Laverty, T., Mozden, N., Misra, S., and Rubin, G. M. (1999). Genetics 153, 135–177. Stillman, D. J., Dorland, S., and Yu, Y. (1994). Genetics 136, 781–788. Sun, X., Zhang, Y., Cho, H., Rickert, P., Lees, E., Lane, W., and Reinberg, D. (1998). Mol. Cell 2, 213–222. Suzuki, Y., and Nishizawa, M. (1994). Mol. Cell. Biol. 14, 3791–3799. Suzuki, Y., Nogi, Y., Abe, A., and Fukasawa, T. (1988). Mol. Cell. Biol. 8, 4991–4999. Svejstrup, J. Q., Li, Y., Fellows, J., Gnatt, A., Bjorklund, S., and Kornberg, R. D. (1997). Proc. Natl. Acad. Sci. USA 94, 6075–6078. Tabtiang, R. K., and Herskowitz, I. (1998). Mol. Cell. Biol. 18, 4707–4718. Thompson, C. M., Koleske, A. J., Chao, D. M., and Young, R. A. (1993). Cell 73, 1361–1375. Thompson, C. M., and Young, R. A. (1995). Proc. Natl. Acad. Sci. USA 92, 4587–4590. Treitel, M. A., and Carlson, M. (1995). Proc. Natl. Acad. Sci. USA 92, 3132–3136. Tudor, M., Murray, P. J., Onufryk, C., Jaenisch, R., and Young, R. A. (1999). Genes Dev. 13, 2365–2360. Urnov, F. D., and Wolffe, A. P. (2001). Oncogene 20, 2991–3006. Vallier, L. G., and Carlson, M. (1991). Genetics 129, 675–684. Wang, G., Cantin, G. T., Stevens, J. L., and Berk, A. J. (2001). Mol. Cell. Biol. 21, 4604–4613. Woods, A., Munday, M. R., Scott, J., Yang, X., Carlson, M., and Carling, D. (1994). J. Biol. Chem. 269, 19509–19515. Xiao, Z., McGrew, J. T., Schroeder, A. J., and Fitzgerald-Hayes, M. (1993). Mol. Cell. Biol. 13, 4691–4702. Yu, G., and Fassler, J. S. (1993). Mol. Cell. Biol. 13, 63–71. Yuan, C. X., Ito, M., Fondell, J. D., Fu, Z. Y., and Roeder, R. G. (1998). Proc. Natl. Acad. Sci. USA 95, 7939–7944. Yudkovsky, N., Ranish, J. A., and Hahn, S. (2000). Nature 408, 225–229. Zhang, H., and Emmons, S. W. (2000). Genes Dev. 14, 2161–2172. Zhu, Y., Qi, C., Jain, S., Rao, M. S., and Reddy, J. K. (1997). J. Biol. Chem. 272, 25500–25506. Zhu, Y., Qi, C., Jia, Y., Nye, J. S., Rao, M. S., and Reddy, J. K. (2000). J. Biol. Chem. 275, 14779–14782. Zilahi, E., Miklos, I., and Sipiczki, M. (2000). Curr. Genet. 38, 227–232.
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STRUCTURE AND FUNCTION OF THE TFIID COMPLEX By ORANART MATANGKASOMBUT, ROY AUTY, AND STEPHEN BURATOWSKI* Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School, Boston, MA
I. TFIID and Transcription Initiation. . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. TFIID Components and Structure. . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. TATA-Binding Protein . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. TBP-Associated Factors . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Alternative TFIIDs . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. TFIID Functions. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Promoter Recognition. . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Mediators of Activation.. . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Enzymatic Activities . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. The Requirement for TFIID In Vivo . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Regulation of TFIID . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VI. The TFIID-Chromatin Connection . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VII. Future Questions . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
67 68 69 72 75 76 77 79 80 82 84 85 87 87
I. TFIID and Transcription Initiation TFIID was first described as an activity in fractionated mammalian cell extracts that is required, together with other fractions, for accurate initiation of RNA Pol II transcription reconstituted in vitro (Matsui et al., 1980; Samuels et al., 1982). An important function of TFIID was revealed when it was shown to bind specifically to the TATA box element of core promoters (Sawadogo and Roeder, 1985). The purification and identification of the protein or proteins responsible for this activity from mammalian cell extracts were difficult because of its complexity and low abundance. A breakthrough was made with the discovery that yeast has a TFIID activity that can function in a reconstituted mammalian transcription system lacking TFIID fraction (Buratowski et al., 1988; Cavallini et al., 1988). This led to the purification of the yeast TATA-binding protein (TBP) and the isolation of its gene (Cavallini et al., 1989; Eisenmann et al., 1989; Hahn et al., 1989; Horikoshi et al., 1989; Schmidt et al., 1989). The cloning of Drosophila and human TBP genes followed closely (Hoey et al., 1990; Hoffman et al., 1990; Kao et al., 1990; Muhich et al., 1990; Peterson et al., * Correspondence to: Stephen Buratowski, Harvard Medical School, Boston, MA 02115 (e-mail:
[email protected]).
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1990). Although yeast TBP was purified as a monomer of 27 kD, native TFIID from mammalian cells migrates as a much larger entity on gel filtration columns (Reinberg et al., 1987). Partially purified TFIID from Drosophila or human cells also shows an extended footprint on promoter DNA (Sawadogo and Roeder, 1985), whereas recombinant TBP shows a much smaller footprint. Moreover, although recombinant TBP can direct basal transcription in vitro, it fails to support activated transcription, unlike the partially purified TFIID (Pugh and Tjian, 1990). These findings indicated that TBP exists in a complex with other proteins. Using anti-TBP antibodies to purify associated proteins from Drosophila and human cells, several polypeptides designated TAFs (TBP-associated factors) were identified (Dynlacht et al., 1991; Tanese et al., 1991). It was later found that TBP actually participates in transcription initiation of all three RNA polymerases and that it is the associated proteins (the TAFs) that confer on the TBP-containing complexes their specificity for each polymerase system (Hernandez, 1993). The purification of the TFIID complex was followed by the cloning of genes for several TAF subunits in Drosophila and human (reviewed in Albright and Tjian, 2000; Burley and Roeder, 1996). Because the basal transcription machinery is highly conserved among eukaryotes, it was surprising that yeast TBP would exist solely as a monomer. This puzzle was solved with the discovery of yeast TAFs. When a gene encoding dmTAF2 was cloned, it was found to be related to the yeast TSM1 gene, and the Tsm1 protein associates with yeast TBP in vivo (Verrijzer et al., 1994). Furthermore, a multisubunit TAF complex was purified from yeast extracts using GST-TBP as a ligand for affinity chromatography (Reese et al., 1994). The use of epitope tag and immunoprecipitation also led to the isolation of TBP-TAFs complex from yeast extract (Poon et al., 1995). Similar to the higher eukaryotic TAFs, yeast TAFs do not support transcription on their own, but can function as coactivators. TFIID and its individual subunits have been subjected to intense study. As will be discussed below, complementary information from biochemical, genetic, and structural studies has shown, and will continue to give, a clearer picture of the forms and functions of this crucial component of the transcription machinery.
II. TFIID Components and Structure The purification of TFIID was soon followed by the identification of its subunits (reviewed in Albright and Tjian, 2000; Burley and Roeder, 1996). The amino acid sequences of many TAFs revealed a high level of conservation. TFIID structure has changed little through evolution, with all
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species containing at least 13 conserved subunits (reviewed in Gangloff et al., 2001b). A consensus on the nomenclature of TAFs has been developed to avoid confusion (Tora, 2002) and is used throughout this review. The currently identified TAFs are listed in Table I. Structural studies of TBP and TAF subcomplexes provided insights into the mechanisms of TFIID function and complex organization. A lowresolution structure of the whole TFIID complex has also been obtained using electron microscopy (EM; Andel et al., 1999; Brand et al., 1999). The complex has a horseshoe shape, consisting of 3–4 lobes connected by flexible regions. It has been suggested that the concave face may represent the DNA binding surface and that DNA might be accommodated in the central channel. TBP was mapped to the central lobe, which is also where TFIIA and TFIIB bind. Further determination of individual subunit positions will be essential for our understanding of how TFIID functions as a complex. TFIID is formed through numerous protein–protein interactions, both TBP–TAF and TAF–TAF (Burley and Roeder, 1996; Gangloff et al., 2001b). Many interactions were identified in vitro, and some have been verified in vivo with genetic studies in yeast. It has been suggested that the largest TAF subunit, TAF1, may be a scaffold for the complex because it interacts directly with TBP and with several other TAFs. Interestingly, multiple TAFs contain a histone fold domain, a conserved protein interaction motif found in the histone proteins. This domain allows tight and extensive contacts between the two partners (Luger et al., 1997). The interactions between the histone-like TAFs are believed to play an important structural role in TFIID and will be discussed in more detail below. A speculative model for TFIID structure is presented in Figure 1.
A. TATA-Binding Protein Crystallographic studies were performed on several forms of TBP, including apoTBP (TBP alone), TBP-DNA, TBP-DNA-TFIIA, and TBPDNA-TFIIB complexes (reviewed in Nikolov and Burley, 1997). These complexes are significant to our mechanistic view of TBP binding to the TATA element and of the functions of TFIIA and TFIIB (Nikolov and Burley, 1997). TBP has a conserved C terminus and a divergent N terminus of variable length, which is not necessary for transcription in certain yeast strains (Burley and Roeder, 1996). The conserved 180 residues of TBP form a quasisymmetric molecular ‘‘saddle’’ with DNA binding at the concave underside and the convex surface available for interaction with TAFs and other transcription factors (Nikolov and Burley, 1997).
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Table I Pol II taf Nomenclature Including the Corresponding Known Orthologues and Paralogues New name
Drosophila melanogaster
Caenorhablitis elegans (Previous name)
Ce New name
Saccharomyces cerevisiae
Saccharomyces pombe
TAFII250 TAFII150 TAFII140 TAFII130/135 TAFII105 TAFII100
TAFII230 TAFII150 TAFII155 or BIP2 TAFII110
taf-1 (W04A8.7) taf-2 (Y37E11B.4) (C11G6.1) taf-5 (R119.6)
taf-1 taf-2 taf-3 taf-4
Taf145/130p Taf150p or TSM1 Taf47p Taf48p or MPT1
TAFII111 (T38673)
TAFII80
taf-4 (F30F8.8)
taf-5
*
*
TAFII72 TAFII73
Cannonball TAFII60 (AAF52013) (AAF54162)
taf-3.1 (W09B6.2) taf-3.2 (Y37E11A.8) taf-8.1 (F54F7.1) taf-8.2 (Y111B2A.16) (ZK1320.7) taf-10 (T12D8.7) taf-11 (K03B4.3)
taf-6.1 taf-6.2 taf-7.1 taf-7.2 taf-8 taf-9 taf-10
*
taf-7.1 (F48D6.1) taf-7.2 (K10D3.3) taf-9 (Y56A4.3) taf-6 (C14A4.10)
taf-11.1 taf-11.2 taf-12 taf-13
PAF65 TAFII80 * PAF65 TAFII55 TAF2Q (BAB71460) * TAFII32/31 * TAFII30 TAFII28 *
TAFII 20/15 TAFII18
Prodos TAFII40 TAFII24 TAFII16 TAFII30 TAFII30 (AAF53875)
Taf90p
Taf60p
(T50183)
(CAA20756)
Taf67p
TAFII62/PTR6
Taf65p Taf17p * Taf25p
(T40895) (S62536) (T39928)
Taf40p
(CAA93543)
*
(T37702) (CAA19300)
*
Taf61/68p Taf19p or FUN81 TAF30
TAFII68 B-TFIID
BTAF1 *
TAFII170/TAF-172
Hel89B
Mot1
denotes TAFs found in histone acetyltransferase complexes: P/CAF in human, and SAGA in S. cerevisiae. Adapted from Tora (2002).
MATANGKASOMBUT ET AL.
TAF1 TAF2 TAF3 TAF4 TAF4B TAF5 TAF5B TAF5L TAF6 TAF6L TAF7 TAF7L TAF8 TAF9 TAF10 TAF10B TAF11 TAF11L TAF12 TAF13 TAF14 TAF15
Homo sapiens
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Fig. 1. Possible subunit structure of TFIID. On the basis of immuno-EM studies (Leurent et al., 2002), yeast two-hybrid data (Andel et al., 1999; Brand et al., 1999; Yatherajam et al., 2003), and in vitro protein interaction data (see text), a model of TFIID interactions was constructed. TAFs that appear in both TFIID and the SAGA HAT complex are shaded. BD1 and BD2 signify the bromodomains found in higher eukaryotic TAF1 or in yeast Bdf1. The exact stoichiometry of histone-like TAFs within each lobe is unclear; some data indicate an octamer-like structure, whereas others support two tetramers.
The co-crystal structure of the TBP-DNA complex shows that TBP binds to the minor groove of the TATA element and introduces a sharp bend in the DNA (Kim and Burley, 1994; Kim et al., 1993a; Kim et al., 1993b). This is mediated by insertion of two pairs of phenylalanine residues in between the first two and between the last two base pairs of the TATAAA sequence. The minor groove is thus widened and fit to the concave surface of the saddle. The particular wrapping of DNA around the nucleosome can affect the disposition of the minor groove and the TATA element, and this can affect the ability of TBP to bind in the context of a nucleosome (Imbalzano et al., 1994). The bend in DNA could also serve other purposes such as
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to bring the activator closer to the core promoter and the preinitiation complex (PIC), or to stabilize PIC assembly (Burley and Roeder, 1996). TFIIB interacts with the C-terminal stirrup of TBP and the DNA both upstream and downstream of the TATA box, whereas TFIIA interacts with the N-terminal stirrup and the DNA upstream of the TATA box on the opposite face of the double helix from TFIIB (Nikolov and Burley, 1997). Therefore, both TFIIA and TFIIB can bind to the TBP-DNA complex simultaneously and synergistically stabilize the complex. Furthermore, TFIIB binding also contributes to the directionality of transcription and forms a bridge between TBP and Pol II and specifies the transcription start site. It will be interesting to see how these interactions fit into the structure of the TFIID complex.
B. TBP-Associated Factors At present, approximately 13 polypeptides have been isolated as TAFs in all species studied, and they are listed in Table I (Tora, 2002). Earlier studies were performed almost exclusively in Drosophila, human cells, and budding yeast, whereas more recent studies in Caenorhabditis elegans indicate that it is also a useful model system, especially for studying the role of TAFs during development (Walker et al., 2001). In some species, additional TAFs have been isolated, but these have not been found universally, and it remains to be seen whether they are bona fide TFIID subunits. Analysis of the primary sequences of the identified TAFs reveals a high level of conservation. Several motifs appear in the homologues of all three species, such as WD40 repeats in TAF5 and the histone fold domains in more than half of the TAFs. However, some motifs are found only in the higher eukaryotic TAFs, but not in the yeast homologs. This is the case for TAF1; the scTAF1 shares homology with dm/hsTAF1 only in the N-terminal half of the protein (Reese et al., 1994). It lacks two copies of a conserved motif termed the bromodomain, and a C-terminal acidic domain. The bromodomain is a conserved motif found in several transcription regulators and is believed to mediate interactions with acetylated histone tails ( Jeanmougin et al., 1997; Marmorstein and Berger, 2001). Recent studies indicate that Bromodomain Factor 1 (Bdf1) associates with TFIID and probably corresponds to this missing piece of yeast TFIID (Matangkasombut et al., 2000). The discovery of the histone fold domain in TAFs began with Drosophila TAF9 and TAF6, which have significant sequence similarity with the histone fold domains of histone H3 and H4, respectively. A cocrystal structure of the histone fold domains of these two TAFs shows that they interact and form a heterotetramer, similar to that of histone H3 and H4
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(Xie et al., 1996). The identification of an H2B homology region in TAF12 led to the hypothesis these three TAFs may form a histone octamer-like structure, despite the lack of an apparent H2A homolog (Hoffmann et al., 1996). According to this proposal, TAF9-TAF6 heterotetramer would interact with two homodimers of TAF12. However, an H2A homology region was identified in TAF4, and this domain has been shown to interact with TAF12 through yeast two-hybrid analysis and a coexpression assay in Escherichia coli (Gangloff et al., 2000). The yeast homolog of hsTAF4 was discovered independently in a purified TFIID preparation (Sanders and Weil, 2000) and in a yeast genetic screen for a high-copy suppressor of a scTAF12 temperature-sensitive (ts) mutant (Reese et al., 2000). TAF9TAF6-TAF12-TAF4 can form a complex with molecular properties consistent with an octamer (Selleck et al., 2001). Point mutations in the interface between H2B-like and H4-like TAFs (TAF12-TAF6) disrupt the octamer formation without disrupting the TAF12-TAF4 or TAF9-TAF6 interaction, indicating structure similar to that of the histone octamer. Genetic studies in yeast provided further support for the specific interactions among these four TAFs. Overexpression of any one of these TAFs can suppress ts alleles of the other TAFs in this group, but similar interactions are not seen with any other TAFs (Michel et al., 1998; Selleck et al., 2001). The ability of the histone-like TAFs to form an octamer-like structure raises the possibility that the TAF octamer may wrap promoter DNA in a manner similar to the nucleosome (Hoffmann et al., 1997; Oelgeschlager et al., 1996). This hypothesis is supported by the resemblance of DNase I footprinting patterns of TFIID on the Adenovirus Major Late (AdML) promoter to those of nucleosomal DNA. However, the arginine side chains in histones that form primary contacts with DNA are not conserved in TAFs (Luger et al., 1997). Therefore, the histone fold domain interaction may be used only for the formation of a compact structure and is not necessarily involved in DNA wrapping. Several more TAFs have been found to contain histone fold domains (Gangloff et al., 2001b). Each shows pairwise interactions: TAF11-TAF13 (Birck et al., 1998; Komarnitsky et al., 1999), TAF3-TAF10 (Gangloff et al., 2001a,c), and TAF8-TAF10 (Gangloff et al., 2001a,b). Genetic interaction has been shown between some of the pairs, indicating that there may be higher-order interactions between these pairs. For instance, genetic interaction between TAF3 and the TAF11-TAF13 pair suggests that the two pairs of TAF3-TAF10 and TAF11-TAF13 might form another subcomplex in TFIID (Gangloff et al., 2001c). Using specific anti-TAF antibodies, individual TAFs have been mapped onto the lobed TFIID structure seen in the electron micrographs (Leurent et al., 2002). Unfortunately, there is no clear assignment of any octamer-like subcomplexes onto specific lobes, and some TAFs
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appear to be in multiple lobes (see Fig. 1). Obviously, much more work needs to be done before the TFIID structure is understood. Interestingly, a subset of TAFs is found in several other transcriptionrelated complexes, such as the histone acetyltransferase (HAT) complexes in yeast and humans and the Polycomb group complex in Drosophila (Grant et al., 1998; Ogryzko et al., 1998; Saurin et al., 2001; Wieczorek et al., 1998). These shared TAFs are specified in Table I. The histone-like TAFs (TAF9, TAF6, TAF12), TAF5, and TAF10 are part of the SAGA (SptAda-Gcn5 acetyltransferase) HAT complex in yeast (Grant et al., 1998). The P/CAF HAT complex in human shares TAF 9, TAF 12, and TAF10 with TFIID, but it also contains homologues of TAF6, called TAF6L, and of TAF5, called TAF5L (Ogryzko et al., 1998). TAF9 was also found as a component of the STAGA (Spt3-TAF31-Gcn5 acetyltransferase) complex, a human HAT complex containing the homolog of Gcn5. Another HAT complex purified from human cells contains TAF4, TAF5, TAF6, TAF7, TAF9, TAF10, and TAF12 and is called TBP-free TAF-containing complex (TFTC) (Wieczorek et al., 1998). The EM structure of TFTC is generally similar to TFIID, consisting of five lobes that form a horseshoe shape (Brand et al., 1999). Surprisingly, this complex could replace TFIID in an in vitro transcription assay. This result indicates either that TAFs can recognize the core promoter and initiate transcription without TBP or that the complex contains an alternative TBP (see following). TFTC also contains hGcn5, which can modify chromatin to facilitate transcription (Brand et al., 1999). Recently, TFTC has been shown to be recruited to UVdamaged DNA, together with nucleotide excision repair proteins, and acetylates histone H3 on the DNA (Brand et al., 2001). It is still unclear how TFTC functions, and further characterization is required. In Drosophila, the polycomb group proteins plays an important role in maintaining transcription repression, whereas the trithorax group proteins maintain the active state of the major developmental regulator genes: the homeotic genes. To study the mechanisms of the polycomb-mediated repression, one of the polycomb group complexes, polycomb repressive complex 1 (PRC1) was purified (Saurin et al., 2001). Unexpectedly, several TAFs, including TAF1, TAF4, TAF5, TAF6, TAF9, and TAF11, were found to tightly associate through purification and coimmunoprecipitate with PRC1. The presence of the histone fold containing TAFs in PRC1 led the authors to speculate that these TAFs may be involved in the interaction with nucleosomes or DNA that help maintain the association of PRC1 through cell division. Therefore, further structural and functional study of TAFs would not only facilitate the understanding of TFIID function but might also shed light on the structure and function of other important transcriptional complexes.
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C. Alternative TFIIDs In multicellular eukaryotes, tissue-specific or stage-specific forms of TFIID have been found. These contain either a variant of TBP or variant TAF subunits. A TBP related factor, TRF1, was identified only in Drosophila and is expressed strongly in neuronal and testicular tissues (Crowley et al., 1993). Although TRF1 is quite similar to TBP in sequence, it binds preferentially to a TC-rich promoter element and directs transcription specifically from these promoters (Holmes and Tjian, 2000). TRF1 appears to form a complex with a distinct set of proteins called nTAFs (neuronal-specific TRF-associated factors; Hansen et al., 1997). Intriguingly, in Drosophila, TRF1 associates with the Pol III TAF, BRF, and substitutes for TBP in Pol III transcription of tRNA and U6 snRNA genes, although this is probably insect-specific, as TRF1 has not been found in other species (Takada et al., 2000). Unlike TRF1, TBP-like factor (TLF/TRF2/TLP/TRP), a second TBPrelated factor, has been identified in multicellular species, including C. elegans, Drosophila, and several vertebrates, and is widely expressed (reviewed in Dantonel et al., 1999). Although TLF shows significant homology to the conserved domain of TBP, it is more distantly related to TBP than TRF1. TLF shares several structure-determining residues with TBP and is thought to adopt a similar saddle-like structure. However, the residues that form the DNA binding surface are not well conserved, and TLF does not bind to the TATA box (Dantonel et al., 1999; Moore et al., 1999). It has been proposed that TLF may play a negative role by competing with TBP for TFIIA binding (Moore et al., 1999; Teichmann et al., 1999). Recently, several approaches were employed to study the role of TLF in vivo. These include the uses of RNA interference (RNAi) in C. elegans (Dantonel et al., 2000; Kaltenbach et al., 2000), antisense oligonucleotides in Xenopus (Veenstra et al., 2000), and 9 dominant negative allele in zebrafish (Muller et al., 2001) to deplete TLF in embryos. These studies indicate that TLF is essential for establishing zygotic transcription and for transcription of a subset of genes during embryonic development. In contrast, TLF knockout mice are viable but have a defect in spermatogenesis (Martianov et al., 2001; Zhang et al., 2001). It has been shown that, unlike C. elegans, Xenopus, and zebrafish, TLF expression in mouse is celltype and stage specific. TLF is highly expressed in testis only at certain stages of spermatogenesis that coincide with the increase in expression of other transcription factors, such as Pol II and TFIIB (Martianov et al., 2001; Zhang et al., 2001). It is therefore proposed that TLF may play an important transcriptional regulatory role in stages of spermatogenesis when a high level of regulated transcription is critical. An unexpected finding
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indicates that TLF could also be involved in maintaining the integrity of the centromeric heterochromatin in spermatids (Martianov et al., 2002). Together, these findings indicate that metazoans may have evolved with two distinct TBP families to add flexibility and control for coordinated gene regulation in a more complex genome. Variant TAFs are also used to regulate tissue-specific or temporal gene expression in metazoans. For instance, a cell-type-regulated subunit of TFIID, hsTAF4B, was found to associate with TFIID in B-lymphocytes (Dikstein et al., 1996b). There is evidence that hsTAF4B may play a coactivator role in NF-kB-mediated antiapoptotic responses and is important for B-lymphocyte and T-lymphocyte development (Silkov et al., 2002; Yamit-Hezi and Dikstein, 1998). A mouse homolog of TAF4B is expressed in ovarian follicles and is essential for ovarian development (Freiman et al., 2001). In humans, a TAF6 isoform, hsTAF6, shows induced expression and caspase-dependent cleavage on apoptotic stimuli and forms a TFIID-like complex lacking TAF9 (Bell et al., 2001). Increased expression of hsTAF6 leads to induction of several apoptotic target genes and is sufficient to induce apoptotic death. Another example of TAF variant usage is found in Drosophila. The cannonball gene product has recently been shown to be a TAF5 variant, called TAF5L, expressed only in the testis and required for spermatogenesis (Hiller et al., 2001). It will be of interest to see whether this testis-specific TAF is associated with TBP or TLF. More extreme variations have also been described, including a TBP-free TAF complex (TFTC) and a TBP-sans-TAFs complex. The TFTC, despite lacking TBP, can function in transcription initiation as discussed above (Wieczorek et al., 1998). A TBP-sans-TAFs complex is found in embryonal carcinoma (EC) cells but not in differentiated cells (Mitsiou and Stunnenberg, 2000). This complex, containing TBP in complex with the unprocessed TFIIA precursor and TFIIA, is called TAC. Although it is shown that this complex can bind to TATA containing DNA, and cotransfection of TBP and TFIIA synergistically increases transcription in EC cells, the transcriptional activity and composition of the complex requires further characterization.
III. TFIID Functions Several functions of TFIID in transcription regulation have been proposed. These roles include promoter recognition, coactivator function, and several enzymatic activities (Pugh, 2000). Each of these is discussed in detail below.
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A. Promoter Recognition TFIID is the only GTF with intrinsic DNA sequence specificity and is responsible for nucleating the assembly of Pol II preinitiation complex at core promoters. The interaction between TFIID and core promoter is mediated by the affinity of several specific subunits for distinct core promoter elements (Verrijzer and Tjian, 1996). Because TBP functions in transcription by all three nuclear RNA polymerases, these specific interactions help TFIID to distinguish different promoter structures and properly direct Pol II transcription machinery to its target genes. In the promoters of Pol II transcribing genes, there are at least three major core promoter elements: the TATA box, the Initiator (Inr), and at least in higher eukaryotes, the downstream promoter element (DPE; Smale, 2001). The TATA box is located upstream of the start site, whereas the initiator is found surrounding the start site and the DPE about 30 base pairs (bp) downstream of the start site (Kutach and Kadonaga, 2000). A promoter may contain any one, two, all three, or none of the identified elements. The structure of the core promoters can determine overall promoter strength and contribute to the combinatorial regulation of gene expression (Smale, 2001). Although either the TATA box or Inr can direct accurate transcription in vitro, the responses to transcriptional activators that regulate in vivo expression patterns are different between the two elements. The lineageor stage-specific or temporal control of expression of some genes requires a particular core element that cannot be substituted with the other (Novina and Roy, 1996). The underlying mechanism could be the intrinsic preferences of some transcription activators for different core promoter elements. This suggests that the rate-limiting steps, essential cofactors, or the mechanisms of initiation from different core promoter elements may be distinct (Butler and Kadonaga, 2001; Smale, 2001). The first core promoter element identified was an A/T-rich sequence located upstream of the transcription start site by 25–30 bp in higher eukaryotes, and at a variable distance between 60 and 120 bp in yeast. This element, with the consensus TATAAA, was named the TATA box. TBP interacts with the minor groove of this element and induces a bend in the promoter DNA (Burley and Roeder, 1996). This bending of DNA may be important for coordinating or stabilizing PIC assembly or may play a role in bringing transcription factors closer to the PIC. Sequences surrounding the TATA box may also contribute to specific TFIID–core promoter interaction. A study on TAF1 indicates that it functions as a core promoter selectivity factor (Shen and Green, 1997). Using promoter-mapping strategies, it has been shown that the region of a core
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promoter that confers TAF1 dependency is the region surrounding the TATA box. However, this may depend on several factors, including the overall core promoter structure and the transcription activators involved, as there is also evidence that activators rather than the core promoter specify TAF1 dependency (Wang et al., 1997; Weissman et al., 2000). The loosely conserved sequence surrounding the start site that can direct accurate transcription in a TATA-less promoter was identified as the Inr (Smale and Baltimore, 1989). In composite promoters with both TATA and Inr, the two elements can synergistically function in transcription initiation. Several proteins have been identified as Inr-binding proteins, including some TAF subunits of TFIID (Novina and Roy, 1996). Several lines of evidence indicate that TAFs may directly contact DNA in this region and may be responsible for Inr function. First, the activity of Inr in the in vitro transcription assays is usually observed with TFIID, but not recombinant TBP (Nakatani et al., 1990; Smale et al., 1990). Second, TFIID also has affinity for specific sequence in the region around the start site. The preferred sequence derived from selection of randomized promoter sequence is similar to the Inr consensus, YYAþ1NT/AYY (Purnell et al., 1994). Furthermore, DNaseI footprinting of TFIID shows an extended footprint over the start site on some promoters, unlike TBP, which shows only a small footprint around the TATA box (Kaufmann and Smale, 1994; Nakatani et al., 1990; Sawadogo and Roeder, 1985). Further studies show that TAF2 has a sequence-specific DNA binding activity and shows a footprint over the start site, overlapping with the Inr (Verrijzer et al., 1994). Photocrosslinking identified TAF1 and TAF2 as subunits in proximity to promoter DNA, both in the context of TFIID and of a trimeric TBP-TAF1-TAF2 complex (Verrijzer et al., 1995). The trimeric complex can also functionally distinguish TATA-containing and Inrcontaining promoters from promoters without an Inr. Although DNA binding-site selection of TAF1 or TAF2 alone does not show preference for the Inr, the Drosophila TAF1-TAF2 complex selects a sequence that matches the Inr consensus (Chalkley and Verrijzer, 1999). These results indicate that TAF1 and TAF2 may be the TFIID subunits that mediate Inr function. Interestingly, TAF2 also shows an affinity for a four-way junction DNA, indicating that TFIID-DNA interaction may also be partly mediated by DNA structure recognition (Chalkley and Verrijzer, 1999). The third type of core promoter element is the DPE. The DPE is found in higher eukaryotes, from Drosophila to human, at approximately 30 bp downstream of the start site with a consensus RGWCGTG (Burke and Kadonaga, 1996, 1997). A photocrosslinking study indicates that Drosophila TAF6 and TAF9 interact with the DPE (Burke and Kadonaga, 1997). TAF6 and TAF9 share homology with histone H4 and H3 in the histone fold
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motifs (Xie et al., 1996). In Drosophila, among 205 examined promoters, the DPE is used as frequently as the TATA box (Kutach and Kadonaga, 2000). Interestingly, there is a strict spacing between the Inr and the DPE, and the sequences of Inr in these DPE-containing promoters adhere more to the consensus than in DPE-less promoters. This is consistent with a cooperation between the Inr and the DPE and may indicate a quite rigid architecture of the part of TFIID that consists of TAF1, TAF2, TAF6, and TAF9. In conclusion, TFIID interacts with the core promoter elements through many specific pairs of protein–DNA interaction. These interactions allow TFIID to distinguish different promoter structures and respond properly to various regulatory signals.
B. Mediators of Activation Since their discovery, TAFs have been implicated in communications between activators and the basal transcription machinery (Dynlacht et al., 1991; Pugh and Tjian, 1991; Tanese et al., 1991). It is believed that activators can increase the rate of transcription by stabilizing binding of the transcription machinery to the promoter through direct interactions (Ptashne and Gann, 1997). Although activators can influence the transcription process by several mechanisms, the recruitment of general transcription factors and Pol II/mediator complexes seems likely (Struhl, 1999). ‘‘Activator bypass’’ experiments, in which various components of the basal transcription machinery are fused to a DNA binding domain, lead to an increase in transcription without the need for the activation domain of an activator (Ptashne and Gann, 1997). Because TBP or TFIID is the first to bind the core promoter and direct PIC assembly, TFIID is likely to be an important target for many activators. It has been shown that direct recruitment of TBP to the promoter by creating a contact with, or fusing it to, a DNA binding domain bypasses the requirement for an activation domain (Chatterjee and Struhl, 1995; Klages and Strubin, 1995). Similarly, fusing a DNA binding domain to several TAFs also leads to increased transcription (Apone et al., 1996; Gonzalez-Couto et al., 1997; Keaveney and Struhl, 1998). Occupancy of TBP at promoters in vivo was examined by chromatin immunoprecipitation (ChIP) assay (Kuras and Struhl, 1999; Li et al., 1999). The results indicate that the level of TBP occupancy at promoters correlates well with transcription level and is stimulated by activators. Interestingly, TBP occupancy is dependent on Srb4, a subunit of the Pol II holoenzyme. This indicates that TBP recruitment is an important mechanism of transcriptional activation in vivo and is interdependent with holoenzyme recruitment.
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Although many direct contacts between various activators and TAFs have been documented in vitro (reviewed in Sauer and Tjian, 1997; Verrijzer and Tjian, 1996), it is less clear which of these interactions occur in the context of the transcription complex in vivo. Using TFIID subcomplexes assembled with recombinant proteins (Chen et al., 1994; Sauer et al., 1995a; 1995b), it was shown that the presence of the specific TAF that makes contact with a particular activator in the TFIID complex is required for activated transcription in vitro. If the activators interact with different TAF subunits, a synergistic increase in transcription level is seen with the subcomplex containing the target TAFs of all activators. Promoter swapping experiments showed that a DNA element that confers TAF1 dependency on a promoter in mammalian cells overlaps with activator binding sites (Wang et al., 1997; Weissman et al., 2000). This provided further support for the role of TAFs as coactivators, although there is also evidence for core promoter element as the determinant of TAF1 dependency in yeast (Shen and Green 1997; Tsukihashi et al., 2000, 2001). These results led some to suggest that TFIID is the major target of activators. However, in some in vitro systems, activated transcription can occur in the absence of TAFs, arguing against an absolute requirement for TAFs (Fondell et al., 1999; Koleske and Young, 1994; Oelgeschlager et al., 1998; Wu et al., 1998). In these cases, transcriptional activation depends on another coactivator complex known as the Mediator complex (see Myers and Kornberg, 2000; Naar et al., 1998 for review). Mediator can also function cooperatively with TFIID for synergistic level of activation (Naar et al., 1999). Therefore, in vivo many more factors are involved in determining the rate-limiting steps that are susceptible to the influences of activators, and the mechanisms of activation may differ from promoter to promoter.
C. Enzymatic Activities 1. Kinase Activity Human TAF1 is claimed to possess two autophosphorylating serine/ threonine kinase activities in the N-terminal and C-terminal domains (Dikstein et al., 1996a). Both kinase domains can also contribute to phosphorylation of the RAP74 subunit of TFIIF in vitro. Phosphorylation of the large subunit of TFIIA on multiple serine residues occurs in both yeast and human cells and can be mediated by human TAF1 in vitro (Solow et al., 2001). However, whether these factors are physiological substrates for hsTAF1 requires further investigation.
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The putative kinase domains are not highly conserved over evolution. The N-terminal kinase domain has weak similarity to a protein kinase family, whereas the C-terminal domains shows no similarity to any known kinase. Further characterization of the N-terminal kinase domain indicates that acidic residues in two small regions are important for the kinase activity (O’Brien and Tjian, 1998). Combined mutation of both regions disrupts kinase activity, and the mutant has reduced ability to rescue the ts 13 cell line, a ts hamster cell line with a mutation in TAF1. This mutant shows a defect in transcription of approximately 6% in a genome-wide analysis (O’Brien and Tjian, 2000), indicating that the kinase activity of the N-terminal domain may be required for expression of a subset of genes in vivo. Less is known about the TAF1 C-terminal kinase domain. The lack of conservation with typical protein kinases and among homologs makes analysis of this domain more difficult. This domain and two bromodomains were thought to be missing from yeast TFIID, as they are not present in the yeast TAF1 homolog. However, it appears that a protein called Bromodomain Factor 1 (Bdf1) can associate with yeast TFIID and corresponds to the missing part of TAF1 (Matangkasombut et al., 2000). Bdf1 purified from yeast is also phosphorylated by an associated kinase activity. However, Bdf1 is not itself a kinase, and phosphorylation is apparently carried out by an associated kinase (C. Sawa and S. Buratowski, unpublished results). Whether this also turns out to be true of TAF1 from higher eukaryotes remains to be seen.
2. Acetyltransferase Activity Recent years have uncovered a critical role of HAT activities in regulating transcription on chromatin templates. Using an activity gel assay, Mizzen et al. (1996) reported evidence for HAT activity in TAF1 homologs from yeast, Drosophila, and human. -TBP antibody immunoprecipitates from HeLa cell extract, and recombinant Drosophila TAF1 and yeast TAF1 exhibit acetyltransferase activity in vitro. The preferred substrates are histone H3, especially at lysine 14, and histone H4. There is also a report that TAF1 can acetylate the large subunit of TFIIE in vitro (Imhof et al., 1997). However, it is not yet clear whether any of these reactions are physiological. In vitro, acetyl-CoA can stimulate both basal and activated transcription in the absence of histones by increasing the affinity of TFIID to promoter DNA in a TAF-dependent fashion (Galasinski et al., 2000). Thus, it is possible that a substrate may be in the TFIID complex itself, or the mere binding of acetyl-CoA to TAF1 induces a conformational change that favors DNA binding of TFIID. Acetylation of histones by TAF1 could
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be a mechanism that helps TFIID gain access to chromatin-bound promoter DNA. Alternatively, acetylated histones may promote TFIID binding via the bromodomains in TAF1 (see below). The HAT domain maps to a highly conserved middle portion of TAF1 (Mizzen et al., 1996). This domain does not show significant similarity to the acetyl-CoA binding sites of other HATs. However, a study on the ts hamster cell line, ts13, containing a point mutation (G716D) in the HAT domain of TAF1, indicates that HAT activity is important for TAF1 function in vivo (Dunphy et al., 2000). The mutant protein has reduced HAT activity in vitro at the nonpermissive temperature but shows no defect in kinase activity or interaction with other TFIID subunits. A putative acetylCoA binding site has been identified, and mutation of two glycine residues in this site gives rise to a HAT-defective mutant that can neither complement the growth arrest phenotype nor rescue transcription from the TAF1dependent cyclin A and D1 promoters in ts13 cell lines. A genome-wide study in the ts13 cell line indicates that approximately 18% of genes are differentially expressed (over two fold) at the nonpermissive temperature (O’Brien and Tjian, 2000).
3. Monoubiquitinating Activity Because monoubiquitination of histones is potentially involved in transcriptional activation, an activity gel assay was employed to identify candidate proteins with this activity (Pham and Sauer, 2000). In Drosophila embryonic nuclear extract, TAF1 was found to be able to monoubiquitinate histone H1 in vitro. This activity maps to the middle portion of the protein overlapping with the HAT domain. Fortuitously, two TAF1 mutants containing point mutations in this putative domain have previously been identified (Wassarman et al., 2000). These mutant proteins show wild-type levels of HAT activity but are unable to ubiquitinate histone H1 in vitro. Moreover, the bulk of H1 from the mutant embryos shows a decreased level of monoubiquitinated form. The in vivo defects in transcriptional activation and embryonic development of these two mutant lines might be explained by the lack of this enzymatic activity. Nevertheless, further investigation will be needed to clarify the role of this potential activity of TAF1.
IV. The Requirement for TFIID In Vivo Genetic analysis in yeast confirms the importance of TAFs, as all (with the exception of yeast TAF14, a protein found in several transcriptionrelated complexes) are essential for viability. However, the requirement for individual TAFs in general Pol II transcription is still unclear. Various
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conditional expression systems have been used to inactivate individual TAFs (Apone et al., 1996, 1998; Durso et al., 2001; Komarnitsky et al., 1999; Michel et al., 1998; Moqtaderi et al., 1996, 1998; Natarajan et al., 1998; Reese et al., 2000; Sanders et al., 1999; Walker et al., 1997). The results have been somewhat confusing. In some cases, widespread loss of transcription is observed. However, in others the transcription effects were quite limited. A consensus seems to be emerging that at least one TAF is required at most promoters and that a broad defect in transcription is seen when the entire TFIID complex is disrupted (Green, 2000; Lee et al., 2000). Therefore, it is very likely that a functional TFIID complex is required for transcription of most genes. However, it is equally clear that at least some promoters can continue transcribing after TAFs have been inactivated. ChIP experiments show that some of these promoters have a reduced TAF/TBP ratio compared with most promoters (Kuras et al., 2000; Li et al., 2000). It is possible that these promoters primarily use free TBP to support transcription. The roles of individual TAFs and TFIID have also been examined in multicellular organisms. In C. elegans, a differential requirement for individual TAFs is also seen. The worm taf-4, but not taf-9 or taf-10, appears to be required globally for transcription (Walker et al., 2001). Studies in Drosophila show that partial loss of function mutations in TAF genes lead to developmental defects (Aoyagi and Wassarman, 2001; Hiller et al., 2001; Pham et al., 1999; Zhou et al., 1998). Because most of the fly TAFs are essential for viability, many experiments were performed with variant TAFs that were discussed in the section on alternative TFIIDs. Mutations in the tissue-specific TAF5L and in TAF6 lead to defective spermatogenesis (Hiller et al., 2001). Interestingly, mice lacking the tissue-specific TAF4B show defects in ovarian development (Freiman et al., 2001). This indicates that the process of gametogenesis may require a high level of regulated transcription and is sensitive to alterations in a variant TFIID. The hamster cell line ts13 has a mutation in the TAF1 gene and has been used to study the role of this TAF in the mammalian system. The cells are ts and arrest in G1 on shifting to the nonpermissive temperature. This is strikingly similar to the phenotype of the yeast TAF1 ts mutant. Intriguingly, there appears to be another link between TAFs and the cell cycle: a high copysuppressor screen of a TAF12 mutant allele identified several genes involved in G2/M control (Reese and Green, 2001). This could be the result of a requirement for a tight transcriptional control during cell cycle progression, similar to gametogenesis. In apparent contrast to these other results, a TAF9 knockout in chicken DT40 cells showed little effect on transcription despite the fact that the cells lost viability (Chen and Manley, 2000).
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In summary, the in vivo data indicate that there may be a differential requirement for individual TAFs, but in general, TFIID complex is probably required at most promoters for transcription initiation. Furthermore, experiments in metazoans indicate an important role for TAFs during developmental processes that require strictly regulated transcription patterns.
V. Regulation of TFIID As discussed above, TFIID binds to the promoter and recruits the rest of the basal factors for transcription initiation. This represents a crucial step that needs to be tightly regulated. Transcription activators may interact directly with TFIID to enhance promoter binding and stabilize this complex. However, to ensure that transcription occurs only at the right promoters at the right time, cells also need to prevent TFIID from binding to nonpromoter sequence or inactive promoters as well. A number of negative regulatory mechanisms exist, with many counteracted by the activity of TFIIA (reviewed in Lee and Young, 1998; Pugh, 1996). These include TBP dimerization, an inhibitory domain of TAF1, Mot1, and NC2, which are briefly described here. TFIIA interacts directly with TBP and also with TAF11 (Kraemer et al., 2001). The interaction stabilizes the TFIID-promoter DNA complex and may induce a conformational change in TFIID that facilitates further PIC assembly (Chi and Carey, 1996). TFIIA is also a target of activators and plays an important role against many of the following inhibitory mechanisms. TBP can form homodimers when not bound to DNA (Coleman et al., 1995). This could inhibit DNA binding, as the dimer formation blocks the DNA binding surface. Mutations in the dimer interface that impair dimerization in vitro lead to an increase in activator-independent transcription and rapid degradation of TBP, indicating that dimerization may be an important mechanism that prevents unregulated transcription and TBP degradation in vivo ( Jackson-Fisher et al., 1999). TFIIA promotes dissociation of TBP and TFIID dimerization, thereby increasing the rate of TBP/TFIID binding to DNA (Coleman et al., 1999). In the context of TFIID, a TAF can also function as an inhibitor of TBP-DNA interactions. The N-terminus of TAF1 has been found to interact with TBP and block TBP-DNA interaction by mimicking the surface of partially unwound minor groove of TATA box when bound to TBP (Kokubo et al., 1993; Liu et al., 1998). This inhibitory interaction is counteracted by TFIIA and by c-Jun, a transcription activator (Kokubo et al., 1998; Lively et al., 2001).
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Another inhibitor of TBP is Mot1, a member of the SNF2 family of DNA-dependent ATPases. Mot1 dissociates the TBP-DNA complex in an ATP-dependent manner (Auble and Hahn, 1993; Auble et al., 1994). ChIP experiments show that this occurs in vivo, as TBP occupancy at several promoters is increased in a mot1 mutant yeast strain (Li et al., 1999). Therefore, Mot1 can negatively regulate a subset of genes by interfering with TBP binding in vivo and could play a positive role in promoter selection by dissociating TBP from non-TATA or weak TATA sequences (Collart, 1996; Muldrow et al., 1999). A human homolog of Mot1 has been identified called BTAF1 (Chicca et al., 1998; van der Knaap et al., 1997). TFIIA can block Mot1 activity (Auble and Hahn, 1993). NC2 is another negative regulator that interacts with the TBP-DNA complex on the underside of the bent DNA. It is composed of two subunits that dimerize via a histone fold interaction. It inhibits TBP-TFIIB interaction and subsequent PIC formation by physically blocking the TBPDNA surface recognized by TFIIB (Goppelt et al., 1996; Kamada et al., 2001). The binding of NC2 to TBP-DNA is also competitive with TFIIA binding. A mutation in TFIIA can suppress the defect caused by reduced dosage of NC2 in yeast (Xie et al., 2000). Interestingly, NC2 can function positively on DPE-containing promoters but represses TATA promoters (Willy et al., 2000). Mutational analysis indicates that the repression and activation functions are separable. Therefore, the effect of NC2 is dependent on the context of core promoters. In summary, TFIID is regulated by many factors, with both negative and positive roles. It is the integrated and counterbalanced effects of these factors that control the many important functions of TFIID. This is necessary to achieve a high level of specific regulation of gene expression in vivo.
VI. The TFIID-Chromatin Connection Both TFIID and chromatin structure play vital roles in transcriptional regulation. For a precise control of gene expression, the combination of transcription activators, core promoter elements, local and large-scale chromatin structures determine the requirements for various factors and choreographs their activity and interactions at each promoter. The coordinated action of chromatin regulators is needed for TFIID to gain access to the promoter in the context of chromatin. Therefore, a close connection between TFIID and chromatin regulators is crucial for the coordinated reaction essential for transcription regulation.
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Examples of this connection are evident. Several components of the HAT complexes interact directly with TBP and TFIID. For instance, the Spt3 and Spt8 subunits of the SAGA complex interact with TBP and are essential for TFIID recruitment to the promoter (Bhaumik and Green, 2001; Dudley et al., 1999;). Moreover, a subset of TAFs is shared between SAGA and TFIID (Grant et al., 1998). Although the functional significance of this is not yet known, it is nevertheless intriguing that the same set of proteins is present in the two functionally related complexes. Another example of this connection is the presence of HAT activity in the TAF1 subunit of TFIID (Mizzen et al., 1996). As discussed earlier, there is evidence indicating TAF1 has HAT activity (Dunphy et al., 2000). Although it is not yet clear what the substrates and the function of this activity are in vivo, a speculative model is that TAF1 acetylates histones in nearby nucleosomes to maintain the active state of the promoter or to create a chromatin structure suitable for subsequent recruitment of RNA Pol II machinery. Alternatively, the acetyltransferase activity in TAF1 may target other members of the transcription machinery and alter their activity or function (Imhof et al., 1997). Furthermore, the HAT activity of TAF1 may alleviate the requirement for the HAT activity of Gcn5 or another HAT with similar substrate specificity (Lee et al., 2000). The histone code hypothesis suggests that transcription factors can in part be targeted to promoters through interactions with specifically modified histone tails ( Jenuwein and Allis, 2001; Strahl and Allis, 2000). In this respect it is very interesting that TAF1 (or Bdf1 in yeast) contains two copies of the bromodomain, a conserved motif that has been proposed to interact specifically with acetylated lysines of histone tails (Dhalluin et al., 1999; Jeanmougin et al., 1997; Winston and Allis, 1999). Acetylation of histone tails at several sites correlates with the transcriptionally active state (Grunstein, 1997). The presence of this motif in TFIID could provide a direct link between TFIID and chromatin modification. The crystal structure of the double bromodomain of TAF1 was solved, and it was shown by isothermal titration calorimetric studies that this module interacts strongly with acetylated histone H4 peptide, especially the di- or tetraacetylated forms (Kd of 1–5 M) but not with the nonacetylated form ( Jacobson et al., 2000). The bromodomains of Bdf1 are essential for its function and proper gene expression (Chua and Roeder, 1995; Ladurner et al., 2003; Matangkasombut and Buratowski, 2003). Recent genetic and biochemical evidence shows that the interaction between the bromodomains and acetylated histone H4 is important in vivo (Ladurner et al., 2003; Matangkasombut et al., 2000). However, Bdf1 is located throughout the genome, not only at promoters, indicating that this protein is not solely a subunit of TFIID.
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VII. Future Questions Although great progress has been made in recent years, there is still a great deal to be discovered about TFIID. The functions of all the individual subunits need to be discovered, not only in the context of TFIID but also in other TAF-containing complexes. This task will be greatly aided when a high-resolution structure of TFIID becomes available. There is still debate about which transcription events are supported by TFIID and which are TAF independent. Only when we understand how TFIID interacts with activators, chromatin, and the rest of the transcription machinery will all these questions be answered.
References Albright, S. R., and Tjian, R. (2000). Gene 242, 1–13. Andel, F., 3rd, Ladurner, A. G., Inouye, C., Tjian, R., and Nogales, E. (1999). Science 286, 2153–2156. Aoyagi, N., and Wassarman, D. A. (2001). Mol. Cell. Biol. 21, 6808–6819. Apone, L. M., Virbasius, C. A., Holstege, F. C., Wang, J., Young, R. A., and Green, M. R. (1998). Mol. Cell 2, 653–661. Apone, L. M., Virbasius, C. M., Reese, J. C., and Green, M. R. (1996). Genes Dev. 10, 2368–2380. Auble, D. T., and Hahn, S. (1993). Genes Dev. 7, 844–856. Auble, D. T., Hansen, K. E., Mueller, C. G., Lane, W. S., Thorner, J., and Hahn, S. (1994). Genes Dev. 8, 1920–1934. Bell, B., Scheer, E., and Tora, L. (2001). Mol. Cell 8, 591–600. Bhaumik, S. R., and Green, M. R. (2001). Genes Dev. 15, 1935–1945. Birck, C., Poch, O., Romier, C., Ruff, M., Mengus, G., Lavigne, A. C., Davidson, I., and Moras, D. (1998). Cell 94, 239–249. Brand, M., Leurent, C., Mallouh, V., Tora, L., and Schultz, P. (1999). Science 286, 2151–2153. Brand, M., Moggs, J. G., Oulad-Abdelghani, M., Lejeune, F., Dilworth, F. J., Stevenin, J., Almouzni, G., and Tora, L. (2001). EMBO J. 20, 3187–3196. Buratowski, S., Hahn, S., Sharp, P. A., and Guarente, L. (1988). Nature 334, 37–42. Burke, T. W., and Kadonaga, J. T. (1996). Genes Dev. 10, 711–724. Burke, T. W., and Kadonaga, J. T. (1997). Genes Dev. 11, 3020–3031. Burley, S. K., and Roeder, R. G. (1996). Annu. Rev. Biochem. 65, 769–799. Butler, J. E., and Kadonaga, J. T. (2001). Genes Dev. 15, 2515–2519. Cavallini, B., Faus, I., Matthes, H., Chipoulet, J. M., Winsor, B., Egly, J. M., and Chambon, P. (1989). Proc. Natl. Acad. Sci. USA 86, 9803–9807. Cavallini, B., Huet, J., Plassat, J. L., Sentenac, A., Egly, J. M., and Chambon, P. (1988). Nature 334, 77–80. Chalkley, G. E., and Verrijzer, C. P. (1999). EMBO J. 18, 4835–4845. Chatterjee, S., and Struhl, K. (1995). Nature 374, 820–822. Chen, J. L., Attardi, L. D., Verrijzer, C. P., Yokomori, K., and Tjian, R. (1994). Cell 79, 93–105. Chen, Z., and Manley, J. L. (2000). Mol. Cell. Biol. 20, 5064–5076. Chi, T., and Carey, M. (1996). Genes Dev. 10, 2540–2550.
88
MATANGKASOMBUT ET AL.
Chicca, J. J., Auble, D. T., and Pugh, B. F. (1998). Mol. Cell. Biol. 18, 1701–1710. Chua, P., and Roeder, G. S. (1995). Mol. Cell. Biol. 15:, 3685–3696. Coleman, R. A., Taggart, A. K., Benjamin, L. R., and Pugh, B. F. (1995). J. Biol. Chem. 270, 13842–13849. Coleman, R. A., Taggart, A. K., Burma, S., Chicca, J. J., and Pugh, B. F. (1999). Mol. Cell 4, 451–457. Collart, M. A. (1996). Mol. Cell. Biol. 16, 6668–6676. Crowley, T. E., Hoey, T., Liu, J. K., Jan, Y. N., Jan, L. Y., and Tjian, R. (1993). Nature 361, 557–561. Dantonel, J. C., Quintin, S., Lakatos, L., Labouesse, M., and Tora, L. (2000). Mol. Cell 6, 715–722. Dantonel, J. C., Wurtz, J. M., Poch, O., Moras, D., and Tora, L. (1999). Trends Biochem. Sci. 24, 335–339. Dhalluin, C., Carlson, J. E., Zeng, L., He, C., Aggarwa, A. K., and Zhou, M. M. (1999). Nature 399, 491–496. Dikstein, R., Ruppert, S., and Tjian, R. (1996a). Cell 84, 781–790. Dikstein, R., Zhou, S., and Tjian, R. (1996b). Cell 87, 137–146. Dudley, A. M., Rougeulle, C., and Winston, F. (1999). Genes Dev. 13, 2940–2945. Dunphy, E. L., Johnson, T., Auerbach, S. S., and Wang, E. H. (2000). Mol. Cell. Biol. 20, 1134–1139. Durso, R. J., Fisher, A. K., Albright-Frey, T. J., and Reese, J. C. (2001). Mol. Cell. Biol. 21, 7331–7344. Dynlacht, B. D., Hoey, T., and Tjian, R. (1991). Cell 66, 563–576. Eisenmann, D. M., Dollard, C., and Winston, F. (1989). Cell 58, 1183–1191. Fondell, J. D., Guermah, M., Malik, S., and Roeder, R. G. (1999). Proc. Natl. Acad. Sci. USA 96, 1959–1964. Freiman, R. N., Albright, S. R., Zheng, S., Sha, W. C., Hammer, R. E., and Tjian, R. (2001). Science 293, 2084–2087. Galasinski, S. K., Lively, T. N., Grebe De Barror, A., and Goodrich, J. A. (2000). Mol. Cell. Biol. 20, 1923–1930. Gangloff, Y. G., Pointud, J. C., Thuault, S., Carre, L., Romier, C., Muratoglu, S., Brand, M., Tora, L., Couderc, J. L., and Davidson, I. (2001a). Mol. Cell. Biol. 21, 5109–5121. Gangloff, Y. G., Romier, C., Thuault, S., Werten, S., and Davidson, I. (2001b). Trends Biochem. Sci. 26, 250–257. Gangloff, Y. G., Sanders, S. L., Romier, C., Kirschner, D., Weil, P. A., Tora, L., and Davidson, I. (2001c). Mol. Cell. Biol. 21, 1841–1853. Gangloff, Y. G., Werten, S., Romier, C., Carre, L., Poch, O., Moras, D., and Davidson, I. (2000). Mol. Cell. Biol. 20, 340–351. Gonzalez-Couto, E., Klages, N., and Strubin, M. (1997). Proc. Natl. Acad. Sci. USA 94, 8036–8041. Goppelt, A., Stelzer, G., Lottspeich, F., and Meisterernst, M. (1996). EMBO J. 15, 3105–3116. Grant, P. A., Schieltz, D., Pray-Grant, M. G., Steger, D. J., Reese, J. C., Yates, J. R., 3rd, and Workman, J. L. (1998). Cell 94, 45–53. Green, M. R. (2000). Trends Biochem. Sci. 25, 59–63. Grunstein, M. (1997). Nature 389, 349–352. Hahn, S., Buratowski, S., Sharp, P. A., and Guarente, L. (1989). Cell 58, 1173–1181. Hansen, S. K., Takada, S., Jacobson, R. H., Lis, J. T., and Tjian, R. (1997). Cell 91, 71–83. Hernandez, N. (1993). Genes Dev. 7, 1291–1308.
STRUCTURE AND FUNCTION OF THE TFIID COMPLEX
89
Hiller, M. A., Lin, T. Y., Wood, C., and Fuller, M. T. (2001). Genes Dev. 15, 1021–1030. Hoey, T., Dynlacht, B. D., Peterson, M. G., Pugh, B. F., and Tjian, R. (1990). Cell 61, 1179–1186. Hoffman, A., Sinn, E., Yamamoto, T., Wang, J., Roy, A., Horikoshi, M., and Roeder, R. G. (1990). Nature 346, 387–390. Hoffmann, A., Chiang, C. M., Oelgeschlager, T., Xie, X., Burley, S. K., Nakatani, Y., and Roeder, R. G. (1996). Nature 380, 356–359. Hoffmann, A., Oelgeschlager, T., and Roeder, R. G. (1997). Proc. Natl. Acad. Sci. USA 94, 8928–8935. Holmes, M. C., and Tjian, R. (2000). Science 288, 867–870. Horikoshi, M., Wang, C. K., Fujii, H., Cromlish, J. A., Weil, P. A., and Roeder, R. G. (1989). Nature 341, 299–303. Imbalzano, A. N., Kwon, H., Green, M. R., and Kingston, R. E. (1994). Nature 370, 481–485. Imhof, A., Yang, X. J., Ogryzko, V. V., Nakatani, Y., Wolffe, A. P., and Ge, H. (1997). Curr. Biol. 7, 689–692. Jackson-Fisher, A. J., Chitikila, C., Mitra, M., and Pugh, B. F. (1999). Mol. Cell 3, 717–727. Jacobson, R. H., Ladurner, A. G., King, D. S., and Tjian, R. (2000). Science 288, 1422–1425. Jeanmougin, F., Wurtz, J. M., Le Douarin, B., Chambon, P., and Losson, R. (1997). Trends Biochem. Sci. 22, 151–153. Jenuwein, T., and Allis, C. D. (2001). Science 293, 1074–1080. Kaltenbach, L., Horner, M. A., Rothman, J. H., and Mango, S. E. (2000). Mol. Cell 6, 705–713. Kamada, K., Shu, F., Chen, H., Malik, S., Stelzer, G., Roeder, R. G., Meisterernst, M., and Burley, S. K. (2001). Cell 106, 71–81. Kao, C. C., Lieberman, P. M., Schmidt, M. C., Zhou, Q., Pei, R., and Berk, A. J. (1990). Science 248, 1646–1650. Kaufmann, J., and Smale, S. T. (1994). Genes Dev. 8, 821–829. Keaveney, M., and Struhl, K. (1998). Mol. Cell 1, 917–924. Kim, J. L., and Burley, S. K. (1994). Nat. Struct. Biol. 1, 638–653. Kim, J. L., Nikolov, D. B., and Burley, S. K. (1993a). Nature 365, 520–527. Kim, Y., Geiger, J. H., Hahn, S., and Sigler, P. B. (1993b). Nature 365, 512–520. Klages, N., and Strubin, M. (1995). Nature 374, 822–823. Kokubo, T., Gong, D. W., Yamashita, S., Horikoshi, M., Roeder, R. G., and Nakatani, Y. (1993). Genes Dev. 7, 1033–1046. Kokubo, T., Swanson, M. J., Nishikawa, J. I., Hinnebusch, A. G., and Nakatani, Y. (1998). Mol. Cell. Biol. 18, 1003–1012. Koleske, A. J., and Young, R. A. (1994). Nature 368, 466–469. Komarnitsky, P. B., Michel, B., and Buratowski, S. (1999). Genes Dev. 13, 2484–2489. Kraemer, S. M., Ranallo, R. T., Ogg, R. C., and Stargell, L. A. (2001). Mol. Cell. Biol. 21, 1737–1746. Kuras, L., Kosa, P., Mencia, M., and Struhl, K. (2000). Science 288, 1244–1248. Kuras, L., and Struhl, K. (1999). Nature 399, 609–613. Kutach, A. K., and Kadonaga, J. T. (2000). Mol. Cell. Biol. 20, 4754–4764. Ladurner, A. G., Inouye, C., Jain, R., and Tjian, R. (2003). Mol. Cell 11, 365–376. Lee, T. I., Causton, H. C., Holstege, F. C., Shen, W. C., Hannett, N., Jennings, E. G., Winston, F., Green, M. R., and Young, R. A. (2000). Nature 405, 701–704. Lee, T. I., and Young, R. A. (1998). Genes Dev. 12, 1398–1408.
90
MATANGKASOMBUT ET AL.
Leurent, C., Sanders, S., Ruhlmann, C., Mallouh, V., Weil, P. A., Kirschner, D. B., Tora, L., and Schultz, P. (2002). EMBO J. 21, 3424–3433. Li, X. Y., Bhaumik, S. R., and Green, M. R. (2000). Science 288, 1242–1244. Li, X. Y., Virbasius, A., Zhu, X., and Green, M. R. (1999). Nature 399, 605–609. Liu, D., Ishima, R., Tong, K. I., Bagby, S., Kokubo, T., Muhandiram, D. R., Kay, L. E., Nakatani, Y., and Ikura, M. (1998). Cell 94, 573–583. Lively, T. N., Ferguson, H. A., Galasinski, S. K., Seto, A. G., and Goodrich, J. A. (2001). J. Biol. Chem. 276, 25582–25588. Luger, K., Mader, A. W., Richmond, R. K., Sargent, D. F., and Richmond, T. J. (1997). Nature 389, 251–260. Marmorstein, R., and Berger, S. L. (2001). Gene 272, 1–9. Martianov, I., Brancorsini, S., Gansmuller, A., Parvinen, M., Davidson, I., and SassoneCorsi, P. (2002). Development 129, 945–955. Martianov, I., Fimia, G. M., Dierich, A., Parvinen, M., Sassone-Corsi, P., and Davidson, I. (2001). Mol. Cell 7, 509–515. Matangkasombut, O., Buratowski, R. M., Swilling, N. W., and Buratowski, S. (2000). Genes Dev. 14, 951–962. Matangkasombut, O., and Buratowski, S. (2003). Mol. Cell 11, 353–363. Matsui, T., Segall, J., Weil, P. A., and Roeder, R. G. (1980). J. Biol. Chem. 255, 11992–11996. Michel, B., Komarnitsky, P., and Buratowski, S. (1998). Mol. Cell 2, 663–673. Mitsiou, D. J., and Stunnenberg, H. G. (2000). Mol. Cell 6, 527–537. Mizzen, C. A., Yang, X. J., Kokubo, T., Brownell, J. E., Bannister, A. J., Owen-Hughes, T., Workman, J., Wang, L., Berger, S. L., Kouzarides, T., Nakatani, Y., and Allis, C. D. (1996). Cell 87, 1261–1270. Moore, P. A., Ozer, J., Salunek, M., Jan, G., Zerby, D., Campbell, S., and Lieberman, P. M. (1999). Mol. Cell. Biol. 19, 7610–7620. Moqtaderi, Z., Bai, Y., Poon, D., Weil, P. A., and Struhl, K. (1996). Nature 383, 188–191. Moqtaderi, Z., Keaveney, M., and Struhl, K. (1998). Mol. Cell 2, 675–682. Muhich, M. L., Iida, C. T., Horikoshi, M., Roeder, R. G., and Parker, C. S. (1990). Proc. Natl. Acad. Sci. USA 87, 9148–9152. Muldrow, T. A., Campbell, A. M., Weil, P. A., and Auble, D. T. (1999). Mol. Cell. Biol. 19, 2835–2845. Muller, F., Lakatos, L., Dantonel, J., Strahle, U., and Tora, L. (2001). Curr. Biol. 11, 282–287. Myers, L. C., and Kornberg, R. D. (2000). Annu. Rev. Biochem. 69, 729–749. Naar, A. M., Beaurang, P. A., Robinson, K. M., Oliner, J. D., Avizonis, D., Scheek, S., Zwicker, S., Kadonaga, J. T., and Tjian, R. (1998). Genes Dev. 12, 3020–3031. Naar, A. M., Beaurang, P. A., Zhou, S., Abraham, S., Solomon, W., and Tjian, R. (1999). Nature 398, 828–832. Nakatani, Y., Horikoshi, M., Brenner, M., Yamamoto, T., Besnard, F., Roeder, R. G., and Freese, E. (1990). Nature 348, 86–88. Natarajan, K., Jackson, B. M., Rhee, E., and Hinnebusch, A. G. (1998). Mol. Cell 2, 683–692. Nikolov, D. B., and Burley, S. K. (1997). Proc. Natl. Acad. Sci. USA 94, 15–22. Novina, C. D., and Roy, A. L. (1996). Trends Genet. 12, 351–355. O’Brien, T., and Tjian, R. (1998). Mol. Cell 1, 905–911. O’Brien, T., and Tjian, R. (2000). Proc. Natl. Acad. Sci. USA 97, 2456–2461. Oelgeschlager, T., Chiang, C. M., and Roeder, R. G. (1996). Nature 382, 735–738. Oelgeschlager, T., Tao, Y., Kang, Y. K., and Roeder, R. G. (1998). Mol. Cell 1, 925–931.
STRUCTURE AND FUNCTION OF THE TFIID COMPLEX
91
Ogryzko, V. V., Kotani, T., Zhang, X., Schiltz, R. L., Howard, T., Yang, X. J., Howard, B. H., Qin, J., and Nakatani, Y. (1998). Cell 94, 35–44. Peterson, M. G., Tanese, N., Pugh, B. F., and Tjian, R. (1990). Science 248, 1625–1630. Pham, A. D., Muller, S., and Sauer, F. (1999). Mech. Dev. 84, 3–16. Pham, A. D., and Sauer, F. (2000). Science 289, 2357–2360. Poon, D., Bai, Y., Campbell, A. M., Bjorklund, S., Kim, Y. J., Zhou, S., Kornberg, R. D., and Weil, P. A. (1995). Proc. Natl. Acad. Sci. USA 92, 8224–8228. Ptashne, M., and Gann, A. (1997). Nature 386, 569–577. Pugh, B. F. (1996). Curr. Opin. Cell. Biol. 8, 303–311. Pugh, B. F. (2000). Gene 255, 1–14. Pugh, B. F., and Tjian, R. (1990). Cell 61, 1187–1197. Pugh, B. F., and Tjian, R. (1991). Genes Dev. 5, 1935–1945. Purnell, B. A., Emanuel, P. A., and Gilmour, D. S. (1994). Genes Dev. 8, 830–842. Reese, J. C., Apone, L., Walker, S. S., Griffin, L. A., and Green, M. R. (1994). Nature 371, 523–527. Reese, J. C., and Green, M. R. (2001). Yeast 18, 1197–1205. Reese, J. C., Zhang, Z., and Kurpad, H. (2000). J. Biol. Chem. 275, 17391–17398. Reinberg, D., Horikoshi, M., and Roeder, R. G. (1987). J. Biol. Chem. 262, 3322–3330. Samuels, M., Fire, A., and Sharp, P. A. (1982). J. Biol. Chem. 257, 14419–14427. Sanders, S. L., Klebanow, E. R., and Weil, P. A. (1999). J. Biol. Chem. 274, 18847–18850. Sanders, S. L., and Weil, P. A. (2000). J. Biol. Chem. 275, 13895–13900. Sauer, F., Hansen, S. K., and Tjian, R. (1995a). Science 270, 1825–1828. Sauer, F., Hansen, S. K., and Tjian, R. (1995b). Science 270, 1783–1788. Sauer, F., and Tjian, R. (1997). Curr. Opin. Genet. Dev. 7, 176–181. Saurin, A. J., Shao, Z., Erdjument-Bromage, H., Tempst, P., and Kingston, R. E. (2001). Nature 412, 655–660. Sawadogo, M., and Roeder, R. G. (1985). Cell 43, 165–175. Schmidt, M. C., Kao, C. C., Pei, R., and Berk, A. J. (1989). Proc. Natl. Acad. Sci. USA 86, 7785–7789. Selleck, W., Howley, R., Fang, Q., Podolny, V., Fried, M. G., Buratowski, S., and Tan, S. (2001). Nat. Struct. Biol. 8, 695–700. Shen, W. C., and Green, M. R. (1997). Cell 90, 615–624. Silkov, A., Wolstein, O., Shachar, I., and Dikstein, R. (2002). J. Biol. Chem. 20, 20. Smale, S. T. (2001). Genes Dev. 15, 2503–2508. Smale, S. T., and Baltimore, D. (1989). Cell 57, 103–113. Smale, S. T., Schmidt, M. C., Berk, A. J., and Baltimore, D. (1990). Proc. Natl. Acad. Sci. USA 87, 4509–4513. Solow, S., Salunek, M., Ryan, R., and Lieberman, P. M. (2001). J. Biol. Chem. 276, 15886–15892. Strahl, B. D., and Allis, C. D. (2000). Nature 403, 41–45. Struhl, K. (1999). Cell 98, 1–4. Takada, S., Lis, J. T., Zhou, S., and Tjian, R. (2000). Cell 101, 459–469. Tanese, N., Pugh, B. F., and Tjian, R. (1991). Genes Dev. 5, 2212–2224. Teichmann, M., Wang, Z., Martinez, E., Tjernberg, A., Zhang, D., Vollmer, F., Chait, B. T., and Roeder, R. G. (1999). Proc. Natl. Acad. Sci. USA 96, 13720–13725. Tora, L. (2002). Genes Dev. 16, 673–675. Tsukihashi, Y., Kawaichi, M., and Kokubo, T. (2001). J. Biol. Chem. 276, 25715–25726. Tsukihashi, Y., Miyake, T., Kawaichi, M., and Kokubo, T. (2000). Mol. Cell. Biol. 20, 2385–2399.
92
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van der Knaap, J. A., Borst, J. W., van der Vliet, P. C., Gentz, R., and Timmers, H. T. (1997). Proc. Natl. Acad. Sci. USA 94, 11827–11832. Veenstra, G. J., Weeks, D. L., and Wolffe, A. P. (2000). Science 290, 2312–2315. Verrijzer, C. P., Chen, J. L., Yokomori, K., and Tjian, R. (1995). Cell 81, 1115–1125. Verrijzer, C. P., and Tjian, R. (1996). Trends Biochem. Sci. 21, 338–342. Verrijzer, C. P., Yokomori, K., Chen, J. L., and Tjian, R. (1994). Science 264, 933–941. Walker, A. K., Rothman, J. H., Shi, Y., and Blackwell, T. K. (2001). EMBO J. 20, 5269–5279. Walker, S. S., Reese, J. C., Apone, L. M., and Green, M. R. (1996). Nature 383, 185–188. Walker, S. S., Shen, W. C., Reese, J. C., Apone, L. M., and Green, M. R. (1997). Cell 90, 607–614. Wang, E. H., Zou, S., and Tjian, R. (1997). Genes Dev. 11, 2658–2669. Wassarman, D. A., Aoyagi, N., Pile, L. A., and Schlag, E. M. (2000). Proc. Natl. Acad. Sci. USA 97, 1154–1159. Weissman, J. D., Howcroft, T. K., and Singer, D. S. (2000). J. Biol Chem. 275, 10160–10167. Wieczorek, E., Brand, M., Jacq, X., and Tora, L. (1998). Nature 393, 187–191. Willy, P. J., Kobayashi, R., and Kadonaga, J. T. (2000). Science 290, 982–985. Winston, F., and Allis, C. D. (1999). Nat. Struct. Biol. 6, 601–604. Wu, S. Y., Kershnar, E., and Chiang, C. M. (1998). EMBO J. 17, 4478–4490. Xie, J., Collart, M., Lemaire, M., Stelzer, G., and Meisterernst, M. (2000). EMBO J. 19, 672–682. Xie, X., Kokubo, T., Cohen, S. L., Mirza, U. A., Hoffmann, A., Chait, B. T., Roeder, R. G., Nakatani, Y., and Burley, S. K. (1996). Nature 380, 316–322. Yamit-Hezi, A., and Dikstein, R. (1998). EMBO J. 17, 5161–5169. Yatherajam, G., Zhang, L., Kraemer, S. M., and Stargell, L. A. (2003). Nucleic Acids Res. 31, 1252–1260. Zhang, D., Penttila, T. L., Morris, P. L., Teichmann, M., and Roeder, R. G. (2001). Science 292, 1153–1155. Zhou, J., Zwicker, J., Szymanski, P., Levine, M., and Tjian, R. (1998). Proc. Natl. Acad. Sci. USA 95, 13483–13488.
TETRATRICOPEPTIDE REPEATS OF TFC4 AND A LIMITING STEP IN THE ASSEMBLY OF THE INITIATION FACTOR TFIIIB By ROBYN D. MOIR AND IAN M. WILLIS Department of Biochemistry, Albert Einstein College of Medicine, Bronx, New York 10461
I. Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Brf1-TFIIIC: A Limiting Interaction in Preinitiation Complex Assembly. .. . . . . . III. Brf1-Tfc4 Interactions . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Structural Domains of Brf1 . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Tetratricopeptide Repeats (TPRs) in Tfc4 . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Brf1 Binding to Tfc4 and Autoinhibition . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Effects of Mutations in Tetratricopeptide Repeat Domains on Brf1 Binding . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Bdp1-Tfc4 Interactions . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Ligand Binding by TPR Arrays in Tfc4 . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Identification of Potential Ligand Binding Sites in the TPR Arrays of Tfc4. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Models for Tetratricopeptide Repeat Organization in Tfc4. . . . . . . . . . . .. . . . . . VI. Ordered Binding of TFIIIB Subunits to TFIIIC-DNA and Dynamic Interactions with Tfc4 . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VII. Tfc4 and Other Pol III Factors.. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VIII. Conclusion . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction Transcription by RNA polymerase (Pol) III is of fundamental importance in all eukaryotes because its products, which include 5S RNA, tRNA, U6 snRNA, RNase P RNA, and 7SL RNA, are essential for protein synthesis, RNA processing, protein transport, and other cellular processes. Pol III genes employ three distinct promoter architectures: type I promoters are represented solely by 5S RNA genes and contain unique intragenic sequence elements for binding the 5S gene-specific factor TFIIIA; type II promoters are found in a functionally diverse group of genes and are distinguished by typically intragenic A and B block elements, initially defined in tRNA genes, that serve as binding sites for the transcription factor TFIIIC; and type III promoters, which are found in metazoans but not lower eukaryotes and that use promoter elements located entirely upstream of the transcription start site. Prototypical members of this group include U6 snRNA and 7SK RNA genes. Recent studies have demonstrated that the upstream promoter structure of these genes and the simple polythymidylate terminator recognized by Pol III provide an efficient system for driving the 93 ADVANCES IN PROTEIN CHEMISTRY, Vol. 67
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expression of small interfering RNAs (for review, see McManus and Sharp, 2002). For further information on the transcription of type III genes and a comprehensive coverage of Pol III transcription factors and polymerase, the reader is referred to several recent reviews (Geiduschek and Kassavetis, 2001; Schramm and Hernandez, 2002). In Saccharomyces cerevisiae, the assembly of transcriptionally competent complexes on type II Pol III genes and on TFIIIA-5S gene complexes requires a stable six-subunit assembly factor, TFIIIC (also known as ), and three additional polypeptides (the TATA-binding protein TBP, Brf1, and Bdp1), which together constitute the initiation factor TFIIIB. The resulting complexes then recruit the 17-subunit Pol III enzyme prior to promoter melting and initiation. With relatively few exceptions, the cisacting promoter elements and the protein components of this transcription system are well conserved from yeast to humans (Geiduschek and Kassavetis, 2001; Huang and Maraia, 2001; Schramm and Hernandez, 2002). The availability of recombinant TFIIIB subunits in the early to mid-1990s allowed TFIIIC-mediated recruitment of the initiation factor to be resolved into a stepwise pathway by native gel electrophoresis and site-specific DNA– protein photocrosslinking (Kassavetis et al., 1994). At the same time, genetic studies identified the subunit of TFIIIC (Tfc4) that participates directly in this process (Marck et al., 1993; Rameau et al., 1994; Willis et al., 1989). The assembly of TFIIIB is mediated initially by protein–protein interactions between the tetratricopeptide repeat (TPR)-containing subunit of TFIIIC (Tfc4) and Brf1. Subsequently, TBP is incorporated through interactions primarily with Brf1. TFIIIB complex assembly is completed with the recruitment of Bdp1 by the other TFIIIB subunits and Tfc4. These binding reactions proceed in a concerted manner that involves a series of conformational changes in the DNA and the proteins (Geiduschek and Kassavetis, 2001; Moir et al., 1997, 2000, 2002b). The structural changes induced by Bdp1 are of particular interest because they lead to kinetic trapping of the DNA within TFIIIB and confer its characteristic high stability and resistance to dissociation by high salt concentrations and polyanions (Cloutier et al., 2001; Kassavetis et al., 1990, 1992). These studies, together with in vivo footprinting (Huibregtse and Engelke, 1989), indicate that TFIIIB assembled onto DNA in the yeast nucleus is stably maintained until it is encountered by a replication fork during S phase (Deshpande and Newlon, 1996). The stability of TFIIIB-DNA permits high levels of transcription by eliminating rate-limiting steps in preinitation complex assembly and by enabling rapid recycling of Pol III after termination (Dieci and Sentenac, 1996). Complex stability is presumably also important for Pol III genes to function as boundaries that block the spread of silent chromatin (Donze and Kamakaka, 2001).
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Pol III genes, particularly those encoding 5S and tRNAs, are highly expressed in growing cells and account for about 15% of nuclear transcription. The synthesis of these transcripts is tightly coupled with the synthesis of other components of the protein synthetic machinery, especially the Pol I–transcribed large ribosomal RNAs (Li et al., 2000; Paule and White, 2000; Warner, 1999). This coordinate regulation has two important consequences: it produces appropriate amounts of the RNAs for ribosome synthesis and function while limiting their energetically costly synthesis at inappropriate times (Li et al., 2000; Warner, 1999). As the transcription of this machinery determines the capacity for cell growth, it is an important target of various tumor suppressors and oncogenes (reviewed in Brown et al., 2000; White, 1998a). The evidence accumulated to date indicates that regulation of Pol III transcription occurs through changes in the amount or activity of the assembly and/or initiation factors rather than through direct effects on the polymerase itself. This chapter focuses on recent advances in our understanding of a critical limiting step in Pol III transcription: the assembly on DNA of the initiation factor TFIIIB by TFIIIC.
II. Brf1-TFIIIC: A Limiting Interaction in Preinitiation Complex Assembly The assembly of TFIIIB-DNA complexes in vitro can be directed by a TBP/TATA box interaction (Margottin et al., 1991). However, most type I and type II pol III promoters (in S. cerevisiae and humans) do not have canonical TATA sequences (Dieci et al., 2000; Huang and Maraia, 2001) and, thus, require TFIIIC for TFIIIB assembly. Notably, even in Schizosaccharomyces pombe, where high-affinity TATA sequences have been retained as an essential promoter element, TFIIIC is still required in vivo (Hamada et al., 2001; Huang et al., 2000). The primary determinant for TFIIIC binding in type II promoters (e.g., tRNA genes) is the B block promoter element (Geiduschek and Tocchini-Valentini, 1988), which interacts with the largest subunit (known in yeast as Tfc3 or 138; Bartholomew et al., 1990; Lefebvre et al., 1994). Mutations in the highly conserved B-block promoter sequence dramatically decrease TFIIIC-DNA binding affinity (Baker et al., 1986), whereas mutations in the A-block have relatively modest effects on this interaction (Baker et al., 1986). Nonetheless, both the A-block and B-block elements exert significant effects on transcriptional efficiency. The identification of extragenic suppressors of an A-block mutation (A19) in the dimeric sup9-eA19-supS1 gene revealed that this mutation makes Pol III gene transcription especially sensitive to the level of
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Brf1 and to the Brf1-binding activity of TFIIIC (Lopez-de-Leon et al., 1992; Rameau et al., 1994; Willis et al., 1989). Gain-of-function mutations in Tfc4 that increase Brf1-binding and overexpression of Brf1, but not the Bdp1 or TBP subunits of TFIIIB, were found to increase sup9-eA19-supS1 expression in vivo (Lopez-de-Leon et al., 1992; Moir et al., 1997; Sethy-Coraci et al., 1998). A broader spectrum of changes, which include the preceding mechanisms, suppresses the conditional phenotype of a mutation (tfc3G439E) in the B-block binding subunit of TFIIIC (Lefebvre et al., 1994; Rozenfeld and Thuriaux, 2001). Finally, overexpression of Brf1 or increasing the Brf1-binding activity of Tfc4 also suppresses the conditional phenotype of a strain deleted for the HMG1-like proteins, Nhp6A and Nhp6B (Kruppa et al., 2001). Nhp6 proteins are essential at elevated temperatures for SNR6 transcription, as they serve to facilitate the binding of TFIIIC to this promoter (Kruppa et al., 2001; Lopez et al., 2001). Although several subunits of TFIIIC are known to interact directly with the subunits of TFIIIB (Chaussivert et al., 1995; Deprez et al., 1999; Hsieh et al., 1999a,b), the preceding genetic observations together with extensive supporting biochemistry indicate that the interaction between the Tfc4 subunit of TFIIIC and Brf1 is a major thermodynamically limiting step in the assembly of TFIIIB (Lopez-de-Leon et al., 1992; Moir et al., 1997; Sethy-Coraci et al., 1998). As might be predicted from the limiting nature of the Brf1-TFIIIC interaction, both components are subject to regulation in vivo. Human TFIIIC (known as TFIIIC2) is targeted by viral transforming proteins that increase the abundance of its subunits (Felton-Edkins and White, 2002) and alter the distribution between transcriptionally active and inactive forms of the factor (Sinn et al., 1995; reviewed in White, 1998a). TFIIIC is also a specific substrate of a poliovirus protease that renders the factor transcriptionally inactive (Shen et al., 1996). All five subunits of mammalian TFIIIC2 are phosphorylated in vivo, as are Tfc1, Tfc3, and Tfc4 in yeast (Conesa et al., 1993; Shen et al., 1996). However, the significance of these modifications for TFIIIC function has not been established. The activity of TFIIIB is subject to mitotic repression in higher eukaryotes (Gottesfeld et al., 1994; White et al., 1995) and responds to Maf1dependent signaling of repression in S. cerevisiae (Upadhya et al., 2002). TFIIIB subunits are also targets of negative regulation by the retinoblastoma protein, RB: RB and the related pocket proteins p107 and p130 bind Brf1 and block its interaction with TFIIIC2 and the polymerase (Chu et al., 1997; Larminie et al., 1997; Sutcliffe et al., 1999). These repressive effects on Brf1 are reversed by the action of cyclin-dependent kinases on both RB and p130 and allows for increased Pol III transcription with cell-cycle entry (Scott et al., 2001). Similar to RB, the p53 tumor suppressor also binds Brf1
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and represses Pol III transcription (Cairns and White, 1998; Chesnokov et al., 1996). The action of viral-transforming proteins serves to release TFIIIB from these inhibitory effects and to thereby increase TFIIIB activity (reviewed in White, 1998b). The existence of a direct mechanism for activating Pol III transcription that is RB independent is indicated by the positive association of c-Myc with Brf1 and TBP (Gomez-Roman et al., 2003). The activity of casein kinase II (CK2) also promotes Pol III transcription and is correlated with an increase in TFIIIB binding to TFIIICDNA. Two TFIIIB factors, HsBrf1 and ScTBP, have been identified as targets of CK2 to date (Ghavidel and Schultz, 1997; Johnston et al., 2002). Together, these data show that the recruitment of Brf1 and TBP by TFIIIC-DNA complexes is a significant regulatory step in TFIIIB complex assembly.
III. Brf1-Tfc4 Interactions A. Structural Domains of Brf1 The most recognizable feature of Brf1 (TFIIB-related factor 1) is its phylogenetically conserved amino-terminal half, which is homologous to the general Pol II factor TFIIB (Buratowski and Zhou, 1992; Colbert and Hahn, 1992; Lopez-de-Leon et al., 1992). In contrast, the carboxy-terminal half of Brf1 is unique and rather poorly conserved: Yeast Brf1 homologues contain three conserved regions, domains I, II and III, (Khoo et al., 1994), only two of which (domains II and III) are preserved in human Brf1 (Mital et al., 1996; Wang and Roeder, 1995). Brf1 is required for transcription of all Pol III genes with internal promoter elements and can accurately direct transcription initiation in vitro when split in two. The TFIIB-like half and a truncated carboxy-terminal region (from domain II to the end of the protein) are, in combination, necessary and minimally sufficient to support TFIIIC-directed transcription in vitro (Kassavetis et al., 1998b). The distinct structural domains of Brf1 are not readily divided into discrete functional units: Multiple regions in Brf1 contribute to interactions with the other TFIIIB subunits as well as to specific subunits of TFIIIC and Pol III. In TATA-directed assembly of TFIIIB, both halves of Brf1 play a role in binding to TBP (on and off the DNA) and in the recruitment of Bdp1, although the predominant energetic contributions are derived from the carboxy terminus (domains II and III; Kassavetis et al., 1997). The TFIIB-like half can form TFIIIB complexes that are unstable yet support transcription, whereas the carboxy-terminal half correctly positions TFIIIB, but the resulting complexes are defective in promoter-opening on
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linear (but not supercoiled) templates (Kassavetis et al., 1997). Conditional mutations in domain II directly affect TBP binding and alter the interaction with Bdp1 in vivo (Andrau et al., 1999). Deletion or mutagenesis of the carboxy-terminal region of Brf1 generates a conditional growth phenotype and results in the assembly of heparin-sensitive TFIIIB complexes in vitro. These latter findings support the functional significance of the cryptic DNA binding domain located in this region (Andrau et al., 1999; Colbert et al., 1998; Huet et al., 1997). In contrast to its minor role in TATA-directed assembly of TFIIIB, the TFIIB-like region of Brf1 plays a major role in Pol III recruitment and promoter melting (Hahn and Roberts, 2000; Kassavetis et al., 1998, 2001). The TFIIB-core region of Brf1 is sufficient for a binary interaction with two Pol III–specific subunits, Rpc17 and Rpc34 (Ferri et al., 2000; Khoo et al., 1994). However, the interaction with full-length Rpc34 is also affected by mutations in domains II and III (Andrau et al., 1999). The Zn2þ-ribbon domain upstream of the TFIIB-core is required for opening the downstream segment of the transcription bubble and for stable binding by the polymerase (Hahn and Roberts, 2000; Kassavetis et al., 2001). Consistent with the requirement for both halves of Brf1 in TFIIICdependent transcription (Kassavetis et al., 1998b), both regions of Brf1 are required to bind Tfc4 (Chaussivert et al., 1995). The TFIIB-like half of Brf1 and the amino terminus of Tfc4 up to TPR5 (Nt-TPR5) represent the minimal regions sufficient for a strong binary interaction. Although mutations in conserved domains II and III do not affect a two-hybrid interaction between full-length Brf1 and Tfc4 (Andrau et al., 1999), the carboxyterminal half of Brf1 is required for binding to the Nt-TPR1 fragment but not to larger regions of Tfc4 (Nt-TPR5 and Nt-TPR9, Chaussivert et al., 1995). The Brf1 interaction with Tfc4 is evidently complex: Tfc4 contains more than a single binding site for Brf1, and intramolecular interactions in both proteins appear to mask the interacting regions (Chaussivert et al., 1995; Moir et al., 2002b).
B. Tetratricopeptide Repeats (TPRs) in Tfc4 The most prominent structural feature of Tfc4 is its 11 tetratricopeptide repeats (Marck et al., 1993; Rameau et al., 1994). TPRs are ubiquitous elements of protein structure that function as sites of protein–protein interaction (Lamb et al., 1995). The motif is defined by a degenerate sequence, usually 34 amino acids in length, that is most often found in tandem arrays (Lamb et al., 1995). Individual TPRs fold into two antiparallel -helices, designated A and B, which are separated by a short turn. The helices within each repeat stack together with helices in adjacent
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TPRs to form a right-handed superhelix (Das et al., 1998, Fig. 1). As revealed in the structures of several TPR protein–ligand complexes, the groove of the TPR superhelix is composed of residues primarily from the A helix of each repeat (Das et al., 1998) and constitutes a ligand-binding site (Gatto et al., 2000; Lapouge et al., 2000; Scheufler et al., 2000). Peptide ligands bind in an extended conformation within this groove. In addition to direct intermolecular interactions, the TPR groove can also serve as a surface for intramolecular binding of domains that, in turn, function as intermolecular ligand-binding sites (Gatto et al., 2000; Lapouge et al., 2000). As noted above, TPRs are usually found as multi-TPR arrays. However, some TPR proteins also contain isolated or solo tetratricopeptide motifs. Other proteins, such as the mitochondrial import receptor protein Tom22, contain just a single tetratricopeptide motif. In this case, the antiparallel alignment of the A and B helix forms an apolar groove, in line with the helical axis, that binds the amino-terminal presequences of proteins destined for the mitochondria (Abe et al., 2000). Currently available crystal structures of TPR arrays show a highly homologous picture for three tandem repeats among unrelated TPR proteins and significant structural similarity to a distantly related protein family, the 14–3–3 helical repeat proteins (Das et al., 1998; Scheufler et al., 2000). Although structures of a TPR protein in both the free and ligand-bound
Fig. 1. A structural model of tetratricopeptide repeats 1–3 of Tfc4 showing the sites of the PCF1-1 and PCF1-2 mutations. Taken from Moir et al. (2002a) with permission. (See Color Insert.)
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state have not yet been reported, the preceding comparisons indicate that ligand binding does not significantly alter TPR structure (Gatto et al., 2000; Lapouge et al., 2000; Scheufler et al., 2000). In this respect, the TPR array, like the 14–3–3 helical repeat, appears to be a stable, rigid structure (Yaffe, 2002). In contrast to this view, different crystal forms of two TPR arrays have been solved in which the terminal repeat forms a single extended helix (i.e., the loop that normally enables antiparallel packing of the A and B helices is absent; Kumar et al., 2001; Taylor et al., 2001). Although these structures involve unique intermolecular interactions in the crystal lattice, they indicate a potential for flexibility in some repeats. A comparison of the structure of seven orthologues of ScTfc4 (four yeast and three higher eukaryotes) shows that the organization of the 11 TPRs is conserved. The first nine TPRs are clustered in two arrays, TPR1–5 and TPR6–9, in the amino-terminal half of the protein, whereas the tenth and eleventh TPRs are isolated motifs, located near the carboxy terminus. The two TPR arrays are separated from each other and from TPR10 by regions of minimal sequence conservation (Dumay-Odelot et al., 2002). Aside from the TPR consensus residues, TPRs 1 through 9 show no sequence homology to each other, and there is only limited homology among specific TPRs across species (Dumay-Odelot et al., 2002; Moir et al., 2000). In contrast, the two single TPRs and adjacent sequences in the carboxyterminal region of Tfc4 are highly conserved (Dumay-Odelot et al., 2002). The conservation of the number of TPRs and their organization in Tfc4, coupled with the low sequence homology elsewhere in the protein, indicates that Tfc4 protein function is based on the preservation of a common TPR-based tertiary structure (Dumay-Odelot et al., 2002). Despite the retention of 11 TPR motifs in Tfc4, not all of the repeats are essential in vivo: Deletion of TPR2, TPR3, TPR4, TPR8, or TPR9 generates a conditional phenotype, whereas deletion of TPR1, TPR5, TPR6, TPR7, TPR10, or TPR11 is lethal (Chaussivert et al., 1995; Dumay-Odelot et al., 2002). The conditional lethal phenotype generated by deletion of TPR2 or TPR8 can be rescued by overexpression of different Pol III factors: TPR2 is rescued by overexpression of only Bdp1 (Dumay-Odelot et al., 2002), whereas TPR8, albeit in a different strain background, is rescued by overexpression of Brf1, TBP, and Tfc1, a TFIIIC subunit associated with Tfc4 and the A block (Chaussivert et al., 1995). Two-hybrid experiments also indicate that individual TPRs exhibit differential importance for specific interactions. For example, deletions of TPR1, TPR2, and TPR3 differentially affect two-hybrid interactions with various partners. Of particular interest, deletion of TPR2 diminishes binding to Brf1 (Chaussivert et al., 1995) while increasing significantly the binding to both Bdp1 and
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RPABC10 (Dumay et al., 1999; Ruth et al., 1996). Thus, TPR2 plays a negative role in the binary interaction between Tfc4 and these two ligands.
C. Brf1 Binding to Tfc4 and Autoinhibition The TPR motifs in Tfc4 were initially suggested to be binding sites for Brf1 on the basis of pull-down assays: Regions of Tfc4 that included the first TPR array, the second TPR array, and a carboxy-terminal fragment containing TPR10 and TPR11 were each able to bind immobilized Brf1 (Khoo et al., 1994). Subsequent two-hybrid analysis of Tfc4 and Brf1 showed that TPR-TPR1-in combination with the amino-terminal region of Tfc4 (NtTPR1) was sufficient for Brf1 binding (Chaussivert et al., 1995). Larger fragments of Tfc4 (e.g., Nt-TPR5 and Nt-TPR9) exhibited incrementally lower two-hybrid interactions and provided the first indication that sites of Brf1 binding in Tfc4 may be masked. Pull-down studies with the human homologue of Tfc4 also demonstrated Brf1 binding for an amino-terminal fragment containing the first two TPRs and, in addition, revealed an independent Brf1 binding site entirely within the second TPR array (Hsieh et al., 1999b). The Nt-TPR9 region of ScTfc4 is well-structured, as revealed by circular dichroism spectroscopy, and is largely resistant to proteolysis by trypsin and chymotrypsin, despite an abundance of potential cleavage sites (Moir et al., 2000). This region supports interactions with Brf1 in solution (Moir et al., 2000), as well as in two-hybrid and far-western assays (Chaussivert et al., 1995; Moir et al., 2000). Solution interactions between Nt-TPR9 and Brf1 competitively inhibit Brf1 recruitment by TFIIIC-DNA and prevent the assembly of TFIIIB. These observations led to the development of a coupled equilibrium-binding assay in which the inhibition of TFIIIB complex assembly on a tRNA gene served as the readout of Brf1 binding to Nt-TPR9 (Moir et al., 2000). The inhibition isotherm obtained by titrating Nt-TPR9 exhibited single-site noncooperative binding (Hill coefficient 1.0) and allowed determination of an apparent binding affinity for the Brf1/Nt-TPR9 interaction (Table I). Subsequently, the contribution of the TPR arrays, individually and in combination, in the Brf1 binding reaction was compared. These data showed that each TPR array is capable of binding to Brf1, albeit with different relative affinities: TPR6–9 bind with a four-fold higher apparent affinity than do TPR1–5. However, the addition of amino-terminal sequences increases the affinity of the TPR1–5 array to create a high-affinity Brf1 binding site in Nt-TPR5. Interestingly, the apparent affinities of both Nt-TPR5 and TPR6–9 are higher than for the larger Nt-TPR9 fragment that contains both of these Brf1 binding sites. Thus, the data demonstrate autoinhibition of
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Table I Apparent Affinity of Wild-Type Tfc4 Fragments for Brf1 Tfc4 (1–1025)
Tfc4 Fragment Nt-TPR9 Nt-TPR5
Global KD TFIIIB Inhibition nM SEa
nH Hill Coefficient
333 21
1.3 0.1
44 6
1.0 0.1
IVR þ TPR6–9
379 61
0.8 0.1
TPR1–5
773 87
1.0b
TPR1–9
210 20
1.8 0.3
TPR6–9
177 27
1.2 0.2
Data reproduced from Moir et al. (2002b) with permission. The standard error was determined during curve fitting of multiple concatenated data sets. b For tetratricopeptide repeats TPR1–5, individual and global fits were performed with the Hill coefficient fixed at 1.0. a
Brf1 binding to Nt-TPR9. It appears that the tertiary structure of Nt-TPR9 limits access of Brf1 to both of its binding sites in this fragment.
D. Effects of Mutations in Tetratricopeptide Repeat Domains on Brf1 Binding An analysis of mutant yeast strains selected for their ability to suppress a promoter defect in the sup9e-A19-supS1 gene and to increase Pol III transcription initially identified three gain-of-function mutations in TPR2 of TFC4 (aka PCF1; Moir et al., 1997; Rameau et al., 1994; Willis et al., 1989). Further mutagenesis and selection, focusing on just the TPR1–9 region, identified additional activating mutations in TPR2 and the helices that flank it (Moir et al., 1997) and thus delimited a critical, functionally limiting region of the protein. Biochemical studies on two of these activating mutations, PCF1-1 (H190Y) and PCF1-2 (T167I), have shown that they both facilitate the recruitment of Brf1 to TFIIIC-DNA without affecting TFIIIC-DNA complex formation (Moir et al., 1997, 2002a). Notably, the common biochemical effect of these mutations on Brf1 binding is achieved despite their location on opposite sides of the TPR superhelix: PCF1-2 (T167I) alters a residue in the ligand-binding groove, whereas PCF1-1 (H190Y) changes a surface accessible residue on the back side of the TPR superhelix (Fig. 1).
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Although the PCF1-1 mutation facilitates Brf1 binding to TFIIIC-DNA complexes by increasing the apparent affinity of this interaction (Moir et al., 2002a), complex assembly inhibition experiments with wild-type and mutant Tfc4 fragments indicate that Brf1 does not make a direct contact with the side chain at amino acid 190. The H190Y substitution has no discernible effect on Brf1 binding to the isolated TPR1–5 array or to the Nt-TPR5 fragment (Table II). However, the larger fragments, Nt-TPR9 and TPR1–9, in which Brf1 binding is autoinhibited, each show a positive effect of the H190Y mutation. Thus, it is thought that PCF1-1 and potentially other dominant substitutions that are solvent-accessible on the back side of the TPR superhelix (Figs. 1 and 2C) overcome the autoinhibited binding of Brf1 by an indirect mechanism. A molecular mechanism consistent with the available data is that the PCF1-1 mutation stabilizes a conformation of Tfc4 that promotes the binding of Brf1 (Moir et al., 1997). Because aromatic or large hydrophobic residues at amino acid 190 are dominant activators of transcription and one of these mutations (tyrosine) indirectly offsets the intrinsic autoinhibition of Brf1 binding to Nt-TPR9 (Moir et al., 2002b), it is plausible that these gain-of-function mutations participate in novel intramolecular interactions with a hydrophobic site in the Nt-TPR9 domain. In contrast to PCF1-1, the location of the PCF1-2 mutation in the ligandbinding channel of the TPR superhelix indicates that it may interact directly with Brf1 (Figs. 1 and 2B). Strong support for this possibility has come from an analysis of mutant (PCF1-2) Tfc4 fragments in the TFIIIB complex inhibition assay. These experiments yielded qualitatively similar results to those involving PCF1-1 with one important exception: the PCF1-2
Table II Apparent Affinity of Mutant Tfc4 Fragments for Brf1
Tfc4 Fragment Nt-TPR9 Nt-TPR5 TPR1–9 TPR1–5
Wild-type (PCF1 þ)
H190Y(PCF1-1)
T167I(PCF1-2)
Global KD TFIIIB inhibition (nM SEa)
Global KDTFIIIB inhibition (nM SEa)
Global KD TFIIIB inhibition (nm SEa)
333 21 44 6 210 20 773 87
192 30 37 8 151 16 722 107
231 24 32 3 nd 465 68
Data taken from Moir et al. (2002a) and Moir et al. (2004) with permission. The standard error was determined during curve fitting of multiple concatenated data sets. a
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Fig. 2. Structural model of potential ligand-binding sites in tetratricopeptide repeats TPR1–3 of Tfc4. (A) The ribbon diagram shows the model of TPR1–3 of Tfc4 (based on the structure of protein phosphatase 5; Das et al., 1998; Moir et al., 2002a) in the same orientation as the surface representation in panel B. The superhelical axis runs vertically. (B) Surface representation of the ligand-binding channel of TPR1–3 in Tfc4. The molecular surfaces shown in green include the highly conserved residues N136, V140, E197, and D195 (Moir et al., 2002b) and the dominant mutations F162L, A164V, and T167I (Moir et al., 1997). Side chain substitutions were introduced using Deep View software, and the image was created in PyMol. (C) The structure in panel B was rotated approximately 180 degrees, as indicated, to reveal the outer surface of the superhelix. The molecular surfaces shown in green include the highly conserved residues L150, E153, K156, L168, L187, and R222 and the sites of dominant mutations E148K, Y172C, H190Y, and W199R (the wild-type side chains were retained in this model). (See Color Insert.)
mutation increased the apparent binding affinity for Brf1 in the context of the TPR1–5 fragment (from 773 to 465 nM; Table II). The increased affinity of Brf1 for PCF1-2 TFIIIC-DNA complexes together with the fragment binding data indicate that interactions of Brf1 in the superhelical groove of the first TPR array play an important role in TFIIIB complex assembly (Moir et al., 2004). Site-directed mutagenesis at both H190 and T167 in TPR2 generates a range of phenotypes with respect to expression of the sup9-eA19-supS1 reporter gene. Collectively, these mutations show that the wild-type residue at either position has an intermediate phenotype; the majority of the mutations at both sites are defective relative to wild-type, whereas only a few mutations (e.g., PCF1-1 and PCF1-2) are better than wild-type (Moir et al., 2002a, 2004). Given the different location of the PCF1-1 and PCF1-2 mutations in the TPR structure (Fig. 1) and the different mechanism by which they increase Brf1 binding to Tfc4, it was expected that their combination would further enhance Pol III transcription. Surprisingly, the PCF1-1, PCF1-2 double mutant is synthetically lethal even though neither single mutation
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has a detectable growth phenotype on rich media (Moir et al., 2004). Overly tight binding of Brf1 by Tfc4 is apparently incompatible with viability, and so the Brf1-Tfc4 interaction must, by necessity, be metastable. This result is most conveniently interpreted in light of the reciprocal changes in photocrosslinking of Tfc4 and Brf1 that are seen on incorporation of Bdp1 into TBP-Brf1-TFIIIC-DNA complexes (Kassavetis et al., 1992). As discussed below, a dynamic (i.e., readily reversible) association of Brf1 with Tfc4 may be required for recruitment of Bdp1. Biochemical experiments showing that Brf1 binds to the second TPR array (TPR6–9) have been reported for the proteins from yeast and humans (Hsieh et al., 1999b; Moir et al., 2002b). However, the biological importance of this interaction has only been demonstrated recently. Radical substitutions at five of eight phylogenetically conserved residues in the A helices of TPR6–9 significantly impair Pol III reporter gene transcription (Liao et al., 2003). In contrast, mutagenesis of two conserved B-helix residues in this array did not affect transcription. The loss of function phenotype generated by three of the A helix mutations could be rescued by overexpression of Brf1, but not TBP or Bdp1, consistent with Brf1 binding to wild-type residues in the ligand-binding channel of this array. Two of the mutations could not be rescued by overexpression of any single TFIIIB subunit, implying that the binding of multiple subunits is defective. Accordingly, biochemical studies on one of these mutations (L469K in TPR7) revealed two defects in TFIIIB-DNA complex assembly, one of which is the recruitment of Brf1. Thus, the genetic and biochemical data support a direct interaction of Brf1 in the ligand-binding groove of TPRs6–9 (see also Section V). The identification of Brf1 as a ligand for the TPR1–5 and TPR6–9 arrays raises a question as to whether this TFIIIB subunit may also bind to the solo TPRs located in the carboxy terminus of Tfc4. Consistent with this possibility, pull-down assays between Brf1 and Tfc4 have shown that a small region of Tfc4, including TPR10 and TPR11, can indeed bind Brf1 (Khoo et al., 1994). However, the contribution of this region of Tfc4 to the Brf1 interaction remains to be defined, as the carboxy-terminal half of Tfc4 is neither required for nor independently binds Brf1 in a two-hybrid assay (Chaussivert et al., 1995) and is inactive (albeit as a refolded protein) in the TFIIIB complex inhibition assay (R. D. Moir, unpublished observations).
IV. Bdp1-Tfc4 Interactions Tfc4 also interacts directly with the Bdp1 subunit of TFIIIB (DumayOdelot et al., 2002; Ruth et al., 1996), and this interaction, like that for Brf1, involves both TPR arrays (Ishiguro et al., 2002; Liao et al., 2003;
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Rozenfeld and Thuriaux, 2001). Deletion of TPR2 first suggested a negative role for this repeat in a two-hybrid interaction with Bdp1 (Ruth et al., 1996). Subsequently, overexpression of Bdp1, but not Brf1 or TBP, was shown to suppress the conditional phenotype of a tfc4 TPR2 deletion strain. On the basis of these findings, it seemed likely that point mutations in TPR2 might have either positive or negative effects on Bdp1 binding. However, a biochemical study of Bdp1 binding to complexes containing wild-type or PCF1-1 TFIIIC revealed no differences in affinity (Moir et al., 2002a). The relationship between this dominant TFC4 allele and Bdp1 is apparently quite subtle but is evident from the synthetic lethal phenotype obtained when PCF1-1 is combined with a conditional Bdp1 deletion (Bdp1355–372; Ishiguro et al., 2002). The basis for the conditional growth phenotype of Bdp1(355–372) is not yet clear and may be complex, as indicated by the fact that overexpression of the mutant Bdp1 protein is lethal (Ishiguro et al., 2002). In vitro, Bdp1(355–372) supports TFIIIC-dependent and TFIIIC-independent transcription on supercoiled templates but forms TFIIIB complexes that are defective in promoter opening on linear templates (Kassavetis et al., 1998a, 2001). A quantitative effect of Bdp1(355–372) on TFIIIC-dependent TFIIIB assembly (as distinct from an effect of PCF1-1 on a post-TFIIIB assembly step) is also possible and may underlie (or contribute to) the synthetic lethal phenotype. The ability of PCF1-1 to increase Brf1 binding and simultaneously antagonize the binding of a Bdp1 protein with diminished capacity would further support the hypothesis that a metastable interaction with Brf1 is necessary for subsequent Bdp1 recruitment. The importance of the second TPR array for the association of Bdp1 with Tfc4 is demonstrated by mutagenesis of residues predicted to lie in ligand-binding channel of TPR6–9 (Fig. 3). Binding of Bdp1 to TPR7 is indicated by a reduced two-hybrid interaction involving a Tfc4 mutation (D468N) that was isolated as a suppressor of a conditional mutation (tfc3-G349E) in the B-block binding subunit of TFIIIC (Rozenfeld and Thuriaux, 2001). This function for TPR7 is strongly supported by mutation of the adjacent residue, L469K. A comparison of Bdp1 binding to wild-type and tfc4 mutant B0 -TFIIIC-DNA complexes revealed that the L469K mutation significantly reduced the affinity of this interaction. In addition, Bdp1 binding to fragments of Tfc4 (TPR1–9 and Nt-TPR9) in two-hybrid and pull-down assays was decreased by the L469K mutation (Liao et al., 2003). These experiments also revealed important differences between the binary Bdp1-Tfc4 and Brf1-Tfc4 interactions. First, in contrast to Brf1, Bdp1 does not detectably bind to the individual TPR arrays; both arrays (TPR1–5 and TPR6–9) are required for the interaction, and thus their tertiary arrangement is likely to be important. Second, Bdp1 is unable to
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Fig. 3. Potential sites of ligand-binding in tetratricopeptide repeats TPR6–9. The ribbon diagram is based on a structure of the cochaperone Hop (domain 2A), which contains three complete TPRs and the A helix of a fourth repeat (Scheufler et al., 2000). Phylogenetically conserved residues from TPR6–9 of Tfc4 (shaded in black; Moir et al., 2002b) that are located at non-TPR consensus positions in the A helix of the repeat were mapped onto the Hop structure using the TPR consensus residues to define their location. The dotted line indicates that in S. cerevisiae Tfc4, proline residues in the middle of TPR7 helix B and TPR8 helix A interrupt the normal TPR fold. These proline residues are not conserved in Tfc4 orthologs.
bind the amino-terminal high-affinity Brf1 binding site contained within Nt-TPR5. Finally, we note that the cumulative genetic and biochemical data indicate that the sites of Bdp1 binding in TPRs1–9 overlap to some degree with those for Brf1.
V. Ligand Binding by TPR Arrays of Tfc4 A TPR structure-based phylogenetic analysis of TPRs1–9 in Tfc4 has identified the invariant and highly conserved residues in each TPR array that are not involved in generating the TPR fold (Moir et al., 2002b). These residues are evenly distributed between the A and B helices in
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TPR1–5, whereas in TPR6–9 they are found mostly in the A helices of the array. The mapping of these sites onto solved TPR crystal structures provides insights into potential regions of ligand binding.
A. Identification of Potential Ligand Binding Sites in the TPR Arrays of Tfc4 Based on the location of dominant activating mutations in TPRs1–3, a structural model of this region was built using the three TPRs of protein phosphatase 5 (Das et al., 1998). From this model (Fig. 2B) it is apparent that a subset of the conserved residues in the A helices of TPR1 and TPR3, a residue in the loop between TPRs2 and 3 and three dominant mutations form a virtually contiguous surface traversing the ligand binding groove. Biochemical studies of the T167I (PCF1-2) mutation indicate that this surface is involved directly in Brf1 binding (discussed in Section III, D). Other dominant mutations and conserved residues in the A helices of TPRs1–3 project toward the back side of the TPR superhelix. Mapping of these sites, together with conserved residues and activating mutations in the B helices of these TPRs, reveals a line of surface-accessible sites that lies across the outer face of the array (Fig. 2C). The location of the wellcharacterized H190Y (PCF1-1) mutation on this surface is indicative that the proposed intramolecular interaction involving this residue may be part of a more extensive network of interactions that influence Brf1 binding. As noted above, conserved, non-TPR consensus residues in TPR6–9 are predominantly in the A helices of the array (Moir et al., 2002b). Structural modeling of this array in S. cerevisiae Tfc4 is problematic, as the presence of two nonconserved proline residues shortens the B helix of TPR7 and the A helix of TPR8 and prevents sequence threading. Presumably, the structure of S. cerevisiae Tfc4 in the interval between these prolines deviates from the canonical TPR structure. To gain some structural insight into the location of conserved A helix residues lying outside this interval, the three-and-ahalf TPR structure of the cochaperone Hop (domain 2A) was used to directly map the positions of conserved residues from TPR6–9 of Tfc4 (Fig. 3). This analysis revealed two clusters of conserved residues, one comprising TPR6 and TPR7 (I434, R437, L438, L440, D468, L469, and E472) and the other comprising TPR9 (D537, V540, S541, L542, and V545). Multicopy suppression of mutations at two positions in TPR9 (S541I and L542G) implicates Brf1 as a ligand for this TPR. Alternatively, or in addition, these mutations may affect the binding of Tfc1 to the array (Hsieh et al., 1999b, see Section VII). Genetic and biochemical experiments
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on the L469K mutation in TPR7 (discussed above) indicate that both Brf1 and Bdp1 may interact in this region. The Bdp1 interaction in TPR7 is also supported by a reduced two-hybrid interaction involving a Tfc4 mutation (D468N) that was isolated initially by its suppression of a conditional tfc3 allele (Rozenfeld and Thuriaux, 2001). The potential sites of ligand binding identified by this analysis indicate that interactions with the two TPR arrays of Tfc4 involve a mode of binding that is distinct from the carboxylate clamp described for the Hop TPR-ligand complexes (Scheufler et al., 2000).
B. Models for Tetratricopeptide Repeat Organization in Tfc4 In some respects, Tfc4-Brf1 binding can be considered a two-site, twoligand interaction. Each TPR array has been shown to contain a binding site for Brf1, and the lack of sequence similarity between the TPR1–5 and TPR6–9 arrays indicates that the TPR channels do not compete for binding to the same region of Brf1 (Moir et al., 2002b). Moreover, twohybrid interactions between fragments of Brf1 and Tfc4 suggest that within Brf1, both the TFIIB-like region and the carboxy-terminal region participate in Tfc4 binding (Chaussivert et al., 1995). However, the autoinhibition of Brf1 binding to each TPR array, in the context of the multi-TPR array structure in Nt-TPR9, indicates that the TPR arrays are not physically independent of one another. Similarly, measurable binding of Bdp1 requires both TPR arrays. Although the relative disposition of the two TPR arrays within Tfc4 has yet to be defined structurally, this is an issue of considerable interest and importance given the limiting nature of the Brf1-Tfc4 interaction and the prevalence of multiarray TPR proteins. The available structures of TPR proteins provide a conceptual foundation to better understand this interaction. The known structures of TPR arrays, whether free or ligand-bound, generate a highly homologous view of the arrangement of the -helices within a three to four TPR-repeat unit (Das et al., 1998; Gatto et al., 2000; Lapouge et al., 2000; Rice and Brunger 1999; Scheufler et al., 2000; Taylor et al., 2001). However, to date, only two models describing the organization of multiple TPR arrays have been reported: The first model involves an extended superhelical array (Das et al., 1998), whereas the second is illustrated by the PEX5/PTS-1 peptide cocrystal structure (Gatto et al., 2000). In the extended superhelix model, TPR arrays that are separated in the primary sequence are proposed to align as if they are adjacent, allowing the periodicity of the superhelix to be maintained. Evidence for this type of organization is provided by the structure of p67phox, which accommodates a 15–amino acid -hairpin element between TPR3
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and TPR4 without perturbation of the superhelical structure (Lapouge et al., 2000). If TPR1–5 and TPR6–9 of Tfc4 were to form an extended superhelix with the intervening 135 amino acids adopting a well-ordered and compact structure (consistent with CD spectroscopy and limited proteolysis (Moir et al., 2000), the molecule would likely be highly asymmetric (Fig. 4A)). Notably, this model places the ligand-binding sites in the groove of TPRs1–3 and in TPRs6–9 at distal locations. In the peroxisomal importer protein, PEX5, the seven consecutive TPRs do not form an extended superhelix but separate into two TPR arrays, each composed of three repeats. These arrays are oriented antiparallel to one
Fig. 4. Organization of multiple tetratricopeptide repeat (TPR) arrays. (A) The model shows an extended superhelix containing nine TPRs. The modeled structure of TPR1–3 of Tfc4 was used to align (using Deep View) TPR3 of one molecule with TPR1 of another molecule. The aligned structures were merged and the process repeated until a nine-TPR structure was built. Similar multi-TPR models have been constructed by Das et al. (1998). The arrow indicates the loop between TPR5 and TPR6 where an insertion of 135 amino acids separating TPR1–5 from TPR6–9 in Tfc4 would occur. The superhelical axis runs vertically. This view shows the back side of TPR1–3 and TPR7–9 and the groove of TPR5 and TPR6. (B) The antiparallel arrangement of two TPR arrays found in the PEX5/PTS1 cocrystal structure (adapted from Gatto et al., 2000). Residues connecting the end of TPR3 with the long hinge helix (the noncannonical fourth TPR) are disordered. The PTS1 peptide (spheres) is sandwiched between the binding grooves of the two TPR arrays.
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another since TPR4 forms a single long helix rather than the usual two-helix structure (Fig. 4B). As a result, the ligand-binding grooves of the arrays face one another and surround the PTS1 peptide. In this structure, residues from both arrays are involved in peptide binding (Gatto et al., 2000). If a similar organization were to be adopted by the two arrays in Tfc4 with the 135 intervening amino acids underlying TPR5 and TPR6 (a position equivalent to the long helix annotated initially as TPR4 in PEX5), the molecule would not be especially asymmetric. Moreover, TPRs at opposite ends of the arrays (e.g., TPR1 and TPR9) would be physically quite close to one another, unlike in the extended superhelix described above. In the absence of a crystal structure, the two models described above may be distinguished by a combination of hydrodynamic methods (e.g., analytical ultracentrifugation) and intramolecular protein–protein crosslinking.
VI. Ordered Binding of TFIIIB Subunits to TFIIIC-DNA and Dynamic Interactions with Tfc4 The clear binary interactions between Tfc4 and both Brf1 and Bdp1 are dramatically and differentially affected by the presence of the other TFIIIC subunits and DNA. As determined by native gel electrophoresis, DNase 1 footprinting and site-specific DNA–protein photocrosslinking, Bdp1 does not detectably bind to TFIIIC-DNA complexes (Kassavetis et al., 1992; Moir et al., 2004). Its incorporation into these complexes requires prior binding by Brf1 and TBP (Kassavetis et al., 1992). Because Brf1 and Bdp1 interact with at least partially overlapping sites in Tfc4 (Fig. 5), it appears that Brf1 binding to TFIIIC-DNA does not simply precede Bdp1 in complex assembly. Rather, through sequential and cumulative changes in Tfc4 and Brf1 structure (reflected in their accessibility to photoprobes in the DNA, changes in circular dichroism, DNase 1 footprinting, etc.), at least some of which are induced by TBP, the Bdp1 binding site in Tfc4 is made available. Together with interactions that include the other TFIIIB subunits and the DNA, Bdp1 has an affinity for the Brf1-TBP-TFIIIC-DNA complex that is approximately two orders of magnitude higher than that for the interaction of Brf1 with TFIIIC-DNA (Moir et al., 2002a). In addition to the structural changes documented by site-specific DNA–protein crosslinking (Kassavetis et al., 1992), it appears that TFIIICdependent assembly of TFIIIB involves dynamic protein–protein interactions between Tfc4 and Brf1. Data supporting this conclusion include the synthetic lethal phenotypes obtained when mutations that increase Brf1 binding are combined (PCF1-1/PCF1-2) and when increased Brf1 binding is coupled with decreased functioning of Bdp1 (PCF1-1/Bdp1 355–372). Together
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with experiments showing that both Brf1 and Bdp1 bind to TPR1–9 via interactions in the ligand-binding grooves of TPR1–3 (at least for Brf1) and TPR6–9, it is evident that Brf1 must be repositioned to allow Bdp1 binding. These and other data described here indicate that the concerted binding of TFIIIB subunits by TFIIIC-DNA can be broken down into the following steps: First, an early step (perhaps the initial step) in the binding of Brf1 to TFIIIC-DNA involves contacts in the ligand-binding grooves provided by TPR1–9, in Tfc4. These interactions with Brf1 are subject to autoinhibition by Tfc4 in the ground state but can be overcome by intramolecular stabilization of an alternative Tfc4 structure (Moir et al., 2002b). The formation of this alternative structure represents a normal step of the assembly process that can be facilitated by mutations such as PCF1-1. Second, Brf1 interactions in TPR1–9 include contacts in the ligandbinding grooves of both TPR1–5 and TPR6–9. Interactions within these individual arrays may be ordered relative to one another. Third, as noted earlier, the interaction between Brf1 and the TPR arrays in Tfc4 is likely to sterically block Bdp1 binding to this same region, and thus, Brf1 must be displaced from this site to allow Bdp1 recruitment. Fourth, the interactions between Brf1 and TPR1–9 are exchanged for interactions involving the high-affinity Brf1 binding site in Nt-TPR5 (whose affinity is not affected by the PCF1-1 or PCF1-2 mutations; Table II). This Brf1 binding-site transition is energetically favorable (Table I) and may be initiated by TBP, which is known to extend the DNase 1 footprint upstream of the start site and to increase Tfc4 and Brf1
Fig. 5. Summary of Tfc4 interactions with RNA polymerase III factors. Saccharomyces cerevisiae Tfc4(1–1025) is represented schematically, with the position of each tetratricopeptide repeat (TPR) highlighted as a gray oval. The gray bar, over a portion of TPR1–5, indicates those repeats that have conserved or functionally important surface accessible residues (highlighted in Fig. 2A and B). Similarly, the gray bar over TPR6–9 illustrates the sequence conservation that occurs across all four repeats, mapped in Figure 3. Fragment interactions: Two-hybrid, pull-down, and TFIIIB complex inhibition assay data are summarized for S. cerevisiae proteins unless otherwise specified. Interactions with S. cerevisiae proteins RPC53 and ABC10 are derived from two-hybrid data (Flores et al., 1999; Dumay-Odelot et al., 2002); data for Bdp1, HsBrf1, and HsTfc1 are from pull-down experiments (Hsieh et al., 1999b; Liao et al., 2003). Where multiple independent interactions are known, the smallest regions or the strongest binding regions are drawn (thin bars and thick bars, respectively). Note: The relative affinity of the Brf1/TPR1–9 interaction, as assayed by complex inhibition, could not be compared directly to data for the other fragments of Tfc4 (see Table I) and so has been included here as a weak interaction. Mutations: Individual mutation sites are noted as arrows, regions where multiple mutations have been identified are drawn as bars. Deletions: A subset of the reported deletions in Tfc4, restricted to specific regions in the protein (e.g., individual repeats), are represented as solid gray triangles.
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photocrosslinking to DNA. DNA interactions involving the cryptic DNA binding site in the carboxy-terminal half of Brf1 (Huet et al., 1997) may also be established at this stage. Fifth, the recruitment of Bdp1 proceeds via contacts in both TPR arrays (as indicated by biochemical studies and the effects of TPR2 and mutations in TPR7). The preceding scheme represents a working model. We should emphasize that our interpretation of events is not exclusive of other possibilities but is consistent with the requirement for both halves of Brf1, with a greater dependence on Bdp1 sequences for proper TFIIIC-dependent assembly of TFIIIB (in contrast to TATA box-mediated, TFIIIC-independent assembly of TFIIIB; Kassavetis et al., 1998b; Kumar et al., 1997), and with the observed changes in Tfc4 photocrosslinking that occur on Brf1, TBP, and subsequently Bdp1 binding to TFIIIC-DNA (Bartholomew et al., 1991; Kassavetis et al., 1992).
VII. Tfc4 and Other Pol III Factors Tfc4 also binds other components of the Pol III transcription apparatus, although a requirement for the TPR arrays and, in particular, specific residues in the TPR channels has been reported for only some of these interactions. Tfc4, Tfc1, and Tfc7 together form A, the domain of TFIIIC associated with the A-block promoter element (Bartholomew et al., 1991; Schultz et al., 1989). Consistent with their physical association in the A complex, genetic suppression data link Tfc4 and Tfc1 in S. cerevisiae. The conditional phenotypes of specific deletions of Tfc4 (TPR8, basic region2, or helix1) can be suppressed by overexpression of Tfc1 (Lefebvre et al., 1992). In addition, a direct interaction between these two proteins has been demonstrated for their human homologues (TFIIIC102 and TFIIIC63): Each TPR array (TPR2–5 and TPR6–9) independently supports binding to both HsBrf1 and TFIIIC63 (human Tfc1), although maximal binding appears to require both TPR arrays (Hsieh et al., 1999b). Although a specific association of Tfc1 with TPR ligand-binding grooves has yet to be demonstrated, it is tempting to consider that the limited extent to which Brf1-binding proceeds on wild-type TFIIIC-DNA complexes in S. cerevisiae may result from a steric effect of Tfc1 in the TPR1–9 array (Moir et al., 1997, 2002a). The TPR arrays of Tfc4 are also involved in interactions with ABC10, a subunit shared by all three nuclear RNA polymerases. Two-hybrid and far-western assays have shown that Tfc4 and ABC10 interact and that Pol III–specific mutations in the carboxy-terminal region of ABC10 decrease binding to Tfc4 (Dumay et al., 1999; Flores et al., 1999). Mutations in Tfc4
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also affect the binary interaction with ABC10: Deletion of TPRs1, 2, or 3 (Dumay et al., 1999) increases binding, whereas the D468N mutation in TPR7 decreases binding (Rozenfeld and Thuriaux, 2001). Moreover, overexpressing the TPR2 deletion mutant of Tfc4 has been shown to suppress a conditional, Pol III–specific transcriptional defect in ABC10 (rpc10–30; Dumay et al., 1999). Together, these data indicate that the TPR arrays in Tfc4 interact with the carboxy-terminal region of ABC10 and that the interaction is biologically significant. A Pol III–specific subunit, RPC53, is also thought to interact with Tfc4. In this case, only a two-hybrid interaction between a fragment of Tfc4 that includes both TPR arrays has been reported (Flores et al., 1999). The association of a TFIIIC subunit, two TFIIIB subunits, and at least one polymerase subunit with the TPR arrays of Tfc4 raises the possibility that all these factors could compete for binding to overlapping sites, at least in vitro. In vivo, however, it seems more reasonable, in light of the studies on Brf1 and Bdp1, that occupancy of the TPR arrays by different components will be temporally resolved and ordered according to the specific function of the ligand at a given stage in the transcription process.
VIII. Conclusion The role of Tfc4 ligand specificity in TFIIIB complex assembly is just beginning to be understood. However, many questions as to the structural changes in both Tfc4 and its substrates associated with ligand binding remain. The significance of these interactions in Tfc4 may not be limited to its function in TFIIIB assembly; the association between Tfc4 and polymerase subunits raises the possibility of an additional role in postassembly functions for TFIIIC, such as polymerase recruitment or recycling. As the list of effectors of Pol III transcription continues to expand in both yeast and mammalian cells (e.g., Maf1, Upadhya et al., 2002 and c-myc, Gomez-Roman et al., 2003), a common theme among known effectors can be recognized: the ability to target TFIIIB and affect its recruitment by TFIIIC or interactions with Pol III. A fundamental understanding of the molecular mechanism or mechanisms by which the cell cycle, cell growth, response to cell stress and damage, oncogenic transformation, and viral infection affect Pol III transcription through TFIIIB is only just beginning to emerge. In particular, the mechanisms by which posttranslational signaling events affect TBP, Brf1, Bpd1, and TFIIIC function in TFIIIB assembly have yet to be defined. Last, understanding the complex mechanism by which Tfc4 interacts with TFIIIB subunits may provide a paradigm for understanding how multiple TPR arrays contribute to the assembly and function of multisubunit complexes.
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References Abe, Y., Shodai, T., Muto, T., Mihara, K., Torii, H., Nishikawa, S., Endo, T., and Kohda, D. (2000). Structural basis of presequence recognition by the mitochondrial protein import receptor Tom20. Cell 100, 551–560. Andrau, J. C., Sentenac, A., and Werner, M. (1999). Mutagenesis of yeast TFIIIB70 reveals C-terminal residues critical for interaction with TBP and C34. J. Mol. Biol. 288, 511–520. Baker, R. E., Gabrielsen, O., and Hall, B. D. (1986). Effects of tRNATyr point mutations on the binding of yeast RNA polymerase III transcription factor C. J. Biol. Chem. 261, 5275–5282. Bartholomew, B., Kassavetis, G. A., Braun, B. R., and Geiduschek, E. P. (1990). The subunit structure of Saccharomyces cerevisiae transcription factor IIIC probed with a novel photocrosslinking reagent. EMBO J. 9, 2197–2205. Bartholomew, B., Kassavetis, G. A., and Geiduschek, E. P. (1991). Two components of Saccharomyces cerevisiae transcription factor IIIB (TFIIIB) are stereospecifically located upstream of a tRNA gene and interact with the second-largest subunit of TFIIIC. Mol. Cell. Biol. 11, 5181–5189. Brown, T. R., Scott, P. H., Stein, T., Winter, A. G., and White, R. J. (2000). RNA polymerase III transcription: its control by tumor suppressors and its deregulation by transforming agents. Gene Expr. 9, 15–28. Buratowski, S., and Zhou, H. (1992). A suppressor of TBP mutations encodes an RNA polymerase III transcription factor with homology to TFIIB. Cell 71, 221–230. Cairns, C. A., and White, R. J. (1998). p53 is a general repressor of RNA polymerase III transcription. EMBO J. 17, 3112–3123. Chaussivert, N., Conesa, C., Shaaban, S., and Sentenac, A. (1995). Complex interactions between yeast TFIIIB and TFIIIC. J. Biol. Chem. 270, 15353–15358. Chesnokov, I., Chu, W. M., Botchan, M. R., and Schmid, C. W. (1996). p53 inhibits RNA polymerase III-directed transcription in a promoter dependent manner. Mol. Cell. Biol. 16, 7084–7088. Chu, W. M., Wang, Z., Roeder, R. G., and Schmid, C. W. (1997). RNA polymerase III transcription repressed by Rb through its interactions with TFIIIB and TFIIIC2. J. Biol. Chem. 272, 14755–14761. Cloutier, T. E., Librizzi, M. D., Mollah, A. K., Brenowitz, M., and Willis, I. M. (2001). Kinetic trapping of DNA by transcription factor IIIB. Proc. Natl. Acad. Sci. USA 98, 9581–9586. Colbert, T., and Hahn, S. (1992). A yeast TFIIB related factor involved in RNA polymerase III transcription. Genes Dev. 6, 1940–1949. Colbert, T., Lee, S., Schimmack, G., and Hahn, S. (1998). Architecture of protein and DNA contacts within the TFIIB-DNA complex. Mol. Cell. Biol. 18, 1682–1691. Conesa, C., Swanson, R. N., Schultz, P., Oudet, P., and Sentenac, A. (1993). On the subunit composition, stoichiometry, and phosphorylation of the yeast transcription factor TFIIIC/tau. J. Biol. Chem. 268, 18047–18052. Das, A. K., Cohen, P. W., and Barford, D. (1998). The structure of the tetratricopeptide repeats of protein phosphatase 5: implications for TPR mediated protein-protein interactions. EMBO J. 17, 1192–1199. Deprez, E., Arrebola, R., Conesa, C., and Sentenac, A. (1999). A subunit of yeast TFIIIC participates in the recruitment of TATA-binding protein. Mol. Cell. Biol. 19, 8042–8051.
TETRATRICOPEPTIDE REPEATS OF Tfc4
117
Deshpande, A. M., and Newlon, C. S. (1996). DNA replication fork pause sites dependent on transcription. Science 272, 1030–1033. Dieci, G., Percudani, R., Giuliodori, S., Bottarelli, L., and Ottonello, S. (2000). TFIIIC independent in vitro transcription of yeast tRNA genes. J. Mol. Biol. 299, 601–613. Dieci, G., and Sentenac, A. (1996). Facilitated recycling pathway for RNA polymerase III. Cell 84, 245–252. Donze, D., and Kamakaka, R. T. (2001). RNA polymerase III and RNA polymerase II promoter complexes are heterochromatin barriers in Saccharomyces cerevisiae. EMBO J. 20, 520–531. Dumay-Odelot, H., Acker, J., Arrebola, R., Sentenac, A., and Marck, C. (2002). Multiple roles of the tau131 subunit of yeast transcription factor IIC (TFIIIC) in TFIIIB assembly. Mol. Cell. Biol. 22, 298–308. Dumay, H., Rubbi, L., Sentenac, A., and Marck, C. (1999). Interaction between yeast RNA polymerase III and transcription factor TFIIIC via ABC10alpha and tau131 subunits. J. Biol. Chem. 274, 33462–33468. Felton-Edkins, Z. A., and White, R. J. (2002). Multiple mechanisms contribute to the activation of RNA polymerase III transcription in cells transformed by papovaviruses. J. Biol. Chem. 277, 48182–48191. Ferri, M. L., Peyroche, G., Siaut, M., Lefebvre, O., Carles, C., Conesa, C., and Sentenac, A. (2000). A novel subunit of yeast RNA polymerase III interacts with the TFIIB related domain of TFIIIB70. Mol. Cell. Biol. 20, 488–495. Flores, A., Briand, J. F., Gadal, O., Andrau, J. C., Rubbi, L., Van, M. V., Boschiero, C., Goussot, M., Marck, C., Carles, C., Thuriaux, P., Sentenac, A., and Werner, M. (1999). A protein-protein interaction map of yeast RNA polymerase III. Proc. Natl. Acad. Sci. USA 96, 7815–7820. Gatto, G. J., Jr., Geisbrecht, B. V., Gould, S. J., and Berg, J. M. (2000). Peroxisomal targeting signal-1 recognition by the TPR domains of human PEX5. Nat. Struct. Biol. 7, 1091–1095. Geiduschek, E. P., and Kassavetis, G. A. (2001). The RNA polymerase III transcription apparatus. J. Mol. Biol. 310, 1–26. Geiduschek, E. P., and Tocchini-Valentini, G. P. (1988). Transcription by RNA polymerase III. Annu. Rev. Biochem. 57, 873–914. Ghavidel, A., and Schultz, M. C. (1997). Casein kinase II regulation of yeast TFIIIB is mediated by the TATA-binding protein. Genes Dev. 11, 2780–2789. Gomez-Roman, N., Grandori, C., Eisenman, R. N., and White, R. J. (2003). Direct activation of RNA polymerase III transcription by c-Myc. Nature 421, 290–294. Gottesfeld, J. M., Wolf, V. J., Dang, T., Forbes, D. J., and Hartl, P. (1994). Mitotic repression of RNA polymerase III transcription in vitro mediated by phosphorylation of a TFIIIB component. Science 263, 81–84. Hahn, S., and Roberts, S. (2000). The zinc ribbon domains of the general transcription factors TFIIB and Brf: conserved functional surfaces but different roles in transcription initiation. Genes Dev. 14, 719–730. Hamada, M., Huang, Y., Lowe, T. M., and Maraia, R. J. (2001). Widespread use of TATA elements in the core promoters for RNA polymerases III, II, and I in fission yeast. Mol. Cell. Biol. 21, 6870–6881. Hsieh, Y. J., Kundu, T. K., Wang, Z., Kovelman, R., and Roeder, R. G. (1999a). The TFIIIC90 subunit of TFIIIC interacts with multiple components of the RNA polymerase III machinery and contains a histone-specific acetyltransferase activity. Mol. Cell. Biol. 19, 7697–7704.
118
MOIR AND WILLIS
Hsieh, Y. J., Wang, Z., Kovelman, R., and Roeder, R. G. (1999b). Cloning and characterization of two evolutionarily conserved subunits (TFIIIC102 and TFIIIC63) of human TFIIIC and their involvement in functional interactions with TFIIIB and RNA polymerase III. Mol. Cell. Biol. 19, 4944–4952. Huang, Y., Hamada, M., and Maraia, R. J. (2000). Isolation and cloning of four subunits of a fission yeast TFIIIC complex that includes an ortholog of the human regulatory protein TFIIICbeta. J. Biol. Chem. 275, 31480–31487. Huang, Y., and Maraia, R. J. (2001). Comparison of the RNA polymerase III transcription machinery in Schizosaccharomyces pombe, Saccharomyces cerevisiae and human. Nucleic Acids Res. 29, 2675–2690. Huet, J., Conesa, C., Carles, C., and Sentenac, A. (1997). A cryptie DNA binding domain at the COOH terminus of TFIIIB70 affects formation, stability, and function of preinitiation complexes. J. Biol. Chem. 272, 18341–18349. Huibregtse, J. M., and Engelke, D. R. (1989). Genomic footprinting of a yeast tRNA gene reveals stable complexes over the 50 -flanking region. Mol. Cell. Biol. 9, 3244–3252. Ishiguro, A., Kassavetis, G. A., and Geiduschek, E. P. (2002). Essential roles of Bdp1, a subunit of RNA polymerase III initiation factor TFIIIB, in transcription and tRNA processing. Mol. Cell. Biol. 22, 3264–3275. Johnston, I. M., Allison, S. J., Morton, J. P., Schramm, L., Scott, P. H., and White, R. J. (2002). CK2 forms a stable complex with TFIIIB and activates RNA polymerase III transcription in human cells. Mol. Cell. Biol. 22, 3757–3768. Kassavetis, G. A., Bardeleben, C., Bartholemew, B., Braun, B. R., Joazeiro, C. A., Pisano, M., and Geiduschek, E. P. (1994). Transcription by RNA Polymerase III. 107–126. Kassavetis, G. A., Bardeleben, C., Kumar, A., Ramirez, E., and Geiduschek, E. P. (1997). Domains of the Brf component of RNA polymerase III transcription factor IIIB (TFIIIB): functions in assembly of TFIIIB-DNA complexes and recruitment of RNA polymerase to the promoter. Mol. Cell. Biol. 17, 5299–5306. Kassavetis, G. A., Braun, B. R., Nguyen, L. H., and Geiduschek, E. P. (1990). S. cerevisiae TFIIIB is the transcription initiation factor proper of RNA polymerase III, while TFIIIA and TFIIIC are assembly factors. Cell 60, 235–245. Kassavetis, G. A., Joazeiro, C. A., Pisano, M., Geiduschek, E. P., Colbert, T., Hahn, S., and Blanco, J. A. (1992). The role of the TATA-binding protein in the assembly and function of the multisubunit yeast RNA polymerase III transcription factor, TFIIIB. Cell 71, 1055–1064. Kassavetis, G. A., Kumar, A., Letts, G. A., and Geiduschek, E. P. (1998a). A postrecruitment function for the RNA polymerase III transcription-initiation factor IIIB. Proc. Natl. Acad. Sci. USA 95, 9196–9201. Kassavetis, G. A., Kumar, A., Ramirez, E., and Geiduschek, E. P. (1998b). Functional and structural organization of Brf, the TFIIB-related component of the RNA polymerase III transcription initiation complex. Mol. Cell. Biol. 18, 5587–5599. Kassavetis, G. A., Letts, G. A., and Geiduschek, E. P. (2001). The RNA polymerase III transcription initiation factor TFIIIB participates in two steps of promoter opening. EMBO J. 20, 2823–2834. Khoo, B., Brophy, B., and Jackson, S. P. (1994). Conserved functional domains of the RNA polymerase III general transcription factor BRF. Genes Dev. 8, 2879–2890. Kruppa, M., Moir, R. D., Kolodrubetz, D., and Willis, I. M. (2001). Nhp6, an HMG1 protein, functions in SNR6 transcription by RNA polymerase III in S. cerevisiae. Mol. Cell 7, 309–318.
TETRATRICOPEPTIDE REPEATS OF Tfc4
119
Kumar, A., Kassavetis, G. A., Geiduschek, E. P., Hambalko, M., and Brent, C. J. (1997). Functional dissection of the B00 component of RNA polymerase III transcription factor IIIB: a scaffolding protein with multiple roles in assembly and initiation of transcription. Mol. Cell. Biol. 17, 1868–1880. Kumar, A., Roach, C., Hirsh, I. S., Turley, S., deWalque, S., Michels, P. A., and Hol, W. G. (2001). An unexpected extended conformation for the third TPR motif of the peroxin PEX5 from Trypanosoma brucer. J. Mol. Biol. 307, 271–282. Lamb, J. R., Tugendreich, S., and Hieter, P. (1995). Tetratrico peptide repeat interactions: to TPR or not to TPR? Trends Biochem. Sci. 20, 257–259. Lapouge, K., Smith, J. S., Walker, A. P., Gamblin, J. S., Smerdon, J. S., and Rittinger, K. (2000). Structure of the TPR domain of p67phox in complex with Rac.GTP. Mol. Cell 6, 899–907. Larminie, C. G., Cairns, C. A., Mital, R., Martin, K., Kouzarides, T., Jackson, S. P., and White, R. J. (1997). Mechanistic analysis of RNA polymerase III regulation by the retinoblastoma protein. EMBO J. 16, 2061–2071. Lefebvre, O., Carles, C., Conesa, C., Swanson, R. N., Bouet, F., Riva, M., and Sentenac, A. (1992). TFC3: gene encoding the B-block binding subunit of the yeast transcription factor IIIC. Proc. Natl. Acad. Sci. USA 89, 10512–10516. Lefebvre, O., Ruth, J., and Sentenac, A. (1994). A mutation in the largest subunit of yeast TFIIIC affects tRNA and 5 S RNA synthesis. Identification of two classes of suppressors. J. Biol. Chem. 269, 23374–23381. Li, Y., Moir, R. D., Sethy-Coraci, I. K., Warner, J. R., and Willis, I. M. (2000). Repression of ribosome and tRNA synthesis in secretion-defective cells is signaled by a novel branch of the cell integrity pathway. Mol. Cell. Biol. 20, 3843–3851. Liao, Y. Willis, I. M., and Moir, R. D. (2003). Two TFIIIB factors, Brf1 and Bdp1, bind to the TFIIIC subunit, Tfc4, in the TPR Arrays. J. Biol. Chem. 278, 44467–44474. Lopez-de-Leon, A., Librizzi, M., Puglia, K., and Willis, I. M. (1992). PCF4 encodes an RNA polymerase III transcription factor with homology to TFIIB. Cell 71, 211–220. Lopez, S., Livingstone-Zatchej, M., Jourdain, S., Thoma, F., Sentenac, A., and Marsolier, M. C. (2001). High-mobility-group proteins NHP6A and NHP6B participate in activation of the RNA polymerase III SNR6 gene. Mol. Cell. Biol. 21, 3096–3134. Marck, C., Lefebvre, O., Carles, C., Riva, M., Chaussivert, N., Ruet, A., and Sentenac, A. (1993). The TFIIIB assembling subunit of yeast transcription factor TFIIIC has both tetratricopoptide repeats and basic helix loop-helix motifs. Proc. Natl. Acad. Sci. USA 90, 4027–4031. Margottin, F., Dujardin, G., Gerard, M., Egly, J. M., Huet, J., and Sentenac, A. (1991). Participation of the TATA factor in transcription of the yeast U6 gene by RNA polymerase C. Science 251, 424–426. McManus, M. T., and Sharp, P. A. (2002). Gene silencing in mammals by small interfering RNAs. Nat. Rev. Genet. 3, 737–747. Mital, R., Kobayashi, R., and Hernandez, N. (1996). RNA polymerase III transcription from the human U6 and adenovirus type 2 VAI promoters has different requirements for human BRF, a subunit of human TFIIIB. Mol. Cell. Biol. 16, 7031–7042. Moir, R. D., Puglia, K., Liao, Y., and Willis, I. M. (2004). A mutation in the TPR channel of Tfc4 directly increases Brf1 binding. Manuscript in preparation. Moir, R. D., Puglia, K. V., and Willis, I. M. (2000). Interactions between the tetratricopeptide repeat-containing transcription factor TFIIIC131 and its ligand, TFIIIB70. Evidence for a conformational change in the complex. J. Biol. Chem. 275, 26591–26598.
120
MOIR AND WILLIS
Moir, R. D., Puglia, K. V., and Willis, I. M. (2002a). A gain-of-function mutation in the second tetratricopeptide repeat of TFIIIC131 relieves autoinhibition of Brfl binding. Mol. Cell. Biol. 22, 6131–6141. Moir, R. D., Puglia, K. C., and Willis, I. M. (2002b). Autoinhibition of TFIIIB70 binding by the tetratricopeptide repeat containing subunit of TFIIIC. J. Biol. Chem. 277, 694–701. Moir, R. D., Sethy-Coraci, I., Puglia, K., Librizzi, M. D., and Willis, I. M. (1997). A tetratricopeptide repeat mutation in yeast transcription factor IIIC131 (TFIIIC131) facilitates recruitment of TFIIB-related factor TFIIIB70. Mol. Cell. Biol. 17, 7119–7125. Paule, M. R., and White, R. J. (2000). Survey and summary: transcription by RNA polymerases I and III. Nucleic Acids Res. 28, 1283–1298. Rameau, G., Puglia, K., Crowe, A., Sethy, I., and Willis, I. (1994). A mutation in the second largest subunit of TFIIIC increases a rate limiting step in transcription by RNA polymerase III. Mol. Cell. Biol. 14, 822–830. Rice, L. M., and Brunger, A. T. (1999). Crystal structure of the vesicular transport protein Sec17: implications for SNAP function in SNARE complex disassembly. Mol. Cell 4, 85–95. Rozenfeld, S., and Thuriaux, P. (2001). Genetic interactions within TFIIIC, the promoter-binding factor of yeast RNA polymerase III. Mol. Genet. Genomics 705–710. Ruth, J., Conesa, C., Dieci, G., Lefebvre, O., Dusterhoft, A., Ottonello, S., and Sentenac, A. (1996). A suppressor of mutations in the class III transcription system encodes a component of yeast TFIIIB. EMBO J. 15, 1941–1949. Scheufler, C., Brinker, A., Bourenkov, G., Pegoraro, S., Moroder, L., Bartunik, H., Hartl, F. U., and Moarefi, I. (2000). Structure of TPR domain-peptide complexes: critical elements in the assembly of the Hsp70-Hsp90 multichaperone machine. Cell 101, 199–210. Schramm, L., and Hernandez, N. (2002). Recruitment of RNA polymerase III to its target promoters. Genes Dev. 16, 2593–2620. Schultz, P., Marzouki, N., Marck, C., Ruet, A., Oudet, P., and Sentenac, A. (1989). The two DNA binding domains of yeast transcription factor tau as observed by scanning transmission electron microscopy. EMBO J. 8, 3815–3824. Scott, P. H., Cairns, C. A., Sutcliffe, J. E., Alzuherri, H. M., McLees, A., Winter, A. G., and White, R. J. (2001). Regulation of RNA polymerase III transcription during cell cycle entry. J. Biol. Chem. 276, 1005–1014. Sethy-Coraci, I., Moir, R. D., Lopez-de-Leon, A., and Willis, I. M. (1998). A differential response of wild type and mutant promoters to TFIIIB70 overexpression in vivo and in vitro. Nucleic Acids Res. 26, 2344–2352. Shen, Y., Igo, M., Yalamanchili, P., Berk, A. J., and Dasgupta, A. (1996). DNA binding domain and subunit interactions of transcription factor IIIC revealed by dissection with poliovirus 3C protease. Mol. Cell. Biol. 16, 4163–4171. Sinn, E., Wang, Z., Kovelman, R., and Roeder, R. G. (1995). Cloning and characterization of a TFIIIC2 subunit (TFIIIC beta) whose presence correlates with activation of RNA polymerase III-mediated transcription by adenovirus E1A expression and serum factors. Genes Dev. 9, 675–685. Sutcliffe, J. E., Cairns, C. A., McLees, A., Allison, S. J., Tosh, K., and White, R. J. (1999). RNA polymerase III transcription factor IIIB is a target for repression by pocket proteins p107 and p130. Mol. Cell. Biol. 19, 4255–4261.
TETRATRICOPEPTIDE REPEATS OF Tfc4
121
Taylor, P., Dornan, J., Carrello, A., Minchin, R. F., Ratajczak, T., and Walkinshaw, M. D. (2001). Two structures of cyclophilin 40. folding and fidelity in the tpr domains. Structure (Camb.) 9, 431–438. Upadhya, R., Lee, J., and Willis, I. M. (2002). Maf1 Is an essential mediator of diverse signals that repress RNA polymerase III transcription. Mol. Cell 10, 1489–1494. Wang, Z., and Roeder, R. G. (1995). Structure and function of a human transcription factor TFIIIB subunit that is evolutionarily conserved and contains both TFIIB- and high-mobility-group protein 2-related domains. Proc. Natl. Acad. Sci. USA 92, 7026–7030. Warner, J. (1999). The economics of ribosome biosynthesis in yeast. Trends Biochem. Sci. 24, 437–440. White, R. J. (1998a). RNA Polymerase III Transcription. 2nd ed., Springer, New York. White, R. J. (1998b). Transcription factor IIIB: An important determinant of biosynthetic capacity that is targeted by tumour suppressors and transforming proteins. Int. J. Oncol. 12, 741–748. White, R. J., Gottlieb, T. M., Downes, C. S., and Jackson, S. P. (1995). Cell cycle regulation of RNA polymerase III transcription. Mol. Cell. Biol. 15, 6653–6662. Willis, I., Schmidt, P., and Soll, D. (1989). A selection for mutants of the RNA polymerase III transcription apparatus. PCF1 stimulates transcription of tRNA and 5S RNA genes. EMBO J. 8, 4281–4288. Yaffe, M. B. (2002). How do 14-3-3 proteins work? Gatekeeper phosphorylation and the molecular anvil hypothesis. FEBS Lett. 513, 53–57.
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MECHANISM OF RNA POLYMERASE I TRANSCRIPTION By LUCIO COMAI Department of Molecular Microbiology and Immunology, Keck School of Medicine, University of Southern California, Los Angeles, CA 90033
I. Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Ribosomal DNA Gene Structure . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Proximal Promoter . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Enhancers . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Space-Promoters and Terminators . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. Species Specificity of Ribosomal RNA Transcription . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Factors Involved in Ribosomal RNA Transcription . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. RNA Polymerase I Core Enzyme. . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. RNA Polymerase I–Associated Factors . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Auxiliary Factors. . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Assembly of an RNA Polymerase I Initiation Complex in Vertebrates. . . . .. . . . . . VI. Factors Required for RNA Polymerase I Transcription in Yeast . . . . . . . . . . . .. . . . . . VII. Regulatory Mechanisms . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Regulation by Posttranslational Modification of Auxiliary Factors . . . .. . . . . . B. Regulation by Tumor Suppressor Proteins. . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Regulation of Transcription Elongation. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Regulation of RNA Polymerase I and Associated Factors. . . . . . . . . . . . . . .. . . . . . VIII. Chromatin and RNA Polymerase I Transcription . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IX. Conclusions . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Acknowledgments . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction Eukaryotic cells possess three nuclear RNA polymerases, each of which is dedicated to the transcription of a specific set of cellular genes. RNA polymerase (Pol) I is devoted to the transcription of the ribosomal DNA genes, which are found in multiple arrayed copies in every eukaryotic cell. These genes encode for the large ribosomal RNA precursor, which is then processed into the three largest subunits of the ribosomal RNA, the 18S, 28S, and 5.8S RNAs (sizes are for human subunits; the sizes of the subunits for other organisms are slightly different). These are very stable structural RNAs that are incorporated into the nascent ribosomes. The concentration of these RNAs can only be regulated at the level of transcription rate and the rate of dilution through cell division. The other ribosome components, the ribosomal proteins and the 5S rRNA, require a parallel regulation. However, the level of regulation for ribosomal proteins 123 ADVANCES IN PROTEIN CHEMISTRY, Vol. 67
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content is often posttranscriptional, involving messenger RNA (mRNA) processing or turnover and protein degradation (Hadjiolov, 1985). Ribosomal DNA gene transcription and ribosome assembly occur in the nucleolus, a highly specialized nuclear compartment. Quantitatively, the ribosomal gene transcription accounts for about 40% of all cellular transcription in living cells. These numbers reflect the relevance of this process for cell function and support the notion that regulation RNA Pol I transcription represents a key step in ribosome production and in determining a cell’s potential for growth and proliferation (Larson et al., 1991; Nurse, 1985). In the last few years, the biochemical purification and the subsequent molecular cloning of several of the factors required to direct promoterspecific RNA Pol I transcription has stimulated a large number of investigative studies on the mechanisms that regulate ribosomal RNA production. In this chapter, I provide an overview of the RNA Pol I transcription system, with particular emphasis on the latest findings on the mechanisms that regulate it. Comprehensive reviews on yeast and vertebrate RNA polymerase I have been published in recent years, and they can provide additional information that is not covered in this chapter (Grummt, 1999; Hannan et al., 1998a; Moss and Stefanovsky, 1995; Paule, 1998; Reeder, 1999).
II. Ribosomal DNA Gene Structure Transcription by RNA Pol I is unique in some respects among these three polymerases: In contrast to RNA polymerases II and III (Pol II and Pol III), each of which is responsible for the transcription of many different genes, RNA Pol I directs RNA synthesis from a single class of genes, the ribosomal RNA (rRNA) genes, which are found in multiple, tandem, head-to-tail arrayed copies in the nucleoli of eukaryotic cells (Hadjiolov, 1985). In Xenopus, all the copies of the rDNA genes are found on a single chromosome, whereas in human cells the rDNA gene clusters are localized on the short arm of the five pairs of the acrocentric chromosomes (Long and Dawid, 1980). In Arabidopsis thaliana, two clusters of rDNA genes are localized near the telomeric region of chromosome 2 (Copenhaver and Pikaard, 1996). Chromosomal regions containing these loci have been named ‘‘nucleolar organizer regions’’ (NORs; Hadjiolov, 1985). NORs stained with colloidal silver techniques (AgNORs) evidence sites of active rRNA transcription. The AgNOR number is known to correlate with the proliferative activity of the cell population, and the AgNOR score of cancer cells (lung, bone, breast, gastric, colorectal) appears to be a good predictor of a patient’s prognosis (Derenzini and Trere, 1991; Derenzini et al., 1998; 2000). NORs and active RNA Pol I
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transcription define a region of the nucleus called the nucleolus, a membraneless organelle dedicated to the assembly of the ribosomes, a process that requires transcripts from all three nuclear RNA polymerases. Nucleoli can be directly visualized by light microscopy as densely staining regions within the nucleus (Scheer and Hock, 1999; Shaw and Jordan, 1995). The rDNA gene was among the earliest eukaryotic genes studied, and the Xenopus rDNA gene was the first isolated (Birniestiel et al., 1968) and the first cloned eukaryotic gene (Morrow et al., 1974). The structure of a single rDNA repeating unit can be divided into two major regions: the rRNA precursor and the intergenic spacer sequences (Fig. 1). The gene for the rRNA precursor varies between 7 and 13 kilobases (kb) in length depending on the species of origin (Hadjiolov, 1985; Long and Dawid, 1980). Once transcribed, the rRNA precursor is processed by endonucleases and exonucleases to yield the final mature rRNA subunits (28S, 18S, and 5.8S in mammals). The intergenic spacer region of the rDNA repeats includes all the sequences responsible for proper RNA Pol I transcription such as proximal promoters, spacer promoters, and terminators (Hadjiolov, 1985). In addition, this region has also been shown to include an origin of replication. Like the rDNA coding unit, this region of the rDNA repeat also varies in length, from about 10 kb in yeast to approximately
Fig. 1. Schematic representation of eukaryotic rDNA gene. IGR, intergenic region; T, terminator; ORI, origin of replication; spP, spacer promoter; pT, promoter-proximal terminator, pP, proximal promoter; ETS, external transcribed sequence; ITS, internal transcribed sequences.
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40 kb in humans. A large number of biochemical studies have established that the proximal promoter is the most important element controlling RNA Pol I transcription.
A. Proximal Promoter The RNA Pol I proximal promoter has been well characterized by deletion, linker scanning, and point mutation analyses in a variety of organisms (Clos et al., 1986b; Grummt, 1981a; Haltiner et al., 1986; Miller et al., 1985; Nagamine et al., 1987; Reeder et al., 1987). The mammalian rRNA promoter has two essential and specially spaced sequences: a CORE element and an upstream control element (UCE, also called UPE). The CORE element of the human promoter overlaps with the transcription start site, extending from þ20 to 45, and is required for specific initiation of transcription. Promoter efficiency is greatly increased by the UCE element, which includes the region from 180 to 107. The UCE is not defined as a classical eukaryotic enhancer, as the function of the UCE is sensitive to changes in orientation and distance to the core promoter element (Haltiner et al., 1986; Jones et al., 1988). CORE and UCE elements share regions of homology, indicating that they may provide contact surfaces to identical or related factors. The current model proposes that the UCE and CORE elements function in a cooperative manner to recruit the Pol I initiation complex to the transcript initiation site. In protozoa, fungi, and plants, the rRNA promoter is much simpler, consisting of only the initiation-site proximal element or CORE (Moss and Stefanovsky, 1995). Among mammals, human and mouse share three conserved sequences in the CORE region of the promoter, between nucleotides at positions 38 to 33, 20 to 12, and 1 to þ18, relative to the transcription initiation site (Financsek et al., 1982; Safrany et al., 1989). Apart from these relatively small regions, comparison of the sequences of the promoter elements between various organisms does not show any obvious sequence similarity, and the available evidence indicates that these differences in nucleotide sequences may result from adaptive evolutionary changes.
B. Enhancers In addition to the proximal promoter, distal enhancer-like elements have been identified in organisms such as mouse, Drosophila, and Xenopus (Reeder, 1984). These sequences consist of multiple repeats of either 61 or 81 bp in length and function in either orientation, either upstream or downstream of the promoter. In Xenopus, these repeated sequences share regions of homology with the rDNA promoter and can compete for basal
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factors with an intact promoter. Enhancers have been proposed to increase the number of stable complexes at the rDNA promoter (Moss and Stefanovsky, 1995; Reeder, 1984, 1999). However, the recent finding that removal of the entire yeast enhancer does not affect either rDNA transcription or cell growth has challenged the model that proposed a stimulatory role for the enhancer (Wai et al., 2001). Therefore, the physiological significance of the enhancer sequences has to be reevaluated.
C. Space-Promoters and Terminators In addition to proximal promoter and enhancers, elements named spacer-promoters are commonly found in the intergenic spacer of the rDNA repeat. These promoters transcribe in the same direction as the proper rDNA promoter; however, the resulting transcripts are very short lived (Kuhn and Grummt, 1987; Moss, 1983). These elements are usually located 160–180 bp upstream of the promoter as well as downstream of the rDNA gene (Baker and Platt, 1986). Interestingly, upstream spacerpromoters are followed by a terminator element (Grummt et al., 1986; Henderson and Sollner-Webb, 1986). A terminator element, in addition to being found between spacer and proximal promoter, is also located at the immediate 30 end of the rDNA gene and is the recognition site for a sequence-specific DNA binding protein (TTF-I, Rib 1), which specifically terminates RNA Pol I transcription. The physiological role of the spacerpromoter/upstream terminator is currently unclear, and two hypotheses have been put forward; first, the terminator proximal to the spacepromoter may terminate transcription immediately upstream of the proximal promoter and therefore function as a ‘‘road block’’ to prevent promoter occlusion by RNA polymerases initiated at spacer-promoters (Henderson et al., 1989). Alternatively, the spacer-promoter/terminator arrangement may function to deliver RNA polymerases to the proximal promoters and therefore increase promoter occupancy (Moss, 1983).
III. Species-Specificity of Ribosomal RNA Transcription Interspecific cell hybrids between human and murine cells do not express the rDNA genes of both genes. Interspecific crosses in plants and between Xenopus species have also shown a gene-specific dominance effect. This phenomenon is known as ‘‘nucleolar dominance’’ and, depending on the chromosomal organization of the hybrid, the rDNA of either one of the species is expressed (Reeder, 1985). These findings indicated that there are some aspects of rDNA transcription that are species specific. The incompatibility between different species has then been
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further observed in cell-free extracts (Grummt, 1981b; Miesfeld and Arnheim, 1984; Miesfeld et al., 1984). For example, although a human nuclear extract can efficiently transcribe the human rDNA gene in vitro, it is incapable of supporting transcription of a mouse ribosomal DNA gene. Nuclear fractionation analyses indicated that, between humans and mice, the selectivity factor SL1 was the species-specific factor (Learned et al., 1985; Schnapp et al., 1991). Addition of mouse SL1 to a human nuclear extract was sufficient to redirect transcription from the cloned mouse rDNA gene, and vice versa (Bell et al., 1990). Because SL1 is sufficient to direct transcription from the homologous promoter in a cell-free transcription assay, failure to express this factor may explain nucleolar dominance in human–mouse somatic hybrids. Interestingly, promoter-swapping experiments have identified a short nucleotide sequence within the CORE element of the mouse promoter that is sufficient, when substituted to the corresponding region of the human promoter, to direct transcription from a mouse extract (Safrany et al., 1989). This result indicated that this element might specify species-specific interactions with the selectivity factor SL1. However, the presence of one species-specific factor is probably not at the basis of nucleolar dominance in other species. For example, the spacing between the UCE and CORE elements is crucial in promoter activity and species-specificity of rRNA transcription between mouse and Xenopus, and a 5-bp insertion between UPE and CORE of the Xenopus promoter, which removes a half-helical turn between these two elements, converts it into a mouse specific promoter in vitro (Pape et al., 1990). In addition, epigenetic mechanisms are also likely to play a role in this phenomenon. Indeed, nucleolar dominance in interspecific plant hybrids appears, at least in part, to be under the control of epigenetic variables such as DNA methylation and histone modifications (Chen and Pikaard, 1997; Chen et al., 1998). Thus, several aspects, such as speciesspecific transcription factors, promoter sequences and spacing, and epigenetic effects, contribute to the species-specific nature of RNA Pol I transcription.
IV. Factors Involved in Ribosomal RNA Transcription The development of a cell-free transcription system, which faithfully transcribes a synthetic rDNA template, was first described about two decades ago (Grummt, 1981b; Haglund and Rothblum, 1987; Learned and Tjian, 1982; Mishima et al., 1981; Wilkinson and Sollner-Webb, 1982). This accomplishment encouraged a large number of biochemical studies on the dissection of the RNA Pol I transcription system. In vitro transcription assays coupled to column fractionation analyses of cell extracts from a
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variety of organisms led to the identification of several fractions containing activities required to direct efficient and accurate initiation by RNA Pol I (Heilgenthal and Grummt, 1991; Learned et al., 1985, 1986; Mishima et al., 1982; Schnapp and Grummt, 1991). Twenty years later, most of these activities have been purified to homogeneity and cloned.
A. RNA Polymerase I Core Enzyme One of the fractions required to direct rDNA transcription contains the RNA polymerase I enzymatic activity. Purification of mammalian RNA polymerase I indicates that the core enzyme is a multisubunit complex with a molecular mass of >500 kDa. (Hannan et al., 1998c; Song et al., 1994). Yeast RNA polymerase I is composed of 15 subunits, five of which are shared by all three yeast nuclear RNA polymerases (Sentenac, 1985). Mouse RNA Pol I is composed of 11 subunits with remarkable correspondence to those of yeast (Song et al., 1994). Several subunits of human RNA Pol I have been cloned, and their sequence analysis shows high homology to the corresponding yeast and mouse subunits (Dammann and Pfeifer, 1998; Hannan et al., 1998c; Seither and Grummt, 1996; Seither et al., 1997a). In addition, complementation experiments in yeast show that four subunits that are shared by all three nuclear RNA polymerase are functionally interchangeable between humans and yeast (Shpakovski et al., 1995). Amino acid sequence deduced from the cDNA clones encoding the two largest subunits of mouse and rat RNA Pol I indicate that these two subunits are homologous to the B and B0 subunits of Escherichia coli. However, the smaller subunits lack bacterial homologues. Thus, it is likely that the RNA polymerase’s largest subunits, which are required for the catalytic function, have evolved from a common ancestral progenitor, whereas the smaller subunits may play a more specialized function required only in eukaryotic cells. The largest subunit of rat RNA polymerase I is phosphorylated by an enzyme whose biochemical properties are reminiscent of casein kinase II. The significance of this phosphorylation remains unclear, although modification RNA polymerase I subunits can affect promoter-specific transcription initiation in Achantamaeba.
B. RNA Polymerase I–Associated Factors Biochemical fractionation studies indicated that purified RNA polymerase I activity can be dissociated into two fractions, both of which possess RNA polymerization activity, but only one of which has promoter-specific activity. The analysis of these different forms of RNA polymerases led to the identification of a set of polypeptides that are loosely associated with
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the polymerase core subunits. On the basis of these findings, it has been proposed that a transcriptionally active, promoter-specific RNA polymerase I holoenzyme requires the presence of one or more of these activities.
1. PAF53 Extensive purification of the mouse RNA polymerase I and comparative analysis of the polypeptide composition of two forms of RNA polymerases indicated that the inactive polymerase fraction was lacking a family of factors of molecular mass around 53 kD. Three polymerase-associated factors (PAF), termed PAF53, PAF51, and PAF49, have been subsequently purified and cloned from mouse cells (Hanada et al., 1996). Sequence analysis of PAF53 shows similarity to one of the yeast RNA Pol I subunits, RPA49. RPA49, similar to PAF53, is easily released from the yeast core enzyme, and it has been shown to be not absolutely required for yeast viability. PAF53 is found to associate with RNA Pol I purified from exponentially growing cells but not with RNA Pol I from quiescent cells. The addition of antibody against PAF53 inhibits promoter-specific, but not random, transcription, and in vitro studies indicated that PAF53 interacts with the auxiliary factor UBF (Seither et al., 1997b). On the basis of findings, it has been proposed that PAF53 plays an essential role in promoter-specific initiation by mediating the recruitment of RNA Pol I to the initiation complex.
2. TIF-IA/RRN3 Several laboratories have identified a factor, termed TIF-IA, TFIC, or Factor C, that is tightly associated with the RNA Pol I core enzyme (Brun et al., 1994; Buttgereit et al., 1985; Mahajan et al., 1990; Mahajan and Thompson, 1990; Schnapp et al., 1990, 1993). Biochemical studies indicated that TIF-IA plays an important role in growth-regulated transcription of rDNA genes, as addition of this factor to extracts from cells in which rRNA transcription is virtually shut off (i.e., serum starvation, cyclohexamide treatment) can restore promoter-specific initiation of RNA Pol I transcription. In vitro reconstituted transcription assays indicated that this factor does not affect the formation of a stable preinitiation complex; however, it appears to be required for the formation of the first phosphodiester bond. In addition, it can stimulate the rate of RNA Poly I transcription reinitiation. TIF-IA has recently been cloned and, it has been shown to be the mammalian homologue of the yeast RRN3 protein, a factor involved in yeast RNA Pol I transcription (Bodem et al., 2000; Miller et al., 2001; Moorefield et al., 2000). Human TIF-IA (subsequently also named hRRN3) and yeast RRN3 share 21% amino acid identity and 43%
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similarity (Moorefield et al., 2000). The amino acid homology between the two proteins is further reflected in their functionality, as human RRN3 is able to complement the lethal phenotype of an RRN3-null yeast mutant. To date, this is the only vertebrate RNA Pol I factor with sequence homology to yeast factors.
3. TIF-IC The factor TIF-IC has been identified as a 65-kDa protein loosely associated with RNA Pol I (Schnapp et al., 1994b). Biochemical characterization of this factor indicated that it is required for the assembly of the preinitiation complex and the formation of the first internucleotide bond. In addition, TIF-IC functions in chain elongation and suppresses RNA polymerase pausing during transcription elongation (Schnapp et al., 1994b). To date, TIF-IC has only been identified in the murine system. Because this factor has not been purified to homogeneity and cloned, its molecular identity remains unknown.
C. Auxiliary Factors 1. Upstream Binding Factor The first cloned auxiliary factor specific for RNA Pol I transcription was the upstream binding factor (UBF). This 97-kDa factor has been purified and cloned from organisms such as human, mouse, rat, and Xenopus (Bachvarov and Moss, 1991; Bell et al., 1990; Jantzen et al., 1990; McStay et al., 1991a,b; O’Mahony and Rothblum, 1991; Pikaard et al., 1989, 1990; Schnapp et al., 1991). UBF is a DNA-binding protein and recognizes both DNA control elements (CORE and upstream control element-UCE) of the rDNA promoter. UBF is the first transcription factor shown to contain a repeated region of 85 amino acids with homology to the nonhistone chromosomal high mobility group proteins 1 and 2 (HMG1 and 2; Bachvarov and Moss, 1991; Jantzen et al., 1990; Fig. 2). On the basis of this homology, these domains have been termed HMG boxes. Since the identification of HMG boxes in UBF, other transcription factors, such as sex-determining factor SRY, the lymphoid enhancer-binding factors LEF-1 and TCF-1, and the mitochondrial transcription factor mtTF1, have been shown to possess one or more HMG boxes (Grosschedl et al., 1994). An important feature of the members of this family of factors is that they all bind to the minor groove of the DNA and have the ability to modulate DNA structure by bending. However, although transcription factors containing one HMG box bind DNA in a sequence-specific fashion, factors with multiple HMG boxes such as UBF recognize irregular
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Fig. 2. Functional domains of human upstream binding factor.
DNA structures in a conformation-specific rather than sequence-specific manner (Copenhaver et al., 1994). For this reason, it has been proposed that UBF plays an architectural role at the ribosomal DNA promoter. In agreement with this model, electron microscopic imaging analysis indicated that DNA is wrapped around a UBF dimer in a structure reminiscent of the nucleosome (Bazett-Jones et al., 1994). Therefore, UBF may function as a scaffold protein, which, by establishing the proper DNA–protein structure, facilitates the formation of the transcription initiation complex at the ribosomal DNA promoter. This model also predicts that the UCE and core elements will cooperate with each other in the recruitment of UBF. Indeed, changes in the spacing between the UCE and core have profound consequences on the ability of UBF to stimulate RNA Pol I transcription. Biochemical analysis of human UBF mutants indicated that the first HMG box is necessary and sufficient for DNA binding, whereas HMG boxes, 2, 3, and 4 modulate the DNA binding efficiency ( Jantzen et al., 1992; Maeda et al., 1992). Biochemical studies have also indicated that UBF forms dimers and that the amino-terminal region of this factor mediates dimerization ( Jantzen et al., 1992a; McStay et al., 1991a; O’Mahony et al., 1992a). The carboxy-terminal region of UBF is rich in serine and acidic amino acids and is required for transcriptional activation ( Jantzen et al., 1992). Serines that reside within the carboxy-terminal domain are potential sites of phosphorylation (O’Mahony et al., 1992a). An important observation regarding UBF function in RNA Pol I transcription comes from studies on the localization of rRNA transcription, rDNA genes, and UBF by confocal microscopy. These studies indicated that UBF is not exclusively bound to the actively transcribed genes but is also associated with inactive genes (Bell et al., 1997; Gebrane-Younes et al., 1997; Junera et al., 1997; Zatsepina et al., 1993;). This finding led to the hypothesis that UBF binds and selects potentially active genes and that covalent modifications of UBF may be a potential mechanism for UBF activation and for
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triggering the transition from potentially active to actively transcribing genes. A natural splicing variant of UBF (UBF2), missing 37 amino acids of HMG box2, has been identified in human, mouse, and frog (Kuhn et al., 1994; O’Mahony and Rothblum, 1991). Biochemical analyses indicated that this variant has reduced DNA binding activity and functions poorly in transcription. Nevertheless, the physiological role of UBF2 remains unclear. Intriguingly, although UBF is quite conserved among vertebrates, a homologous protein is not found in Saccaromyces cerevisiae and Acanthamoeba castellanii although an activity with biochemical properties similar to UBF may be present in yeast (see following).
2. UBF and the Enhancers Biochemical studies using Xenopus extracts have revealed that UBF, in addition to recognizing the proximal promoter, binds cooperatively to the enhancer sequences (Pikaard et al., 1989; Putnam et al., 1994). These analyses indicated that enhancers compete quite efficiently with promoter sequences for UBF binding, possibly suggesting a ‘‘hand-over’’ process, in which UBF that is recruited to enhancers will then translocate to the proximal promoter (Osheim et al., 1996). However, more recent results have suggested a revised mechanism, in which enhancer-bound UBF stimulates the formation of a productive initiation complex at the proximal promoter by increasing the local density of essential factors, such as SL1, to this promoter (Sullivan and McStay, 1998; Sullivan et al., 2001). Interestingly, both UBF1 and UBF2 bind to the enhancer DNA, indicating a potential role for UBF2 at the enhancer sequences (McStay et al., 1997).
3. Selectivity Factor 1 An essential factor required for accurate RNA Pol I transcription is the selectivity factor SL1 (SL1; Clos et al., 1986a; Learned et al., 1985). As mentioned earlier, SL1 is also a species-specific factor that directs promoter-specific transcription in the presence of its cognate template. SL1 does not bind specifically to the rRNA promoter; however, in the presence of UBF, it forms a strong cooperative DNA-binding complex at the ribosomal DNA promoter that is critical for initiation of transcription (Bell et al., 1988, 1989; Learned et al., 1986). Mutations in the promoter sequences that affect either the binding of UBF to the DNA template or the interaction of UBF with SL1 result in drastic reduction of transcription activity (Bell et al., 1988). These findings indicate that the interactions between UBF and its DNA recognition sequence, and between UBF and SL1, play a major role in RNA Pol I transcription. For many years the molecular identification of SL1 has proven a difficult task. The
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recognition that this factor is a multisubunit complex containing the TATA-binding protein (TBP) led to its purification and full characterization (Comai et al., 1992). Mammalian SL1, like the RNA polymerase II factor TFIID and the RNA polymerase III factor TFIIIB, is a protein complex composed of TBP and TBP-associated factors (TAFs; Fig. 3). SL1 contains three TAFs of molecular mass 48 (TAFI48), 63 (TAFI63), and 110 kD (TAFI110; Comai et al., 1994; Zomerdijk et al., 1994). The deduced amino acid sequences revealed that all three Pol I TAFs are novel proteins and do not display any sequence homology to either the RNA Pol II TAFs or any other known protein. TAFI48 contains two tandem motifs whose role is currently unknown. TAFI63 contains two putative Zn fingers that may be involved in binding to the rDNA promoter. Protein–protein interaction assays indicate that each subunit of SL1 makes contact with each other, supporting the idea that this strong complex is held together by a multitude of protein interaction surfaces. Once at the promoter, SL1 makes contact with DNA through TAFI63 and TAFI110, whereas TBP and TAFI48 are likely to mediate the interaction of SL1 with UBF. Mouse SL1 is also a multiprotein complex of TBP and TAFIs (Eberhard et al., 1993), and the corresponding murine factors have been subsequently cloned and characterized (Heix et al., 1997). This generated much interest because it was thought that comparison of the primary amino acid sequences between the human and mouse TAFIs could provide some clues on the molecular basis of the species-specific properties of SL1. However, human and mouse TAFIs share an extremely high degree of amino acid identity, and there is not any obvious differences between the hortologous factors. Thus, if the species specificity resides within one or more of the TAFs, it is likely that subtle differences may be at the base of this phenomenon.
Fig. 3. TATA-binding protein/TATA-binding protein–associated factor complexes.
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The finding that a similar TBP/TAF arrangement is part of the RNA Pol I factor SL1, the RNA Pol II transcription factor TFIID, and the RNA Pol III transcription factor TFIIIB (Hernandez, 1993; Rigby, 1993) led to the conclusion that TBP is a universal transcription factor involved in transcription by all three nuclear RNA polymerases. A consequence of this type of TBP/TAF complex is that TBP, the common subunit, is not likely to be responsible for selectivity between RNA Pol I, II, and III promoters. Instead, the TAFs within each complex must direct assembly of class-specific initiation complexes. Therefore, unique specificities and protein–protein interactions will dictate the selective association of TBP/TAFs complexes. Indeed, in vitro experiments indicated that when TBP is bound first by TAFI48, 63, or 110, subunits of TFIID such as TAFII150 and 250, which are known to directly associate with free TBP in solution, are incapable of associating with the precomplexed TBP. Similarly, if TBP first forms a complex with TAFII250 or 150, TAFIs were prevented from binding with TBP. These results indicate that, at least in vitro, mutually exclusive binding prevents the formation of hybrid TBP/TAF complexes. However, we cannot rule out that in the cell, other factors, such as cellular compartmentalization, may contribute to the formation of promoter-selective TBP/TAFs complexes.
4. Additional Factors Involved in RNA Polymerase I Transcription In addition to the aforementioned set of transcription factors, several laboratories have identified a number of other cellular proteins that can modulate the activity of RNA Pol I in vitro. One of these factors, EIBF (enhancer I binding factor), is related to the Ku antigen (Ku70/80) and has been shown to interact with the proximal promoter as well as the enhancer elements of the rat rDNA gene (Ghosh et al., 1993; Zhang and Jacob, 1990). Treatment of cell extract with Ku antibody resulted in inhibition of rDNA transcription, and addition of purified rat EIBF/Ku to in vitro transcription reaction can reverse the inhibition (Hoff et al., 1994). The inhibitory effect is only observed when the antibody is added to the reaction before preinitiation complex formation. Intriguingly, the Ku complex (Ku70/80 and DNA-PKcs) has also been reported to repress RNA Pol I transcription (Kuhn et al., 1993, 1995). Therefore, the relationship between the EIBF/Ku-related antigen and the Ku complex remains to be further investigated. A core promoter-binding factor (CPBF) has been described in rats. This is a dimeric factor that binds to the rDNA promoter sequences and increases the rate of transcription initiation (Liu and Jacob, 1994). CPBF appears to exert its function by physically interacting with EIBF/Ku at the rDNA promoter (Liu and Jacob, 1994; Niu et al., 1995). CPBF is highly
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homologous to human USF (upstream stimulating factor; consisting of two subunits USF1 and USF2), a basic helix–loop–helix (bHLH) zipper protein involved that binds to E-boxes of RNA Pol II–transcribed genes. Cotransfection assays indicated that a homodimer of USF1 or USF2 inhibits ribosomal DNA transcription, whereas a USF1/2 heterodimer stimulates transcription (Ghosh et al., 1997). Another cellular factor that has been proposed to stimulate ribosomal RNA synthesis is topoisomerase I. The treatment of cells with a DNA topoisomerase I–specific inhibitor, camptothecin, can greatly reduce transcription from supercoiled rDNA templates in vitro (Garg et al., 1987; Zhang et al., 1988), and studies in yeast demonstrated that topoisomerase I is required for the efficient elongation of rRNA chains (Schultz et al., 1992). More recently, mouse basonuclein has been shown to bind to the rDNA promoter and to stimulate Pol I transcription in vitro (Tian et al., 2001; Tseng et al., 1999). Basonuclein is a Zn finger protein that is localized to the nucleolus of the growing mouse oocytes. Unlike Xenopus oocytes, which amplify their rDNA to generate enough ribosomes to sustain their massive growth, mouse oocytes do not go through rDNA amplification. On the basis of these findings, mouse oocytes are probably using factors such as basonuclein, rather than gene amplification, to boost rDNA transcription to support the increased demand for cell growth. This finding indicates that, in addition to the normal set of auxiliary factors, cell-specific or developmentally regulated factors can regulate ribosomal DNA transcription according to the biosynthetic needs of the cell.
V. Assembly of an RNA Polymerase I Initiation Complex in Vertebrates The identification of the cis-acting sequences and protein factors involved in ribosomal DNA transcription strongly suggested that the network of protein–protein and protein–DNA interactions among UBF, SL1, and the promoter elements plays a major role in the assembly of a stable preinitiation complex. A number of studies have established that UBF and SL1 play a key role in this process and are necessary to direct a high level of RNA Pol I transcription in vitro. The current stepwise model of factors assembly predicts that the binding of UBF dimer to the UCE and CORE elements is a prerequisite for the recruitment of the selectivity factor SL1 to the rDNA promoter (Fig. 4, step I). Biochemical studies have indicated that two subunits of SL1, TAFI48 and TBP, interact directly with the carboxy-terminal acidic domain of UBF (Beckmann et al., 1995; Tuan et al., 1999; step II). This finding demonstrates that the function of the carboxyl-terminal activation domain of UBF is to recruit the essential
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Fig. 4. Stepwise model of transcription complex assembly. UBF, upstream binding factor. (See Color Insert.)
factor SL1 to the promoter. Once bound to UBF, SL1 interacts with the DNA on the promoter region, as observed by the extension of the UBF footprinting pattern upstream from the UCE and downstream from the CORE region (Bell et al., 1988; Jantzen et al., 1992; Tuan et al., 1999). In vivo DNA cross-linking studies have determined that two subunits of SL1, TAFI63 and TAFI110, are in close contact with the DNA (Beckmann et al., 1995). The subsequent recruitment of the RNA polymerase core enzyme appears to be mediated by multiple protein–protein interactions (step III). The RNA Pol I–associated factor PAF53 has been shown to bind directly to UBF (Schnapp et al., 1994a), whereas TIF-IA/RRN3 bridges RNA Pol I to the SL1 complex (Miller et al., 2001). The assembly of the initiation complex on the promoter and the transition from a closed to an open complex is then followed by promoter clearance and transcription elongation by RNA Pol I (step IV). Unlike the RNA polymerase II system, RNA polymerase I transcription does not require a form of energy such as ATP for initiation and elongation (Gokal et al., 1990; Schnapp and
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Grummt, 1991). The fate of each component of the preinitiation complex once transcription is initiated has been recently investigated in vitro using immobilized DNA templates (Aprikian et al., 2001; Panov et al., 2001). In the human system, UBF and SL1 appear to remain bound to the promoter, ready to recruit a new RRN3/Pol I complex for a new round of transcription. Similar experiments in yeast have indicated that UAF is the only factor that remains bound to the promoter once the polymerase initiates transcription, whereas the core factor (CF), TBP, and RRN3 dissociate from the promoter-bound upstream activator factor (UAF) once elongation of transcription by RNA Pol I occurs. Whether the differences in the mechanisms of RNA Pol I postassembly reflect species-related differences remains to be further investigated. The stepwise assembly model has recently been challenged by a number of studies that have identified RNA Pol I holoenzymes complexes (Albert et al., 1999; Hannan et al., 1999; Saez-Vasquez and Pikaard, 1997; Seither et al., 1998). These complexes, which have been purified from mouse, frog, and plant extracts by either conventional or affinity chromatography, contain the RNA Pol I core enzyme associated with a variety of other factors. Interestingly, although some of the associated proteins are bona fide RNA Pol I factors, such as UBF and SL1, others are proteins with kinase and histone acetyltransferase activities or are members of DNA repair and replication pathways. The significance of these findings is not clear yet. However, the RNA Pol I holoenzyme may mediate different cellular functions linked to rDNA transcription, such as chromatin modification, DNA repair, and DNA replication. It is also interesting to note that each of the isolated complexes have a different set of associated factors, and in at least two reports, UBF appears to be excluded from the RNA Pol I holoenzyme complex (Albert et al., 1999; Hannan et al., 1999).
VI. Factors Required for RNA Polymerase I Transcription in Yeast An unexpected finding in the RNA Pol I field was the realization that yeast do not have factors that resemble the two essential vertebrate RNA Pol I auxiliary factors, UBF and SL1. Indeed, many attempts to clone yeast RNA Pol I factors by homology screening did not yield any results. Moreover, analysis of the yeast genomic sequence does not show any protein with obvious similarity to UBF or SL1. This was unexpected, as the RNA Pol II and Pol III machineries are rather conserved between vertebrates and yeast. Consequently, the identification of yeast factors involved in RNA Pol I transcription has evolved separately from the
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vertebrate field. The development of an in vitro transcription assay for yeast (Riggs and Nomura, 1990; Schultz et al., 1991), in combination with genetic approaches, has resulted in the isolation and characterization of two multiprotein complexes required for RNA Pol I transcription. One of these complexes, the CORE factor, is composed of four polypeptides: RRN6, RRN7, RRN11, and TBP (Keys et al., 1994; Lalo et al., 1996; Lin et al., 1996). The second factor, the UAF, consists of six subunits: RRN5, RRN9, RRN10, histone H3, histone H4, and a 30-kD factor (Keys et al., 1996; Siddiqi et al., 2001b). Each of these proteins has been cloned, and apart from TBP, sequence analysis does not show any motif that resembles the mammalian TAFIs or UBF. Therefore, from the RNA Pol machinery from yeast, it appears that vertebrates have evolved separately. Nevertheless, functional studies indicate that the yeast and vertebrate factors may share some of their biochemical properties. For example, UAF binds strongly to the upstream activating sequences (UAS) and is necessary for the formation of a stable preinitiation complex at the yeast rDNA promoter, a function that is reminiscent of vertebrate UBF. In addition, the core factor interacts with the core region, and likewise, SL1 selects the site of transcription initiation and recruits RNA Pol I to the promoter. The precise role of TBP in yeast RNA Pol I transcription remains controversial. Unlike SL1, the association of yeast TBP with the core factors RRN6, 7, and 11 is weak; therefore, it is a matter of debate whether yeast TBP is a true component of the CORE factor. In addition, in vitro transcription assays have yielded different results in regard to TBP’s role in yeast RNA Pol I transcription initiation. One of the models proposes that TBP is not required for basal RNA Pol I transcription and is exclusively necessary to recruit CF and RNA polymerase to the promoter site in a UAF-dependent manner (Siddiqi et al., 2001a; Steffan et al., 1996). Another view is that TBP can also stimulate transcription independent of UAF (Aprikian et al., 2000). Once the CORE and UAF factors are brought to the promoter, the recruitment of the RNA Pol I is mediated, at least in part, by the factor RRN3 (Yamamoto et al., 1996). The recent discovery of high homology between RRN3 and mammalian TIF-IA has generated a lot of interest. As its mammalian counterpart, yeast RRN3 is loosely associated with the polymerase core enzyme, and this association appears to be regulated during growth. As discussed above, human RRN3/TIF-IA can rescue the growth disadvantage of RRN3-null yeast strains, indicating similar functionalities between the homologue proteins. Recent studies show that RNN3 binds to one of the subunits of the core factor CF, indicating an important role in mediating the recruitment of the RNA Pol core enzyme to the promoter (Peyroche et al., 2000).
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VII. Regulatory Mechanisms RNA Pol I transcription has been shown to be regulated in a growth rate–and differentiation-dependent manner (Grummt, 1999; Reeder, 1999), and a number of extracellular stimuli, such as serum deprivation, glucocorticoids, insulin, viral infection, and phorbol esters, affect the rate of rRNA synthesis (Cavanaugh and Thompson, 1985, 1986; Cavanaugh et al., 1984; Grummt et al., 1976; Hannan et al., 1998b; Learned et al., 1983; Mahajan and Thompson, 1990; Mahajan et al., 1990). In addition, RNA Pol I transcription is regulated during the progression of the cell cycle, with the maximal amount of rRNA synthesis attained during G2 and subsequent silencing at mitosis (Hadjiolov, 1985). Thus, mechanisms that repress or stimulate RNA Pol I transcription provide a mean to adjust protein synthesis according to the demand for cell growth. In principle, ribosomal RNA synthesis can be regulated at several levels: first, as there are several hundred rDNA genes in a genome, rRNA synthesis can be adjusted by modulating the number of genes that are actively transcribed. This regulatory process will influence the number of productive preinitiation complexes formed at the rDNA promoters. Second, regulation of the active recruitment of the RNA polymerase to the preinitiation complex can also affect the level of rRNA synthesis. Third, modulation of the rate of transcription elongation of the RNA polymerase I catalytic activity can influence the amount of rRNA produced at a given time. Although the published work has provided some evidence in favor of one or more of these regulatory processes, the mechanisms that control preinitiation complex formation have been the subject of the largest number of investigations. Overall, these studies have revealed that posttranslational modifications of UBF and SL1 play an important role in the formation of a productive preinitiation complex. In addition, several studies have indicated that global cellular regulators such as the tumor suppressor proteins pRb and p53 can also directly regulate RNA Pol I transcription.
A. Regulation by Posttranslational Modification of Auxiliary Factors 1. Regulation by Phosphorylation UBF, the first cloned RNA polymerase I specific factor, has been the focus of a large number of studies addressing the mechanism of RNA Pol I transcriptional regulation. Analysis of UBF primary amino acids sequence indicates that the carboxy-terminal activation domain is extremely rich in serine. This observation led many investigators to postulate that these
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serine residues might be the targets of phosphorylation by cellular protein kinases. Indeed, metabolic labeling studies of cells in culture have indicated that UBF is heavily phosphorylated under normal growth conditions. Moreover, UBF phosphorylation and RNA polymerase I transcription increase on serum stimulation of quiescent cells (O’Mahony et al., 1992a; Voit et al., 1992). Subsequent studies on the characterization of UBF in serum-starved and serum-stimulated cells provided evidence that UBF is differentially phosphorylated, and this correlated with difference in activity (O’Mahony et al., 1992b). Rapid stimulation of ribosomal RNA synthesis during lymphocytes stimulation by phytohemagglutinin has also been shown to be accompanied by a drastic increase in the level of UBF phosphorylation, indicating a functional relationship between mitogenic stimulation of rRNA synthesis and phosphorylation of this nucleolar factor (Kalousek and Krizkova, 2000). In addition, stimulation of rat vascular smooth muscle by angiotensin II induces a rapid phosphorylation of UBF, which is paralleled by a significant increase in rRNA synthesis (Hershey et al., 1995). Taken together, these findings strongly indicate that phosphorylation plays an important role in the regulation of UBF activity and established the basis for a series of biochemical studies on the identification of the phosphorylation sites and the cellular kinases involved in this process. Casein kinase II (CKII) was the first kinase shown to phosphorylate UBF at multiple sites, primarily located in the carboxy-terminal tail (Voit et al., 1992). However, site-directed mutagenesis indicated that although CKII-mediated phosphorylation of UBF contributes to UBF activity, it is not sufficient to restore full UBF transactivation (Voit et al., 1995). Because the carboxy-terminal activation domain of UBF interacts directly with SL1, and the phosphorylation status of UBF appears to modulate its transcriptional activity, it was important to determine whether UBF phosphorylation/dephosphorylation has an effect on the interaction with SL1. In vitro protein–protein interaction and DNase I footprinting assays showed that the phosporylation status of UBF is critical for allowing the interaction between UBF and SL1, and it plays an important role in the recruitment of SL1 to the CORE and UCE elements of the ribosomal RNA promoter (Tuan et al., 1999). The key role of UBF phosphorylation in modulating the interaction between UBF and SL1 has been demonstrated by other studies. First, mitogen-induced phosphorylation of UBF has been shown to promote its association with TBP, one of the SL1 subunits (Kihm et al., 1998). Then, work on viral regulation of host transcription has pointed out that SV40 large T antigen, a viral oncogene that promotes cell growth, stimulates Pol I transcription by recruiting a cellular kinase to the rDNA promoter that phosphorylates UBF (Zhai and Comai, 1999; Zhai et al., 1997). Importantly, phosphorylation of UBF by a large T
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antigen-associated kinase enhances the ability of UBF to interact with SL1 (Zhai and Comai, 1999). Although these results clearly established that phosphorylation of multiple serine residues within the carboxy-terminal tail of UBF is required for SL1 recruitment, it is not known which specific phosphorylated residues are important for this interaction and which protein kinase or kinases are involved in this process. What it is clear is that phosphorylation of UBF by CKII is not sufficient to reconstitute the interaction between UBF and SL1 (Zhai and Comai, 1999). The importance of serine phosphorylation for UBF activity has further been demonstrated in studies on the mechanism of RNA Pol I transcriptional silencing and reactivation during the transition from mitosis to the G1 phase of the cell cycle (Klein and Grummt, 1999; Voit et al., 1999). UBF is inactive at mitosis, and it is reactivated early during the G1 phase (Klein and Grummt, 1999). Reactivation of UBF requires the phosphorylation of a serine residue at position 484 by the cell cycle–regulated kinases cdk4/ cyclin D1 and ckd2/cyclin E (Voit et al., 1999). Replacement of serine 484 decreases rDNA transcriptional activation several folds, indicating that phosphorylation of this serine residue is required for UBF-mediated activation of RNA Pol I transcription. However, phosphorylation/dephosphorylation of UBF at Ser 484 does not regulate DNA binding nor influence the interaction between UBF and SL1. So the functional significance of this modification remains unknown. In conclusion, although much remains to be learned about the location of many important UBF phosphorylation sites and the functional consequences of these modifications, the picture that emerges is consistent with an essential role for covalent protein modification in the modulation of UBF activity. Changes in the phosphorylation status of the selectivity factor SL1 have also been proposed as a mechanism of regulation of RNA Pol I activity. Studies on the mitotic inactivation of RNA Pol I transcription indicated that the SL1 factor found in mitotic cells is transcriptionally inactive (Kuhn et al., 1998). A more detailed examination revealed that TBP and TAFI110 are phosphorylated during prometaphase in a process that is mediated by the cdc2/cyclin B complex (Heix et al., 1998). Although the relevance of TBP phosphorylation is unclear, phosphorylation of TAFI110 appears to be important for the inactivation of SL1 and inactivation of RNA Pol I transcription. Moreover, protein binding assays indicated that phosphorylation of TAFI110 disrupts the interactions between SL1 and UBF, providing further evidence for the central role that this protein– protein interaction plays in activation of RNA Pol I transcription. In agreement with these findings, it has been found that inhibition of the cdc2/cycB kinase in mammalian cells prevents the shutdown of RNA Pol I
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transcription at mitosis (Sirri et al., 1999). Altogether, these studies emphasize the prominent role that covalent modification of SL1 by a G2specific kinase plays in the repression of RNA Pol I transcription on entry into mitosis. Studies on the regulation of Pol I transcription during cell differentiation have also revealed a potential regulatory pathway involving posttranslational modifications of SL1. Differentiation of the human promyelocytic leukemic cell line U937 is accompanied by a drastic decrease in Pol I transcriptional activity (Comai et al., 2000). Complementation assays and fractionation experiments indicated that SL1 activity is several folds lower in differentiated than undifferentiated U937 cells, indicating that the activity of the TBP/TAFs complex SL1 is severely repressed in differentiated U937 cells. Because abundance of SL1 does not change during differentiation, it is likely that modification of one or more subunits of SL1 may be at the basis of its inactivation during lymphoid cell differentiation.
2. Regulation by Acetylation Acetylation of lysine residues in the carboxy-terminal tail of histones by cellular histone acetyltransferases has been proposed as a major mechanism to modulated nucleosome structure, and many studies have revealed a close relationship between this histone modification and regulation of gene expression. In addition, other studies have indicated that nonhistone proteins can be substrates of acetyltransferases. Transcription factors such as p53, GATA-1, and MyoD can be acetylated in vivo, and biochemical studies indicated that acetylation enhances their DNA binding activity (Bayle and Crabtree, 1997; Boyes et al., 1998; Gu and Roeder, 1997; Sartorelli et al., 1999). On the basis of these findings, it has been speculated that acetyltransferases might also regulate the activity of RNA Pol I transcription factors. Indeed, two recent studies have revealed that UBF and one of the SL1 subunits are acetylated in vivo (Muth et al., 2001; Pelletier et al., 2000). UBF appears to be acetylated at several sites in the HMG box 1 and 2 and within the carboxy-proximal region. Functional studies indicated that acetylation of these sites regulates UBF activity, as acetylated UBF is transcriptionally more active than deacetylated UBF. However, acetylation of UBF does not affect its DNA binding activity, as shown for other transcription factors, and it remains unclear how this posttranslational modification modulates UBF activity. In addition to UBF, SL1 has been shown to be the substrate of cellular acetylases. This study was prompted by the indication that PCAF, a p300/ CBP associated factor with acetyltransferase activity, is recruited to the
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rDNA promoter by interaction with the termination factor TTF-1 (Muth et al., 2001). Analysis of the functional consequence of this recruitment indicated that PCAF specifically acetylates TAFI63 and, to a lesser extent, TAFI48. Although the significance of TAFI48 acetylation remains to be further investigated, the physiological relevance of TAFI63 acetylation has been demonstrated by in vivo labeling experiments, both in human and in mouse cells. Intriguingly, a significant difference is observed in the level of acetylation between human and mouse TAFI63 (human << mouse), possibly underlying a species-specific effect. In vitro analyses indicate that acetylation of TAFI63 by PCAF is likely to increase the activity of SL1, as the addition of increasing amounts of wild-type PCAF, but not of an acetylasedefective mutant PCAF, to in vitro transcription reactions stimulates ribosomal RNA synthesis several folds. Electrophoretic mobility shift assays indicated that TAFI63 acetylation facilitates the interaction of this subunit of SL1 with DNA. Interestingly, PCAF-mediated enhancement of RNA Pol I transcription is eliminated by the addition of Sir2, a factor with NADdependent deacetylase activity and a member of the silent information regulators, a group of proteins implicated in gene silencing. These data indicate that the residues in TAFI63, and possibly TAFI48, which are acetylated by PCAF, are possible substrates of Sir2 deacetylase. In agreement with this hypothesis, the acetyltransferase cofactor NAD is required for efficient inhibition of PCAF-mediated transcriptional activation by Sir2.
B. Regulation by Tumor Suppressor Proteins Tumor suppressor proteins such as the retinoblastoma protein (pRb) and p53 monitor cell proliferation by regulating the activity of a variety of cellular genes involved in growth and cell cycle progression. Because ribosomal RNA transcription is closely linked to cell growth, it is plausible that pRb and p53 may directly modulate rRNA synthesis. Evidence supporting this hypothesis was first obtained in 1995, when pRb was shown to be involved in the repression of RNA Pol I transcription in human cells that are induced to differentiate by the addition of 12-O-tetradecanoylphorbol-13-acetate (TPA) (Cavanaugh et al., 1995). This study indicated that as soon as cells begin to differentiate, there is an accumulation of pRb in the nucleoli and a sharp decrease in rRNA synthesis. Transcription assays and protein interaction studies indicated that pRB binds directly to UBF and inhibits the activity of this transcription factor. Although the inhibitory effect of pRb on RNA polymerase I transcription has been confirmed by another laboratory (Voit et al., 1997), the region of pRb
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that interacts with UBF and the mechanism of inhibition are still contentious. The initial study indicated that UBF binds to a region of pRb named the Rb pocket, which has been shown to mediate the interactions with a number of other cellular and viral proteins (Cavanaugh et al., 1995). However, the second study indicated that the carboxy-terminal domain of pRb is responsible for this molecular interaction (Voit et al., 1997). In addition, although the data from one laboratory indicated that pRb prevents the binding of UBF to DNA (Voit et al., 1997), others have indicated that, by binding to UBF, pRb inhibits the association of this factor with SL1 (Hannan et al., 2000). As mentioned earlier, differentiation of myeloid leukemia cells results also in the inactivation of SL1 (Comai et al., 2000), and this raises the possibility of a functional link between the binding of pRB to UBF and the inactivation of SL1. It is conceivable, for example, that the interaction between UBF and pRb represents one step in a process that eventually leads to the down regulation of SL1 activity and repression of RNA Pol I transcription. In summary, the involvement of pRb in RNA Pol I transcription is consistent with the regulatory role of pRb in cell growth, which is documented for multiple sets of genes transcribed by RNA Pol II and III. Thus, repression of rRNA synthesis by pRb offers an additional level of control in the regulation of cell growth and proliferation. The tumor suppressor p53, an important factor in maintaining genomic stability, has also been involved in the regulation of RNA Pol I transcription. Using cotransfection assays, it has been demonstrated that wild-type, but not a point mutant form of p53 associated with cancer, can repress Pol I transcription from a human rRNA gene promoter (Budde and Grummt, 1999; Zhai and Comai, 2000). Furthermore, recombinant p53 inhibits rRNA transcription in a cell-free transcription system, whereas the mutant p53 fails to do so (Zhai and Comai, 2000). In agreement with these results, p53-null human epithelial cells display increased RNA Pol I transcription as compared with epithelial cells that express p53. Western blot analysis showed that both cell lines have comparable amounts of RNA polymerase and auxiliary factors, indicating that the observed decrease in transcriptional activity is not a consequence of differences in the abundance of these transcription factors (Zhai and Comai, 2000). Further in vitro biochemical analysis showed that p53 binds to SL1 and prevents it from interacting with UBF. Template commitment assays confirmed that the formation of an UBF-SL1 complex can partially relieve the inhibition of transcription. However, only the assembly of a complete initiation complex (UBF/SL1/Pol I) on the rDNA promoter confers substantial protection against p53 inhibition (Zhai and Comai, 2000). Thus, p53 represses RNA Pol I transcription by directly interfering
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with the assembly of a stable transcriptional complex on the rDNA promoter.
C. Regulation of Transcription Elongation Early work with mouse and human cells pointed to variations in the transcription elongation rates as a potential mechanism of regulation of rRNA synthesis. Studies with isolated rat liver that has been starved and refed indicated that rDNA transcription rates reflect changes in RNA Pol I elongation rates rather than changes in the relative number of template-bound RNA polymerases (Coupar and Chesterton, 1975). Similarly, studies with lymphocytes from peripheral blood indicated that activation of rRNA synthesis following stimulation with phytohemagglutinin is related to an increase in RNA polymerase I elongation rates, rather than an increase in the number of transcribed rDNA genes or in the rate of transcription initiation (Dauphinais, 1981). In contrast, more recent studies with starved and serum-stimulated cells(O’Mahony et al., 1992b), and with mitogen-stimulated lymphocytes (Kalousek and Krizkova, 2000), indicated that differences in the abundance and in the phosphorylation level of UBF, two variables that can influence the efficiency of assembly of the preinitiation complex, correlate with enhanced transcriptional activity. New indications for potentially important regulatory steps at the level of transcription elongation came from cell cycle studies. These analyses indicated that nascent rRNA transcripts are released from the nucleosome during metaphase, consistent with a model in which mitotic shutdown of RNA Pol I transcription is associated with an arrest of initiated polymerase within the first 100–200 bp downstream of the rDNA promoter (Weisenberger and Scheer, 1995). However, as discussed earlier, other studies have indicated that repression of RNA Pol I transcription during mitosis is, at least in part, directly linked to the inactivation of transcription initiation factors such as SL1 (Heix et al., 1998; Kuhn et al., 1998). A possible explanation to reconcile these data is that regulatory mechanisms at the level of transcription initiation and elongation operate concurrently to adjust the level of rRNA synthesis to the need of the cell. Notwithstanding the described reports, very little is currently known about the factors regulating elongation by RNA Pol I elongation, and further analysis of this process has lagged behind. To date, a single RNA Pol I elongation factor, TIF-IC, has been identified. TIF-IC purified from mouse extracts stimulates RNA Pol I elongation in vitro (Schnapp et al., 1994b). In addition, it can prevent pausing of the RNA Pol I during the early stages of elongation. The
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nature of this factor and its role in regulation of RNA Pol I transcription elongation await its molecular cloning.
D. Regulation of RNA Polymerase I and Associated Factors Biochemical studies of RNA polymerase I transcription from yeast to mammals have identified at least two forms of RNA polymerase I, only one of which is capable of initiating promoter-specific transcription (Grummt, 1999). In yeast, further analysis has shown that the fraction of cellular polymerase that is active in transcription is associated with the growthregulated factor RRN3 (Milkereit and Tschochner, 1998) and that this form of the polymerase I is efficiently recruited to the preinitiation complex on the rDNA promoter. Interestingly, significant amounts of free RRN3 and RNA polymerase I can be found in stationary phase yeast cells, indicating that the formation of a strong complex between RRN3 and RNA Pol I is regulated in a growth-dependent fashion. The mechanisms that regulate this molecular interaction are currently unknown, but it is likely that modifications of either RRN3 or the RRN3-interacting RNA Pol I subunit play a role in this process. Additional evidence for a role of the Pol I enzyme in regulation of transcription comes from studies on the protozoa Achantamoeba castellanii. These studies indicated that the activity of the RNA Pol I core enzyme is regulated during the transition from a growing amoeba to a dormant cyst (Bateman and Paule, 1986; Paule et al., 1984). On cyst formation, there is a drastic decrease in RNA Pol I activity, and biochemical fractionation of extracts made from the two stages indicated that the decrease in activity is a direct consequence of changes in promoter-specific RNA Pol I activity. RNA Pol I from free-living cells and cysts have the same catalytic activity, but the polymerase from encysting cells is unable to direct promoter-specific transcription. In addition, the polymerase from cysts is unable to bind to DNA. Direct comparison of the subunit composition of the RNA Pol I core enzyme showed a difference in the apparent molecular mass of one subunit, indicating that a posttranslational modification is directly responsible for the altered activity. To date, the nature of this modification remains unknown.
VIII. Chromatin and RNA Polymerase I Transcription The chromatin structure of rDNA genes has been investigated in yeast and rat cells using the intercalating agent psoralen (Conconi et al., 1989; Dammann et al., 1995; Lucchini and Sogo, 1995). These studies have determined that rDNA genes are found either in densely packed nucleosomes or completely free of nucleosomes. Nucleosomal-packed genes are
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transcriptionally inactive, whereas nucleosome-free ribosomal DNA genes are actively transcribed. Interestingly, both conformations can be found within the same ribosomal rDNA locus, indicating that active copies of rDNA genes are randomly distributed along the locus. The transition from a nucleosome-packed to a nucleosome-free chromatin structure is probably related to the recruitment to the rDNA promoter and enhancer region of factors that disrupt the nucleosome arrays. In vitro studies have identified at least two factors that may participate in this process: the UBF and the termination factor TTF-I/Reb-1 (Kermekchiev et al., 1997; Kuhn and Grummt, 1992). In vitro, UBF can counteract the negative effect of histone H1 on RNA Pol I transcription. In addition, biochemical analysis shows that UBF can displace histone H1 from an in vitro assembled nucleosome. On the basis of these data, UBF may modify the chromatin structure to facilitate the transition from silent to potentially active and transcribed genes. The second factor thought to be involved in the remodeling of chromatin on rDNA genes is TTF-I/Reb1 (Mason et al., 1997). TTF-I/Reb1 is a transcription termination factor that binds to the terminator element immediately downstream from the rDNA gene and, in cooperation with the release factor PTRF, induces transcription termination and causes the dissociation of the elongating complex from the DNA template (Mason et al., 1997; Jansa and Grummt, 1999; Jansa et al., 1998). The presence of this terminator element just upstream of the rDNA promoter led investigators to propose that TTF-I/Reb1 may also be involved in local chromatin disruption (Lucchini and Sogo, 1995). This hypothesis prompted biochemical studies with in vitro reconstituted chromatin, which showed that TTF-I activates transcription from nucleosome templates in vitro by recruiting ATP-dependent remodeling factors to the rDNA promoter (Langst et al., 1997). More recently, the nucleosome remodeling factor that is recruited to the rDNA promoter by TTF-I has been purified and shown to be a member of the ISWI-chromatin remodeling complexes family (Strohner et al., 2001). This factor, named nucleolar remodeling complex, or NoRC, is composed of at least four subunits: TIP5, which interacts directly with TTF-I; SNF2h, the mammalian homologue of Drosophila ISWI; and two still undefined proteins. Experiments with in vitro reconstituted chromatin indicate that purified NoRC, as with other nucleosome remodeling complexes (Flaus and Owen-Hughes, 2001), facilitates the sliding of nucleosomes on DNA in an ATP-dependent manner. Taken together, these studies emphasize the important role of TTF-I and the recruitment of nucleosome remodeling factors to the rDNA locus. It remains to be established, however, whether NoRC has a positive or negative effect on rDNA gene expression. Moreover, it is currently unclear whether modifications of histone tails by histone acetyltranferases
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are also involved in chromatin disruption and activation of rDNA gene transcription. A report has indicated that a histone acetyltransferase activity copurifies with Xenopus RNA polymerase I holoenzyme (Albert et al., 1999). However, the functional significance of this finding remains to be determined.
IX. Conclusions In the last few years the research efforts of many laboratories have dissected the plethora of molecular interactions between DNA and proteins that regulate RNA Pol I, demonstrating their essential role in the establishment of a functional and promoter-specific transcription complex. An important conclusion from these studies is that de novo synthesis of transcription factors plays a secondary role in the regulation of rRNA synthesis in eukaryotes. Rather, covalent modification of preexisting factors mediates the assembly of a stable transcription initiation complex and regulates the efficiency of transcription by RNA Pol I. An important implication of the finding that phosphorylation of UBF and SL1 regulates transcription is that signaling molecules are likely to have a significant influence on rRNA synthesis. Therefore, one of the next challenges is to identify the signal transduction pathways that modulate RNA Pol I transcription and to characterize how diverse signals converge and cross talk on the RNA Pol I transcriptional machinery.
Acknowledgments I thank the members of my laboratory for helpful discussion and Dr. Luca Comai for critical reading of the manuscript. Research in my laboratory is supported by grants from the National Institute of Health and the American Cancer Society.
References Albert, A. C., Denton, M., Kermekchiev, M., and Pikaard, C. S. (1999). Mol. Cell. Biol. 19, 796–806. Aprikian, P., Moorefield, B., and Reeder, R. H. (2000). Mol. Cell. Biol. 20, 5269–5275. Aprikian, P., Moorefield, B., and Reeder, R. H. (2001). Mol. Cell. Biol. 21, 4847–4855. Bachvarov, D., and Moss, T. (1991). Nucleic Acids Res. 19, 2331–2335. Baker, S. M., and Platt, T. (1986). Cell 47, 839–840. Bateman, E., and Paule, M. R. (1986). Cell 47, 445–450. Bayle, J. H., and Crabtree, G. R. (1997). Chem. Biol. 4, 885–888. Bazett-Jones, D. P., Leblanc, B., Herfort, M., and Moss, T. (1994). Science 264, 1134–1137. Beckmann, H., Chen, J. L., O’Brien, T., and Tjian, R. (1995). Science 270, 1506–1509.
150
COMAI
Bell, S. P., Jantzen, H. M., and Tjian, R. (1990). Genes Dev. 4, 943–954. Bell, S. P., Learned, R. M., Jantzen, H. M., and Tjian, R. (1988). Science 241, 1192–1197. Bell, P., Mais, C., McStay, B., and Scheer, U. (1997). J. Cell Sci. 110, 2053–2063. Bell, S. P., Pikaard, C. S., Reeder, R. H., and Tjian, R. (1989). Cell 59, 489–497. Birniestiel, M., Speirs, J., Purdom, I., Jones, K., and Loening, U. E. (1968). Nature 219, 454–459. Bodem, J., Dobreva, G., Hoffmann-Rohrer, U., Iben, S., Zentgraf, H., Delius, H., Vingron, M., and Grummt, I. (2000). EMBO Rep. 1, 171–175. Boyes, J., Byfield, P., Nakatani, Y., and Ogryzko, V. (1998). Nature 396, 594–598. Brun, R. P., Ryan, K., and Sollner-Webb, B. (1994). Mol. Cell. Biol. 14, 5010–5021. Budde, A., and Grummt, I. (1999). Oncogene 18, 1119–1124. Buttgereit, D., Pflugfelder, G., and Grummt, I. (1985). Nucleic Acids Res. 13, 8165–8180. Cavanaugh, A. H., Gokal, P. K., Lawther, R. P., and Thompson, E. A. J. (1984). Proc. Natl. Acad. Sci. USA 81, 718–721. Cavanaugh, A. H., Hempel, W. M., Taylor, L. J., Rogalsky, V., Todorov, G., and Rothblum, L. I. (1995). Nature 374, 177–180. Cavanaugh, A. H., and Thompson, E. A. J. (1985). Nucelic Acids Res. 13, 3357–3369. Cavanaugh, A. H., and Thompson, E. A. (1986). J. Biol. Chem. 261, 12738–12744. Chen, Z. J., Comai, L., and Pikaard, C. S. (1998). Proc. Natl. Acad. Sci. USA 95, 14891–14896. Chen, Z. J., and Pikaard, C. S. (1997). Proc. Natl. Acad. Sci. USA 94, 3442–3447. Clos, J., Buttgereit, D., and Grummt, I. (1986a). Proc. Natl. Acad. Sci. USA 83, 604–608. Clos, J., Normann, A., Ohrlein, A., and Grummt, I. (1986b). Nucleic Acids Res. 14, 7581–7595. Comai, L., Tanese, N., and Tjian, R. (1992). Cell 68, 965–976. Comai, L., Song, Y., Tan, C., and Bui, T. (2000). Cell Growth Differ. 11, 63–70. Comai, L., Zomerdijk, J. C., Beckmann, H., Zhou, S., Admon, A., and Tjian, R. (1994). Science 266, 1966–1972. Conconi, A., Widmer, R. M., Koller, T., and Sogo, J. M. (1989). Cell 57, 753–761. Copenhaver, G. P., and Pikaard, C. S. (1996). Plant J. 9, 259–272. Copenhaver, G. P., Putnam, C. D., Denton, M. L., and Pikaard, C. S. (1994). Nucleic Acids Res. 22, 2651–2657. Coupar, B. E. H., and Chesterton, C. J. (1975). Eur. J. Biochem. 59, 25–30. Dammann, R., Lucchini, R., Koller, T., and Sogo, J. M. (1995). Mol. Cell. Biol. 15, 5294–5303. Dammann, R., and Pfeifer, G. P. (1998). Biochem. Biophys. Acta 1396, 153–157. Dauphinais, C. (1981). Eur. J. Biochem. 114, 487–492. Derenzini, M., and Trere, D. (1991). Virchows Arch. B Cell Pathol. Incl. Mol. Pathol. 61, 1–8. Derenzini, M., Trere, D., Pession, A., Govoni, M., Sirri, V., and Chieco, P. (2000). J. Pathol. 191, 181–186. Derenzini, M., Trere, D., Pession, A., Montanaro, L., Sirri, V., and Ochs, R. L. (1998). Am. J. Pathol. 152, 1291–1297. Eberhard, D., Tora, L., Egly, J. M., and Grummt, I. (1993). Nucleic Acids Res. 21, 4180–4186. Financsek, I., Mizumoto, K., Mishima, Y., and Muramatsu, M. (1982). Proc. Natl. Acad. Sci. USA 79, 3092–3096. Flaus, A., and Owen-Hughes, T. (2001). Curr. Opin. Genet. Dev. 11, 148–154. Garg, L. C., DiAngelo, S., and Jacob, S. T. (1987). Proc. Natl. Acad. Sci. USA 84, 3185–3188.
MECHANISM OF RNA POLYMERASE I TRANSCRIPTION
151
Gebrane-Younes, J., Fomproix, N., and Hernandez-Verdun, D. (1997). J. Cell Sci. 110, 2429–2440. Ghosh, A. K., Datta, P. K., and Jacob, S. T. (1997). Oncogene 14, 589–594. Ghosh, A. K., Hoff, C. M., and Jacob, S. T. (1993). Gene 125, 217–222. Gokal, P. K., Mahajan, P. B., and Thompson, E. A. (1990). J. Biol. Chem. 265, 16234–16243. Grosschedl, R., Giese, K., and Pagel, J. (1994). Trends Genet. 10, 94–100. Grummt, I., Smith, V. A., and Grummt, F. (1976). Cell 7, 439–445. Grummt, I. (1981a). Nucleic Acids Res. 9, 6093–6102. Grummt, I. (1981b). Proc. Natl. Acad. Sci. USA 78, 727–731. Grummt, I. (1999). Prog. Nucleic. Acid Res. Mol. Biol. 62, 109–154. Grummt, I., Kuhn, A., Bartsch, I., and Rosenbauer, H. (1986). Cell 47, 901–911. Gu, W., and Roeder, R. G. (1997). Cell 90, 595–606. Hadjiolov, A. A. (1985). The nucleolus and ribosome biogenesis, in Cell Biology Monographs, Vol. 12. New York: Springer. Haglund, R. E., and Rothblum, L. I. (1987). Mol. Cell. Biochem. 73, 11–20. Haltiner, M. M., Smale, S. T., and Tjian, R. (1986). Mol. Cell. Biol. 6, 227–235. Hanada, K., Song, C. Z., Yamamoto, K., Yano, K., Maeda, Y., Yamaguchi, K., and Muramatsu, M. (1996). EMBO J. 15, 2217–2226. Hannan, R. D., Cavanaugh, A., Hempel, W. M., Moss, T., and Rothblum, L. (1999). Nucleic Acids Res. 27, 3720–3727. Hannan, K. M., Hannan, R. D., and Rothblum, L. I. (1998a). Front. Biosci. 3, d376–d398. Hannan, K. M., Hannan, R. D., Smith, S. D., Jefferson, L. S., Lun, M., and Rothblum, L. I. (2000). Oncogene 19, 4988–4999. Hannan, R. D., Hempel, W. M., Cavanaugh, A., Arino, T., Dimitrov, S. I., Moss, T., and Rothblum, L. (1998b). J. Biol. Chem. 273, 1257–1267. Hannan, K. M., Rothblum, L. I., and Jefferson, L. S. (1998c). Am. J. Physiol. 275, C130–C138. Heilgenthal, G., and Grummt, I. (1991). J. Chromatogr. 587, 25–32. Heix, J., Vente, A., Voit, R., Budde, A., Michaelidis, T. M., and Grummt, I. (1998). EMBO J. 17, 7373–7381. Heix, J., Zomerdijk, J. C., Ravanpay, A., Tjian, R., and Grummt, I. (1997). Proc. Natl. Acad. Sci. USA 94, 1733–1738. Henderson, S. L., Ryan, K., and Sollner-Webb, B. (1989). Genes Dev. 3, 212–223. Henderson, S., and Sollner-Webb, B. (1986). Cell 47, 891–900. Hernandez, N. (1993). Genes Dev. 7, 1291–1308. Hershey, J. C., Hautmann, M., Thompson, M. M., Rothblum, L. I., Haystead, T. A., and Owens, G. K. (1995). J. Biol. Chem. 270, 25096–25101. Hoff, C. M., Ghosh, A. K., Prabhakar, B. S., and Jacob, S. T. (1994). Proc. Natl. Acad. Sci. USA 91, 762–766. Jansa, P., and Grummt, I. (1999). Mol. Gen. Genet. 262, 508–514. Jansa, P., Mason, S. W., Hoffmann-Rohrer, U., and Grummt, I. (1998). EMBO J. 17, 2855–2864. Jantzen, H.-M., Admon, A., Bell, S. P., and Tjian, R. (1990). Nature 344, 830–836. Jantzen, H.-M., Chow, A. M., King, D. S., and Tjian, R. (1992). Genes Dev. 6, 1950–1963. Jones, M. H., Learned, R. M., and Tjian, R. (1988). Proc. Natl. Acad. Sci. USA 85, 669–673. Junera, H. R., Masson, C., Geraud, G., Suja, J., and Hernandez-Verdun, D. (1997). Mol. Biol. Cell 8, 145–156. Kalousek, I., and Krizkova, P. (2000). Cell. Mol. Biol. (Noisy-le-grand) 46, 1163–1171.
152
COMAI
Kermekchiev, M., Workman, J. L., and Pikaard, C. S. (1997). Mol. Cell. Biol. 17, 5833–5842. Keys, D. A., Vu, L., Steffan, J. S., Dodd, J. A., Yamamoto, R. T., Nogi, Y., and Nomura, M. (1994). Genes Dev. 8, 2349–2362. Keys, D. A., Lee, B. S., Dodd, J. A., Nguyen, T. T., Vu, L., Fantino, E., Burson, L. M., Nogi, Y., and Nomura, M. (1996). Genes Dev. 10, 887–903. Kihm, A. J., Hershey, J. C., Haystead, T. A., Madsen, C. S., and Owens, G. K. (1998). Proc. Natl. Acad. Sci. USA 95, 14816–14820. Klein, J., and Grummt, I. (1999). Proc. Natl. Acad. Sci. USA 96, 6096–6101. Kuhn, A., Gottlieb, T. M., Jackson, S. P., and Grummt, I. (1995). Genes Dev. 9, 193–203. Kuhn, A., and Grummt, I. (1987). EMBO J. 6, 3487–3492. Kuhn, A., and Grummt, I. (1992). Proc. Natl. Acad. Sci. USA 89, 7340–7344. Kuhn, A., Stefanovsky, V., and Grummt, I. (1993). Nucleic Acids Res. 21, 2057–2063. Kuhn, A., Vente, A., Doree, M., and Grummt, I. (1998). J. Mol. Biol. 284, 1–5. Kuhn, A., Voit, R., Stefanovsky, V., Evers, R., Bianchi, M., and Grummt, I. (1994). EMBO J. 13, 416–424. Lalo, D., Steffan, J. S., Dodd, J. A., and Nomura, M. (1996). J. Biol. Chem. 271, 21062–21067. Langst, G., Blank, T. A., Becker, P. B., and Grummt, I. (1997). EMBO J. 16, 760–778. Larson, D. E., Zahradka, P., and Sells, B. H. (1991). Biochem. Cell. Biol. 69, 5–22. Learned, R. M., Cordes, S., and Tjian, R. (1985). Mol. Cell. Biol. 5, 1358–1369. Learned, R. M., Learned, T. K., and Tjian, M. M. H. a. R. (1986). Cell 45, 847–857. Learned, R. M., Smale, S. T., Haltiner, M. M., and Tjian, R. (1983). Proc. Natl. Acad. Sci. USA 80, 3558–3562. Learned, R. M., and Tjian, R. (1982). J. Mol. Appl. Genet. 1, 575–584. Lin, C. W., Moorefield, B., Payne, J., Aprikian, P., Mitomo, K., and Reeder, R. H. (1996). Mol. Cell. Biol. 16, 6436–6443. Liu, Z., and Jacob, S. T. (1994). J. Biol. Chem. 269, 16618–16625. Long, E. O., and Dawid, I. B. (1980). Ann. Rev. Biochem. 49, 727–764. Lucchini, R., and Sogo, J. M. (1995). Nature 374, 276–280. Maeda, Y., Hisatake, K., Kondo, T., Hanada, K., Song, C. Z., Nishimura, T., and Muramatsu, M. (1992). EMBO J. 11, 3695–3704. Mahajan, P. B., Gokal, P. K., and Thompson, E. A. (1990). J. Biol. Chem. 265, 16244–16247. Mahajan, P. B., and Thompson, E. A. (1990). J. Biol. Chem. 265, 16225–16233. Mason, S. W., Wallisch, M., and Grummt, I. (1997). J. Mol. Biol. 268, 229–234. McStay, B., Frazier, M. W., and Reeder, R. H. (1991A). Genes Dev. 5, 1957–1968. McStay, B., Hu, C. H., Pikaard, C. S., and Reeder, R. H. (1991B). EMBO J. 10, 2297–2303. McStay, B., Sullivan, G. J., and Cairns, C. (1997). EMBO J. 16, 396–405. Miesfeld, R., and Arnheim, N. (1984). Mol. Cell. Biol. 4, 221–227. Miesfeld, R., Sollner-Webb, B., Croce, C., and Arnheim, N. (1984). Mol. Cell. Biol. 4, 1306–1312. Milkereit, P., and Tschochner, H. (1998). EMBO J. 17, 3692–3703. Miller, G., Panov, K. I., Friedrich, J. K., Trinkle-Mulcahy, L., Lamond, A. I., and Zomerdijk, J. C. (2001). EMBO J. 20, 1373–1382. Miller, K. G., Tower, J., and Sollner-Webb, B. (1985). Mol. Cell. Biol. 5, 554–562. Mishima, Y., Financsek, I., Kominami, R., and Muramatsu, M. (1982). Nucleic Acids Res. 10, 6659–6670.
MECHANISM OF RNA POLYMERASE I TRANSCRIPTION
153
Mishima, Y., Yamamoto, O., Kominami, R., and Muramatsu, M. (1981). Nucleic Acids Res. 9, 6773–6785. Moorefield, B., Greene, E. A., and Reeder, R. H. (2000). Proc. Natl. Acad. Sci. USA 97, 4724–4729. Morrow, J., Cohen, S., Chang, A., Boyer, H., Goodman, H., and Hellings, R. (1974). Proc. Natl. Acad. Sci. USA 71, 1743–1747. Moss, T. (1983). Nature 302, 223–228. Moss, T., and Stefanovsky, V. Y. (1995). Prog. Nucleic. Acid Res. Mol. Biol. 50, 25–66. Muth, V., Nadaud, S., Grummt, I., and Voit, R. (2001). EMBO J. 20, 1353–1362. Nagamine, M., Kishimoto, T., Aono, J., Kato, H., Kominami, R., and Muramatsu, M. (1987). Mol. Cell. Biol. 7, 1486–1495. Niu, H., Zhang, J., and Jacob, S. T. (1995). Gene Expr. 4, 111–124. Nurse, P. (1985). The genetic control of cell volume. In ‘‘The Evolution of Genome Size,’’ (T. Cavalier-Smith, Ed.), pp. 185–196. John Wiley and Sons, Chichester, U.K. O’Mahony, D. J., and Rothblum, L. I. (1991). Proc. Natl. Acad. Sci. USA 88, 3180–3184. O’Mahony, D. J., Smith, S. D., Xie, W., and Rothblum, L. I. (1992a). Nucleic Acids Res. 20, 1301–1308. O’Mahony, D. J., Xie, W. Q., Smith, S. D., Singer, H. A., and Rothblum, L. I. (1992b). J. Biol. Chem. 267, 35–38. Osheim, Y. N., Mougey, E. B., Windle, J., Anderson, M., O’Reilly, M., Miller, O. L., Jr., Beyer, A., and Sollner-Webb, B. (1996). J. Cell. Biol. 133, 943–954. Panov, K. I., Friedrich, J. K., and Zomerdijk, J. C. (2001). Mol. Cell Biol. 21, 2641–2649. Pape, L. K., Windle, J. J., and Sollner-Webb, B. (1990). Genes Dev. 4, 52–62. Paule, M. (1998). Transcription of ribosomal genes by eukaryotic RNA polymerase I. Austin, TX; Landes Bioscience. Paule, M. R., Iida, C. T., Perna, P. J., Harris, G. H., Knoll, D. A., and D’Alessio, J. M. (1984). Nucleic Acids Res. 12, 8161–8180. Pelletier, G., Stefanovsky, V. Y., Faubladier, M., Hirschler-Laszkiewicz, I. I., Savard, J., Rothblum, L. I., Cote, J., and Moss, T. (2000). Mol. Cell 6, 1059–1066. Peyroche, G., Milkereit, P., Bischler, N., Tschochner, H., Schultz, P., Sentenac, A., Carles, C., and Riva, M. (2000). EMBO J. 19, 5473–5482. Pikaard, C. S., McStay, B., Schultz, M. C., Bell, S. P., and Reeder, R. H. (1989). Genes Dev. 3, 1779–1788. Pikaard, C. S., Smith, S. D., Reeder, R. H., and Rothblum, L. (1990). Mol. Cell. Biol. 10, 3810–3812. Putnam, C. D., Copenhaver, G. P., Denton, M. L., and Pikaard, C. S. (1994). Mol. Cell. Biol. 14, 6476–6488. Reeder, R. H. (1984). Cell 38, 349–351. Reeder, R. H. (1985). J. Cell. Biol. 101, 2013–2016. Reeder, R. H. (1999). Prog. Nucleic. Acid Res. Mol. Biol. 62, 293–327. Reeder, R. H., Pennock, D., McStay, B., Roan, J., Tolentino, E., and Walker, P. (1987). Nucleic Acids Res. 15, 7429–7441. Rigby, P. W. (1993). Cell 72, 7–10. Riggs, D. L., and Nomura, M. (1990). J. Biol. Chem. 265, 7596–7603. Saez-Vasquez, J., and Pikaard, C. S. (1997). Proc. Natl. Acad. Sci. USA 94, 11869–11874. Safrany, G., Tanaka, N., Kishimoto, T., Ishikawa, Y., Kato, H., Kominami, R., and Muramatsu, M. (1989). Mol. Cell. Biol. 9, 349–353. Sartorelli, V., Puri, P. L., Hamamori, Y., Ogryzko, V., Chung, G., Nakatani, Y., Wang, J. Y., and Kedes, L. (1999). Mol. Cell 4, 725–734.
154
COMAI
Scheer, U., and Hock, R. (1999). Curr. Opin. Cell. Biol. 11, 385–390. Schnapp, A., Clos, J., Hadelt, W., Schreck, R., Cvekl, A., and Grummt, I. (1990). Nucleic Acids Res. 18, 1385–1393. Schnapp, A., and Grummt, I. (1991). J. Biol. Chem. 266, 24588–24595. Schnapp, A., Rosenbauer, H., and Grummt, I. (1991). Mol. Cell. Biochem. 104, 137–147. Schnapp, A., Schnapp, G., Erny, B., and Grummt, I. (1993). Mol. Cell. Biol. 13, 6723–6732. Schnapp, G., Santori, F., Carles, C., Riva, M., and Grummt, I. (1994a). EMBO J. 13, 190–199. Schnapp, G., Schnapp, A., Rosenbauer, H., and Grummt, I. (1994b). EMBO J. 13, 4028–4035. Schultz, M. C., Choe, S. Y., and Reeder, R. H. (1991). Proc. Natl. Acad. Sci. USA 88, 1004–1008. Schultz, M. C., Brill, S. J., Ju, Q., Sternglanz, R., and Reeder, R. H. (1992). Genes Dev. 6, 1332–1341. Seither, P., Coy, J. F., Pouska, A., and Grummt, I. (1997). Mol. Gen. Genet. 255, 180–186. Seither, P., and Grummt, I. (1996a). Genomics 37, 135–139. Seither, P., Iben, S., and Grummt, I. (1998). J. Mol. Biol. 275, 43–53. Seither, P., Zatsepina, O., Hoffmann, M., and Grummt, I. (1997b). Chromosoma 106, 216–225. Sentenac, A. (1985). CRC Crit. Rev. Biochem. 18, 31–90. Shaw, P. J., and Jordan, E. G. (1995). Annu Rev. Cell. Dev. Biol. 11, 93–121. Shpakovski, G. V., Acker, J., Wintzerith, M., Lacroix, J. F., Thuriaux, P., and Vigneron, M. (1995). Mol. Cell. Biol. 15, 4702–4710. Siddiqi, I. N., Dodd, J. A., Vu, L., Eliason, K., Oakes, M. L., Keener, J., Moore, R., Young, M. K., and Nomura, M. (2001a). EMBO J. 20, 4512–4521. Siddiqi, I., Keener, J., Vu, L., and Nomura, M. (2001b). Mol. Cell. Biol. 21, 2292–2297. Sirri, V., Roussel, P., and Hernandez-Verdun, D. (1999). J. Cell. Sci. 112, 3259–3268. Song, C. Z., Hanada, K., Yano, K., Maeda, Y., Yamamoto, K., and Muramatsu, M. (1994). J. Biol. Chem. 269, 26976–26981. Steffan, J. S., Keys, D. A., Dodd, J. A., and Nomura, M. (1996). Genes Dev. 10, 2551–2563. Strohner, R., Nemeth, A., Jansa, P., Hofmann-Rohrer, U., Santoro, R., Langst, G., and Grummt, I. (2001). EMBO J. 20, 4892–4900. Sullivan, G. J., Bridger, J. M., Cuthbert, A. P., Newbold, R. F., Bickmore, W. A., and McStay, B. (2001). EMBO J. 20, 2867–2874. Sullivan, G. J., and McStay, B. (1998). Nucleic Acids Res. 26, 3555–3561. Tian, Q., Kopf, G. S., Brown, R. S., and Tseng, H. (2001). Development 128, 407–416. Tseng, H., Biegel, J. A., and Brown, R. S. (1999). J. Cell. Sci. 112, 3039–3047. Tuan, J. C., Zhai, W., and Comai, L. (1999). Mol. Cell. Biol. 19, 2872–2879. Voit, R., Hoffmann, M., and Grummt, I. (1999). EMBO J. 18, 1891–1899. Voit, R., Kuhn, A., Sander, E. E., and Grummt, I. (1995). Nucleic Acids Res. 23, 2593–2599. Voit, R., Schafer, K., and Grummt, I. (1997). Mol. Cell. Biol. 17, 4230–4237. Voit, R., Schnapp, A., Kuhn, A., Rosenbauer, H., Hirschmann, P., Stunnenberg, H. G., and Grummt, I. (1992). EMBO J. 11, 2211–2218. Wai, H., Johzuka, K., Vu, L., Eliason, K., Kobayashi, T., Horiuchi, T., and Nomura, M. (2001). Mol. Cell. Biol. 21, 5541–5553. Weisenberger, D., and Scheer, U. (1995). J. Cell. Biol. 129, 561–5675. Wilkinson, J. K., and Sollner-Webb, B. (1982). J. Biol. Chem. 257, 14375–14383.
MECHANISM OF RNA POLYMERASE I TRANSCRIPTION
155
Yamamoto, R. T., Nogi, Y., Dodd, J. A., and Nomura, M. (1996). EMBO J. 15, 3964–3973. Zatsepina, O. V., Voit, R., Grummt, I., Spring, H., Semenov, M. V., and Trendelenburg, M. F. (1993). Chromosoma 102, 599–611. Zhai, W., Tuan, J. A., and Comai, L. (1997). Genes Dev. 11, 1605–1617. Zhai, W., and Comai, L. (1999). Mol. Cell. Biol. 19, 2791–2802. Zhai, W., and Comai, L. (2000). Mol. Cell. Biol. 20, 5930–5938. Zhang, H., Wang, J. C., and Liu, L. F. (1988). Proc. Natl. Acad. Sci. USA 85, 1060–1064. Zhang, J., and Jacob, S. T. (1990). Mol. Cell. Biol. 10, 5177–5186. Zomerdijk, J. C., Beckmann, H., Comai, L., and Tjian, R. (1994). Science 266, 2015–2018.
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FUNCTIONAL PROPERTIES OF ATP-DEPENDENT CHROMATIN REMODELING ENZYMES By ANTHONY N. IMBALZANO* AND HENGYI XIAO Department of Cell Biology, University of Massachusetts Medical School, Worcester, Massachusetts 01655
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. History of Nucleosome-Remodeling Complexes . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Isolation and Characterization of SWI and SNF Genes . . . . . . . . . . . . . . . . . . .. . . . . . B. Multisubunit Complexes that Alter Nucleosome Structure . . . . . . . . . . . . . .. . . . . . III. Requirements for Nucleosome-Remodeling Enzymes.. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Interaction with DNA/Chromatin. . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. ATP Binding and Hydrolysis . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Cofactor Requirements . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Nucleosome Remodeling is an Enzymatic Process . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Mechanisms of ATP-Dependent Chromatin Remodeling . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Nucleosome Sliding . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Nucleosome Disruption. . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Formation of an Altered Dinucleosome-Like Species . . . . . . . . . . . . . . . . . . . . .. . . . . . D. Octamer Transfer in trans. . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . V. Initiation of ATP-Dependent Chromatin Remodeling . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Spooling or Peeling Model. .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Twisting Model . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Bulging or Looping Model . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . VI. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Acknowledgments. . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction In eukaryotic cells, DNA is packaged into a highly compacted and condensed nucleoprotein structure called chromatin. Biochemical studies and electron microscopy indicated that DNA in eukaryotic chromatin is folded as regular units, each of which contains 146 base pairs of DNA and a core of histone proteins. Structurally, DNA makes approximately 1.8 turns around a central histone octamer that consists of two molecules of each of the four core histones: H2A, H2B, H3, and H4. The combination of a histone core and associated DNA makes up the nucleosome. Nucleosomes are linked by 20–100 base pairs (bp) of linker DNA, so as to form a beadlike nucleosomal array. In conjunction with the linker histone, H1, *Correspondence to: Anthony N. Imbalzano, Department of Cell Biology, University of Massachusetts Medical School, 55 Lake Avenue North, Worcester, MA 01655 (e-mail
[email protected]). 157 ADVANCES IN PROTEIN CHEMISTRY, Vol. 67
Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00
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which is present stoichiometrically with histone octamers, nucleosomal arrays fold into higher-order structures that are only just beginning to be structurally characterized in detail. The packaging of DNA into nucleosomes and higher-order chromatin structures presents cells with a significant barrier to DNA utilization and necessitates mechanisms by which the chromatin structure can be modified and the disparate nuclear processes that require access to DNA sequence, including DNA replication, repair, recombination, and RNA transcription, can occur and be regulated. Although the mechanisms concerned with modification of higher-order chromatin structure are poorly understood, it has become widely accepted that altering the structure of, or remodeling, nucleosomes is an important regulatory step for most of the nuclear DNA transactions. In general, nucleosome structure can be remodeled in two ways: via covalent modifications, including acetylation, phosphorylation, ubiquitination, ADP-ribosylation, and methylation, that predominantly occur at the histone terminal tails, and via direct, energy-dependent structural alterations that may change the nucleosome position with respect to the DNA sequence or, not on the path of the DNA around the nucleosome or that may displace histone subunits. Like most cellular reactions, histone modifications and energy-dependent nucleosome structural alterations require enzymatic proteins. Two classes of highly conserved nucleosome remodeling enzymes, also refered to as chromatin remodeling enzymes, have been identified and have been the subject of intense study in recent years. The first class includes enzymes that covalently modify the nucleosomal histones, such as histone acetyltransferases (HATs) and deacetylases (HDACs), which, respectively, add and remove acetyl groups from the amino termini of the four core histones. The second class of enzymes are those that use the energy of ATP hydrolysis to alter or disrupt nucleosome structure. These ATP-dependent chromatin remodeling enzymes are multiprotein complexes that alter nucleosome structure by affecting histone–DNA interactions, thereby changing the location or conformation of the nucleosome. In this chapter, we focus our discussion on the ATP-dependent chromatin-remodeling enzymes. A number of excellent recent reviews examining enzymes that posttranslationally modify the histone proteins are available (Berger, 2002; Roth et al., 2001; Turner, 2002).
II. History of Nucleosome-Remodeling Complexes A. Isolation and Characterization of SWI and SNF Genes The yeast SWI (switch) and SNF (sucrose non-fermenting) genes were isolated in genetic screens searching for genes affecting expression of the HO endonuclease that is required for mating-type switching and for genes
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required for expression of the SUC2 gene, which encodes an enzyme required for sucrose utilization (Breeden and Nasmyth, 1987; Neigeborn and Carlson, 1984; Stern et al., 1984). Several of the identified SNF genes subsequently were shown to be required for activation of reporter gene expression by synthetic activator fusion proteins in yeast (Laurent and Carlson, 1992; Laurent et al., 1990, 1991). In addition, the SNF gene products themselves could activate reporter gene transcription when fused to a heterologous DNA binding domain (Laurent et al., 1990, 1991). Similarly, multiple SWI genes were demonstrated to be required for optimal expression of several inducible yeast loci, as well as for the transcriptional competence of mammalian nuclear hormone receptors introduced into yeast (Yoshinaga et al., 1992). The conclusion from these studies was that SWI and SNF gene products played a general role in activator-mediated transcription, perhaps by acting as coactivators necessary for optimizing the activity of sequence-specific DNA binding activator proteins. Genetic analyses of the SWI and SNF alleles indicated that deletion of or mutations in any one of the alleles generated phenotypes similar to those resulting from deletion or mutation of multiple alleles (Laurent and Carlson, 1992; Laurent et al., 1991; Peterson and Herskowitz, 1992). These studies indicated that the gene products functioned in the same pathway, perhaps as part of a multisubunit complex. The discovery of the identity of the SWI2 and SNF2 alleles and the presence of a functional ATPase domain within the coding sequence that showed significant similarity to known DNA-stimulated ATPases (Laurent et al., 1993; Peterson and Herskowitz, 1992) provided an unexpected twist to the putative coactivator function of the SWI and SNF gene products. Concurrent with this body of work were efforts to further understand SWI gene function through identification of suppressor alleles that would rescue SWI gene mutants. Suppressor mutations included mutations in the coding sequence of core histone proteins, mutations in the promoters of core histones that altered expression levels and thereby histone stoichiometry between octamer subunits, and mutations in SIN1, a nonhistone chromatin protein with similarity to mammalian HMG1 (Hirschhorn et al., 1995; Kruger and Herskowitz, 1991; Kruger et al., 1995). Thus, the general coactivator function of the SWI and SNF genes’ products was linked to chromatin structure, as the suppressor studies argued that the normal function of SWI and SNF proteins was to alter chromatin structure. A second strong link between SWI and SNF proteins and chromatin structure came from promoter analyses of SWI/SNF dependent genes. Increases in in vivo nuclease sensitivity at the promoters correlated with activation of gene expression; in SWI and SNF mutant strains, the lack of
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gene expression correlated with a lack of increased nuclease sensitivity (Hirshhorn et al., 1992; Matallana et al., 1992). The Drosophila Brahma gene was isolated in a screen for suppressors of polycomb group (Pc-G) mutations (Tamkun et al., 1992). The gene showed high levels of similarity with the yeast SWI2/SNF2 gene, indicating that at least some of the SWI and SNF genes were evolutionarily conserved. Because Pc-G proteins are thought to maintain DNA in an inactive state, rescue of Pc-G mutations by Brahma mutations implicated Brahma in antagonizing stable chromatin structure (Tamkun et al., 1992). Thus a connection between SWI and SNF gene products and chromatin structure was evident in higher eukaryotes. Subsequent efforts indicated that SWI2/ SNF2 homologues existed in mammalian cells as well. Two highly related but distinct genes, human Brahma (hBRM) and Brahma-related gene 1 (BRG1) were isolated and shown to enhance activation by mammalian steroid hormone receptors (Chiba et al., 1994; Khavari et al., 1993; Muchardt and Yaniv, 1993). Importantly, mutations in the ATP binding site abolished this function (Khavari et al., 1993; Murchardt and Yaniv, 1993). Thus the role of SWI and SNF gene products as coactivators of transcription, perhaps via the alteration of chromatin structure, was shown to be conserved in higher eukaryotes. Complementing these studies of SWI2/SNF2 homologues in higher eukaryotes were important in vitro experiments analyzing the regulation of the Drosophila hsp70 promoter. Plasmids encoding the Drosophila hsp70 locus could be assembled into chromatin by mixing the template with Drosophila embryo extracts. These extracts contain histones and chromatin assembly factors that permit formation of long nucleosome arrays (Becker and Wu, 1992). Addition of recombinant GAGA protein, a transcription factor critical for hsp70 activation in vivo, to the assembled template resulted in the appearance of DNAse I hypersensitivity at the TATA box and heat-shock element and a rearrangement of adjacent nucleosomes (Tsukiyama et al., 1994). However, these changes in chromatin disruption were not observed when apyrase, an ATP-hydrolyzing enzyme, was included in the reaction following nucleosome assembly. This study, therefore, clearly demonstrated that, in vitro, transcriptional activation at an inducible locus incorporated into chromatin was associated with an ATP-dependent process. Although the authors did not identify the ATPase potentially responsible for the chromatin remodeling observed in this work, their efforts linking transcription-associated changes in chromatin structure with ATP hydrolysis intensified the search for factors that could alter chromatin structure in an ATP-dependent manner.
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B. Multisubunit Complexes that Alter Nucleosome Structure Biochemical evidence that the SWI and SNF gene products could be isolated from yeast extract as a multisubunit complex was provided by two groups that also demonstrated that the purified complex contained ATPase activity that could be stimulated by single-stranded as well as double-stranded DNA (Cairns et al., 1994; Coˆte´ et al., 1994; Peterson et al., 1994). Similarly, the human BRG1 protein eluted from sizing columns at a high molecular weight (Khavari et al., 1993), indicating that human cells contained large complexes containing SWI/SNF proteins. Subsequent purification from human cell lines of two large complexes that contained BRG1 or BRM and that possessed DNA-stimulated ATPase activity confirmed the existence of the complexes (Imbalzano et al., 1994; Kwon et al., 1994). Concurrent work showed that the yeast complex, termed SWI/SNF, and the human complexes, termed hSWI/SNF ‘‘A’’ and ‘‘B,’’ could directly alter reconstituted chromatin templates in vitro in an ATP-dependent manner (Coˆte´ et al., 1994; Imbalzano et al., 1994; Kwon et al., 1994). This was demonstrated most directly by mixing purified complex with in vitro assembled, rotationally phased, mononucleosome particles. Limiting cleavage of uniquely end-labeled, rotationally phased mononucleosomes with DNAse I results in a 10-bp ladder of cleavage products, as DNAse I cleaves nucleosome particles only at the point of the DNA helix farthest from the surface of the histone octamer. In the absence of ATP, no change in the DNAse I cleavage pattern of mononucleosomes was observed in the presence of yeast or human SWI/SNF complexes. However, in the presence of ATP, a stark change was observed. The 10-bp ladder cleavages decreased in intensity, whereas new cleavage sites appeared throughout the rest of the molecule. The simplest explanation for these changes was that the SWI/SNF complexes, in an ATP-dependent manner, directly altered the nucleosome structure such that the accessibility of the particle to DNAse I was increased. Two other lines of experimentation supported a direct, ATP-dependent effect on nucleosome structure by SWI/SNF complexes. When GAL4 fusion proteins or a combination of the TATA-binding protein (TBP) and TFIIA were added to nucleosomes containing binding sites for these proteins, the transcription factors bound poorly or not at all. Similar results were obtained when SWI/SNF enzymes were added in the absence of ATP. However, in the presence of ATP, the affinity of GAL4 proteins for their binding sites on the nucleosome was drastically increased (Coˆte´ et al., 1994; Kwon et al., 1994), and the TBP/TFIIA complex was able to stably bind to the nucleosome (Imbalzano et al., 1994). Experiments done by
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DNAse I footprinting demonstrated that the SWI/SNF-mediated structural alterations were occurring, as the altered DNAse I digestion pattern was evident. These data indicated that the structural alterations caused by SWI/SNF facilitated the binding of the transcription factors. Importantly, two pieces of evidence indicated that the SWI/SNF-mediated changes occurred without displacing the histone octamer. The first indication was from the DNAse I footprinting experiments. Though the altered digestion pattern more closely resembled that of naked DNA than that of the unaltered, rotationally phased nucleosome, it was not the same, indicating that the histones had not been entirely displaced (Coˆte´ et al., 1994; Imbalzano et al., 1994; Kwon et al., 1994). Stronger evidence came from gel-shift experiments that demonstrated an increased affinity of GAL4 proteins for SWI/SNF-altered nucleosomes (Coˆte´ et al., 1994). The GAL4-nucleosome complex exhibited a mobility in native gels that was equivalent to that exhibited by high concentrations of GAL4 bound to unaltered nucleosomes and that was reduced compared to GAL4-naked DNA complexes. In addition, hSWI/SNF complexes could structurally alter nucleosome plasmid templates in an ATP-dependent manner (Imbalzano et al., 1996; Kwon et al., 1994). Introduction of a nucleosome to a closed circular DNA introduces one negative supercoil. A plasmid that contained 14–15 nucleosomes therefore migrated on a two-dimensional agarose gel with a superhelical density of 14–15 negative supercoils. Analysis of hSWI/SNF-treated templates in the presence but not the absence of ATP indicated that the average template molecule lost five to six negative supercoils. This structural change demonstrated that SWI/SNF complexes could alter both mononucleosome and plasmid nucleosomal templates. The mechanism responsible for the alterations remains undetermined. The simplest explanation is that each loss of one negative supercoil is the result of the complete loss of a histone octamer from the template. Given that octamers are not lost on mononucleosome and linear array templates treated with SWI/SNF complexes, there is a belief that octamers also are not displaced on circular nucleosomal templates; however, this has not been directly demonstrated, and the basis for the loss of superhelical density remains undefined. In the years following the description of the nucleosome-remodeling properties of yeast and human SWI/SNF complexes, other highly related complexes containing the previously identified ATPase subunits or subunits closely related to the SWI2/SNF2 ATPase were identified and grouped into the SWI/SNF family of complexes (Cairns et al., 1996; Nie et al., 2000; Papoulas et al., 1998). In addition, other complexes containing central ATPase subunits that were similar to the SWI2/SNF2 ATPase only in the ATPase domain but not elsewhere were identified in yeast, flies,
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and mammals (Bochar et al., 2000; Guschin et al., 2000; Ito et al., 1997; LeRoy et al., 1998, 2000; Poot et al., 2000; Shen et al., 2000; Strohner et al., 2001; Tsukiyama and Wu, 1995; Tsukiyama et al., 1999; Varga-Weisz et al., 1997; Wade et al., 1998; Xue et al., 1998; Zhang et al., 1998). In fact, the ATP-dependent activity originally described in Drosophila extracts was attributed to a complex defined as NURF, for nucleosome remodeling factor (Tsukiyama and Wu, 1995). NURF contained an ATPase termed ISWI, which is a member of the SWI2/SNF2 ATPase family based on the similarity of its ATPase domain but that has distinct domains elsewhere in its coding sequence (Tsukiyama et al., 1995). At present, there are four families of ATP-dependent chromatin-remodeling enzymes recognized, grouped on the relative similarity of the central ATPase subunit: the SWI/SNF family, the ISWI family, the CHD/Mi2 family, and the Ino80 family (Fig. 1). Though the different enzymes show some differences in cofactor requirement and substrate recognition, they all function in multisubunit complexes to alter chromatin structure in an ATP-dependent manner (Fig. 2). Some studies comparing the activities of different family members have supported the idea that some, if not many, of the basic mechanisms behind nucleosome remodeling are the same (Boyer et al., 2000). Thus, further discussion of nucleosome remodeling will draw on mechanistic studies performed with different family members. Differences between various family members will be pointed out where appropriate.
III. Requirements for Nucleosome-Remodeling Enzymes A. Interaction with DNA/Chromatin An enzyme that causes structural changes in chromatin must interact with the DNA in some manner. Biochemical fractionation of cells to isolate human SWI/SNF used DNA affinity columns, (Kwon et al., 1994), indicating that the complexes possessed nonspecific DNA binding activity. This was also confirmed by early gel shift studies with the yeast SWI/SNF enzyme and mononucleosome templates, which revealed that under some experimental conditions, the radiolabeled nucleosome particles tended to remain in the gel wells and not migrate into the matrix, indicating an interaction between SWI/SNF and the nucleosome particle (Coˆte´ et al., 1994). A study examining the DNA binding properties of yeast SWI/SNF revealed that SWI/SNF bound to naked DNA nonspecifically, but with a relatively high affinity of 109 M (Quinn et al., 1996). This report also showed that the enzyme could bind to synthetic four-way-junction DNA. It has been argued that four-way-junction DNA resembles
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Fig. 1. Subfamilies of SWI2/SNF2 ATPases.
the DNA at the entry/exit point of a nucleosome particle where the helices form a cross-over structure (Lilley, 1992). It was thus suggested that SWI/SNF might bind to the nucleosome at the entry/exit point of DNA. The protein surface or surfaces of ATP-dependent chromatinremodeling enzymes that interact with nucleosomes remain undefined. None of the members of the extended family of enzymes contains subunits that have obvious DNA binding domains. The only exception is the BAF57 subunit of human SWI/SNF enzymes, which contains a single HMG-like domain (Wang et al., 1998). Because HMG proteins bind bent DNA, this
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Fig. 2. Summary of the complexes formed by different SWI2/SNF2 ATPases.
appeared to be a likely candidate for mediating SWI/SNF binding. However, the BAF57 subunit has no homologue in the yeast complexes, and mutational analysis in which the HMG domain was deleted resulted in no effect on enzyme function. Though it remains possible that the domain contributes to chromatin interaction, it is clearly not required. Initial attempts at microscopic visualization of remodeling enzymes with DNA and linear polynucleosome arrays using electron spectroscopic techniques revealed that yeast SWI/SNF was capable of forming loops via the simultaneous interaction with more than one nucleosome (Bazett-Jones et al., 1999). In addition to indicating that SWI/SNF might induce topological constraints on substrate molecules, these data indicated that the initial interaction of the complex with chromatin likely was not specifically mediated by one individual subunit. Cross-linking studies of yeast SWI/SNF and RSC with nucleosome particles before and after addition of ATP revealed that subunits that were in close enough proximity to the crosslinking agent incorporated into the nucleosomal DNA were displaced on addition of ATP (Sengupta et al., 2001). Interestingly, in the case of
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SWI/SNF, the ATPase subunit SWI2/SNF2 could be cross-linked to the unremodeled nucleosome but not the remodeled nucleosome or to naked DNA, indicating that the ATPase subunit was involved in nucleosome recognition. However, Sth1, the corresponding ATPase of the RSC complex, was not cross linked to either nucleosomal or free DNA, raising questions about whether direct interaction with the ATPase subunit is universally required for substrate recognition. More recently, two structural analyses of yeast RSC and yeast SWI/SNF have been performed using three-dimensional reconstruction from individual electron micrographs. The RSC studies reveal a ring of protein density around a central cavity, the dimensions of which are consistent with its occupation by a single nucleosome particle (Asturias et al., 2002). The SWI/SNF structure also contained a ring of protein density that formed a rim around a pocket of sufficient size to interact with a single nucleosome particle (Smith et al., 2003). Though different, the two structures share the apparent capacity to interact with a single nucleosome in an environment largely surrounded by enzymatic subunits. Furthermore, though the experiments do not reveal the positions of individual subunits in the structure, they strongly support the idea that substrate recognition is mediated via surfaces comprising multiple subunit proteins.
B. ATP Binding and Hydrolysis As the name implies, there is a requirement for ATP hydrolysis so that ATP-dependent chromatin-remodeling enzymes mediate changes in chromatin structure. Isolation and sequencing of the yeast SWI2/SNF2 gene showed that it possessed a putative ATPase domain that resembled that of known helicases. Expression of the domain as a bacterial fusion protein showed that the domain did, in fact, hydrolyze ATP (Laurent et al., 1993). Later work with purified yeast SWI/SNF complex demonstrated that the complex itself possessed ATPase activity (Cairns et al., 1994; Coˆte´ et al., 1994). The ATP requirement for SWI/SNF-mediated chromatin remodeling was demonstrated by biochemical experiments showing that observed structural alterations in nucleosome and polynucleosome structure occurred in the presence, but not the absence, of added ATP (Coˆte´ et al., 1994; Imbalzano et al., 1994; Kwon et al., 1994). Similar findings were made for the activities in the Drosophila extract system that could affect nucleosome positioning (Ito et al., 1997; Tsukiyama and Wu, 1995; Varga-Weisz et al., 1997). Experiments showing that apyrase, which cleaves ATP, blocked chromatin remodeling confirmed the requirement for ATP, whereas use of nonhydrolyzable ATP analogs demonstrated the
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requirement not just for ATP but for actual hydrolysis to support chromatin remodeling. Finally, evaluation of ADP and a panel of nucleotide and deoxynucleotide triphosphates revealed a clear specificity for ATP; of the related analogs tested, only dATP supported any nucleosome altering activity, and higher concentrations were required to observe activity (Imbalzano et al., 1996; Kwon et al., 1994; Tsukiyama and Wu, 1995; Tsukiyama et al., 1994; Varga-Weisz et al., 1995). The ATPase domain shared by the catalytic subunits that form the core of ATP-dependent chromatin remodeling enzymes comprises seven regions of homology, termed blocks I–VII, and closely resembles ATPase domains of known helicases (Eisen et al., 1995). Helicases unwind DNA helices. Thus, the prospect that these enzymes possessed helicase activity that might be used to alter chromatin structure was attractive. Yet the recombinant fusion protein containing the yeast SWI2/SNF2 ATPase did not possess measurable helicase activity (Laurent et al., 1993), and extensive efforts to show that the purified yeast SWI/SNF enzyme had helicase activity failed (Coˆte´ et al., 1994). Despite the sequence similarity to known helicases, to date, none of the members of the ATP-dependent chromatinremodeling enzymes have demonstratable helicase activities. However, the yeast Ino80 complex has been shown to be associated with two subunits that are related to known DNA helicases, and the complex itself possesses DNA helicase activity (Shen et al., 2000). It remains possible that the other ATP-dependent remodeling enzymes associate with helicases that are not integral subunits of the enzymes.
C. Cofactor Requirements All of the family members exhibit measurable ATPase activity in the absence of cofactors; however, the presence of DNA or chromatin greatly stimulates ATPase activity. Early studies with yeast SWI/SNF enzyme examined a host of possible cofactors, with double-stranded and single-stranded DNA and nucleosome particles providing the best stimulation of ATPase activity (Cairns et al., 1994; Coˆte´ et al., 1994). Similarly, the human SWI/ SNF complexes exhibited ATPase activity that could be stimulated by both single-stranded and double-stranded DNA (Kwon et al., 1994). In contrast, the first of the ISWI-containing factors to be examined in detail, Drosophila NURF, showed minimal DNA-stimulated ATPase activity but a much greater stimulation mediated by nucleosome particles (Tsukiyama and Wu, 1995). This indicated that the different classes of ATP-dependent chromatin-remodeling enzymes might recognize their substrates differently or, alternatively, might differently translate that physical interaction with the substrate toward stimulation of ATP hydrolysis.
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D. Nucleosome Remodeling is an Enzymatic Process Initial studies of chromatin-remodeling complexes in assays examining changes in nucleosome structure were often performed under conditions in which the remodeling complex and substrate were present at similar ratios. Although the capacity of these complexes to hydrolyze ATP clearly occurred via an enzymatic process, the actual reaction that resulted in an alteration to nucleosome structure was not rigorously demonstrated to be enzymatic. One nonenzymatic explanation for chromatin remodeling involved chromatin-remodeling complexes stably associating with substrate molecules and using the energy of ATP hydrolysis to keep the template in an altered state. Such a scenario could require maintenance of a 1:1 association between substrate and remodeler. However, development of a unique assay coupling the activities of nucleosome-remodeling complexes with restriction enzyme cleavage finally definitively determined that chromatin-remodeling activities were, in fact, enzymatic (Logie and Peterson, 1997). In this study, the authors used an array of repeat sequences known to position nucleosomes, and they modified the middle repeat to contain a unique restriction-enzyme cleavage site. When the template was assembled into a nucleosomal array, the site was largely resistant to restriction-enzyme digestion. However, in the presence of ySWI/SNF and ATP, the site became accessible, and the template was cleaved into two shorter fragments. This enabled the authors to perform several remodeling ‘‘cycles,’’ in which ySWI/SNF, ATP, and excess amounts of template were added to the reaction; the template was remodeled and digested; and then, with little or no full-length template left in the reaction, more template was added. The subsequent digestion of all of these template molecules and of those added repeatedly on complete or nearly complete cleavage of the previously added aliquot of template provided evidence that the remodeling factor was altering nucleosome structure on multiple template molecules. Thus the chromatin remodeler was acting catalytically, and efforts to investigate the mechanism or mechanisms by which nucleosome structures were altered intensified.
IV. Mechanisms of ATP-Dependent Chromatin Remodeling Chromatin remodeling, nucleosome remodeling, and nucleosome disruption are general terms that have been used to summarize a number of changes in chromatin structure. Often, these terms are used to define any event that alters the nuclease sensitivity of a region of chromatin, as nuclease sensitivity is a frequently used marker of changes in chromatin structure. However, there have been other in vitro assays devised to
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examine ATP-dependent chromatin remodeling activity as well, including assays monitoring changes in nucleosomal plasmid supercoiling, histone octamer transfer, and nucleosome–nucleosome interactions (reviewed in Narlikar et al., 2002). Not every remodeling enzyme functions in every in vitro remodeling assay, but each one works in multiple assays. It remains unclear as to whether each of the assays is measuring aspects of the same activity or whether there is more than one distinct activity.
A. Nucleosome Sliding Passive movement of nucleosome along DNA, also referred to as translational repositioning, was noted previously in response to elevated temperature and changes in ionic conditions (Meersseman et al., 1992; Pennings et al., 1991). Such changes in octamer positioning imply that histone–DNA contacts are broken and subsequently reformed, resulting in DNA that was previously associated with the histone octamer being rendered nonnucleosomal (Fig. 3a). Given the stability of the octamer interaction with DNA, it is easy to imagine that such movement would be energetically unfavorable. Thus, the solution to generating such movement on genomic chromatin in cells might be an enzyme that is capable of disrupting histone–DNA contacts via energy generated by a coupled ATP hydrolysis reaction. Support for the idea that chromatin remodeling is essentially the act of creating nucleosome mobility is the fact that all four classes of ATPdependent enzymes are capable of changing the translational position of nucleosomes on DNA. This was first demonstrated in some of the initial studies using undefined ATP-dependent activities in Drosophila extracts, in which the authors saw changes in the translational position of nucleosomes on array template (Tsukiyama et al., 1994; Varga-Weisz et al., 1995). Later studies used the ISWI-containing complexes NURF and CHRAC (chromatin accessibility complex) and recombinant ISWI protein to examine unidirectional changes in translational positioning of single mononucleosome particles assembled on DNA fragments of greater than 200 bp in length (Hamiche et al., 1999; La¨ngst et al., 1999). Similarly, yeast and human SWI/SNF complexes as well as the Mi2 complex were shown to reposition nucleosomes on short linear DNAs as well as on small, circular plasmid DNA (Brehm et al., 2000; Gavin et al., 2001; Goschin et al., 2000; Guyon et al., 2001; Jaskelioff et al., 2000; Whitehouse et al., 1999). In addition, the ISWI family of enzymes possesses the ability to create regularly spaced nucleosome arrays from templates containing randomly spaced nucleosomes (reviewed in La¨ngst and Becker, 2001). This nucleosome spacing function, though apparently not shared by the other families
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Fig. 3. Models for changes in chromatin structure mediated by ATP-dependent remodeling enzymes.
of ATP-dependent remodelers, further confirms that the ISWI enzymes can translationally reposition nucleosomes and indicates that the ISWI complexes may have a role in chromatin assembly. The obvious consequences of nucleosome sliding in vivo include exposing or shielding regulatory sequences to permit or restrict the interaction of DNA binding factors important in transcription, replication, or recombination with the chromatin. Few definitive examples of nucleosome sliding affecting one of these processes in vivo exist; the best example is perhaps the changes that occur on the interferon beta promoter following activation by infection of cells with RNA viruses (Agalioti et al., 2000; Lomvardas and Thanos, 2001). On activation of the promoter, a cascade of transcription factor and chromatin-remodeling enzyme interactions occurs, culminating in the assembly of a complete enhancesome and a preinitiation complex lacking only TBP on the promoter. However, transcription was not initiated until the SWI/SNF complex interacted with the promoter (Agalioti et al., 2000). Detailed examination of nucleosome positioning before and after transcriptional induction revealed that a nucleosome partially obscuring the TATA sequence was moved via the activity of the SWI/SNF enzyme to a position approximately 35 bp downstream, thereby permitting TBP to bind and transcription to commence (Lomvardas and Thanos, 2002). Undoubtedly, future analyses of activated
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promoters will seek to emphasize whether and how nucleosome positioning is changed to determine how frequently nucleosome sliding contributes to regulatable events in vivo. Though there are no existing data, nucleosome sliding also potentially could affect higher-order chromatin structure by increasing or decreasing the spacing between individual nucleosomes in an array. Such changes might affect the potential for a specific stretch of genomic DNA to bend or fold into higher-order structure and might therefore affect enzymes or factors that interact with chromatin.
B. Nucleosome Disruption Though the ISWI class of enzymes likely mediates chromatin remodeling via a sliding mechanism, the other classes of ATP-dependent remodelers may possess a second, distinct activity. SWI/SNF family enzymes can cause a significant increase in DNAse I or in restriction-enzyme accessibility on mononucleosome particles, even when there is no appreciable amount of flanking DNA on which the nucleosome can translationally reposition (Fig. 3b). In contrast, ISWI enzymes show a minimal requirement for flanking DNA to mediate sliding (Brehm et al., 2000). As discussed, rotationally phased mononucleosome particles with a minimal length of DNA around the octamer show a significantly different DNAse I digestion pattern after remodeling by SWI/SNF enzymes. Although this observation could be explained by sliding, this would require extrusion, or ‘‘peeling’’ of part of the DNA off the histone octamer. Such events have been documented on nucleosome particles (Anderson and Widom, 2000; Polach and Widom, 1995), and there is a direct relationship between the fraction of time that a particular DNA sequence spends ‘‘peeled,’’ or ‘‘off’’ the octamer, and its proximity to the end of the DNA fragment. Thus, the end of the DNA sequence wrapped around a histone octamer is predicted to be free more frequently than a DNA sequence near the middle of the fragment. However, analyses of restriction enzyme cleavage of unique sequences at various positions on a mononucleosome particle in the presence of ATP and human SWI/SNF enzymes revealed that restriction sites near the center of the DNA fragment were made more accessible than sites near the end of the DNA fragment (Narlikar et al., 2001). These results are exactly the opposite of what would have been predicted if sliding were occurring via a mechanism in which the DNA were peeled or extruded from around the histone core. Furthermore, reductions in superhelical density observed on circular nucleosomal plasmid templates in the presence of SWI/SNF enzymes cannot easily be explained by a sliding mechanism. Finally, experiments in which the DNA was crosslinked
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to the histone octamer did not show inhibited SWI/SNF-mediated nucleosome remodeling, as judged by changes in DNAse I sensitivity (Lee et al., 1999). This observation is particularly difficult to reconcile with a sliding mechanism for the observed changes in nucleosome structure. Collectively, these observations indicate that a remodeling event distinct from nucleosome sliding can occur in the presence of the SWI/SNF family of remodeling enzymes. Key to building a model to explain such changes in nucleosome structure is a requirement that nucleosomal DNA sequences be made accessible without involving translational repositioning of the DNA. Such changes could occur via changes in histone octamer conformation, via changes in the path of the DNA around the nucleosome, such that different histone–DNA contacts are formed, via changes in the conformation of the DNA itself, or via a combination of these possibilities. Recently, a model proposing conformational change as an alternative to sliding was proposed (Narlikar et al., 2002). These authors suggested that ATP hydrolysis drives conformational changes in the DNA, octamer, or both that result in high-energy intermediate structures that stochastically collapse into one of multiple, distinct, final (remodeled) states. Some of these remodeled states could be achieved via a sliding mechanism, whereas others would maintain some aspects of the conformational changes in nucleosome structure without associated sliding.
C. Formation of an Altered Dinucleosome-Like Species Two groups analyzing alteration of mononucleosome particles by the SWI/SNF family members human SWI/SNF and yeast RSC (remodels the structure of chromatin) came to the conclusion that the altered nucleosome species had characteristics of a nucleosome dimer, even after the remodeling enzyme was no longer associated with the substrate (Fig. 3c; Lorch et al., 1998; Schnitzler et al., 1998). The altered, dinucleosome-like structure retained the expected properties of the remodeled nucleosome in that it was hypersensitive to nucleases and bound transcription factors with increased affinity. However, it exhibited mobility on gels and in gradient centrifugation experiments similar to that exhibited by dinucleosomes. Importantly, the enzymes were capable of converting the remodeled species back to the original unremodeled state, giving the first indication that these enzymes could catalyze changes in nucleosome structure in both directions. Subsequent experiments indicated that the dimer formed was not covalently linked, and it was proposed that the enzymes mediated a release of 60–80 bp from the end to promote formation of the remodeled, dimeric species (Lorch et al., 2001). How
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creation of dimeric remodeled nucleosomes relates to remodeling on nucleosomal arrays and in vivo function awaits further research.
D. Octamer Transfer in trans Years ago, it was proposed that nucleosome-free regions might be formed by transfer of histone octamers from the DNA to an acceptor molecule in trans (reviewed in Adams and Workman, 1993). This idea, termed nucleosome displacement, was shown to occur in vitro by challenging a single mononucleosome particle containing radiolabeled DNA and bound by five dimers of the yeast GAL4-derived transcription factor with excess quantities of unlabeled DNA fragments (Workman and Kingston, 1993). The nucleosome bound by the GAL4 derivative dissociated into separate complexes of five GAL4 dimers bound to naked, radiolabeled DNA, and a histone octamer bound to the unlabeled DNA. Because octamers from radiolabeled nucleosomes unbound by the GAL4 factor could not be displaced when challenged with excess free DNA, the authors suggested that the transcription factor–bound nucleosome was ‘‘metastable’’ and that in the presence of a histone acceptor such as the free DNA, the octamer was capable of being displaced in trans. Further work using similar assays demonstrated that histone-binding proteins could similarly act as acceptors for octamers displaced in trans (Chen et al., 1994). Evidence that ATP-dependent remodeling enzymes could mediate octamer transfer in trans came from work examining the properties of the yeast SWI/SNF family member RSC (Lorch et al., 1999). Nucleosome particles mixed with RSC, ATP, and radiolabeled free DNA fragments gave rise to nucleosome particles containing radiolabeled DNA (Fig. 3d), indicating that octamer exchange had occurred. It appeared that the free DNA interacted with a RSC containing nucleosome intermediate product with the characteristics of the nucleosome dimer described above. Subsequent work demonstrated that human SWI/SNF complexes also could mediate octamer transfer in trans (Phelan et al., 2000). There is no direct evidence that either dimer formation or octamer transfer in trans occurs in vivo as a mechanism to alter chromatin structure and facilitate any aspect of gene regulation or other use of the DNA. However, these mechanisms represent a means by which DNA could be rendered nucleosome free without forcing tighter packing of nucleosomes in adjacent regions. In addition, there is evidence that regions of the DNA do become nucleosome free or, perhaps better stated, free of classical nucleosome structure. For instance, the yeast PHO5 promoter, when inactive, contains four positioned nucleosomes across its promoter.
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On activation of the gene by phosphate starvation, nucleosome positioning is lost, and the region appears to be free of defined nucleosomes (Almer et al., 1986). Although sliding could account for the accessibility of this region during gene activation, it would seem likely that repositioning of these nucleosomes might affect transcriptional activation, especially if the nucleosomes were more tightly packed across the transcription initiation site or immediately downstream, where the transcribing polymerase might have to contend with increased nucleosome density. A mechanism by which the octamers were transferred in trans to DNA more distal to the promoter could facilitate the process by which the gene becomes activated.
V. Initiation of ATP-Dependent Chromatin Remodeling Despite the significant amounts of effort spent investigating the process by which nucleosome remodeling occurs, exactly how the discussed models relate to each other and relate to chromatin remodeling in vivo remains unanswered. Not surprisingly, there also exist multiple, nonexclusive models for how changes in nucleosome structure are initiated, irrespective of how the change in nucleosome structure is ultimately manifested. The three best developed of these models can be described as the spooling/peeling model, the twist model, and the bulging/looping model. At present there is no indication that any of these models are correct, nor that the remodeling enzymes employ only one mechanism to initiate structural changes. Ultimately, aspects of different models might be combined to explain how the energy generated by ATP hydrolysis translates into structural change.
A. Spooling or Peeling Model Variations of this model exist. One basis for this model is the idea that the enzymes might track along the DNA, breaking histone–DNA bonds and causing release of the DNA from the end (Pazin and Kadonaga, 1997). Though this possibility has not been ruled out, the sheer size of the remodeling enzymes and potential concerns regarding the buildup of positive and negative supercoiled indicate that this model is unlikely. A variation envisions that the enzyme itself interacts specifically with the mononucleosome particle and mediates a transient unpeeling of the DNA from the edge (Anderson and Widom, 2000; Polach and Widom, 1995). As discussed above, evidence indicates that this kind of behavior exists with low frequency in the absence of chromatin-remodeling enzymes; potentially, the remodeling enzyme lowers the energy barrier and facilitates the process.
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A peeling or spooling model can explain all possible outcomes seen following ATP-dependent remodeling enzyme activity, but is particularly useful for explaining formation of dinucleosome products and for octamer transfer in cis. Release of large segments of DNA from the octamer to generate free DNA might mediate invasion of a neighboring or distal nucleosome to permit formation of dinucleosomes or transfer of the octamer.
B. Twisting Model Examination of the crystal structure of a nucleosome particle revealed that the nucleosome could accommodate small changes in helix geometry (reviewed in Luger and Richmond, 1998). Years ago, van Holde and colleagues proposed that thermal energy could alter the twist of the DNA by disrupting a small number of histone–DNA contacts at the edge of a nucleosome (van Holde, 1998). If means to introduce such twist changes to the end of the DNA could be identified, the twist could be propagated, 1 bp at a time, through the length of the nucleosomal DNA, thus effecting a change in nucleosome position, and at least transiently, a change in nucleosome structure. This possibility has been termed ‘‘twist diffusion’’ (Luger et al., 1997). Findings that members of three classes of enzymes could introduce superhelical torsion into linear DNA provide support for this model, as the enzymes themselves could repeatedly induce a small twist that could thus be propagated and tuned into a substantial change in nucleosome positioning (Havas et al., 2000). Importantly, disruption of histone– DNA contacts would occur locally instead of over large tracts of DNA, and the path of the DNA around the octamer would not have to change in any significant manner. Elegant models invoking this mechanism have been proposed (reviewed in Flaus and Owen-Hughes, 2001). Despite the model’s recent popularity, there are data to argue against a simple twist model. First, a simple twist model implies that as DNA twists along the octamer, specific sequences will rotate through every position on the helix. However, early studies examining the interaction of TBP to nucleosomes containing TATA sequences showed that human SWI/SNF could facilitate TBP binding, but only if the TATA sequence were in one specific rotational position (Imbalzano et al., 1994). This was explained by the requirement for simultaneous bending of the DNA by both the octamer and the TBP. Implicit in the observation is that not all orientations of the binding site were equivalent; had the sequence simply been twisted through the helix by the chromatin remodeling enzyme, at some point the TATA sequence should have rotated through the orientation that was accessible for TBP binding. The fact that this was not observed
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indicates that a base-pair-by-base-pair twist does not occur. Second, there are data demonstrating that DNA containing nicks is not inhibitory to efficient nucleosome sliding by ISWI (La¨ngst and Becker, 2001). A nick should be inhibitory to propagation of altered twist, thereby arguing against such a model. However, a recent study using yeast RSC indicated that remodeling on nucleosomes containing nicked DNA was less efficient (Saha et al., 2002).
C. Bulging or Looping Model Bulging, or looping indicates that a (presumably) small region of the DNA forms a loop or bulge on the surface of the octamer. How such a bulge is generated is an interesting problem. It has been suggested that the remodeling enzyme would need to contact the nucleosome at two positions: one for anchoring purposes, the other to contact and lift the DNA via the disruption of histone–DNA contacts (reviewed in Becker and Horz, 2002). The presence of a bulge suggests that the helix is distorted, or stretched, to accommodate the displacement or, more likely, that DNA is translationally moved into the nucleosome, from linker regions if available, to compensate for the length of DNA displaced on the bulge. One could imagine models in which the bulge is formed at specific or random locations, collapses, and is reformed at the same or different locations. Alternatively, the bulge could be formed and propagated throughout the length of the nucleosomal DNA. Such a mechanism could account for ATPdependent remodeling with or without sliding. It could also, via displacement of DNA from the octamer surface, lead to the initiation of octamer transfer or dinucleosome formation. The exact nature of how bulges might form and translate to remodeled chromatin remains to be determined.
VI. Conclusions Intense efforts have yielded a wealth of data regarding changes in nucleosome structure by ATP-dependent chromatin remodeling enzymes; however, linking all of the data points into cohesive models has proven elusive. Further work will be necessary to elucidate precisely how energy from ATP hydrolysis initiates structural changes on nucleosomal templates. In addition, a more complete understanding of what physical form or forms remodeled chromatin assumes is needed, as is greater investigation of how the structural changes are maintained. Finally, whether all classes of ATP-dependent remodeling enzymes function via the same mechanisms will need to be determined. In short, there currently exist decent working models to guide further in-depth studies of the mechanisms
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of ATP-dependent chromatin remodeling. Work over the next 5–10 years will likely yield novel ideas that will alter and further develop these current hyotheses into more precisely defined models.
Acknowledgments We thank Ivana de la Serna and David Hill for comments on the manuscript. Work in the authors’ laboratory is supported by grants from the National Institutes of Health, the National Cancer Institute, and the Leukemia and Lymphoma Society.
References Adams, C. C., and Workman, J. L. (1993). Cell 72, 305–308. Agalioti, T., Lomvardas, S., Parekh, B., Yie, J., Maniatis, T., and Thanos, D. (2000). Cell 103, 667–678. Almer, A., Rudolph, H., Hinnen, A., and Horz, W. (1986). EMBO J. 5, 2689–2696. Anderson, J. D., and Widom, J. (2000). J. Mol. Biol. 296, 979–987. Asturias, F. J., Chung, W. H., Kornberg, R. D., and Lorch, Y. (2002). Proc. Natl. Acad. Sci. USA 99, 13477–13480. Bazett-Jones, D. P., Coˆte´, J., Landel, C. C., Peterson, C. L., and Workman, J. L. (1999). Mol. Cell. Biol. 19, 1470–1478. Becker, P. B., and Horz, W. (2002). Annu. Rev. Biochem. 71, 247–273. Becker, P. B., and Wu, C. (1992). Mol. Cell. Biol. 12, 2241–2249. Berger, S. L. (2002). Curr. Opin. Genet. Dev. 12, 142–148. Bochar, D. A., Savard, J., Wang, W., Lafleur, D. W., Moore, P., Cote, J., and Shiekhattar, R. (2000). Proc. Natl. Acad. Sci. USA 97, 1038–1043. Boyer, L. A., Logie, C., Bonte, E., Becker, P. B., Wade, P. A., Wolffe, A. P., Wu, C., Imbalzano, A. N., and Peterson, C. L. (2000). J. Biol. Chem. 275, 18864–18870. Breeden, L., and Nasmyth, K. (1987). Cell 48, 389–397. Brehm, A., Langst, G., Kehle, J., Clapier, C. R., Imhof, A., Eberharter, A., Muller, J., and Becker, P. B. (2000). EMBO J. 19, 4332–4341. Coˆte´, J., Quinn, J., Workman, J. L., and Peterson, C. L. (1994). Science 265, 53–60. Cairns, B. R., Kim, Y.-J., Sayre, M. H., Laurent, B. C., and Kornberg, R. D. (1994). Proc. Natl. Acad. Sci. USA 91, 1950–1954. Cairns, B. R., Lorch, Y., Li, Y., Zhang, M., Lacomis, L., Erdjument-Bromage, H., Tempst, P., Du, J., Laurent, B., and Kornberg, R. D. (1996). Cell 87, 1249–1260. Chen, H., Li, B., and Workman, J. L. (1994). EMBO J. 13, 380–390. Chiba, H., Muramatsu, M., Nomoto, A., and Kato, H. (1994). Nucleic Acids Res. 22, 1815–1820. Eisen, J. A., Sweder, K. S., and Hanawalt, P. C. (1995). Nucleic Acids Res. 23, 2715–2723. Flaus, A., and Owen-Hughes, T. (2001). Curr. Opin. Genet. Dev. 11, 148–154. Gavin, I., Horn, P. J., and Peterson, C. L. (2001). Mol. Cell 7, 97–104. Guschin, D., Geiman, T. M., Kikyo, N., Tremethick, D. J., Wolffe, A. P., and Wade, P. A. (2000). J. Biol. Chem. 275, 35248–35255. Guyon, J. R., Narlikar, G. J., Sullivan, E. K., and Kingston, R. E. (2001). Mol. Cell. Biol. 21, 1132–1144. Hamiche, A., Sandaltzopoulos, R., Gdula, D. A., and Wu, C. (1999). Cell 97, 833–842.
178
IMBALZANO AND XIAO
Havas, K., Flaus, A., Phelan, M., Kingston, R., Wade, P. A., Lilley, D. M., and Owen-Hughes, T. (2000). Cell 103, 1133–1142. Hirschhorn, J. N., Bortvin, A. L., Ricupero-Hovasse, S. L., and Winston, F. (1995). Mol. Cell. Biol. 15, 1999–2009. Hirschhorn, J. N., Brown, S. A., Clark, C. D., and Winston, F. (1992). Genes Dev. 6, 2288–2298. Imbalzano, A. N., Kwon, H., Green, M. R., and Kingston, R. E. (1994). Nature 370, 481–485. Imbalzano, A. N., Schnitzler, G. R., and Kingston, R. E. (1996). J. Biol. Chem. 271, 20726–20733. Ito, T., Bulger, M., Pazin, M. J., Kobayashi, R., and Kadonaga, J. (1997). Cell 90, 145–155. Jaskelioff, M., Gavin, I., Peterson, C. L., and Logie, C. (2000). Mol. Cell. Biol. 20, 3058–3068. Khavari, P. A., Peterson, C. L., Tamkun, J. W., and Crabtree, G. R. (1993). Nature 366, 170–174. Kruger, W., Peterson, C. L., Sil, A., Coburn, C., Arents, G., Moudrianakis, E. N., and Herskowitz, I. (1995). Genes Dev. 9, 2770–2779. Kruger, W., and Herskowitz, I. (1991). Mol. Cell. Biol. 11, 4135–4146. Kwon, H., Imbalzano, A. N., Khavari, P. A., Kingston, R. E., and Green, M. R. (1994). Nature 370, 477–481. La¨ngst, G., Bonte, E. J., Corona, D. F., and Becker, P. B. (1999). Cell 97, 843–852. La¨ngst, G., and Becker, P. B. (2001). J. Cell. Sci. 114, 2561–2568. La¨ngst, G., and Becker, P. B. (2001). Mol. Cell 8, 1085–1092. Laurent, B. C., Treich, I., and Carlson, M. (1993). Genes Dev. 7, 583–591. Laurent, B. C., Treitel, M. A., and Carlson, M. (1990). Mol. Cell. Biol. 10, 5616–5625. Laurent, B. C., Treitel, M. A., and Carlson, M. (1991). Proc. Natl. Acad. Sci. USA 88, 2687–2691. Laurent, B. C., and Carlson, M. (1992). Genes Dev. 6, 1707–1715. Lee, K. M., Sif, S., Kingston, R. E., and Hayes, J. J. (1999). Biochemistry 38, 8423–8429. LeRoy, G., Loyola, A., Lane, W. S., and Reinberg, D. (2000). J. Biol. Chem. 275, 14787–14790. LeRoy, G., Orphanides, G., Lane, W. S., and Reinberg, D. (1998). Science 282, 1900–1904. Lilley, D. M. J. (1992). Nature 357, 282–283. Logie, C., and Peterson, C. L. (1997). EMBO J. 16, 6772–6782. Lomvardas, S., and Thanos, D. (2001). Cell 106, 685–696. Lomvardas, S., and Thanos, D. (2002). Cell 110, 261–271. Lorch, Y., Cairns, B. R., Zhang, M., and Kornberg, R. D. (1998). Cell 94, 29–34. Lorch, Y., Zhang, M., and Kornberg, R. D. (1999). Cell 96, 389–392. Lorch, Y., Zhang, M., and Kornberg, R. D. (2001). Mol. Cell 7, 89–95. Luger, K., Ma¨der, A. W., Richmond, R. K., Sargent, D. F., and Richmond, T. J. (1997). Nature 389, 251–260. Luger, K., and Richmond, T. J. (1998). Curr. Opin. Struct. Biol. 8, 33–40. Matallana, E., Franco, L., and Pe´rez-Ortı´n, J. E. (1992). Mol. Gen. Genet. 231, 395–400. Meersseman, G., Pennings, S., and Bardbury, E. M. (1992). EMBO J. 11, 2951–2959. Muchardt, C., and Yaniv, M. (1993). EMBO. J. 12, 4279–4290. Narlikar, G. J., Fan, H. Y., and Kingston, R. E. (2002). Cell 108, 475–487. Narlikar, G. J., Phelan, M. L., and Kingston, R. E. (2001). Mol. Cell 8, 1219–1230. Neigeborn, L., and Carlson, M. (1984). Genetics 108, 845–858. Nie, Z., Xue, Y., Yang, D., Zhou, S., Deroo, B. J., Archer, T. K., and Wang, W. (2000). Mol. Cell. Biol. 20, 8879–8888.
PROPERTIES OF ATP-DEPENDENT CHROMATIN REMODELING ENZYMES
179
Papoulas, O., Beek, S. J., Moseley, S. L., McCallum, C. M., Sarte, M., Shearn, A., and Tamkun, J. W. (1996). Development 125, 3955–3966. Pazin, M. J., and Kadonaga, J. T. (1997). Cell 88, 737–740. Pennings, S., Meersseman, G., and Bradbury, E. M. (1991). J. Mol. Biol. 220, 101–110. Peterson, C. L., Dingwall, A., and Scott, M. P. (1994). Proc. Natl. Acad. Sci. USA 91, 2905–2908. Peterson, L., and Herskowtiz, I. (1992). Cell 68, 573–583. Phelan, M. L., Schnitzler, G. R., and Kingston, R. E. (2000). Mol. Cell. Biol. 20, 6380–6389. Polach, K. J., and Widom, J. (1995). J. Mol. Biol. 254, 130–149. Poot, R. A., Dellaire, G., Hulsmann, B. B., Grimaldi, M. A., Corona, D. F., Becker, P. B., Bickmore, W. A., and Varga-Weisz, P. D. (2000). EMBO J. 19, 3377–3387. Quinn, J., Fyrberg, A. M., Ganster, R. W., Schmidt, M. C., and Peterson, C. L. (1996). Nature 379, 844–847. Roth, S. Y., Denu, J. M., and Allis, C. D. (2001). Annu. Rev. Biochem. 70, 81–120. Saha, A., Wittmeyer, J., and Cairns, B. R. (2002). Genes Dev. 16, 2120–2134. Schnitzler, G., Sif, S., and Kingston, R. E. (1998). Cell 94, 17–27. Sengupta, S. M., VanKanegan, M., Persinger, J., Logie, C., Cairns, B. R., Peterson, C. L., and Bartholomew, B. (2001). J. Biol. Chem. 276, 12636–12644. Shen, X., Mizuguchi, G., Hamiche, A., and Wu, C. (2000). Nature 406, 541–544. Smith, C. L., Horowitz-Scherer, R., Flanagan, J. F., Woodcock, C. L., and Peterson, C. L. (2003). Nat. Struct. Biol. 10, 141–145. Stern, M. J., Jensen, R., and Herskowitz, I. (1984). J. Mol. Biol. 178, 853–868. Strohner, R., Nemeth, A., Jansa, P., Hofmann-Rohrer, U., Santoro, R., Langst, G., and Grummt, I. (2001). EMBO J. 20, 4892–4900. Tamkun, J. W., Deuring, R., Scott, M. P., Kissinger, M., Pattatucci, A. M., Kaufman, T. C., and Kennison, J. A. (1992). Cell 68, 561–572. Tsukiyama, T., Becker, P. B., and Wu, C. (1994). Nature 367, 525–531. Tsukiyama, T., Daniel, C., Tamkun, J., and Wu, C. (1995). Cell 83, 1021–1026. Tsukiyama, T., Palmer, J., Landel, C. C., Shiloach, J., and Wu, C. (1999). Genes Dev. 13, 686–697. Tsukiyama, T., and Wu, C. (1995). Cell 83, 1011–1020. Turner, B. M. (2002). Cell 111, 285–291. van Holde, K. E. (1988). Chromatin. Springer, New York. Varga-Weisz, P. D., Blank, T. A., and Becker, P. B. (1995). EMBO J. 14, 2209–2216. Varga-Weisz, P. D., Wilm, M., Bonte, E., Dumas, K., Mann, M., and Becker, P. B. (1997). Nature 388, 598–602. Wade, P. A., Jones, P. L., Vermaak, D., and Wolffe, A. P. (1998). Curr. Biol. 8, 843–846. Wang, W., Chi, T., Xue, Y., Zhou, S., Kuo, A., and Crabtree, G. R. (1998). Proc. Natl. Acad. Sci. USA 95, 492–498. Whitehouse, I., Flaus, A., Cairns, B. R., White, M. F., Workman, J. L., and OwenHughes, T. (1999). Nature 400, 784–787. Workman, J. L., and Kingston, R. E. (1992). Science 258, 1780–1784. Xue, Y., Wong, J., Moreno, G. T., Young, M. K., Cote, J., and Wang, W. (1998). Mol. Cell 2, 851–861. Yoshinaga, S. K., Peterson, C. L., Herskowitz, I., and Yamamoto, K. R. (1992). Science 258, 1598–1604. Zhang, Y., LeRoy, G., Seelig, H. P., Lane, W. S., and Reinberg, D. (1998). Cell 95, 279–289.
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HISTONE ACETYLTRANSFERASE PROTEINS CONTRIBUTE TO TRANSCRIPTIONAL PROCESSES AT MULTIPLE LEVELS By MICHAEL S. TOROK AND PATRICK A. GRANT* Department of Biochemistry and Molecular Genetics, University of Virginia School of Medicine, Charlottesville, VA
I. Introduction . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. HATs and HAT Complexes: New Insights into HAT Regulation and Effects on Transcription . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. HAT Proteins Regulate Transcription . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. SAGA HAT Complex Consists of Modular Functional Domains . . . . . .. . . . . . C. The MYST Family of Acetyltransferases. . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . D. HAT Complexes are Regulated in Multiple Ways. . . . . . . . . . . . . . . . . . . . . . . .. . . . . . III. HAT Complexes Functionally Interact with Chromatin-Remodeling Complexes and Influence Transcription . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Specific Genes Recruit HAT Complexes and Chromatin Remodelers . . . . . B. Bromo Domain–Containing Proteins Function in Modifiers and Remodelers . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. HAT Complex Proteins Contain RING, SANT, and PHD Domains . .. . . . . . IV. HAT Proteins Function in Regulating Transcriptional Elongation . . . . . . . .. . . . . . A. The HAT Elp3 is a Component of the Elongator Complex . . . . . . . . . . .. . . . . . B. The HATs MOF, PCAF, and Sas3 Function in Transcriptional Elongation . . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Transcriptional Elongation is Regulated by Other Chromatin Modifiers . . V. The Histone Code: Insights into Epigenetic Regulation of Transcription. . . . . . A. The Histone Code Proposes Mechanisms of Transcriptional Regulation . . B. The Histone Code and the Endpoint to Signaling Pathways . . . . . . . . . .. . . . . . VI. HATs: New Insight into Transcription and DNA Repair . . . . . . . . . . . . . . . . . . . .. . . . . . VII. Conclusion and Perspectives . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . Acknowledgments. . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction In eukaryotes, transcriptional regulation is mediated within the context of nucleosomes, the repeating subunits of chromatin. The core nucleosome particle consists of 147 base pairs of DNA wrapped around the histone octamer composed of two copies of each of the four core histones: H2A, H2B, H3, and H4. Each core histone is composed of a globular domain and an unstructured amino-terminal tail of 25–40 residues. This *Correspondence to: Patrick A. Grant, University of Virginia Health System, Box 800733, Charlottesville, VA 22908 (e-mail:
[email protected]). 181 ADVANCES IN PROTEIN CHEMISTRY, Vol. 67
Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00
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unstructured tail extends through the DNA gyres and beyond the nucleosomal space (Grant, 2001). DNA packaged into nucleosomes is compacted as much as ten- to 20,000 fold from its naked state, and this topological complexity poses a significant obstacle to DNA-templated processes such as transcription, replication, repair, and recombination. Importantly, the structure of chromatin is dynamically regulated, permitting localized decondensation and remodeling that facilitates the progress of nuclear machinery (Grant, 2001). Chromatin structure is altered by multiple chromatin-modifying and chromatin-remodeling complexes. Chromatin-modifying complexes (modifiers) use posttranslational histone modifications such as acetylation, methylation, ubiquitination, and phosphorylation to dynamically regulate chromatin structure by establishing and maintaining distinct, functional domains (Fischle et al., 2003). The most well-studied modifiers are the histone acetylatransferase (HAT) complexes (Roth et al., 2001). HATs acetylate the "-amino groups of conserved lysine residues within the histone tails. The historic histone acetylation model is that acetylation contributes to the formation of a transcriptionally competent environment by ‘‘opening’’ condensed chromatin and allowing general transcription factors access to the DNA template (Grunstein, 1997; Struhl et al., 1998). Conversely, histone hypoacetylation, mediated by histone deacetylases (HDACs), is associated with transcriptional repression and silencing (Grunstein, 1997). Exactly how histone acetylation relieves chromatin condensation is poorly understood, but one hypothesis is that histone lysine acetylation may weaken adjacent nucleosome–nucleosome interactions (Luger and Richmond, 1998). Recently, striking advances in the field indicate that HATs play a much larger role in transcription than previously believed.
II. HATs and HAT Complexes: New Insights into HAT Regulation and Effects on Transcription The first type A (nuclear) HAT to be discovered was the Tetrahymena thermophila p55 protein (Brownell and Allis, 1995). This protein was found to have high sequence homology to the yeast transcriptional coactivator protein, Gcn5, indicating a direct relationship between histone acetylation and transcriptional activation (Grant and Berger, 1999). HATs also function in the cytoplasm, where acetylation events on newly synthesized histones occur before chromatin assembly. The first type B (cytoplasmic) HAT protein identified was Hat1 (Kleff et al., 1995; Parthun et al., 1996). Strikingly, the first HDAC was identified in mammals and found to be homologous to the yeast corepressor, Rpd3 (Taunton et al., 1996). Since these discoveries, a multitude of HAT enzymes have been described in
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numerous eukaryotic organisms and are implicated in a varietv of nuclear functions, many of which are transcriptional coactivators (see Table I) (Grant and Berger, 1999). Furthermore, it has become increasingly clear that these HAT enzymes function as part of larger multisubunit complexes. In fact, transcriptional events are known to involve exchange of complexes containing HDAC functions for those containing HAT activities (Roth et al., 2001) and vice versa (Deckert and Struhl, 2002). The focus of this review will be to describe the current understanding of HAT complex functions in transcription.
A. HAT Proteins Regulate Transcription The most well-studied HAT complexes include those containing the catalytic HAT subunit Gcn5, such as the SAGA, SLIK/SALSA, ADA, STAGA, and TFTC complexes (see Table I, Brand et al., 1999; Grant et al., 1997; Martinez et al., 1998; Pray-Grant et al., 2002; Sendra et al., 2000; Sterner et al., 2002a). In addition to Gcn5, these HAT complexes can contain up to 20 additional subunits apparently arranged in a modular fashion of subcomplexes within the larger complex. Why HAT complexes contain so many subunits is unclear, but recently, some subunits have been shown to be required for regulating Gcn5 HAT activity. For instance, recombinant Gcn5 can only acetylate nucleosomes as part of a HAT complex. In fact, specificity of lysine acetylation is different for Gcn5 when associated with different HAT complexes. For example, recombinant Gcn5 predominantly acetylates free histone H3 (K14) and to some extent H4 (K8/16), but not nucleosomal histones. In the context of the SAGA and SLIK complexes, Gcn5 acetylates nucleosomal H3 (K14/ 18 > 9/23) and H2B, whereas Gcn5 in the ADA complex acetylates H3 (K14/18) and H2B in nucleosomes (Grant and Berger, 1999; Zhang et al., 1998). Taken together, HAT complex subunits confer expanded substrate and lysine acetylation specificity and may play additional roles in regulating transcription at specific promoters (Grant et al., 1999). Studies of SAGA subunit function reveal that the Spt, Tral, and Ada subunits play roles in SAGA function beyond HAT activity. Spt (suppressors of Ty transcription) proteins 3, 7 and 8 interact with TATA boxbinding protein (TBP), linking SAGA function with that of the basal transcriptional machinery (Sterner et al., 1999). Tra1 (yeast TRRAP homologue) and Ada (alteration/deficiency in activation) proteins have been shown to interact with acidic activators, thus linking SAGA Gcn5 with upstream activators and basal transcriptional machinery (Brown et al., 2001; Drysdale et al., 1998; Utley et al., 1998). It is important to note that
HAT group
HAT complex
Catalytic subunit
Organism
Histone acetylation substrates
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Table I Histone Acetyltransferases and Transcription Function
SAGA SLIK STAGA ADA A2 TFTC PCAF Elongator HATB HAT-A3 HATB
Gcn5 Gcn5 Gcn5 Gcn5 Gcn5 Gcn5L PCAF Elp3 HPA2 HAT1 HAT1
Yeast Yeast Human Yeast Yeast Human Human Yeast Yeast Yeast–human Yeast–human
H3 and H2B H3 and H2B H3 H3 and H2B H3 and H2B H3 H3 H3, H4, H2B, H2A H4 H4 and H2A H4
Coactivator Coactivator Coactivator Coactivator Coactivator Coactivator Coactivator Transcriptional elongation Transcriptional elongation Histone deposition Histone deposition
MYST
NuA3 NuA4
ORC
Sas3 Esa1 Sas2 Tip60 MOF MOZ MORF HBO1
Yeast Yeast Yeast Human Fly Human Human Human
H3 H4 and H2A H2A, H3, H4 H3, H4, H2A H4 H2A, H2B, H3, H4 H3, H4 H3 and H4
Silencing Cell cycle, DNA repair Silencing DNA repair, apoptosis Dosage compensation Leukemogenesis Leukemogenesis Origin recognition/replication
TFIID TFIIIC Mediator
TAF1 p90/110/220 Nut1
Yeast–human Human Yeast
H3 and H4 H3, H2A, H4 H3, H4
TBP-associated factor RNAPII initiation Links activators with RNAPII
p300/CBP
p300/CBP
Human
H3, H4, H2B, H2A
Co-activator
Nuclear receptor
SRC1/ACTR SCR-3 TIF-2 ATF-2
Mice–human Mice–human Mice–human Yeast–human
H3 and H4
Steroid receptor co-activator Steroid receptor co-activator Steroid receptor co-activator DNA binding activator
MSL
Basal factors
H2B, H4
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SAGA subunits may also function in gene repression, as shown by the finding that Gcn5 HAT activity is required for repression of the yeast gene ARG1 in rich media (Ricci et al., 2002).
B. SAGA HAT Complex Consists of Modular Functional Domains A new model for SAGA function has emerged with recent evidence that SAGA, as well as other HAT complexes, may be composed of multiplefunctional domains. In brief, Gcn5 was shown to form a catalytic trimer with coactivators Ada2 and Ada3 in the context of the SAGA and ADA complexes. This catalytic trimer was demonstrated to have nucleosome acetylation activity (Balasubramanian et al., 2002). In fact, a minimal complex composed of these proteins has been described in yeast that can associate with nucleosomes (Grant and Berger, 1999) In keeping with the model of complex subunits conferring specificity of lysine acetylation, it was found that an Ada2-Gcn5 dimer (K14) and the Ada2-Gcn5-Ada3 trimer (K18 > K14 > K9) each have different lysine acetylation specificity (Balasubramanian et al., 2002). These indicate that certain SAGA subunits could play important roles in directing acetylation by Gcn5. Another functional domain of SAGA is comprised of a subset of TBP-associated factors (TAFs), originally described as components of the general transcriptionfactor complex TFIID. The SAGA complex contains histone-like TAFs: 5, 6, 9, 10, and 12. TAF 12 has been shown to be required for SAGA-mediated nucleosomal acetylation (Grant et al., 1998). Interestingly, the histone-like TAFs 6, 9, and 12, along with Ada1, another SAGA component, form a TAF octamer, similar in organization to the histone octamer (Selleck et al., 2001). This octameric structure can also be reconstituted from TAF components of TFIID; the role of this TAF octamer is currently under investigation. Collectively, the different functional components of SAGA participate in mechanisms of Gcn5 HAT regulation, implicating the importance of fine-tuning HAT activities. It is, therefore, important to further examine the interrelationships between these subcomplexes, which may yield significant insights into transcriptional regulation. Furthermore, these types of studies may be applicable to TFIID and perhaps other chromatin-modifying complexes.
C. The MYST Family of Acetyltransferases MYST acetyltransferase domains are named for its founding members MOZ (monocytic leukemia zinc finger protein) Ybf2/Sas3, Sas2, and Tip60 (Tat interacting protein). All are HATs, but they have varying
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functions implicated in transcriptional activation, silencing, development, and human disease. MOZ, when fused in frame to CBP, is associated with acute myeloid leukemias (Roth, 1996). Tip60 is a HAT capable of acetylating core histones (Kimura and Horikoshi, 1998). Esa1 (essential SAS-related acetyltransferase) is another MYST family member and is found in the yeast coactivator HAT complex NuA4 (Allard et al., 1999). Interestingly, the Drosophila melanogaster protein Chameau is a MYST HAT implicated in transcriptional repression of HOX genes, which regulate organism development, and the telomeric position effect, where genes located proximal to heterochromatic chromatin become silenced (Grienenberger et al., 2002). Similar data have been described in yeast with the HATs Ybf2/Sas3 and Sas2 (something about silencing 2) involvement in regulation of transcriptional silencing at HML, telomeres, and rDNA (Howe et al., 2001; John et al., 2000; Osada et al., 2001; Reifsnyder et al., 1996). More interesting is the finding that Sas2 antagonizes the yeast histone deacetylase Sir2, which together with other Sir proteins is involved in the nucleation of telomeric heterochromatin. A lysine acetylation gradient exists at telomeres where Sas2 hyperacetylates lysines until a boundary is reached that is regulated by deacetylation by Sir2 (Kimura et al., 2002; Suka et al., 2002). The MYST family also includes the HAT HBO1. This enzyme interacts with the ORC1 subunit of the human origin recognition complex (ORC; Iizuka and Stillman, 1999). This finding is interesting because the ORC is involved in both DNA replication and transcriptional silencing in yeast and D. melanogaster (Munshi et al., 2001). These examples support a role for HAT modifications through MYST acetyltransferases in both activating and silencing transcription, as well as offering clues about the mechanisms behind the establishment of transcriptional boundaries.
D. HAT Complexes are Regulated in Multiple Ways HAT complex subunits modulate HAT activity, and there is growing evidence that some subunits are posttranslationally modified. These modifications can potentially alter a subunit’s association with other proteins and, thus, alter subsequent functions of the subunit-binding partners (Roth et al., 2001). For example, the HAT CREB-binding protein (CBP) when phosphorylated exhibits increased HAT activity (Ait-Si-Ali et al., 1998, 1999, 2000). Conversely, modifications on subunits can down-regulate HAT activity, as seen when protein kinase C phosphorylates serine 89 of p300, inhibiting p300’s intrinsic HAT activity (Yuan et al., 2002). Inhibition also occurs in the case of DNA-dependent protein kinase, which
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phosphorylates the mammalian short-form of Gcn5, and consequently inhibits its HAT activity (Barlev et al., 1998). HAT proteins also mediate transcriptional regulation through modification of chromatin-related factors such as high-mobility-group (HMG) proteins. The mammalian HMG I/Y nucleates formation of an enhanceosome at the promoter/enhancer region of the interferon-beta (INF-) gene, leading to gene expression (Reeves and Beckerbauer, 2001). Formation of the enhanceosome is regulated through the acetylation of specific HMG I/Y lysine residues. CBP acetylation of HMG I/Y lysine 65 destabilizes the enhanceosome at the INF- locus. Conversely, Gcn5/PCAF acetylation of lysine 71 stabilizes the enhanceosome preventing CBP acetylation (Munshi et al., 2001). This evidence indicates that modification of chromatin-related factors, along with HAT subunits, can either potentiate or terminate transcription processes. In addition to the examples of enzymatic modifications directly affecting the catalytic mechanism of HATs, there is also the possibility for a model indicating that HAT complex subunit modifications affect protein–protein interactions that are mediated through special domains. These domains will be discussed later in this review, but it is interesting to note that these domain-containing proteins are found in both chromatinmodifying and chromatin-remodeling complexes. This observation might provide insight into the mechanisms by which chromatin-modifying and chromatin-remodeling complexes interact and cooperate to regulate transcription.
III. HAT Complexes Functionally Interact with Chromatin-Remodeling Complexes and Influence Transcription There are multiple examples of chromatin-modifying complexes found in association with chromatin-remodeling complexes (remodelers). SWI/SNF is a remodeler that uses adenosine triphosphate-hydrolysis to alter histone–DNA interactions, resulting in the histone octamer sliding along DNA, or resulting in a complete removal of the octamer from DNA (Vignali et al., 2000). Associations between chromatin modifiers and remodelers raise questions about functional interaction between complex types while pointing toward a model of sequential occupancy or cooperativity at specific promoters. The model of sequential occupancy attempts to answer the question of which type of complex, modifier or remodeler, is required first at a specific promoter to initiate transcription.
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A. Specific Genes Recruit HAT Complexes and Chromatin Remodelers Transcriptional regulation of the yeast genes INO1, HO, SUC2, and HIS3 has been found to be dependent on both modifiers and remodelers (Neely and Workman, 2002). Dependence on both types of chromatin-altering complexes might suggest a requirement for ordered recruitment of each type of complex at select promoters. In an ordered recruitment model, remodelers and modifiers could both interact with the same activator proteins in a particular order and, therefore, be recruited to the same promoters. For example, SAGA (modifier) and SWI/SNF (remodeler) both have been shown to interact with the acidic activators Gcn4, Swi5, Hap4, Pho4, and VP16 (Ikeda et al., 1999; Neely et al., 1999; Utley et al., 1998). New evidence indicates another model whereby modifiers may nucleate interaction platforms to recruit remodelers. This model was suggested by data in which histone acetylation by the HAT complexes SAGA and NuA4, at certain promoters, lead to an increased retention of SWI/SNF at those promoters (Hassan et al., 2001). Further support of this model is found in ordered recruitment of bromo domain– containing transcriptional complexes to the INF- promoter. In this case, acetylation of H4 K8-mediated recruitment of the SWI/SNF complex and acetylation of H3, K9, and K14 is critical for recruitment of TFIID, a bromo domain–containing complex (Agalioti et al., 2002). Conversely, there is also evidence that chromatin remodeling may precede HAT recruitment (Fry and Peterson, 2001).
B. Bromo Domain–Containing Proteins Function in Modifiers and Remodelers New evidence indicates that the major mediators in remodeler recruitment may be bromo and chromo domain–containing proteins. Bromo domains mediate protein binding to acetyl-lysines in histones and other proteins ( Jacobson et al., 2000), and chromo domains have been shown to bind methyl-lysines ( Jacobs et al., 2002). For instance, TAF1, a component of TFIID, contains two tandem bromo domains that bind selectively to multiply acetylated H4 peptides ( Jacobson et al., 2000). Furthermore, the Gcn5 bromo domain preferentially binds acetylated H4 K16 (Owen et al., 2000). Bromo domains are present in the SAGA subunits Spt7 and Gcn5 and in the SWI/SNF subunit Swi2/Snf2. The Gcn5 bromo domain has been found to be required for nucleosome remodeling and transcriptional activation (Syntichaki et al., 2000). It is therefore likely that Gcn5 may play two roles in transcriptional activation. The first role is the establishment
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Table II Specialized Domain-Containing Proteinsa SANT
Bromo
PHD
RING
Swi3 Ada2 N-CoR TFIIIB C-myb Rsc8 CoREST MTA-L1
Gcn5 PCAF hBRG1 Swi2 TAF1 Bdf1 Sth1 Rsc1 Rsc2 TIF1 TIF1 TIF1 Brahma Bonus hBrm CBP/p300 Spt7 hCcg1
Bre1 Rad5 Rad6 Rad18 Ur1 hRUL138 MAT1 Kap1 Tif1- Z Mel18 BRCA1 BARD1 mahoganoid NI RF CBP
Acf1 CBP P300 Kap1 ZMOX1-a SPBP AF10 Cti6 M153R MEKK1 HBXAP TAF3 PF1 TRIP-Br SHL PCCX1 PHF2
a
Acetyltransferase proteins are shown in bold.
of an ‘‘open’’ chromatin conformation via histone acetylation. The potential second role of Gcn5 is the maintenance of that open conformation via its bromo domain and the subsequent binding of acetylated lysines and stabilization of modifiers or remodelers (Hassan et al., 2001). Interestingly, a SAGA complex lacking the Gcn5 bromo domain has reduced nucleosomal HAT activity in vitro, yet a deleted Gcn5 bromo domain in the ADA complex retains nucleosome acetylation activity (Sterner et al., 1999). Table II provides a list of bromo domain–containing proteins, and Figure 1 illustrates a model of bromo domain–containing proteins in the establishment and maintenance of transcriptionally active chromatin. Bromo domains are found in multiple additional proteins implicated in transcriptional activation, including the RSC (remodels the structure of chromatin) complex proteins: Sth1, Rsc1, and Rsc2. Deletion of either bromo domain in double-bromo domain proteins Rsc1 and Rsc2 causes different phenotypic effects (Cairns et al., 1999). This finding indicates that all bromo domains are not alike and that they may mediate distinct biological functions even when found within the same protein.
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Fig. 1. Illustrates a model of processive nucleosomal acetylation via the Gcn5 bromo domain.
C. HAT Complex Proteins Contain RING, SANT, and PHD Domains Interestingly, most bromo domain–containing proteins also contain other functional domains. There are a growing number of proteins containing both bromo domains and other catalytic domains. Gcn5 and PCAF (p300/CBP-associated factor) contain bromo domains and HAT catalytic domains. Bromo domain proteins hBRG1, human brahma-related gene 1, and yeast Swi2 also contain chromatin remodeling ATPase domains. In addition to a bromo domain, TAF1 also has a kinase domain, as does Bdf1, a component of TFIID (Matangkasombut et al., 2000). TAF1 has also been reported to exhibit ubiquitin-activating/conjugating activity in D. melanogaster (Pham and Sauer, 2000). The p300/CBP family of proteins contain plant homeodomain (PHD), ZZ, or TAZ (transcriptional coactivator with PDZ-binding motif) domains, in addition to bromo domains (Marmorstein, 2001). Many bromo domain–containing proteins also contain metal binding domains. The functions of these domains are being investigated and may reveal insights into transcriptional regulation. Finding the bromo domain in so many chromatin-regulating proteins reveals a potential central regulation mechanism that mediates
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recruitment to acetyl-lysines or retention of modifiers and remodelers. In particular, bromo domain–containing proteins may help link specific enzymes, such as HATs or remodeler ATPases, to the proper substrate. In this way, bromo domain–containing proteins may also mediate progressiveacetylation on nucleosomal arrays. Figure 1 provides a model whereby an acetylated lysine could be bound by the Gcn5 bromo domain and result in the retention of Gcn5 allowing the local acetylation of additional lysines/histone tails along an array. Additional regulation can be mediated through other prominent domains, such as RING (really interesting, new gene) PHD, and SANT, found in components of many modifiers and remodelers. Table II illustrates the list of proteins containing these domains. The RING domain of Bre1 (sensitive to the drug brefeldin A) has been shown to be required for other modifications of histones: monoubiquintination of H2B and subsequent H3 K4 methylation (Hwang and Worman, 2002). These data indicate that the RING domain may play an important role in transcriptional regulation. PHD domains are found in many subunits of modifiers and remodelers, though the function of the domain itself remains unclear. New data indicate that the PHD domain is needed for HAT activity. As an example, the PHD domain of CBP has been shown to be required for CBP HAT activity (Kalkhoven et al., 2002). Interestingly, recent evidence shows that a novel PHD domain–containing protein, Cti6, is required for the recruitment of the SAGA HAT activity to the Cyc8-Tup1 corepressor complex at the GAL1 gene in yeast (Papamichos-Chronakis et al., 2002). Cyc8 (Ssn6) and Tup1 form a corepressor complex that, when tethered to DNA, inhibits transcription. Cti6 physically associates with both the Cyc8-Tup1 transcription corepressor and the SAGA coactivator complex, leading to derepression, indicating that Cti6 is needed for reactivation of certain silenced genes. This evidence indicates that PHD domains may be part of a mechanism for derepression through coactivator recruitment to promoters. SANT domains were named for the founding members: Swi3, Ada2, N-CoR, and TFIIB. These members are all transcriptional regulators. This domain is a putative DNA-binding motif similar to that of the myb-related proteins. SANT domains are found in both coactivator and corepressor complexes, indicating that this domain might play a role in both transcriptional activation and repression (Neely and Workman, 2002). Recent evidence indicates that the Ada2 SANT domain is required for histone tail binding and enzymatic catalysis of Gcn5 within Gcn5 containing HAT complexes (Boyer et al., 2002; Sterner et al., 2002b).
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IV. HAT Proteins Function in Regulating Transcriptional Elongation A. The HAT Elp3 is a Component of the Elongator Complex The role of HATs in transcriptional initiation is well characterized, yet recent data reveal that HATs also play important functions in transcriptional elongation. The conserved HAT protein Elp3 (elongator protein 3) is a subunit of the elongator complex, a component of RNA polymerase II (RNAPII). Elp3, in vitro, can acetylate the four core histones, indicating that HAT activity may be involved in the elongation process (Wittschieben et al., 1999). This finding proposes a model in which HATs function downstream of transcriptional initiation and in which RNAPII requires HAT modifiers to assist in transcription through condensed chromatin environments at gene coding regions (Workman and Kingston, 1998). Figure 2 illustrates a model of HAT protein association to RNAPII during transcriptional elongation. The yeast Elp3 null phenotype reveals late activation of the PHO5 promoter, indicating functional importance of Elp3 for elongation in vivo (Wittschieben et al., 2000). Recent data reveal that double mutants of CTK1, the gene encoding the kinase subunit of RNAPII carboxy-terminal kinase I (CTDK-I), and Elp3 are synthetically lethal. CTDK-1 phosphorylation of RNAPII is associated with transcriptional elongation. Interestingly,
Fig. 2. Illustrates the functional interactions of HAT complexes and remodelers implicated in facilitating transcriptional elongation.
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this lethality can be rescued by an ELP3 mutant capable of associating with the elongator complex, but conferring greatly reduced HAT activity ( Jona et al., 2001). These findings indicate that the functional overlap of CTDK-I and Elp3 is in the assembly of RNAPII elongating complexes. In addition, Elp3 has also been shown to be synthetically lethal with gcn5 knockout yeast strains, indicating functional overlap between the activities of transcription-related HAT complexes in vivo (Svejstrup, 2002). These data are further supported with the finding that ELP3 deletion is also synthetically lethal with deletion of the H3 tail (Wittschieben et al., 2000).
B. The HATs MOF, PCAF, and Sas3 Function in Transcriptional Elongation Recent studies provide a role for particular HAT proteins in transcriptional elongation. Male X-chromosome dosage compensation in D. melanogaster is accomplished by increasing gene transcription twofold (Park and Kuroda, 2001). This is mediated by MOF (males only on the first) HAT activity, whereby H4 lysine 16 is hyperacetylated (Hilfiker et al., 1997). An immunoreagent against H4, acetyl-lysine 16 revealed that acetylation on D. melanogaster genes is higher in the middle and 30 ends of coding regions than at promoter regions (Smith et al., 2001). These data indicate that acetylation, mediated by certain HATs, occurs on chromatin beyond the promoter regions of D. melanogaster genes (Hartzog et al., 2002). Furthermore, studies on human -globin locus transcription reveal hyperacetylation of H3 lysines over the length of the entire 100-kb locus (Hartzog et al., 2002). This study indicates that acetylation occurs within gene coding regions, not solely at gene promoters. Beyond the Elongator complex, there are other examples of HATs associating with transcriptional elongation machinery. In humans, the HAT protein PCAF (p300/CBP-associated factor) has been shown to physically associate with RNAPII (Cho et al., 1998). It is interesting to note that p300 interacts specifically with the nonphosphorylated, initiation-competent form of RNAPII, whereas PCAF interacts with the elongation-competent form of RNAPII (Cho et al., 1998). Yeast NuA3, an H3-specific HAT complex, interacts with the elongation factor Spt16 in vivo and in vitro (Hartzog et al., 2002). Spt16 has been shown to be a component of the remodeler FACT (facilitates activation on chromatin templates) (Orphanides et al., 1999). These data indicate that multiple HAT complexes, as well as remodelers, mediate important functions in transcriptional elongation.
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C. Transcriptional Elongation is Regulated by Other Chromatin Modifiers Transcriptional initiation has been shown to be regulated by multiple histone modifications (Grant, 2001). Though beyond the scope of this review, it is worth highlighting that recently evolving data indicate that transcriptional elongation may also be regulated by multiple posttranslational modifications. New data reveal that the Set2 histone methyltransferase associates with RNAPII during transcriptional elongation (Li et al., 2002). It has been further demonstrated that SET2-RNAPII association is modulated through interactions with RNAPII’s phosphorylated CTD domain (Li et al., 2003) and the Set2 WW protein domain (Li et al., 2002). Adding to the complexity of this model is striking homology data indicating that homologs of the HAT Elp3 may potentially exhibit histone demethylation activity (Chinenov, 2002). This indicates that different modifiers and remodelers may coordinately regulate transcriptional elongation.
V. The Histone Code: Insights into Epigenetic Regulation of Transcription A. The Histone Code Proposes Mechanisms of Transcriptional Regulation Transcriptional regulation may be conducted through mechanisms proposed by the histone code hypothesis, which states that specific combinatorial profiles of histone posttranslational modifications confer specific biological functions (Strahl and Allis, 2000). The most well-studied modifications to data are histone acetylation, methylation, ubiquitination, and phosphorylation; however, it is probable that additional modifications may also play important roles in epigenetic (beyond primary DNA sequence) regulation. The histone code hypothesis further proposes specific protein interactions between modified histone residues and specialized chromatin-interacting domains, such as the aforementioned bromo and chromo domains. In this way, it is believed that these histone tail-protein domain interactions facilitate events such as protein recruitment and enzyme and substrate interactions. Growing evidence strongly indicates that transcription is tightly regulated by dynamic interplay of histone modifications, the removal of specific modifications, and downstream recruitment of other chromatin-interacting proteins. Interestingly, recent studies indicate that regulation as proposed by the histone code may occur on the same or on different histones (Fischle et al., 2003).
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Histone tail and nontail domains contain many modifiable amino acid residues. For instance, serine residues are well-documented phosphorylation sites; lysine residues can be modified by acetylation, monoubiquitination, or mono-, di-, and trimethylation (Fischle et al., 2003); and arginine residues can be mono- or dimethylated by histone methyltransferases (Zhang et al., 2002). Some individual residues are substrates for multiple modifications. An example of this is H3 K14, which can be acetylated by the HATs Gcn5 and Sas3 or mono-, di-, or trimethylated by histone methyltransferases (Zhang et al., 2002). In either case, each type of modification relays distinct biological functions, while at the same time inhibiting another modification on that particular residue. As a consequence, some modifications are removed and new ones laid down. This dynamic interplay relies on histone demodifying enzymes and complexes. HDACs Hda1 and Rpd3 remove the acetyl moiety from acetylated lysines and thereby ‘‘reset’’ lysine substrates for subsequent remodification. In this way, HAT and HDAC activities contribute to the dynamic equilibrium of global histone acetylation (Robyr et al., 2002; Vogelauer et al., 2000). It has been proposed that histone deacetylation may ‘‘reset’’ lysine residues for future lysine methylation and that it possibly nucleates a proximal change from euchromatin to heterochromatin (Rice and Allis, 2001).
B. The Histone Code and the Endpoint to Signaling Pathways The histone code hypothesis is applicable to any chromatin-templated process and may describe the end point for many upstream signaling events, much like the signaling paradigm of receptor tyrosine kinases (Schreiber and Bernstein, 2002). Furthermore, chromatin modifier and remodeler complexes may be regulated by upstream signaling events and subsequently transmit signals for gene regulation. Evidence for this is found in the p42/p44 (ERK 1/2) and p38 mitogen-activated protein kinase (MAPK) pathways, which both induce phosphorylation of H3 (Cheung et al., 2000). In fact, MAPK-activated Rsk-2 kinase is directly involved in H3 phosphorylation in vivo (Sassone-Corsi et al., 1999). These data indicate a direct link between signal transduction and the generation of patterns of histone modification.
VI. HATs: New Insight into Transcription and DNA Repair DNA damage also signals to chromatin and has been studied in mammalian cells in which a histone variant of H2A, H2A.X, is rapidly phosphorylated at its C-terminal tail after exposure to ionizing radiation
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(Rogakou and Sekeri-Pataryas, 1999). Strikingly, the HAT Esa1, a component of the NuA4 HAT complex, is required for DNA double-strand break repair (Bird et al., 2002). It has not yet been shown what upstream signaling may regulate Esa1 HAT activity in DNA damage; however, there are multiple examples of histone acetylation linking transcription and DNA repair. For instance, Gcn5 has been shown to play a role in photoreactivation and nucleotide excision repair (Teng et al., 2002), and CBP/ p300 associates with thymine DNA glycosylase (Tini et al., 2002). In these cases, it is evident that multiple HAT proteins play a variety of functions from transcription to DNA repair.
VII. Conclusion and Perspectives HATs and histone acetylation have historically been implicated in transcriptional initiation; however, new evidence reveals additional important roles for HATs at multiple levels of transcriptional regulation. Specific HAT and HAT-associated proteins contain important functional domains: RING, SANT, PHD, bromo, and chromo domains, which confer varying levels of HAT regulation. These domains are found in many chromatinremodeling complexes and may serve as a central element in chromatinmodifying complex recruitment of chromatin-remodeling complexes. Furthermore, research on these specialized domains may lead to deeper insights into coordinated modifier and remodeler activity at many promoters. New evidence reveals a role for HATs in additional cellular processes beyond transcriptional intiation. HAT activity in transcriptional elongation likely facilitates RNAPII during promoter clearance and transcription through coding regions. Also, data indicate that HATs play an active role in DNA repair, revealing that modifiers may play an important part in any chromatin-templated process.
Acknowledgments We thank Anastasia Mitchell for stimulating discussions and thorough proofreading as well as Chris Gallo and present members of the Grant laboratory for helping us formulate some of the concepts presented here. We acknowledge the many colleagues and collaborators with whom we have worked or discussed ideas. Because of space limitations, not all the original work could be cited, and wherever possible we refer the reader to excellent reviews in the current literature. P.A.G. is the recipient of a Burroughs Wellcome Fund Career Award in Biomedical Sciences. Work in the laboratory is supported by NIH grant DK-58646.
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References Agalioti, T., Chen, G., and Thanos, D. (2002). Cell 111, 381–392. Ait-Si-Ali, S., Carlisi, D., Ramirez, S., Upegui-Gonzalez, L. C., Duquet, A., Robin, P., Rudkin, B., Harel-Bellan, A., and Trouche, D. (1999). Biochem. Biophys. Res. Commun. 262, 157–162. Ait-Si-Ali, S., Polesskaya, A., Filleur, S., Ferreira, R., Duquet, A., Robin, P., Vervish, A., Trouche, D., Cabon, F., and Harel-Bellan, A. (2000). Oncogene 19, 2430–2437. Ait-Si-Ali, S., Ramirez, S., Barre, F. X., Dkhissi, F., Magnaghi-Jaulin, L., Girault, J. A., Robin, P., Knibiehler, M., Pritchard, L. L., Ducommun, B., Trouche, D., and Harel-Bellan, A. (1998). Nature 396, 184–186. Allard, S., Utley, R. T., Savard, J., Clarke, A., Grant, P., Brandl, C. J., Pillus, L., Workman, J. L., and Cote, J. (1999). EMBO J. 18, 5108–5119. Balasubramanian, R., Pray-Grant, M. G., Selleck, W., Grant, P. A., and Tan, S. (2002). J. Biol. Chem. 277, 7989–7995. Barlev, N. A., Poltoratsky, V., Owen-Hughes, T., Ying, C., Liu, L., Workman, J. L., and Berger, S. L. (1998). Mol. Cell. Biol. 18, 1349–1358. Bird, A. W., Yu, D. Y., Pray-Grant, M. G., Qiu, Q., Harmon, K. E., Megee, P. C., Grant, P. A., Smith, M. M., and Christman, M. F. (2002). Nature 419, 411–415. Boyer, L. A., Langer, M. R., Crowley, K. A., Tan, S., Denu, J. M., and Peterson, C. L. (2002). Mol. Cell 10, 935–942. Brand, M., Yamamoto, K., Staub, A., and Tora, L. (1999). J. Biol. Chem. 274, 18285–18289. Brown, C. E., Howe, L., Sousa, K., Alley, S. C., Carrozza, M. J., Tan, S., and Workman, J. L. (2001). Science 292, 2333–2337. Brownell, J. E., and Allis, C. D. (1995). Proc. Natl. Acad. Sci. USA 92, 6364–6368. Cairns, B. R., Schlichter, A., Erdjument-Bromage, H., Tempst, P., Kornberg, R. D., and Winston, F. (1999). Mol. Cell 4, 715–723. Cheung, P., Allis, C. D., and Sassone-Corsi, P. (2000). Cell 103, 263–271. Chinenov, Y. (2002). Trends Biochem. Sci. 27, 115–117. Cho, H., Orphanides, G., Sun, X., Yang, X. J., Ogryzko, V., Lees, E., Nakatani, Y., and Reinberg, D. (1998). Mol. Cell. Biol. 18, 5355–5363. Deckert, J., and Struhl, K. (2002). Mol. Cell. Biol. 22, 6458–6470. Drysdale, C. M., Jackson, B. M., McVeigh, R., Klebanow, E. R., Bai, Y., Kokubo, T., Swanson, M., Nakatani, Y., Weil, P. A., and Hinnebusch, A. G. (1998). Mol. Cell. Biol. 18, 1711–1724. Fischle, W., Wang, Y., and Allis, C. D. (2003). Curr. Opin. Cell. Biol. 15, 172–183. Fry, C. J., and Peterson, C. L. (2001). Curr. Biol. 11, R185–R197. Grant, P. A. (2001). Genome. Biol. 2, 0003–0006. Grant, P. A., and Berger, S. L. (1999). Semin. Cell. Dev. Biol. 10, 169–177. Grant, P. A., Duggan, L., Cote, J., Roberts, S. M., Brownell, J. E., Candau, R., Ohba, R., Owen-Hughes, T., Allis, C. D., Winston, F., Berger, S. L., and Workman, J. L. (1997). Genes Dev. 11, 1640–1650. Grant, P. A., Eberharter, A., John, S., Cook, R. G., Turner, B. M., and Workman, J. L. (1999). J. Biol. Chem. 274, 5895–5900. Grant, P. A., Schieltz, D., Pray-Grant, M. G., Steger, D. J., Reese, J. C., Yates, J. R., 3rd, and Workman, J. L. (1998). Cell 94, 45–53. Grienenberger, A., Miotto, B., Sagnier, T., Cavalli, G., Schramke, V., Geli, V., Mariol, M. C., Berenger, H., Graba, Y., and Pradel, J. (2002). Curr. Biol. 12, 762–766. Grunstein, M. (1997). Nature 389, 349–352.
198
TOROK AND GRANT
Hartzog, G. A., Speer, J. L., and Lindstrom, D. L. (2002). Biochim. Biophys. Acta 1577, 276–286. Hassan, A. H., Neely, K. E., and Workman, J. L. (2001). Cell 104, 817–827. Hilfiker, A., Hilfiker-Kleiner, D., Pannuti, A., and Lucchesi, J. C. (1997). EMBO J. 16, 2054–2060. Howe, L., Auston, D., Grant, P., John, S., Cook, R. G., Workman, J. L., and Pillus, L. (2001). Genes Dev. 15, 3144–3154. Hwang, K. K., and Worman, H. J. (2002). Biochem. Biophys. Res. Commun. 293, 1217–1222. Iizuka, M., and Stillman, B. (1999). J. Biol. Chem. 274, 23027–23034. Ikeda, K., Steger, D. J., Eberharter, A., and Workman, J. L. (1999). Mol. Cell. Biol. 19, 855–863. Jacobs, S. A., Harp, J. M., Devarakonda, S., Kim, Y., Rastinejad, F., and Khorasanizadeh, S. (2002). Nat. Struct. Biol. 9, 833–838. Jacobson, R. H., Ladurner, A. G., King, D. S., and Tjian, R. (2000). Science 288, 1422–1425. John, S., Howe, L., Tafrov, S. T., Grant, P. A., Sternglanz, R., and Workman, J. L. (2000). Genes Dev. 14, 1196–1208. Jona, G., Wittschieben, B. O., Svejstrup, J. Q., and Gileadi, O. (2001). Gene. 267, 31–36. Kalkhoven, E., Teunissen, H., Houweling, A., Verrijzer, C. P., and Zantema, A. (2002). Mol. Cell. Biol. 22, 1961–1970. Kimura, A., and Horikoshi, M. (1998). Genes Cells 3, 789–800. Kimura, A., Umehara, T., and Horikoshi, M. (2002). Nat Genet 32, 370–377. Kleff, S., Andrulis, E. D., Anderson, C. W., and Sternglanz, R. (1995). J. Biol. Chem. 270, 24674–24677. Li, B., Howe, L., Anderson, S., Yates, J. R., 3rd, and Workman, J. L. (2003). J. Biol. Chem. 278, 8897–8903. Li, J., Moazed, D., and Gygi, S. P. (2002). J. Biol. Chem. 277, 49383–49388. Luger, K., and Richmond, T. J. (1998). Curr. Opin. Struct. Biol. 8, 33–40. Marmorstein, R. (2001). Cell. Mol. Life Sci. 58, 693–703. Martinez, E., Kundu, T. K., Fu, J., and Roeder, R. G. (1998). J. Biol. Chem. 273, 23781–23785. Matangkasombut, O., Buratowski, R. M., Swilling, N. W., and Buratowski, S. (2000). Genes Dev. 14, 951–962. Munshi, N., Agalioti, T., Lomvardas, S., Merika, M., Chen, G., and Thanos, D. (2001). Science 293, 1133–1136. Neely, K. E., and Workman, J. L. (2002). Mol. Genet. Metabol. 76, 1–5. Neely, K. E., Hassan, A. H., Wallberg, A. E., Steger, D. J., Cairns, B. R., Wright, A. P., and Workman, J. L. (1999). Mol. Cell 4, 649–655. Orphanides, G., Wu, W. H., Lane, W. S., Hampsey, M., and Reinberg, D. (1999). Nature 400, 284–288. Osada, S., Sutton, A., Muster, N., Brown, C. E., Yates, J. R., 3rd, Sternglanz, R., and Workman, J. L. (2001). Genes Dev. 15, 3155–3168. Owen, D. J., Ornaghi, P., Yang, J. C., Lowe, N., Evans, P. R., Ballario, P., Neuhaus, D., Filetici, P., and Travers, A. A. (2000). EMBO J. 19, 6141–6149. Papamichos-Chronakis, M., Petrakis, T., Ktistaki, E., Topalidou, I., and Tzamarias, D. (2002). Mol. Cell 9, 1297–1305. Park, Y., and Kuroda, M. I. (2001). Science 293, 1083–1085. Parthun, M. R., Widom, J., and Gottschling, D. E. (1996). Cell 87, 85–94. Pham, A. D., and Sauer, F. (2000). Science 289, 2357–2360. Pray-Grant, M. G., Schieltz, D., McMahon, S. J., Wood, J. M., Kennedy, E. L., Cook, R. G., Workman, J. L., Yates, J. R., 3rd, and Grant, P. A. (2002). Mol. Cell. Biol. 22, 8774–8786.
HAT PROTEINS CONTRIBUTE TO TRANSCRIPTIONAL PROCESSES
199
Reeves, R., and Beckerbauer, L. (2001). Biochim. Biophys. Acta 1519, 13–29. Reifsnyder, C., Lowell, J., Clarke, A., and Pillus, L. (1996). Nat. Genet. 14, 42–49. Ricci, A. R., Genereaux, J., and Brandl, C. J. (2002). Mol. Cell. Biol. 22, 4033–4042. Rice, J. C., and Allis, C. D. (2001). Curr. Opin. Cell. Biol. 13, 263–273. Robyr, D., Suka, Y., Xenarios, I., Kurdistani, S. K., Wang, A., Suka, N., and Grunstein, M. (2002). Cell 109, 437–446. Rogakou, E. P., and Sekeri-Pataryas, K. E. (1999). Exp. Gerontol. 34, 741–754. Roth, S. Y. (1996). Nat. Genet. 14, 3–4. Roth, S. Y., Denu, J. M., and Allis, C. D. (2001). Annu. Rev. Biochem. 70, 81–120. Sassone-Corsi, P., Mizzen, C. A., Cheung, P., Crosio, C., Monaco, L., Jacquot, S., Hanauer, A., and Allis, C. D. (1999). Science 285, 886–891. Schreiber, S. L., and Bernstein, B. E. (2002). Cell 111, 771–778. Selleck, W., Howley, R., Fang, Q., Podolny, V., Fried, M. G., Buratowski, S., and Tan, S. (2001). Nat. Struct. Biol. 8, 695–700. Sendra, R., Tse, C., and Hansen, J. C. (2000). J. Biol. Chem. 275, 24928–24934. Smith, E. R., Allis, C. D., and Lucchesi, J. C. (2001). J. Biol. Chem. 276, 31483–31486. Sterner, D. E., Belotserkovskaya, R., and Berger, S. L. (2002a). Proc. Natl. Acad. Sci. USA 99, 11622–11627. Sterner, D. E., Grant, P. A., Roberts, S. M., Duggan, L. J., Belotserkovskaya, R., Pacella, L. A., Winston, F., Workman, J. L., and Berger, S. L. (1999). Mol. Cell. Biol. 19, 86–98. Sterner, D. E., Wang, X., Bloom, M. H., Simon, G. M., and Berger, S. L. (2002b). J. Biol. Chem. 277, 8178–8186. Strahl, B. D., and Allis, C. D. (2000). Nature 403, 41–45. Struhl, K., Kadosh, D., Keaveney, M., Kuras, L., and Moqtaderi, Z. (1998). Cold Spring Harb. Symp. Quant. Biol. 63, 413–421. Suka, N., Luo, K., and Grunstein, M. (2002). Nat. Genet. 32, 378–383. Svejstrup, J. Q. (2002). Curr. Opin. Genet. Dev. 12, 156–161. Syntichaki, P., Topalidou, I., and Thireos, G. (2000). Nature 404, 414–417. Taunton, J., Hassig, C. A., and Schreiber, S. L. (1996). Science 272, 408–411. Teng, Y., Yu, Y., and Waters, R. (2002). J. Mol. Biol. 316, 489–499. Tini, M., Benecke, A., Um, S. J., Torchia, J., Evans, R. M., and Chambon, P. (2002). Mol. Cell 9, 265–277. Utley, R. T., Ikeda, K., Grant, P. A., Cote, J., Steger, D. J., Eberharter, A., John, S., and Workman, J. L. (1998). Nature 394, 498–502. Vignali, M., Hassan, A. H., Neely, K. E., and Workman, J. L. (2000). Mol. Cell. Biol. 20, 1899–1910. Vogelauer, M., Wu, J., Suka, N., and Grunstein, M. (2000). Nature 408, 495–498. Wittschieben, B. O., Fellows, J., Du, W., Stillman, D. J., and Svejstrup, J. Q. (2000). EMBO J. 19, 3060–3068. Wittschieben, B. O., Otero, G., de Bizemont, T., Fellows, J., Erdjument-Bromage, H., Ohba, R., Li, Y., Allis, C. D., Tempst, P., and Svejstrup, J. Q. (1999). Mol. Cell 4, 123–128. Workman, J. L., and Kingston, R. E. (1998). Annu. Rev. Biochem. 67, 545–579. Yuan, L., Soh, J., and Weinstein, I. (2002). Biochim. Biophys. Acta 1592, 205–211. Zhang, K., Tang, H., Huang, L., Blankenship, J. W., Jones, P. R., Xiang, F., Yau, P. M., and Burlingame, A. L. (2002). Anal. Biochem. 306, 259–269. Zhang, W., Bone, J. R., Edmondson, D. G., Turner, B. M., and Roth, S. Y. (1998). EMBO J. 17, 3155–3167.
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POSTTRANSLATIONAL MODIFICATIONS OF HISTONES BY METHYLATION By ADAM WOOD* AND ALI SHILATIFARDÀ1
*Department of Biochemistry, Saint Louis University School of Medicine, À St. Louis, Missouri; The Saint Louis University Cancer Center, Saint Louis University School of Medicine, St. Louis, Missouri
I. Introduction . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . II. Lysine Methyltransferases . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. SET Domain Containing Lysine-Specific Histone Methyltransferases . .. . . . . . B. Non-SET Domain Containing Lysine Methyltransferases. . . . . . . . . . . . . . . . .. . . . . . III. Arginine Methyltransferases . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . A. Structure . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . B. Catalytic Mechanism. . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . C. Identified Arginine Target Residues . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . IV. Epilogue . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . References .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . . . . . . . . . . . . . . . . . . . . .. . . . . .
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I. Introduction During development, cells become committed to different fates, in part through heritable, quasi-stable changes in gene expression. The mechanisms by which the 2-m-long DNA of eukaryotic organisms is packaged into the cell nucleus while remaining functional still remains not well understood. However, the last few years have been a watershed for understanding the first stage in this packaging process: the formation of the nucleosome core particle. Nucleosomes were first observed by viewing electron micrographs of lysed nuclei (Kornberg, 1974). The chromatin appears as a series of ‘‘beads on a string,’’ the beads being the individual nucleosomes and the ‘‘string’’ the linker DNA. Since the discovery of the ‘‘beads on a string,’’ it has been demonstrated that each nucleosome consists of eight core histone proteins (two each of H3, H4, H2A, and H2B; Luger et al., 1997). Each of the core histones contains a shared region of homology termed the histone fold, and it is this motif that allows for histone–DNA contacts and for dimerization of histones (Arents and Moudrianakis, 1995). Two H3/H4 heterodimers interact via a four-helix bundle formed between the two H3 histone folds, and this tetramer is then
1 Correspondence to: Ali Shilatifard, Saint Louis University School of Medicine, Department of Biochemistry, 1402 South Grand Blvd. St. Louis, MO 63104 (e-mail:
[email protected]).
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flanked by two H2B/H2A heterodimers with two four-helix bundles linking histones H2B and H4, thus bringing the core octamer together (Luger et al., 1997). The core histones are wrapped by 147 base pairs of DNA (1.65 turns around the histone octamer) in a left-handed superhelix, forming the intact nucleosome. The nucleosome structure is stabilized by about 142 hydrogen bonds between the DNA and the core histones, and several hydrophobic interactions are present between DNA and histones within the core (Luger et al., 1997). The core histones are rich in lysine and arginine residues, whose positive charge can form stable interactions with the negatively charged phosphodiester backbone of DNA. Taken together, these interactions allow for a highly stable complex to form between DNA and the core histone proteins. The nucleosome structure, when further stabilized by the linker histone H1, can compact the linear DNA about 30–40-fold, which is a significant reduction in length (Luger et al., 1997). This compaction affects the binding of nonhistone proteins, such as transcription factors, by restricting access to the binding sites within the DNA. Reduction in DNA length produced by histone-induced supercoiling in a nucleosome is significant and is an essential first step in the formation of higher-order chromatin structures. During the last decade, there has been an explosion of information regarding the role of nucleosomes in the regulation of gene expression (Workman and Kingston, 1998). Just as exciting has also been the growing possibility that the nucleosomes can transmit epigenetic information from one cell generation to the next (Jenuwein and Allis, 2001; Turner, 2002). The histone amino termini, or tails, are the sites of many of the covalent modifications that alter the nucleosome structure (Fig. 1). These tails extend away from the core of the nucleosome and are available for interactions with the DNA or with other proteins (Luger et al., 1997). There are multiple modification sites on each histone tail, and some amino acids in the histone tail can be modified in two or more ways (Schreiber and Bernstein, 2002; Turner, 2002; Figs. 1, 2). Because the sequence of the N-terminal tails is rich in arginine and lysine residues, the histone tails are very basic; covalent modification of these histone residues such as acetylation can alter the charge and structural properties of the nucleosome and can serve to restrict or allow access to the nucleosomal DNA. Recently, it has been demonstrated that modifications can also occur within the globular domain of histone H3. Gottschling and coworkers and Struhl and colleagues independently demonstrated that methylation on lysine 79 of histone H3, which occurs within the globular domain, restricts the binding of silencing proteins such as Sir2 to specific regions of the genome (Ng et al., 2002a; Van Leeuwen et al., 2002). The deletion of Dot1, which
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Fig. 1. Nucleosome structure and histone tail modification sites. (See Color Insert.)
result in the loss of histone H3 methylation on lysine 79, as will be discussed later, abolishes the interaction of the silencing proteins in the body of genes and causes silencing defects at the telomers (Ng et al., 2002a; Van Leewen et al., 2002). Several known covalent modifications have been observed so far that can modify the amino acid residues in the histone tails. There include acetylation, phosphorylation, ubiquitination, and methylation. Although some of these modifications have been known for many years, only recently have functional roles for these modifications begun to surface (Workman and Kingston, 1998). Each histone can undergo numerous modifications, and the combinatorial effect of these serves to elicit a multitude of different responses. This combinatorial modification of histone tails has been referred to as the ‘‘histone code’’ (Jenuwein and Allis, 2001; Strahl and Allis, 2000) and has been proposed to play a pivotal role in the regulation of gene expression (Fig. 3). Histone acetylation was one of the first posttranslational histone proteins to be described and demonstrated to function in the regulation of transcriptional activation (Braunstein et al., 1993). To date, this modification of histone is the one best characterized (Workman and Kingston, 1998). Acetylation of a lysine residue on a histone tail can neutralize the positive charge, thus weakening the interaction of the nucleosome with the DNA backbone. This then allows for the ‘‘remodeling’’ of the nucleosome so that the transcriptional machinery, as well as other proteins, can gain access to previously restricted sites within the DNA
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Fig. 2. Histone residues modified by methylation and the enzymes responsible for each.
(Kristjuhan et al., 2002). The acetylation of histone tail has come to be known as a hallmark of gene-specific transcriptional activation, and conversely, histone deacetylation, catalyzed by HDACs, play a pivotal role in transcriptional repression (Strahl and Allis, 2000; Utley et al., 1998; Wolffe, 2001; Zhou et al., 2002). HDACs are also found in large protein complexes that function specifically at promoters. Phosphorylation of histone tails has recently been discovered as another covalent modification of histones. The covalent modification of histones by phosphorylation has been demonstrated to be involved in DNA repair, chromosomal condensation, apoptotic signaling, and heat shock–induced pathways (Ballal et al., 1975; Berger, 2001; Hunt and Dayhoff, 1977; Nowak and Corces, 2000; Rogakou et al., 2000; Van Hooser et al., 1998). Interestingly, phosphorylation of serine 10 of histone H3, catalyzed by the protein kinases Ipl1, Rsk2, and Msk1 (Berger, 2001), also alter chromatin structure and function and affect transcriptional activation, condensation of chromosomes during mitosis and meiosis, and regulation of cell division.
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Fig. 3. General information for known SET domain and PRMT proteins (* undefined).
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Histone ubiquitination was discovered as a modification of histone H2A more than 20 years ago (Ballal et al., 1975; Hunt and Dayhoff, 1977). Only recently, though, has a functional role for the ubiquitination of histone proteins been described. In Saccharomyces cerevisiae, it has been demonstrated that Rad6 is involved in the attachment of ubiquitin to lysine 123 of histone H2B (Robzyk et al., 2000). This modification of histone H2B is required as a signal for histone methylation on lysines 4 and 79 of histone H3 (Dover et al., 2002; Ng et al., 2002a, 2002b; Sun and Allis, 2002). Rad6 has been demonstrated to be involved in several different pathways such as DNA repair (Jentsch et al., 1987), proteolytic degradation (Dohmen et al., 1991), and transcriptional silencing (Huang et al., 1997). Therefore, Rad6 requires an specific E3 ligase to direct it only to its role in transcription (Wood et al., 2003). Several lines of evidence have demonstrated that Bre1 is the E3 ligase for Rad6 in transcription. First, Bre1 is found in a macromolecular complex with Rad6; second, Bre1 is required for the ubiquitination of histone H2B in vivo, which seems to be a signal for methylation of histone H3 at its 4th and 79th lysines; third, Bre1 is required for telomeric silencing; fourth, Bre1 is recruited with Rad6 to a promoter; fifth, Bre1 is essential for the recruitment of Rad6 to chromatin at a promoter; and sixth, Bre1 is dedicated to the transcriptional regulatory role of Rad6. Much like histone ubiquitination, histone methylation was discovered several decades ago, but the biological significance of this modification remained elusive. However, its functional role in the regulation of gene expression is now becoming clearer. The attachment of methyl groups to histone proteins occurs predominantly at specific lysine or arginine residues on histones H3 and H4 (Jenuwein and Allis, 2001; Schreiber and Bernstein, 2002; Stallcup, 2001; Turner, 2002; Figs. 1, 2). This modification, unlike acetylation, phosphorylation, and ubiquitination, is stable. Although deacetylases, phosphatases, and deubiquitinating enzymes have all been described, as of yet there is no known demethylase that can function on methylated histone. We have recently demonstrated that an RNA polymerase II elongation factor, the Paf1 complex, is required for the recruitment of histone methyltransferases to the elongating RNA polymerase, and have therefore suggested that it is possible that modification by methylation may serve as a mark of transcriptional memory for transcribed genes. To date, two types of histone methylation have been observed: methylation of the "-amino group of lysine, and methylation of the N nitrogens of the arginine side chain guanido group (Aletta et al., 1998; Rea et al., 2000).
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II. Lysine Methyltransferases A. SET Domain Containing Lysine-Specific Histone Methyltransferases The first histone methyltransferase to be identified was the product of the Drosophila suppressor of position-effect variegation gene, Su(var)3-9, Clr4+ (cryptic locus regulator) in Schizosaccharomyces pombe, and SUV39H1 and SUV39H2 in human (Rea et al., 2000). Su(var)3-9 contains a SET domain, a highly conserved motif originally identified at the C-terminal ends of Su(var)3-9, enhancer of zeste (a member of the Polycomb family), and Trithorax, three gene-regulatory factors in Drosophila (Dorn et al., 1993; Jones and Gelbart, 1993; Tschiersch et al., 1994; reviewed in Alvarez-Venegas and Avramova, 2002). In yeast, the product of the SET1 gene is found in a macromolecular complex called COMPASS (complex of proteins associated with Set1; Miller et al., 2001; Nagy et al., 2002; Roguev et al., 2001). COMPASS can methylate the fourth lysine of histone H3 and thereby effect regulation of transcription of genes located near chromosome telomeres. To date, COMPASS is the only histone methyl transferase that was biochemically isolated and that functions in a macromolecular complex (Krogan et al., 2002). The yeast Set1 protein alone is not capable of methylating histone H3 and requires the presence of other components of COMPASS for its enzymatic function. It has been proposed that components of COMPASS may function either in properly positioning the catalytic domain of Set1 or in proper substrate recognition by Set1 (Krogan et al., 2002). On the basis of homology of the conserved SET domains, this family of proteins is classified into four families: Su(var)3-9, E(z), TRX, and ASH1 (Alvarez-Venegas and Avramova, 2002; Jenuwein et al., 1998). Since its initial discovery, the SET domain has been described as part of several proteins involved in modulating gene activity in eukaryotes ranging from yeast to humans (Alvarez-Venegas and Avramova, 2002; Jenuwein et al., 1998; Nislow et al., 1997). This modulation happens via the histone methyltransferase activity found in many SET domain proteins and the complexes containing them. These SET domain-containing protein complexes have a number of identified lysine residue targets, and as we will describe later, the methylation of each different residue can elicit an array of different cellular responses. Because of their fairly recent discovery, only a few SET domain-containing enzymes and their target residues have been identified and crystallized (Jacobs et al., 2002; Min et al., 2002; Wilson et al., 2002; Zhang et al., 2002). To date, all of the characterized SET containing proteins have been
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demonstrated to have a highly conserved active site and a binding site for S-adenosyl-L-methionine (SAM domain).
1. Structure The histone methltransferase activity (HMTase) of the SET-domaincontaining protein resides within the SET domain. The SET domain itself is approximately 130 amino acids in length (Fig. 4; Jenuwein et al., 1998). Although some areas of the sequence are highly conserved between species, others are much more varied. The pre-SET and post-SET domains can differ in sequence and structure between different HMTases. However, members of the same SET domain protein families have high
Fig. 4. Sequence alignment of some known lysine methyltransferases. The residues involved in the catalytic domain ‘‘knot’’ are indicated by red bars. The conserved active site tyrosine residue is denoted by *. (See Color Insert.)
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homology in these pre-SET and post-SET domains. In each case, however, the pre-SET and post-SET domains are required for methyltransferase activity (Jacobs et al., 2002; Min et al., 2002; Rea et al., 2000; Wilson et al., 2002; Zhang et al., 2002). In many cases, these pre-SET and post-SET domains intertwine with the SET domain itself to alter the tertiary structure of the enzymes (Min et al., 2002). These variations lend different site specificities to each enzyme for methyl group attachment. However, the catalytic core of the enzyme and the SAM substrate binding site remain highly conserved. In the Su(var)3-9 and E(z) families of HMTases, the pre-SET region is rich in cysteine residues that form a triangular zinc cluster that acts to bind zinc ions very tightly, an interaction that serves to stabilize the structure (Min et al., 2002). This stabilization and positioning of the zinc cluster is essential for activity, as mutations in conserved residues that serve to position the cluster result in an inactive enzyme (Min et al., 2002). Enzymes containing this zinc cluster are also sensitive to metal chelating agents such as EDTA (Min et al., 2002; Zhang et al., 2002). Although some TRX family members also contain cysteine-rich regions N-terminal to the SET domain, these regions are at a considerable distance from the SET domain itself and may serve other functions. The members of the Su(var)3-9 family of HMTs are the only SET-domain-containing enzymes shown to have a cysteine-rich post-SET domain. Apart from the zinc clusters, many of the identified HMTases have similarities in the region N-terminal to the SET domain. The crystal structure of SET7/9 reveals the presence of an acidic groove on the pre-SET domain surface of the protein, which extends to the SET domain itself (Wilson et al., 2002). This groove may serve as the binding site for the basic N-terminal tails of histone proteins, and mutations of residues within the groove result in aberrant substrate binding. The place in which pre-SET groove intersects the SET domain also happens to be where the adenine moiety of AdoMet binds (Wilson et al., 2002). However, it should be noted that several other grooves on the surface of the SET domain surface approach the AdoMet binding site and that, as of yet, it is unclear which is responsible for histone tail binding. Although the adenine ring of AdoMet makes several contacts with the enzyme, it appears that the sulfonium component does not interact with the binding site (Wilson et al., 2002). However, the ribose ring forms Van der Waals interactions with a conserved residue in the core of the SET domain, as will be discussed later. The rest of the AdoMet binding site is a region of high sequence conservation, and the contacts between the site residues and AdoMet rely solely on hydrogen bonding. In fact, the amine nitrogen of AdoMet is hydrogen bonded to a highly conserved histidine residue on the ‘‘loop’’ of the active site knot (discussed later), and mutation of this residue abolishes enzyme activity (Jacobs et al., 2002; Min et al., 2002; Wilson et al., 2002).
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The catalytic core of the SET domain is rich in -strands that comprise several regions of -sheets. The actual number of -strands may vary between different enzymes (eight and nine in Clr4 and SET7/9, respectively), but the number of alpha helices remains constant at two. In many cases, -strands found in the pre-SET region will form -sheets with the -strands of the SET domain, imparting slight variations to the SET domain structure. These small changes alter the target residue site specificity for methylation and allow the SET domain methyltransferases to target many different residues. This interplay between the pre-SET domain and the catalytic core is critical for enzyme function, and it has been demonstrated that both the pre-SET and post-SET regions that flank the SET domain are required for histone methyltransferase activity. The C-terminus region of the SET domain makes an unusual turn that threads back through a loop formed between a -strand and an alpha helix at the core of the domain to form a ‘‘knot’’ (Jacobs et al., 2002). This knot structure is the catalytic core of the enzyme. Conserved residues in the loop and C-terminal ‘‘thread’’ combine to establish a hydrophobic core in the knot, which serves to stabilize the structure. Further reinforcement of the active site structure is achieved by hydrogen bonding between several conserved residues within the knot (Fig. 4; Jacobs et al., 2002; Min et al., 2002; Wilson et al., 2002; Zhang et al., 2002). Both the hydrophobic core of the active site and the residues involved in the hydrogen bonding constitute regions of high sequence conservation. This ‘‘knotted’’ catalytic core is unique to SET domain containing enzymes and does not resemble any other previously characterized methyltransferases that use AdoMet as a methyl group donor. Within this core, AdoMet and the substrate lysine are brought into close proximity. The core of each SET domain protein contains a completely conserved tyrosine residue, which is critical to the active site function. The conserved tyrosine forms Van der Waals interactions with the ribose of AdoMet and also serves to deprotonate the "-amino group of the target lysine residue (Jacobs et al., 2002). This is critical for activity, as mutation of the tyrosine results in an inactive enzyme (Wilson et al., 2002). Note once again that the conserved residues that are critical for the catalytic ability of the SET domain are derived from the knot itself; thus, the knot structure is crucial for the formation of the active site and the activity of the SET domain itself.
2. Catalytic Mechanism of the SET Domain When AdoMet and the lysine of the substrate histone tail are bound and properly oriented in the catalytic pocket of the SET domain, the conserved tyrosine residue acts to deprotonate the "-amino group of the lysine
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Fig. 5. Catalytic data of characterized histone methyltransferases (* undefined value).
residue (Trievel et al., 2002; Fig. 6). This primes the lysine side chain to make a nucleophilic attack on the methyl group of the AdoMet molecule. The methyl group is transferred to the lysine side chain, resulting in the formation of methylated histone and the byproduct of AdoMet demethylation, AdoHcy (Jacobs et al., 2002; Min et al., 2002; Wilson et al., 2002; Zhang et al., 2002). In vitro catalytic data have been obtained for only a few of the characterized SET domain containing proteins. These data are summarized in Fig. 5. Most of the examined HKMTs have optimal methyltransferase activity between a pH of 8 and 10. The mechanism for methyl group attachment to lysine residues is illustrated in Fig. 6.
3. Identified Lysine Target Residues To date, the lysine residue methylation of histone proteins by SETdomain-containing enzymes occurs specifically at Lys4, Lys9, Lys27, and Lys36 of histone H3, and Lys20 of H4, and each lysine residue can be methylated up to three times. As more SET-domain proteins are characterized, it is possible that more histone and nonhistone targets will be found as well. For the residues that have been established as targets for KHMTs, though, a number of biological consequences for their methylation such as transcriptional silencing and activation, have been observed (Fig. 3). What is interesting about some of these methyltransferases is that they are capable of methylating the same lysine residue once, twice, or three times. Studies performed in Kouzarides’s laboratory have demonstrated that dimethylated K4 of histone H3 is associated with active euchromatic regions, but not in silent heterochromatic sites (Santos-Rosa et al., 2002). They have also demonstrated that Set1 subunit of COMPASS can catalyze dimethylation and trimethylation of K4. This correlates with the activity of many genes. They have also developed antibodies that discriminate between the di- and trimethylated state of lysine 4 of histone H3 and have demonstrated that dimethylation occurs at both inactive and active euchromatic genes, whereas trimethylation is present exclusively at active genes.
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Fig. 6. Mechanism of methyl group transfer to lysine residues.
What remains to be determined is how COMPASS dimethylates at some sites and trimethylates at other sites. It is possible that either the subunits of COMPASS or other proteins within the yeast genome can participate in this process. Lysine 9 of histone H3 is the target of many SET-domain-containing enzymes (Fig. 2). Most of the HKMTs that methylate this residue belong to the Su(var)3-9 family. In Drosophila and humans, methylation of this lysine provides a binding site for the chromo domain of heterochromatin protein 1 (HP1; Aagaard et al., 2000; Loyola et al., 2001). Work done in Drosophila has demonstrated that the interaction of HP1 and methylated
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H3 K9 is localized to regions of heterochromatin on polytene chromosomes, and alteration of HP1 recruitment affects heterochromatic gene silencing (Jacobs et al., 2001). Methylation of H3 lysine 9 then may serve to mediate heterochromatin assembly and gene silencing through its recruitment of HP1 (Hall et al., 2002; Schotta et al., 2002). The methylation of lysine 4 of histone H3 has also been extensively studied. Although the methylation of lysine 9 in higher eukaryotes is strongly related to transcriptional repression, methylation of histone H3 lysine 4 seems to antagonize this effect. In mammals, SET7/9 specifically methylates lysine 4, and this methylation coincides with transcriptional activation (Nishioka et al., 2002). SET7/9 also lacks the cysteine-rich domains found in lysine 9-specific methyltransferases, once again demonstrating the role of the pre-SET and post-SET domains in substrate specificity (Rea et al., 2000; Wilson et al., 2002). In S. cerevisiae, the methylation of K4 is carried out by COMPASS, a large SET-domain-containing protein. In contrast to the Lys4 methylation in higher eukaryotes, SET1-dependent methylation is involved with transcriptional repression at the telomeres and with mating type and rDNA loci (Bryk et al., 2002; Krogan et al., 2002; Miller et al., 2001). Recently, it was demonstrated that methylation of lysine 79 of histone H3 can regulate telomere-associated gene silencing by restricting the binding of silencing proteins to the telomeres and other ‘‘silenced’’ regions of chromatin (Van Leeuwen et al., 2002). It is also very possible that methylation of histone H3 on lysine 4 may function via a similar mechanism.
B. Non-SET Domain Containing Lysine Methyltransferases A novel histone methyltransferase was recently described as having specificity for lysine 79 of histone H3 (Ng et al., 2002a; Van Leeuwen et al., 2002). This enzyme, named Dot1 (disruptor of telomeric silencing 1) was originally discovered based on a high–copy suppressor screen of telomeric silencing (Singer et al., 1998). It was recently demonstrated that the target—a lysine residue by Dot1—is not found in the tail region of the histone but, rather, in the globular core of the histone itself. The residues are located on the top and bottom of the nucleosome core (Van Leeuwen et al., 2002). To date, Dot1 is the only enzyme known to methylate residues within the histone core. In vitro assays have revealed another interesting characteristic of Dot1. Almost all other HMTs can methylate free histone, some even being able to methylate peptides having homology to their respective target sequence. Dot1, however, will only methylate if the nucleosome is intact. Most interestingly, it was recently demonstrated that
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histone methylation by COMPASS requires histone ubiquitination by Rad6 and its E3 ligase Bre1 (Ng et al., 2002a, 2002b; Wood et al., 2003). This modification of histone H2B by ubiquitination is also a required modification for Dot1’s function: linking the molecular function of a non-SET domain and a SET domain containing methyltransferase.
1. Structure Dot1, unlike other known lysine methyltransferases, does not contain a SET domain. Rather, the methyltransferase fold resembles those of known arginine methyltransferases. In fact, many of the conserved residues found in PRMTs that are important for methylation activity are found in Dot1, and like the other known HMTs, Dot1 uses AdoMet as a methyl donor. Very recently, the crystal structure of human Dot1 in complex with AdoMet was resolved (Min et al., 2003). The N terminus of hDot1 contains the active site, and the N-terminal and C-terminal domains of the Dot1 catalytic domain are linked by a loop that also serves as part of the AdoMet binding site. The loop, and thus the SAM binding site, is highly conserved between Dot1 and other AdoMet binding proteins. Mutations within this conserved domain disrupt AdoMet binding and methyltransferase activity. The SAM binding site also contains a channel positioned near the methyl group, possibly for the binding of substrate lysine residues. Mutations within this channel do in fact abolish methyltransferase activity without affecting the overall structure of the protein (Min et al., 2003). The structure of the C terminus of Dot1 was not resolved. However, because of its distance from the active site, it is believed to be important for the substrate specificity and binding of Dot1. This region carries a positive charge, which would allow for a favorable interaction with the negatively charged backbone of DNA (Min et al., 2003). This may also explain why Dot1 is only able to methylate histone H3 in the context of an intact nucleosome.
2. Identified Lysine Target Residues The methylation of H3 K79 catalyzed by Dot1 is important for silencing at the telomeres and is believed be involved in the recruitment of Sir2 and Sir4 to these regions of DNA. A proposed mechanism is that histone methylation by Dot1 restricts Sir protein binding to the telomeres, and loss or overexpression of Dot1 function allows for promiscuous binding of the Sir proteins to chromatin within the genome, effectively titrating them away from the telomeres (Ng et al., 2002a; Van Leeuwen et al., 2002). The loss of the Sir proteins from the telomeres confers silencing defects to Dot1 deficient strains, and thus, methylation of Lysine 79 serves to restrict silencing to specific regions of the genome.
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III. Arginine Methyltransferases The methylation of arginine residues of histone proteins has also received much attention in recent years. However, not much is known about the biological significance of this process as of yet. Recent experiments have identified three distinct types of methylation that occur at arginine residues on histone tails along with two types of PRMTs that accomplish this task. To date, the three known products of arginine methylation are NG-monomethylarginine, NGNG-symmetric dimethylarginine (in which both guanido nitrogens are methylated), or NGN’G-asymmetric dimethylarginine (in which only one guanido nitrogen receives two methyl groups; McBride and Silver, 2001; Stallcup, 2001). The two types of PRMTs are classified by the type of methylation they accomplish. Type I PRMTs (PRMT1, PRMT3, CARM1/PRMT4, and Rmt1/Hmt1) produce monomethylarginine and asymmetric dimethylarginine (Chen et al., 1999; Frankel and Clarke, 2000; Gary et al., 1996; McBride et al., 2000) whereas type II methyltransferases (JBP1/PRMT5), have the ability to form monomethyl or symmetric dimethylarginine (Branscombe et al., 2001). These differences arise from variations in the arginine binding pocket, which can accommodate a previously methylated guanido group (type II) and then allow the other to be modified (symmetric), or a pocket that only allows one of the guanido nitrogens to be modified (asymmetric type 1; Branscombe et al., 2001). Like the SET domain containing methyltransferases, PRMTs require AdoMet as a cofactor and methyl group donor. The PRMTs identified so far exist as members of multisubunit complexes, the formation of which is critical for in vivo methyltransferase activity (Teyssier et al., 2002). Many of the arginine methyltransferases also form homo-dimers or homooligomers, a step that is required for their catalytic activity (Weiss et al., 2000). This need to oligomerize has not been demonstrated for any known lysine methyltransferases. The identities of several protein arginine methyltransferases are now known, but only a few have been shown to have specificity for histone proteins. The mammalian PRMT1, JBP1, and CARM1, as well as the Saccharomyces Rmt1, have histone methyltransferase activity (McBride and Silver, 2001). However, the catalytic mechanism for the methyl group transfer as well as the makeup of the active sites of PRMTs differ somewhat from SET domain proteins.
A. Structure The catalytic domain of the PRMTs is highly conserved and consists of about 310 amino acids constituting a SAM binding domain and substrate
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binding domain (Branscombe et al., 2001; Weiss et al., 2000; Zhang et al., 2000). In addition to this conserved region, each PRMT has a unique region N-terminal to the core. Although not much is known about the function of these unique domains, it is believed that they are involved in substrate specificity and targeting. The N-terminal region of the core contains a mixed / SAM binding and methyltransferase domain. The mixed / SAM binding domain, known as the Rossman fold (Rossmann et al., 1974), is conserved among many known methyltransferase structures. Within this domain, SAM makes extensive contacts with several highly conserved residues in the I and post-I motifs (Fig. 7). The methionine group of AdoMet interacts with a conserved arginine and aspartate within the catalytic core (Zhang et al., 2000). Also, the ribose ring makes Van der Waals interactions with a glycine found in the loop between the first beta strand and the second
Fig. 7. Sequence alignment of known protein arginine methyltransferases (PRMTs). Sequences essential for AdoMet and substrate binding are indicated by blue and red bars. (See Color Insert.)
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helix of the core (Zhang et al., 2000). One of the most critical interactions for PRMT activity concerns the orientation of AdoMet in the binding pocket. The AdoMet molecule is secured in this pocket by a hydrophobic interaction between the adenine ring and the phenyl ring of a phenylalanine from the N terminus of the core (Zhang et al., 2000). Aside from the conserved N-terminal region of the core found in many methyltransferases, PRMTs contain two additional -helices that precede the mixed / structure (Zhang et al., 2000). One of these helices contains the phenylalanine residue that secures AdoMet in the binding site, indicating that these extra helices may allow access for AdoMet and AdoHcy binding and exchange (Zhang et al., 2000). Also contained in the AdoMet binding region is a 12-residue loop, termed the double-E loop, which contains two completely conserved glutamates that are part of the active site (Zhang et al., 2000). The C terminus of the core contains a -barrel believed to be involved in substrate binding. Within this barrel is a highly conserved loop (named the THW loop for the conserved residues at its center), which is in very close proximity to the double-E loop of the N-terminal core (Zhang et al., 2000). These two loops are the most conserved regions of the enzymes and constitute the active site. The loops are further stabilized by a region of -sheet that makes several contacts with the loops, forming a ‘‘floor’’ for the active site (Zhang et al., 2000). These contacts and the residues involved are absolutely conserved between known histone PRMTs (Fig. 7), and are essential for the formation of the active site. Also contained in the -barrel structure of many PRMTs is a helix-turnhelix ‘‘antenna’’ motif that is involved in the formation of PRMT homodimers or homo-oligomers (Weiss et al., 2000). This dimerization usually occurs via hydrophobic contacts between the helix-turn-helix motif of one monomer and the N-terminal SAM binding domain of another (Weiss et al., 2000; Zhang et al., 2000). Several PRMTs have been shown to form either dimers or oligomers through this interaction, a process that is essential to their methyltransferase activity (Teyssier et al., 2002).
B. Catalytic Mechanism The conserved glutamates that give rise to the double-E loop interact with the guanido nitrogens of the target arginine residue. This interaction serves to redistribute the positive charge of the -nitrogens, leading to the deprotonation of one group and to priming it as a nucleophile (McBride and Silver, 2001). The deprotonated nitrogen makes a nucleophilic attack on the nearby methyl group of AdoMet. As mentioned earlier, differences between the type I and type II PRMTs determine the next methylation
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step, either catalyzing the dimethylation of one nitrogen or allowing the symmetric methylation of both guanido groups (Branscombe et al., 2001). In both types of enzymes, however, the proton stripped from the nitrogen is dispersed through a histidine–aspartate proton relay system and released into the surrounding matrix (Fersht and Sperling, 1973). The residues involved in this mechanism are completely conserved in PRMTs and are essential to the function of the enzymes. This importance is underscored by the finding that mutation of either glutamate in the double-E loop results in an inactive enzyme.
C. Identified Arginine Target Residues The known histone arginine methylation targets known so far are R17 and R26 of histone H3, and R3 of H4. JBP1/PRMT5 has been shown to methylate histone H2A and H4, although the sites of these modifications remain unknown (Pollack et al., 1999). CARM1 (cofactor associated arginine methyltransferase 1) is the enzyme responsible for the methylation of R17 and R26 of histone H3. Experiments have shown that CARM1 is recruited to the promoter regions of specific genes, and the ensuing methylation of histone arginine residues in these regions coincides with transcriptional activation (Chen et al., 1999; Schurter et al., 2001). Thus, CARM1 is recognized as a transcriptional coactivator. Histone H4 is the substrate for the type I enzymes Rmt1 (yeast) and PRMT1 (mammals). Both PRMTs methylate R3 in the histone tail. In mammals, this modification also results in transcriptional activation, as R3 methylation of histone H4 results in subsequent acetylation of the H4 tail (Wang et al., 2001). It should also be noted that both CARM1 and PRMT1 have other substrates besides histone proteins, and both of these enzymes have been shown to function as coactivators of nuclear hormone receptors, further reinforcing their role in transcriptional activation. As mentioned before, Janus kinase-binding protein 1 (JBP1/PRMT5) has been shown to methylate histones H2A and H4. Unfortunately, the sites of these modifications are as of yet unknown. However, it is known that JBP1 is the only PRMT yet capable of performing symmetric dimethylation of substrate arginine residues (Branscombe et al., 2001). It will be interesting to see the functional significance of this modification.
IV. Epilogue Histone modifications by methylation have been demonstrated to play a pivotal role in the regulation of chromatin dynamics and gene expression. Several proteins to date have been identified as functioning as histone
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methyltransferases with different substrate specificity. The wealth of other uncharacterized Set-domain containing proteins in the database has set the stage for characterization of the role of such enzymes. What remains unclear? Are histone proteins the only substrate for these Setdomain-containing proteins? Furthermore, we do not know how monomethylation, dimethylation, and trimethylation function of such methyl transferases are molecularly regulated. Most important, what are the biological consequences of histone or protein methylation in developmental regulation or even in pathogenesis of human diseases? What is clear is that the next several years will also bring a watershed of information regarding the molecular regulation and the consequences of protein modification by methylation.
References Aagaard, L. et al. (2000). Mitotic phosphorylation of SUV39II1, a novel component of active centromeres, coincides with transient accumulation at mammalian centromeres. J. Cell. Sci. 113(Pt 5), 817–829. Aletta, J. M., Cimato, T. R., and Ettinger, M. J. (1998). Protein methylation: a signal event in post-translational modification. Trends Biochem. Sci. 23(3), 89–91. Alvarez-Venegas, R., and Avramova, Z. (2002). SET domain proteins of the Su(var) 3-9, E(z) and trithorax families. Gene 285(1–2), 25–37. Arents, G., and Moudrianakis, E. N. (1995). The histone fold: a ubiquitous architectural motif utilized in DNA compaction and protein dimerization. Proc. Nat. Acad. Sci. USA 92(24), 11170–11174. Ballal, N. R. et al. (1975). Changes in nucleolar proteins and their phosphorylation patterns during liver regeneration. J. Biol. Chem. 250(15), 5921–5925. Berger, S. L. (2001). An embarrassment of niches: the many covalent modifications of histones in transcriptional regulation. Oncogene 20(24), 3007–3013. Branscombe, T. L. et al. (2001). PRMT5 (Janus kinase-binding protein 1) catalyzes the formation of symmetrie dimethylarginine residues in proteins. J. Biol. Chem. 276(35), 32971–32976. Braunstein, M. et al. (1993). Transcriptional silencing in yeast is associated with reduced nucleosome acetylation. Genes Dev. 7(4), 592–604. Bryk, M. et al. (2002). Evidence that Set1, a factor required for methylation of histone H3, regulates rDNA silencing in S. cerevisiae by a Sir2 independent mechanism. Current Biol. 12(2), 165–170. Chen, D. et al. (1999). Regulation of transcription by a protein methyltransferase. Science 284(5423), 2174–2177. Dohmen, R. J. et al. (1991). The N-end rule is mediated by the UBC2(RAD6) ubiquitin conjugating enzyme. Proc. Natl. Acad. Sci. USA 88(16), 7351–7355. Dorn, R. et al. (1993). The enhancer of position-effect variegation of Drosophila, E(var) 3-93D, codes for a chromatin protein containing a conserved domain common to several transcriptional regulators. Proc. Natl. Acad. Sci. USA 90(23), 11376–11380. Dover, J. et al. (2002). Methylation of histone H3 by COMPASS requires ubiquitination of histone H2B by Rad6. J. Biol. Chem. 277(32), 28368–28371.
220
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Fersht, A. R., and Sperling, J. (1973). The charge relay system in chymotrypsin and chymotrypsinogen. J. Mol. Biol. 74(2), 137–149. Frankel, A., and Clarke, S. (2000). PRMT3 is a distinct member of the protein arginine N-methyltransferase family. Conferral of substrate specificity by a zinc-finger domain. J. Biol. Chem. 275(42), 32974–32982. Gary, J. D. et al. (1996). The predominant protein-arginine methyltransferase from Saccharomyces cerevisiae. J. Biol. Chem. 271(21), 12585–12594. Hall, I. M. et al. (2002). Establishment and maintenance of a heterochromatin domain. Science 297(5590), 2232–2237. Huang, H. et al. (1997). The ubiquitin conjugating enzyme Rad6 (Ubc2) is required for silencing in Saccharomyces cerevisiae. Mol. Cell. Biol. 17(11), 6693–6699. Hunt, L. T., and Dayhoff, M. O. (1977). Amino terminal sequence identity of ubiquitin and the nonhistone component of nuclear protein A24. Biochem. Biophys. Res. Comm. 74(2), 650–655. Jacobs, S. A. et al. (2001). Specificity of the HP1 chromo domain for the methylated N-terminus of histone H3. EMBO J. 20(18), 5232–5241. Jacobs, S. A. et al. (2002). The active site of the SET domain is constructed on a knot. Nat. Struct. Biol. 9(11), 833–838. Jentsch, S., McGrath, J. P., and Varshavsky, A. (1987). The yeast DNA repair gene RAD6 encodes a ubiquitin-conjugating enzyme. Nature 329(6135), 131–134. Jenuwein, T., and Allis, C. D. (2001). Translating the histone code. Science 293(5532), 1074–1080. Jenuwein, T. et al. (1998). SET domain proteins modulate chromatin domains in euand heterochromatin. Cell. Mol. Life Sci. 54(1), 80–93. Jones, R. S., and Gelbart, W. M. (1993). The Drosophila Polycomb-group gene Enhancer of zeste contains a region with sequence similarity to trithorax. Mol. Cell. Biol. 13(10), 6357–6366. Kornberg, R. D. (1974). Chromatin structure: a repeating unit of histones and DNA. Science 184(139), 868–871. Kristjuhan, A. et al. (2002). Transcriptional inhibition of genes with severe histone h3 hypoacetylation in the coding region. Mol. Cell 10(4), 925–933. Krogan, N. J. et al. (2002). COMPASS, a histone H3 (Lysine 4) methyltransferase required for telomeric silencing of gene expression. J. Biol. Chem. 277(13), 10753–10755. Loyola, A. et al. (2001). Reconstitution of recombinant chromatin establishes a requirement for histone-tail modifications during chromatin assembly and transcription. Genes Dev. 15(21), 2837–2851. Luger, K. et al. (1997). Crystal structure of the nucleosome core particle at 2.8 A resolution. Nature 389(6648), 251–260. McBride, A. E., and Silver, P. A. (2001). State of the arg: protein methylation at arginine comes of age. Cell 106(1), 5–8. McBride, A. E. et al. (2000). Analysis of the yeast arginine methyltransferase Hmt1p/ Rmt1p and its in vivo function. Cofactor binding and substrate interactions. J. Biol. Chem. 275(5), 3128–3136. Miller, T. et al. (2001). COMPASS: a complex of proteins associated with a trithorax related SET domain protein. [erratum appears in Proc. Natl. Acad. Sci. USA 2001 Dec 18, 98(26): 15393-4]. Proc. Natl. Acad. Sci. USA 98(23), 12902–12907. Min, J. et al. (2002). Structure of the SET domain histone lysine methyltransferase Clr4. Nat. Struct. Biol. 9(11), 828–832.
POSTTRANSLATIONAL MODIFICATIONS OF HISTONES BY METHYLATION
221
Min, J. et al. (2003). Structure of the catalytic domain of human DOT1L, a non-SET domain Nucleosomal Histone Methyltransferase. Cell 112(5), 711–723. Nagy, P. L. et al. (2002). A trithorax-group complex purified from Saccharomyces cerevisiae is required for methylation of histone H3. Proc. Natl. Acad. Sci. USA 99(1), 90–94. Ng, H. H. et al. (2002a). Lysine methylation within the globular domain of histone H3 by Dot1 is important for telomeric silencing and Sir protein association. Genes Dev. 16(12), 1518–1527. Ng, H. H. et al. (2002b). Ubiquitination of histone H2B by Rad6 is required for efficient Dot1-mediated methylation of histone H3 lysine 79. J. Biol. Chem. 277(38), 34655–34657. Nishioka, K. et al. (2002). Set9, a novel histone H3 methyltransferase that facilitates transcription by precluding histone tail modifications required for heterochromatin formation. Genes Dev. 16(4), 479–489. Nislow, C., Ray, E., and Pillus, L. (1997). SET1, a yeast member of the trithorax family, functions in transcriptional silencing and diverse cellular processes. Mol. Biol. Cell 8(12), 2421–2436. Nowak, S. J., and Corces, V. G. (2000). Phosphorylation of histone H3 correlates with transcriptionally active loci. Genes Dev. 14(23), 3003–3013. Pollack, B. P. et al. (1999). The human homologue of the yeast proteins Skb1 and Hsl7p interacts with Jak kinases and contains protein methyltransferase activity. J. Biol. Chem. 274(44), 31531–31542. Rea, S. et al. (2000). Regulation of chromatin structure by site specific histone H3 methyltransferases. Nature 406(6796), 593–599. Robzyk, K., Recht, J., and Osley, M. A. (2000). Rad6-dependent ubiquitination of histone H2B in yeast. Science 287(5452), 501–504. Rogakou, E. P. et al. (2000). Initiation of DNA fragmentation during apoptosis induces phosphorylation of H2AX histone at serine 139. J. Biol. Chem. 275(13), 9390–9395. Roguev, A. et al. (2001). The Saccharomyces cerevisiae Set1 complex includes an Ash2 homologue and methylates histone 3 lysine 4. EMBO J. 20(24), 7137–7148. Rossmann, M. G., Moras, D., and Olsen, K. W. (1974). Chemical and biological evolution of nucleotide binding protein. Nature 250(463), 194–199. Santos-Rosa, H. et al. (2002). Active genes are tri-methylated at K4 of histone H3. Nature 419(6905), 407–411. Schotta, G. et al. (2002). Central role of Drosophila SU(VAR)3–9 in histone H3 K9 methylation and heterochromatic gene silencing. EMBO J. 21(5), 1121–1131. Schreiber, S. L., and Bernstein, B. E. (2002). Signaling network model of chromatin. Cell 111(6), 771–778. Schurter, B. T. et al. (2001). Methylation of histone H3 by coactivator-associated arginine methyltransferase 1. Biochemistry 40(19), 5747–5756. Singer, M. S. et al. (1998). Identification of high copy disruptors of telomeric silencing in Saccharomyces cerevisiae. Genetics 150(2), 613–632. Stallcup, M. R. (2001). Role of protein methylation in chromatin remodeling and transcriptional regulation. Oncogene 20(24), 3014–3020. Strahl, B. D., and Allis, C. D. (2000). The language of covalent histone modifications. Nature 403(6765), 41–45. Sun, Z. W., and Allis, C. D. (2002). Ubiquitination of histone H2B regulates H3 methylation and gene silencing in yeast. Nature 418(6893), 104–108.
222
WOOD AND SHILATIFARD
Teyssier, C., Chen, D., and Stallcup, M. R. (2002). Requirement for multiple domains of the protein arginine methyltransferase CARM1 in its transcriptional coactivator function. J. Biol. Chem. 277(48), 46066–46072. Trievel, R. C. et al. (2002). Structure and catalytic mechanism of a SET domain protein methyltransferase. Cell 111(1), 91–103. Tschiersch, B. et al. (1994). The protein encoded by the Drosophila position-effect variegation suppressor gene Su(var)3-9 combines domains of antagonistic regulators of homeotic gene complexes. EMBO J. 13(16), 3822–3831. Turner, B. M. (2002). Cellular memory and the histone code. Cell 111(3), 285–291. Utley, R. T. et al. (1998). Transcriptional activators direct histone acetyltransferase complexes to nucleosomes. Nature 394(6692), 498–502. Van Hooser, A. et al. (1998). Histone H3 phosphorylation is required for the initiation, but not maintenance, of mammalian chromosome condensation. J. Cell Sci. 111(23), 3497–3506. Van Leeuwen, F., Gafken, P. R., and Gottschling, D. E. (2002). Dot1p modulates silencing in yeast by methylation of the nucleosome core. Cell 109(6), 745–756. Wang, H. et al. (2001). Methylation of histone H4 at arginine 3 facilitating transcriptional activation by nuclear hormone receptor. Science 293(5531), 853–857. Weiss, V. H. et al. (2000). The structure and oligomerization of the yeast arginine methyltransferase, Hmtl. Nat. Struct. Biol. 7(12), 1165–1171. Wilson, J. R. et al. (2002). Crystal structure and functional analysis of the histone methyltransferase SET7/9. Cell 111(1), 105–115. Wolffe, A. P. (2001). Chromatin remodeling: why it is important in cancer. Oncogene 20(24), 2988–2990. Wood, A. et al. (2003). Bre1, an e3 ubiquitin ligase required for recruitment and substrate selection of rad6 at a promoter. Mol. Cell 11(1), 267–274. Workman, J. L., and Kingston, R. E. (1998). Alteration of nucleosome structure as a mechanism of transcriptional regulation. Ann. Rev. Biochem. 67, 545–579. Zhang, X. et al. (2002). Structure of the Neurospora SET domain protein DIM-5, a histone H3 lysine methyltransferase. Cell 111(1), 117–127. Zhang, X., Zhou, L., and Cheng, X. (2000). Crystal structure of the conserved core of protein arginine methyltransferase PRMT3. EMBO J. 19(14), 3509–3519. Zhou, Y., Santoro, R., and Grummt, I. (2002). The chromatin remodeling complex NoRC targets HDAC1 to the ribosomal gene promoter and represses RNA polymerase I transcription. EMBO J. 21(17), 4632–4640.
AUTHOR INDEX
A Aagard, L., 212 Aalfs, J. D., 56 Abe, A., 49 Abe, Y., 99 Abraham, S., 45, 52, 53, 80 Acker, J., 100, 106, 113, 129 Adams, C. C., 173 Adelman, K., 24 Admon, A., 131, 134 Aebersold, R., 54 Agah, H., 52 Agalioti, T., 170, 186, 187, 188 Agarwal, K., 11, 25 Aggarwa, A. K., 86 Aguilera, A., 26 Ait-Si-Ali, S., 186 Albert, A. C., 138 Albert, T., 17 Albright, S. R., 68, 76, 83 Albright-Frey, T. J., 83 Aletta, J. M., 206 Allard, S., 186 Alley, S. C., 183 Allis, C. D., 81, 82, 86, 158, 182, 183, 186, 192–195, 202–206 Allison, S. J., 96, 97 Almer, A., 174 Almouzni, G., 74 Alvarez-Venegas, R., 206, 207 Alzuherri, H. M., 96 Andel, F., III, 69, 71 Anderson, C. W., 182 Anderson, J. D., 171, 174 Anderson, M., 133 Anderson, S., 194 Anderson, S. F., 52 Andrau, J. C., 28, 98, 113, 115 Andrulis, E. D., 182
Anikin, M., 30 Ansari, A. Z., 56 Aono, J., 126 Aoyagi, N., 82, 83 Apone, L. M., 68, 72, 79, 83 Aprikian, P., 138, 139 Archbault, J., 18, 27 Archer, T. K., 162 Arents, G., 159, 201 Arino, T., 140 Armache, K.-J., 2, 4, 10, 11, 12, 15, 22, 25, 27, 28 Arnheim, N., 128 Arrebola, R., 96, 100, 106, 113 Artsimovitch, I., 17, 20, 24 Asturias, F. A., 27 Asturias, F. J., 1, 3, 4, 8, 9, 15, 27, 58–62, 166 Attardi, L. D., 80 Auble, D. T., 85 Auerbach, S. S., 82, 86 Auston, D., 186 Avizonis, D., 80 Avramova, Z., 206, 207 Awery, D. E., 29 Awrey, D. E., 23
B Bachvarov, D., 131 Bagby, S., 84 Bai, Y., 68, 83, 183 Baker, R. E., 95 Baker, S. M., 127 Balasubramanian, R., 185 Balciunas, D., 47, 48, 56, 60 Ballal, N. R., 204, 205 Ballario, P., 188 Baltimore, D., 78 223
224 Bannister, A. J., 81, 82, 86 Barabino, S. M., 26 Baraznenok, V., 48, 59 Bardbury, E. M., 169 Bardeleben, C., 94, 97, 98 Barford, D., 99, 104, 108, 109, 110 Barlev, N. A., 187 Barre, F. X., 186 Bartholomew, B., 24, 25, 94, 95, 114, 165 Bartsch, I., 127 Bartunik, H., 99, 100, 107, 109 Baskaran, R., 18 Bass, I., 25 Bateman, E., 147 Bayle, J. H., 143 Bazett-Jones, D. P., 132, 165 Beard, W. A., 30 Beaton, A., 54, 55 Beaurang, P. A., 45, 52, 53, 80 Becker, P. B., 148, 160, 163, 166, 167, 169, 171, 176 Beckerbauer, L., 187 Beckmann, H., 134, 136, 137 Beek, S. J., 162 Bell, B., 76 Bell, S. D., 15, 29 Bell, S. P., 128, 131, 132, 133, 137 Belotserkovskaya, R., 183, 189 Benecke, A., 196 Benjamin, L. R., 84 Bentley, D., 26, 27, 52 Berenger, H., 186 Berg, J. M., 99, 100, 109, 110, 111 Berger, S. L., 72, 81, 82, 86, 158, 182, 183, 185, 187, 189, 191, 204, 205 Bergstrom, J., 46 Berk, A. J., 52, 53, 54, 67, 78, 96 Bernstein, B. E., 195, 206 Besnard, F., 78 Beve, J., 44, 46, 49, 51 Beyer, A., 133 Bhagat, S., 29 Bhaumik, S. R., 83, 86 Bianchi, M., 133 Bickmore, W. A., 133, 163 Biegel, J. A., 136 Birck, C., 73 Bird, A. W., 196 Birniestiel, M., 125
AUTHOR INDEX
Bischler, N., 9, 15, 28, 139 Bjorklund, S., 44, 45, 48, 50, 56, 60, 68 Blackwell, T. K., 72, 83 Blanco, J. A., 94, 105, 111, 114 Blank, T. A., 148, 167, 169 Blankenship, J. W., 195 Blau, J., 27 Bloom, M. H., 191 Boayke, K. A., 49 Bochar, D. A., 163 Bodem, J., 130 Bone, J. R., 183 Bonte, E., 163, 166, 169 Borst, J. W., 85 Bortvin, A. L., 159 Borukhov, S., 8, 12, 20, 22, 24 Boschiero, C., 28, 113, 115 Botchan, M. R., 97 Bottarelli, L., 95 Bouet, F., 114 Bourenkov, G., 99, 100, 107, 109 Boyer, H., 125 Boyer, L. A., 163, 191 Boyer, T. G., 52, 53 Boyes, J., 143 Bradbury, E. M., 169 Brancorsini, S., 76 Brand, M., 69, 71, 73, 74, 76, 183 Brandl, C. J., 185, 186 Branscombe, T. L., 215, 216, 218 Braun, B. R., 94, 95 Braunstein, M., 203 Brautigam, C. A., 30 Breeden, L., 159 Brehm, A., 169, 171 Brenner, M., 78 Brenowitz, M., 94 Brent, C. J., 114 Brent, R., 4 Briand, J. F., 28, 113, 115 Brick, P., 1, 2, 4, 9 Bridger, J. M., 133 Brill, S. J., 139 Brinker, A., 99, 100, 107, 109 Brino, L., 9, 15, 28 Bromleigh, V., 52, 53 Brophy, B., 97, 98, 101, 105 Brown, C. E., 183, 186 Brown, P. O., 48, 50, 52, 59
AUTHOR INDEX
Brown, R. S., 136 Brown, S. A., 160 Brown, T. A., 49 Brown, T. R., 95, 97 Brownell, J. E., 81, 82, 86, 182, 183 Brun, I., 24 Brun, R. P., 130 Brunger, A. T., 109 Brunning, A., 29 Bryk, M., 213 Buchnell, D. A., 1, 2, 3, 4 Budde, A., 142, 145, 146 Budovskaya, Y. V., 47 Bui, T., 143, 145 Bulger, M., 162, 167 Burakov, D., 53, 56 Buratowski, R. M., 72, 81, 86, 190 Buratowski, S., 14, 18, 67, 72, 73, 81, 83, 86, 97, 185, 190 Burgess, R. R., 1–3, 8, 11, 19 Burke, T. W., 14, 78 Burley, S. K., 14, 68, 69, 71–73, 77, 79, 85 Burlingame, A. L., 195 Burma, S., 84 Burson, L. M., 139 Burton, Z. F., 25 Bushnell, D. A., 1–4, 8, 10–13, 15, 16, 19, 21, 22, 26, 27, 44, 48, 49, 58 Bustamante, C., 24 Butler, J. E., 14, 77 Buttgereit, D., 130, 133 Byfield, P., 143
C Cabon, F., 186 Cadena, D. L., 18, 44 Cagas, P. M., 8 Cairns, B. R., 161, 162, 165, 166, 167, 169, 172, 176, 188, 189 Cairns, C. A., 96, 97, 133 Callebaut, I., 9, 15, 28 Campbell, A. M., 68, 85 Campbell, E. A., 17, 20, 28, 29 Campbell, S., 75 Candau, R., 183 Cantin, G. T., 54 Carey, M., 15, 84
225
Carles, C., 9, 15, 28, 94, 98, 113, 114, 115, 137, 139 Carling, D., 47 Carlisi, D., 186 Carlson, J. E., 86 Carlson, M., 47, 49, 50, 56, 159, 166, 167 Carre, L., 73 Carrello, A., 100, 109 Carrozza, M. J., 183 Causton, H. C., 83, 86 Cavalli, G., 186 Cavallini, B., 67 Cavanaugh, A. H., 138, 140, 144, 145 Cedin, S., 28 Chacon, P., 29 Chafin, D. R., 22 Chait, B. T., 73, 75, 79 Chalkley, G. E., 78 Chamberlin, M. J., 24 Chambon, P., 67, 72, 79, 86, 196 Chang, A., 125 Chang, C. P., 53 Chang, W. H., 3, 4, 8, 9, 15, 19, 27 Chang, Y. W., 47 Chao, D. M., 44, 46, 47, 52 Chatterjee, S., 78 Chaussivert, N., 94, 96, 98, 100, 101, 105, 109 Chaves, R. S., 49 Chedin, S., 28 Chen, D., 215, 217, 218 Chen, G., 186, 187, 188 Chen, H., 14, 85, 173 Chen, J. L., 68, 78, 80, 136, 137 Chen, Y., 17 Chen, Z., 83 Chen, Z. J., 128 Cheng, K. K., 51 Cheng, X., 216, 217 Chesnokov, I., 97 Chesterton, C. J., 146 Cheung, P., 195 Chi, T., 84, 164 Chiang, C. M., 73, 80 Chiang, G. G., 18 Chiba, H., 160 Chibazakura, T., 50 Chicca, J. J., 84, 85 Chieco, P., 125 Chinenov, P., 194
226 Chipoulet, J. M., 67 Chitikila, C., 84 Chlenov, M., 29 Cho, E. J., 18 Cho, H., 27, 52, 54, 58, 193 Choder, M., 11 Chow, A. M., 132, 137 Christman, M. F., 196 Chu, W. M., 96, 97 Chua, P., 86 Chung, G., 143 Chung, W. H., 166 Cimato, T. R., 206 Clapier, C. R., 169, 171 Clark, C. D., 160 Clarke, A., 186 Clarke, S., 215 Clos, J., 126, 130, 133 Cloutier, T. E., 94 Coburn, C., 159 Cohen, P. W., 99, 104, 108, 109, 110 Cohen, S., 73, 79, 125 Colbert, T., 94, 97, 98, 105, 111, 114 Cole, C. N., 26 Coleman, R. A., 84 Collart, M. A., 85 Comai, L., 128, 134, 136, 137, 141, 142, 143, 145 Conaway, J. W., 25, 52, 54 Conaway, R. C., 25, 52, 54 Conconi, A., 147 Conesa, C., 9, 15, 28, 96, 98, 100, 101, 105, 106, 109, 114 Cook, E. H., 54 Cook, P. R., 26 Cook, R. G., 183, 186 Copenhaver, G. P., 124, 132, 133 Corden, J. L., 8, 18 Cordes, S., 128, 129, 133 Corona, D. F., 163, 169 Cosma, M. P., 57 Cote, J., 143, 161–163, 165, 166, 167, 183, 186, 188 Couderc, J. L., 73 Coulombe, B., 14 Coupar, B. E. H., 146 Court, M., 9, 15 Covitz, P. A., 50 Cox, J. M., 14 Coy, J. F., 129
AUTHOR INDEX
Crabtree, G. R., 143, 160, 161, 164 Craighead, J. L., 4, 8, 9, 15, 27 Cramer, P., 1–4, 8, 10–13, 15, 16, 19, 21–22, 25–30 Croce, C., 128 Cromlish, J. A., 67 Crosio, C., 195 Crowe, A., 94, 96, 98, 102 Crowley, K. A., 191 Crowley, T. E., 75 Cuthbert, A. P., 133 Cvekl, A., 130
D Dahmus, M. E., 13, 18, 44 D’Alessio, J. M., 147 Dammann, R., 129 Damschroder-Williams, P., 54 Dang, T., 96 Daniel, C., 163 Dantonel, J. C., 75 Darcy, T. J., 29 Darnell, R. B., 54 Darst, S. A., 1, 3, 8, 17, 19, 20, 22, 28, 29, 33 Das, A., 24 Das, A. K., 99, 104, 108, 109, 110 Dasgupta, A., 96 Datta, P. K., 136 Dauphinais, C., 146 Davenport, R. J., 24 David, P. R., 1–3, 8, 11, 19 Davidson, A. R., 18, 27 Davidson, I., 69, 73, 75, 76 Davis, J. A., 27 Davis, R. J., 27 Dawid, I. B., 124, 125 Dayhoff, M. O., 204, 205 Dea, C., 54 de Bizemont, T., 192 Deckert, J., 183 DeDecker, B. S., 14 Delius, H., 130 Dellaire, G., 163 del Rio-Portilla, F., 2 Denissova, L., 5 Dennis, J. H., 14
227
AUTHOR INDEX
Denton, M. L., 132, 133, 138 Denu, J. M., 158, 182, 183, 186, 191 Deprez, E., 96 Derenzini, M., 124, 125 Deroo, B. J., 162 Deshpande, A. M., 94 Deuring, R., 160 Devarakonda, S., 188 deWalque, S., 110 Dhalluin, C., 86 DiAngelo, S., 136 Dieci, G., 94, 95, 101, 106 Dierich, A., 75 Dikstein, R., 76, 80 Dilworth, F. J., 74 Dimitrov, S. I., 140 Dingwall, A., 161 Dkhissi, F., 186 Dobreva, G., 130 Dodd, J. A., 139 Dohmen, R. J., 205 Dollard, C., 68, 79 Donze, D., 94 Doree, M., 142, 146 Dorland, S., 50 Dorn, R., 206 Dornan, J., 100, 109 Dotson, M. R., 59, 62 Doublie, S., 21, 30 Dover, J., 205 Downes, C. S., 96 Drysdale, C. M., 183 Du, J., 162 Du, W., 192, 193 Ducommun, B., 186 Dudley, A. M., 86 Duggan, L. J., 183, 189 Dujardin, G., 95 Dumas, K., 163, 166 Dumay, H., 101, 115 Dumay-Odelot, H., 100, 106, 113 Dunphy, E. L., 82, 86 Duquet, A., 186 Durso, R. J., 83 Dusterhoft, A., 101, 106 Dvir, A., 16, 17 Dye, M. J., 26 Dynlacht, B. D., 67, 68, 79
E Eberhard, D., 134 Eberharter, A., 169, 171, 183, 188 Ebright, R. H., 14, 16, 27, 29 Ederth, J., 17 Edmondson, D. G., 183 Edwards, A. M., 1–3, 8–9, 11, 19, 23, 29 Egly, J. M., 67, 79, 95, 134 Eick, D., 17 Eisen, J. A., 167 Eisenman, R. N., 97, 115 Eisenmann, D. M., 68, 79 Eliason, K., 127, 139 Ellenberger, T., 21, 30 Eloranta, J. J., 9, 28 Emami, K. H., 14 Emanuel, P. A., 78 Emmons, S. W., 55 Endo, T., 99 Engelke, D. R., 94 Erdjument-Bromage, H., 44, 48, 49, 51–54, 58, 74, 162, 189, 192 Erie, D. A., 23 Ernest, I., 48, 59 Erny, B., 130 Ettinger, M. J., 206 Evangelista, C. C., Jr., 49 Evans, P. R., 188 Evans, R. M., 196 Evers, R., 133
F Fan, H. Y., 51, 169 Fang, Q., 73, 185 Fantino, E., 139 Fassler, J. S., 50 Faubladier, M., 143 Faus, I., 67 Feaver, W. J., 3, 45 Fellows, J., 45, 56, 192, 193 Felton-Edkins, Z. A., 96 Ferguson, H. A., 84 Ferreira, R., 186 Ferri, M. L., 9, 15, 28, 98 Fersht, A. R., 218 Fiedler, U., 15, 16
228
AUTHOR INDEX
Filetici, P., 188 Filleur, S., 186 Fimia, G. M., 75 Financsek, I., 126, 129 Fire, A., 67 Fischle, W., 182, 194, 195 Fisher, A. K., 83 Fitzgerald-Hayes, M., 50 Flanagan, J. F., 166 Flanagan, P. M., 44 Flaus, A., 148, 169, 175 Fleming, K. G., 14 Flores, A., 28, 113, 115 Flores, O., 27 Fomproix, N., 132 Fondell, J. D., 51–53, 55, 56, 58, 80 Forbes, D. J., 96 Forde, N. R., 24 Franco, L., 160 Frankel, A., 215 Franklin, M. C., 21 Frazier, M. W., 131, 132 Freedman, L. P., 52, 53, 56 Freese, E., 78 Freiman, R. N., 76, 83 Fried, M. G., 73, 185 Friedrich, J. K., 130, 131, 137, 138 Fry, C. J., 188 Fu, J., 1–3, 8, 11, 12, 16, 19, 21, 22, 183 Fu, Z. Y., 53, 55, 56 Fujii, H., 67 Fukasawa, T., 49, 50 Fuller, M. T., 76, 83 Funk, J. O., 17 Furger, A., 26 Furter, R., 17 Furter-Graves, E. M., 17 Fyrberg, A. M., 163
G Gabrielsen, O., 95 Gadal, O., 28, 113, 115 Gadbois, E. L., 52, 60 Gafken, P. R., 202, 203, 213, 214 Galasinski, S. K., 81, 84 Galman, C., 48, 56, 60 Gamble, M., 52, 53 Gamblin, J. S., 99, 100, 109, 110
Gangloff, Y. G., 69, 73 Gann, A., 79 Gansmuller, A., 76 Ganster, R. W., 163 Garabedian, M. J., 56 Garg, L. C., 136 Garraway, I. P., 14 Gary, J. D., 215 Gaskell, A., 2 Gatto, G. J., Jr., 99, 100, 109, 110, 111 Gavin, I., 169 Gdula, D. A., 169 Ge, H., 52, 53, 81, 86 Gebrane-Younes, J., 132 Gegnas, L. D., 14 Geiduschek, E. P., 94, 95, 97, 98, 105, 106, 111, 114 Geiger, J. H., 14, 25, 71 Geiman, T. M., 163, 169 Geisbrecht, B. V., 99, 100, 109, 110, 111 Gelbert, W. M., 206 Geli, V., 186 Genereaux, J., 185 Gentz, R., 85 Gerard, M., 95 Geraud, G., 132 Gerstein, M., 3 Ghavidel, A., 97 Ghosh, A. K., 135, 136 Ghosh, G., 14 Giese, K., 131 Gilbert, D., 2 Gileadi, O., 18, 27, 193 Gilmour, D. S., 17, 78 Gim, B. S., 48, 49, 51, 54, 55 Ginns, E. I., 54 Girault, J. A., 186 Giuliodori, S., 95 Gnatt, A. L., 1–3, 8, 11, 12, 16, 19, 21, 22, 45, 56 Gokal, P. K., 130, 137, 140 Goldfarb, A., 5, 18, 19, 24, 25 Golemis, E. A., 4 Golub, T. R., 46, 47, 51, 52 Gomez-Roman, N., 97, 115 Gong, D. W., 84 Gong, X. Q., 25 Gonzalez-Couto, E., 79 Goodman, H., 125 Goodrich, J. A., 16, 81, 84
AUTHOR INDEX
Goppelt, A., 85 Gotschling, D. E., 202, 203, 213, 214 Gottesfeld, J. M., 96 Gottlieb, T. M., 96, 135 Gottschling, D. E., 182 Gould, S. J., 99, 100, 109, 110, 111 Goussot, M., 28, 113, 115 Govoni, M., 125 Graba, Y., 186 Gralla, J. D., 15 Grandori, C., 97, 115 Grant, P. A., 74, 86, 182–183, 185, 186, 188, 189, 194, 196 Grebe De Barror, A., 81 Green, M. R., 46, 47, 51, 52, 68, 71, 72, 77, 79, 80, 83, 85, 86, 161, 162, 163, 166, 167, 175 Greenblatt, J., 14, 15, 18, 27 Greene, E. A., 131 Greenleaf, A. L., 13, 17, 18, 29 Grienenberger, A., 186 Griffin, L. A., 68, 72 Grimaldi, M. A., 163 Gromoller, A., 47 Gross, S., 26 Grosschedl, R., 131 Groudine, M., 25 Grummt, F., 140 Grummt, I., 15, 124, 126–135, 137, 138, 140–146, 148, 163, 204 Grunstein, M., 86, 182, 186, 195 Gu, W., 23, 24, 51–54, 58, 143 Guarente, L., 14, 67 Guermah, M., 80 Guilfoyle, T. J., 9 Guo, H., 22 Guschin, D., 163, 169 Gustafsson, C. M., 44, 46, 48–52, 58–62 Guyon, J. R., 169 Gygi, S. P., 194
H Hadelt, W., 130 Hadjiolov, A. A., 124, 125, 140 Haglund, R. E., 128 Hahn, S., 14, 27, 54, 57, 67, 71, 85, 94, 97, 98, 105, 111, 114 Hain, J., 29
Halay, E. D., 14 Hall, B. D., 17, 95 Hall, I. M., 213 Haltiner, M. M., 126, 140 Hamada, M., 95 Hambalko, M., 114 Hamiche, A., 163, 167, 169 Hammamori, Y., 143 Hammell, C. M., 26 Hammer, R. E., 76, 83 Hampsey, M., 193 Han, M., 54, 55 Han, S. J., 48–51, 54, 55, 60 Hanada, K., 129, 130, 132 Hanauer, A., 195 Hanawalt, P. C., 167 Handa, H., 25 Hannan, K. M., 124, 129, 145 Hannan, R. D., 124, 138, 140, 145 Hannett, N., 83, 86 Hansen, J. C., 183 Hansen, K. E., 85 Hansen, S. K., 75, 80 Hardin, S., 13, 18 Harel-Bellan, A., 186 Harmon, K. E., 196 Harp, J. M., 188 Harris, G. H., 147 Hartl, F. U., 99, 100, 107, 109 Hartl, P., 96 Hartzog, G. A., 193 Hassan, A. H., 187, 188, 189 Hassig, C. A., 182 Hausner, W., 29 Hautmann, M., 141 Havas, K., 175 Hawley, D. K., 24, 25 Hayashibara, K. C., 48, 50, 52, 59 Hayes, J. J., 172 Haystead, T. A., 141 Hayward, M. M., 14 He, C., 86 Health, C. V., 26 Heilgenthal, G., 129 Heix, J., 134, 142, 146 Hellings, R., 125 Hempel, W. M., 138, 140, 144, 145 Henderson, S. L., 127 Hengartner, C. J., 46, 47, 51, 52 Henry, N. L., 45
229
230 Henry, S., 44, 46 Herfort, M., 132 Herman, P. K., 47 Hernandez, N., 9, 28, 68, 94, 97, 135 Hernandez-Verdun, D., 132, 143 Herrero, P., 49 Hershey, J. C., 141 Herskowitz, I., 49, 159 Heumann, H., 5 Hieter, P., 98 Hilfiker, A., 193 Hilfiker-Kleiner, D., 193 Hiller, M. A., 76, 83 Hinnebusch, A. G., 83, 84, 183 Hinnen, A., 174 Hiraoka, Y., 49 Hirose, Y., 26 Hirsch, J. P., 44, 46 Hirschhorn, J. N., 159, 160 Hirschler-Laskiewicz, I. I., 143 Hirschmann, P., 141 Hirsh, I. S., 110 Hisatake, K., 14, 132 Hishinuma, F., 50 Hittelman, A. B., 56 Hock, R., 125 Hoey, T., 67, 68, 75, 79 Hoff, C. M., 135 Hoffmann, A., 67, 73, 79 Hoffmann, M., 130, 142 Hoffmann-Rohrer, U., 15, 130, 148, 163 Hol, W. G., 110 Holmberg, S., 48, 59 Holmes, M. C., 75 Holstege, F. C., 15, 18, 27, 46, 47, 51, 52, 83, 86 Horikoshi, M., 67, 68, 78, 84, 186 Horiuchi, T., 127 Horn, P. J., 169 Horner, M. A., 75 Horowitz-Scherer, R., 166 Hortnagel, K., 17 Horz, W., 174, 176 Houweling, A., 191 Hovde, S. L., 25 Howard, B. H., 74 Howard, S. C., 47 Howard, T., 74 Howcroft, T. K., 78, 80
AUTHOR INDEX
Howe, L., 183, 186, 194 Howley, R., 73, 185 Hsieh, Y. J., 96, 101, 105, 109, 113, 114 Hu, C., 131 Hu, P., 9, 28 Huang, H., 205 Huang, L., 195 Huang, Y., 94, 95 Huet, J., 67, 95, 98, 114 Huibregtse, J. M., 94 Hull, W. W., 17 Hulsmann, B. B., 163 Hunt, L. T., 204, 205 Hunter, T., 18 Hurt, E., 26 Huylebroeck, D., 48, 59 Hwang, K. K., 191 Hwang, M. S., 55
I Iassan, J.-P., 14 Iben, S., 130, 138 Igo, M., 96 Iida, C. T., 67, 147 Iizuka, M., 186 Ikeda, K., 183, 188 Ikura, M., 84 Imbalzano, A. N., 71, 161–163, 166, 167, 175 Imhof, A., 81, 86, 169, 171 Iniguez-Lluhi, J. A., 56 Inouye, C., 69, 71, 86 Isaksson, L. A., 17 Ishiguro, A., 106 Ishihama, A., 9, 27 Ishikawa, Y., 126, 128 Ishima, R., 84 Ito, M., 51–56, 58 Ito, T., 162, 167 Izban, M. G., 23, 24, 25 Izhaky, D., 24
J Jackson, B. M., 83, 183 Jackson, S. P., 15, 29, 96, 97, 98, 101, 105, 135 Jackson-Fisher, A. J., 84
231
AUTHOR INDEX
Jacob, S. T., 135, 136 Jacobs, S. A., 188, 207, 209, 210, 211, 213 Jacobson, R. H., 75, 86, 188 Jacq, X., 74, 76 Jacquot, S., 195 Jaenisch, R., 54, 55 Jager, J., 30 Jain, A., 14 Jain, R., 86 Jain, S., 56 Jan, G., 75 Jan, L. Y., 75 Jan, Y. N., 75 Jansa, P., 148, 163 Jansma, D. B., 18, 27 Jantzen, H. M., 128, 131, 132, 133, 137 Jaskelioff, M., 169 Jeanmougin, F., 72, 86 Jefferson, L. S., 129, 145 Jeffery, D. A., 47 Jennings, E. G., 18, 27, 46, 47, 51, 52, 60, 83, 86 Jensen, G. J., 1, 3, 8, 15, 19 Jensen, R., 159 Jentsch, S., 205 Jenuwein, T., 86, 202, 203, 206, 207 Jeon, C.-J., 11, 25 Jia, Y., 54, 55 Jiang, Y. W., 50, 52, 54, 58–62 Joazeiro, C. A., 94, 105, 111, 114 John, S., 183, 186, 188 Johnson, T., 82, 86 Johnston, I. M., 97 Johzuka, K., 127 Jokerst, R. S., 29 Jona, G., 193 Jones, D. N., 2 Jones, K., 125 Jones, M. H., 126 Jones, P. L., 163 Jones, P. R., 195 Jones, R. S., 206 Jordan, E. G., 125 Jourdain, S., 96 Ju, Q., 139 Junera, H. R., 132
K Kadonaga, J. T., 14, 77, 78, 79, 80, 85, 162, 167, 174 Kadosh, D., 182 Kahl, B. F., 16 Kaine, B. P., 2 Kalkhoven, E., 191 Kalousek, I., 141, 146 Kaltenbach, L., 75 Kamada, K., 85 Kamakaka, R. T., 94 Kane, C. M., 9, 11, 23, 24 Kang, J. S., 60 Kang, Y. K., 52, 80 Kao, C. C., 67 Kapanidis, A. N., 14 Karlsson, K. A., 46 Kashlev, M., 5, 19, 20, 23, 24 Kassavetis, G. A., 94, 95, 97, 98, 105, 106, 111, 114 Kato, H., 126, 128, 160 Kaufman, T. C., 160 Kaufmann, J., 14, 78 Kawaichi, M., 80 Kay, L. E., 84 Keaveney, M., 79, 83, 182 Kedes, L., 143 Keener, J., 139 Kehle, J., 169, 171 Kelleher, R. J., 44 Keller, W., 26 Kelly, G., 2 Kennedy, E. L., 183 Kennison, J. A., 160 Kermekchiev, M., 138, 148 Kershnar, E., 80 Kettenberger, H., 2, 4, 10, 11, 12, 15, 22, 25, 27, 28 Keys, D. A., 139 Khavari, P. A., 160–163, 166, 167 Khazak, V., 4 Khoo, B., 97, 98, 101, 105 Khorasanizadeh, S., 188 Kieselbach, T., 48, 59 Kihm, A. J., 141 Kikyo, N., 163, 169 Killeen, M., 14 Kim, H. S., 55 Kim, J. L., 14, 71
232
AUTHOR INDEX
Kim, J. M., 57 Kim, T. K., 16, 27 Kim, Y., 14, 71, 188 Kim, Y. J., 44, 45, 46, 48–51, 54, 55, 57, 60, 68, 161, 166, 167 Kimura, A., 186 Kimura, M., 9, 27 Kim Y. J., 50 King, B. H., 54 King, D. S., 86, 132, 137, 188 Kingston, R. E., 56, 71, 74, 161, 162, 163, 166, 167, 169, 171–173, 175, 192, 202, 203 Kireeva, M. L., 19, 20 Kirschner, D., 71, 73 Kishimoto, T., 126, 128 Kissinger, M., 160 Klages, N., 79 Klebanow, E. R., 83, 183 Kleff, S., 182 Klein, H. L., 51 Klein, J., 142 Knibiehler, M., 186 Knoll, D. A., 147 Kobayashi, R., 9, 28, 85, 97, 162, 167 Kobayashi, T., 127 Kobor, M. S., 18, 27 Koh, S. S., 47, 56, 60 Kohda, D., 99 Kokubo, T., 73, 79–82, 84, 86, 183 Koleske, A. J., 14, 46, 47, 80 Koller, T., 147 Kolodrubetz, D., 96 Komarnitsky, P., 18, 73, 83 Kominami, R., 126, 128, 129 Komissarova, N., 19, 20, 23, 24 Kondo, T., 132 Kondou, S., 50 Kopf, G. S., 136 Korkhin, Y., 14 Kornberg, R. D., 1–4, 8–16, 19, 21, 22, 26–27, 44–45, 48–52, 54, 56, 58–62, 68, 80, 161, 162, 166, 167, 172, 173, 189, 201 Korolev, S., 22 Korzheva, N., 18, 19, 20 Kosa, P., 14, 83 Kotani, T., 74 Kouyoumdjian, F., 18, 27 Kouzarides, T., 81, 82, 86, 96
Kovelman, R., 96, 101, 105, 109, 113, 114 Kozlov, M., 5, 19 Kraemer, S. M., 71, 84 Krapp, S., 2 Kraut, J., 21 Kristjuhan, A., 203 Krizkova, P., 141, 146 Kruger, W., 159 Kruh, G. D., 18 Krumm, A., 25 Kruppa, M., 96 Ktistaki, E., 191 Kubalek, E. W., 1, 3 Kuchin, S., 47, 49, 56 Kugel, J. F., 16 Kuhn, A., 127, 133, 135, 141, 142, 146, 148 Kumar, A., 21, 97, 98, 106, 110, 114 Kundu, T. K., 96, 183 Kuo, A., 164 Kuras, L., 79, 83, 182 Kurdistani, S. K., 195 Kuroda, M. I., 193 Kurpad, H., 73, 83 Kutach, A. K., 14, 77, 79 Kuznedelov, K., 20 Kwon, H., 71, 161, 162, 163, 166, 167, 175 Kwon, J. Y., 48, 54, 55
L Labouesse, M., 75 Lacomis, L., 162 Lacroix, J. F., 129 Ladbury, J. E., 14 Ladias, J. A., 2 Ladurner, A. G., 52, 54, 69, 71, 86, 188 Lafleur, D. W., 163 Lagrange, T., 14 Lakatos, L., 75 Lalo, D., 139 Lamb, J. R., 98 Lamond, A. I., 130, 131, 137 Landel, C. C., 163, 165 Lander, E. S., 46, 47, 51, 52 Landick, R., 17, 20, 24 Lane, W. S., 27, 49, 50, 51, 52, 54, 58, 60, 85, 163, 193 Lange, U., 29 Langer, D., 29
233
AUTHOR INDEX
Langer, M. R., 191 Langst, G., 148, 163, 169, 171, 176 La Porta, A., 24 Lapouge, K., 99, 100, 109, 110 Laptenko, O., 8, 12, 20, 22 Larkin, R. M., 9 Larminie, C. G., 96 Larson, D. E., 124 Larsson, T., 46 Laurent, B. C., 159, 161, 162, 166, 167 Laverty, T., 54, 55 Lavigne, A. C., 73 Lawther, R. P., 140 Learned, R. M., 126, 128, 129, 133, 137, 140 Leblanc, B., 132 Le Douarin, B., 72, 86 Lee, B. S., 139 Lee, D. K., 14, 52 Lee, J., 8, 12, 20, 22, 48, 54, 55, 60, 96 Lee, K. M., 172 Lee, S., 98 Lee, T. I., 46, 47, 51, 52, 60, 83, 84, 86 Lee, Y. C., 46, 48–51, 55, 60 Lee, Y. H., 54 Lees, E., 52, 53, 54, 58, 193 Lefebvre, O., 15, 28, 94, 95, 96, 98, 101, 106, 114 Lehming, N., 47 Lejeune, F., 74 Lemon, B. D., 52, 53, 56 LeRoy, G., 163 Lester, W., 18, 27 Letts, G. A., 98, 106 Leurent, C., 69, 71, 73, 74 Leuther, K. K., 1, 3 Levillain, E., 9, 28 Levine, M., 83 Levivier, E., 9, 15 Li, B., 17, 173, 194 Li, H., 16 Li, J., 194 Li, X. Y., 79, 83, 85 Li, Y., 8, 22, 44, 45, 48, 50, 56, 59, 60, 62, 95, 162, 192 Liao, S. M., 47 Liao, Y., 103–107, 111, 113 Librizzi, M. D., 94, 96, 97, 102, 103, 104, 114 Lieberman, P. M., 67, 75, 80
Lieu, H. M., 54 Lilley, D. M., 164, 175 Lin, C. W., 139 Lin, T. Y., 76, 83 Linask, K. L., 27 Lindstrom, D. L., 193 Lis, J. T., 13, 18, 24, 52, 57, 75 Littlefield, O., 14 Liu, D., 84 Liu, J. K., 75 Liu, L. F., 136, 187 Liu, Y., 54 Liu, Z., 135 Lively, T. N., 81, 84 Livingstone-Zatchej, M., 96 Lo, K., 14 Loeb, L. A., 30 Loening, U. E., 125 Logie, C., 163, 165, 168, 169 Lomvardas, S., 170, 186, 187 Long, A. M., 21 Long, E. O., 124, 125 Lopes, J., 44, 46 Lopez, S., 96 Lopez-de-Leon, A., 96, 97 Lorch, Y., 44, 49, 162, 166, 172, 173 Losson, R., 72, 86 Lottspeich, F., 85 Lowe, N., 188 Lowe, T. M., 95 Lowell, J., 186 Loyola, A., 163, 212 Lucchesi, J. C., 193 Lucchini, R., 147, 148 Luger, K., 69, 73, 175, 182, 201, 202 Lui, M., 44, 48, 49, 51, 58 Lukhtanov, E., 19, 24, 25 Lun, M., 145 Luo, K., 186 Luse, D. S., 16, 18, 23, 24, 25
M Mader, A. W., 69, 73, 175 Madsen, C. S., 141 Maeda, Y., 129, 130, 132 Magill, C. P., 15 Magnaghi-Jaulin, L., 186 Mahajan, P. B., 130, 137, 140
234 Maier-Davis, B., 1, 2, 3, 8, 11, 19 Mais, C., 132 Malhotra, A., 19 Malik, S., 51–55, 58, 62, 80, 85 Mallouh, V., 9, 15, 28, 69, 71, 73, 74 Mancebo, H., 27 Mango, S. E., 75 Maniatis, T., 26, 170 Manley, J. L., 26, 83 Mann, M., 163, 166 Maraia, R. J., 94, 95 Marck, C., 28, 94, 98, 100, 101, 106, 113, 114, 115 Marc Timmers, H. T., 16 Margottin, F., 95 Mariol, M. C., 186 Markovtsov, V., 5 Marmorstein, R., 72, 190 Marsolier, M. C., 96 Martianov, I., 75, 76 Martin, B. M., 54 Martin, E., 5 Martin, K., 96 Martin, M. E., 52, 53 Martinez, E., 51, 52, 53, 58, 75, 183 Marzouki, N., 114 Mason, P., 26 Mason, S. W., 148 Masson, C., 132 Masuda, S., 17, 20, 26 Matallana, E., 160 Matangkasombut, O., 72, 81, 86, 190 Matsui, T., 67 Matthes, H., 67 Matthews, S., 2 Mautner, J., 17 Maxon, M. E., 16 McAllister, W. T., 30 McBride, A. E., 215, 217 McCallum, C. M., 162 McGrath, J. P., 205 McGrew, J. T., 50 McKune, K., 9, 11, 17 McLees, A., 96 McMahon, S. J., 183 McManus, M. T., 94 McNeil, J. B., 52 McStay, B., 126, 131–133 McVeigh, R., 183 Meersseman, G., 169
AUTHOR INDEX
Megee, P. C., 196 Meisterernst, M., 54, 85 Mencia, M., 83 Mengus, G., 73 Meredith, G. D., 1, 3, 8, 15, 19 Merika, M., 186, 187 Michaelidis, T. M., 142, 146 Michel, B., 73, 83 Michels, P. A., 110 Miesfeld, R., 128 Mihara, K., 99 Miklos, I., 49 Milkereit, P., 15, 139, 147 Miller, G., 130, 131, 137 Miller, K. G., 126 Miller, O. L., Jr., 133 Miller, T., 207, 213 Min, J., 207, 209, 210, 211, 214 Min, S., 48, 49, 50 Minakhin, L., 28, 29 Minchin, R. F., 100, 109 Miotto, B., 186 Mirza, U. A., 73, 79 Mishima, Y., 126, 128, 129 Misra, S., 54, 55 Mital, R., 96, 97 Mitchell, A. P., 50 Mitomo, K., 139 Mitra, M., 84 Mitsiou, D. J., 76 Mitsuzawa, H., 9 Miyake, T., 80 Mizuguchi, G., 163, 167 Mizumoto, K., 126 Mizzen, C. A., 81, 82, 86, 195 Moarefi, I., 99, 100, 107, 109 Moazed, D., 194 Moggs, J. G., 74 Moir, R. D., 94–96, 98–111, 113, 114 Mollah, A. K., 94 Monaco, L., 195 Montanaro, L., 125 Moore, C., 26 Moore, P., 163 Moore, P. A., 75 Moore, R., 139 Moorefield, B., 131, 138, 139 Moorow, J., 125 Moqtaderi, Z., 83, 182 Moras, D., 73, 75, 216
235
AUTHOR INDEX
Moreno, F., 49 Moreno, G. T., 163 Moroder, L., 99, 100, 107, 109 Morris, P. L., 75 Morton, J. P., 97 Moseley, S. L., 162 Moss, T., 124, 126, 127, 131, 132, 138, 140, 143 Mote, J., 23 Moudrianakis, E. N., 159, 201 Mougey, E. B., 133 Mozden, N., 54, 55 Muchardt, C., 160 Mueller, C. G., 85 Muhandiram, D. R., 84 Muhich, M. L., 67 Muldrow, T. A., 85 Muller, F., 75 Muller, J., 169, 171 Muller, S., 83 Munday, M. R., 47 Munshi, N., 186, 187 Murakami, K. S., 17, 20 Muramatsu, M., 126, 128, 129, 130, 132, 160 Muratoglu, S., 73 Murray, P. J., 52, 54, 55 Musfeldt, M., 29 Mustaev, A., 5, 18, 19, 20, 24, 25 Muster, N., 186 Muth, V., 143, 144 Muto, T., 99 Muzzin, O., 17, 20 Myer, V. E., 14, 47 Myers, L. C., 44, 45, 48–52, 58–62, 80 Myers, M. P., 9, 28 Mysliwiec, T., 18
N Naar, A. M., 45, 52, 53, 80 Nadaud, S., 143, 144 Nagamine, M., 126 Nagy, P. L., 207 Nakatani, S. K., 73 Nakatani, Y., 73, 74, 78, 79, 81, 82, 84, 86, 143, 183, 193 Narlikar, G. J., 169, 171, 172 Nasmyth, K., 57, 159
Natarajan, K., 83 Nedialkov, Y. A., 25 Neely, K. E., 187, 188, 189, 191 Neigeborn, L., 159 Nemeth, A., 148, 163 Neuhaus, D., 188 Newbold, R. F., 133 Newlands, J., 24 Newlon, C. S., 94 Ng, H. H., 202–205, 213, 214 Nguyen, L. H., 94 Nguyen, T. T., 139 Nie, Z., 162 Nigg, E. A., 14 Nikiforov, V., 5, 18, 19, 25 Nikolov, D. B., 14, 69, 71, 72 Nilsson, L., 54, 55 Nishikawa, J. I., 84 Nishikawa, S., 99 Nishimura, T., 132 Nishioka, K., 213 Nishizawa, M., 50 Nislow, C., 207 Niu, H., 135 Nogales, E., 69, 71 Nogi, Y., 49, 139 Nomoto, A., 160 Nomura, M., 127, 139 Nonet, M. L., 29, 46 Normann, A., 126 Novina, C. D., 77, 78 Nudler, E., 18, 24, 25, 197 Nurse, P., 124 Nye, J. S., 54, 55
O Oakes, M. L., 139 O’Brien, R., 13, 14, 18 O’Brien, T., 81, 82, 136, 137 Ochs, R. L., 125 Oelgeschlager, T., 52, 73, 80 Ogg, R. C., 84 Ogryzko, V. V., 74, 81, 86, 143, 193 Ohba, R., 183, 192 Ohishi, T., 50 Ohkuma, Y., 16 Ohrlein, A., 126 Okano, H. J., 54
236
AUTHOR INDEX
Oliner, J. D., 80 Olsen, K. W., 216 O’Mahony, D. J., 131, 132, 133, 141, 146 Onesti, S., 1, 2, 4, 9 Onufryk, C., 54, 55 Opalka, N., 29 Oppizzi, M., 26 O’Reilly, M., 133 Orlicky, S. M., 9 Orlova, M., 24 Ornaghi, P., 188 Orphanides, G., 26, 163, 193 Osada, S., 186 Osheim, Y. N., 133 Osley, M. A., 204, 205 Ossipow, V., 14 Otero, G., 192 Ottonello, S., 95, 101, 106 Oudet, P., 96, 114 Oulad-Abdelghani, M., 74 Owen, D. J., 188 Owen-Hughes, T., 81, 82, 86, 148, 169, 175, 183, 187 Owens, G. K., 141 Ozer, J., 75
P Pacella, L. A., 183, 189 Pagel, J., 131 Pal, M., 16, 18 Palangat, M., 24 Palmer, J., 163 Palotie, A., 54 Pan, G., 15 Panizza, S., 57 Pannuti, A., 193 Panov, K. I., 130, 131, 137, 138 Papamichos-Chronakis, M., 191 Pape, L. K., 128 Papoulas, O., 162 Parekh, B., 170 Park, J. M., 48, 49, 50, 54, 55, 57 Park, S. J., 49, 51 Park, Y., 193 Parker, C. S., 67 Parthun, M. R., 182 Parvin, J. D., 52 Parvinen, M., 75, 76
Pata, J. D., 30 Patel, P. H., 30 Patlan, V., 30 Pattatucci, A. M., 160 Paule, M. R., 16, 95, 124, 147 Payne, J., 139 Pazin, M. J., 162, 167, 174 Pegoraro, S., 99, 100, 107, 109 Pei, R., 67 Pelletier, G., 143 Pelletier, H., 21 Pennings, S., 169 Pennock, D., 126 Penttila, T. L., 75 Percudani, R., 95 Pe´rez-Ortı´n, J. E., 160 Perna, P. J., 147 Persinger, J., 165 Pession, A., 125 Peterson, C. L., 159–163, 165–169, 188, 191 Peterson, L., 159 Peterson, M. G., 67 Petrakis, T., 191 Peyroche, G., 9, 15, 28, 98, 139 Pfannstiel, J., 26 Pfeifer, G. P., 129 Pflugfelder, G., 130 Pham, A. D., 82, 83, 190 Phelan, M. L., 169, 173, 175 Philibert, R. A., 54 Pikaard, C. S., 124, 128, 131–133, 138, 148 Pile, L. A., 82 Pillus, L., 186, 207 Pisano, M., 94, 105, 111, 114 Plassat, J. L., 79 Platas, A. A., 25 Platt, T., 127 Poch, O., 73, 75 Podolny, V., 73, 185 Poglitsch, C. L., 1, 3, 19 Pointud, J. C., 73 Polach, K. J., 171, 174 Polesskaya, A., 186 Pollack, B. P., 218 Poltoratsky, V., 187 Poon, D., 68, 83 Poot, R. A., 163 Pouska, A., 129
AUTHOR INDEX
Powell, W., 23, 24, 25 Prabhakar, B. S., 135 Pradel, J., 186 Prasad, R., 21 Pray-Grant, M. G., 74, 86, 183, 185, 196 Prelich, G., 18 Price, D. H., 22 Pritchard, L. L., 186 Proudfoot, N., 26 Ptashne, M., 56, 79 Pugh, B. F., 14, 67, 68, 76, 79, 84, 85 Puglia, K. V., 94, 96–111, 113, 114 Pullner, A., 17 Purdom, I., 125 Puri, P. L., 143 Purnell, B. A., 78 Purton, T., 27 Putnam, C. D., 132, 133
Q Qi, C., 54, 55, 56 Qian, N., 47, 49, 56 Qin, J., 51–55, 58, 74 Qiu, Q., 196 Quinn, J., 161, 162, 163, 166, 167 Quintin, S., 75
R Rachez, C., 45, 52, 53 Rameau, G., 94, 96, 98, 102 Ramirez, E., 97, 98, 114 Ramirez, S., 186 Ranallo, R. T., 84 Ranish, J. A., 27, 54, 57 Rao, M. S., 54, 55, 56 Raschke, E. E., 17 Rastinejad, F., 188 Ratajczak, T., 100, 109 Ravanpay, A., 134 Ray, E., 207 Rea, S., 206, 209, 213 Recht, J., 204, 205 Reddy, J. K., 54, 55, 56 Reed, R., 26 Reeder, R. H., 124, 126, 127, 131, 132, 133, 138–140
237
Reese, J. C., 68, 72, 73, 74, 79, 83, 86, 185 Reeve, J. N., 29 Reeves, R., 187 Reifsnyder, C., 186 Reinberg, D., 14, 16, 26, 27, 52, 54, 58, 68, 163, 193 Reines, D., 23, 24, 25 Reischl, J., 2 Remacle, J. E., 48, 59 Rhee, E., 83 Rhem, E. J., 54, 55 Rhodes, L., 49 Ricci, A. R., 185 Ricciardi, R. P., 52, 53 Rice, J. C., 195 Rice, L. M., 109 Rice, W. J., 29 Richards, K. L., 9, 11 Richardson, C. C., 21 Richmond, R. K., 69, 73, 175 Richmond, T. J., 69, 73, 175, 182 Richter, C., 28 Rickert, P., 52, 54, 58 Ricupero-Hovasse, S. L., 159 Rigby, P. W., 135 Riggs, D. L., 139 Rine, J., 47 Rittinger, K., 99, 100, 109, 110 Riva, M., 9, 15, 28, 94, 98, 114, 137, 139 Roach, C., 110 Roan, J., 126 Roberts, J. W., 24 Roberts, S., 27, 98 Roberts, S. M., 183, 189 Robin, P., 186 Robinson, K. M., 80 Robyr, D., 195 Robzyk, K., 204, 205 Rodriguez-Navarro, S., 26 Roeder, G. S., 86 Roeder, R. G., 14, 51–56, 58, 59, 62, 67–69, 72, 73, 75, 77–80, 84, 85, 96, 97, 101, 105, 109, 113, 114, 143, 183 Roederg, R. G., 80 Rogakou, E. P., 196 Rogalsky, V., 144, 145 Roguev, A., 207 Romier, C., 69, 73 Rondon, A. G., 26 Ronne, H., 47, 48, 56, 60
238
AUTHOR INDEX
Rosenbauer, H., 127, 128, 131, 141, 146 Rosenblum-Vos, L. S., 49 Rossmann, M. G., 216 Roth, S. Y., 158, 182, 183, 186 Rothblum, L. I., 124, 128, 129, 131, 133, 138, 140, 141, 143–146 Rothman, J. H., 72, 75, 83 Rougeulle, C., 86 Roussel, P., 143 Roy, A. L., 54, 77, 78 Rozenfeld, S., 96, 106, 109, 115 Rubbi, L., 28, 101, 113, 115 Rubin, G. M., 54, 55 Rudd, M. D., 23, 25 Rudkin, B., 186 Rudolph, H., 174 Ruet, A., 94, 98, 114 Ruff, M., 73 Ruhlmann, C., 71, 73 Ruppert, S., 80 Ruth, J., 95, 96, 101, 106 Ruy, S., 52, 54 Ryan, K., 127, 130 Ryan, R., 80 Ryu, G. H., 49, 51
S Sadhale, P. P., 4, 9, 28 Saez-Vasquez, J., 138 Safrany, G., 126, 128 Sagnier, T., 186 Saha, A., 176 Sakai, A., 50 Sakurai, H., 9, 49, 50 Salunek, M., 75, 80 Samkurashvili, I., 18, 24 Samuels, M., 67 Samuelsen, C. O., 48, 59 Samuelsson, T., 48, 59 Sanchez, J. F., 14 Sandaltzopoulos, R., 169 Sander, E. E., 141 Sanders, S. L., 71, 73, 83 Santangelo, T. J., 24 Santori, F., 137 Santoro, R., 148, 163, 204 Santos-Rosa, H., 211 Sargent, D. F., 69, 73, 175
Sarte, M., 162 Sartorelli, V., 143 Sassone-Corsi, P., 75, 76, 195 Sauer, F., 80, 82, 83, 190 Saurin, A. J., 74 Savard, J., 143, 163, 186 Sawadogo, M., 26, 67, 68, 78 Sawaya, M. R., 21, 30 Sayre, M. H., 9, 44, 45, 48, 161, 166, 167 Scafe, C., 44, 46 Schafer, K., 144, 145 Scheek, S., 80 Scheer, E., 76 Scheer, U., 125, 132, 146 Schepartz, A., 14 Scheufler, C., 99, 100, 107, 109 Schibler, U., 14 Schieltz, D., 74, 86, 183, 185 Schiltz, R. L., 74 Schimmack, G., 98 Schlag, E. M., 82 Schlichter, A., 189 Schmid, C. W., 97 Schmidt, M. C., 67, 78, 163 Schmidt, P., 94, 96, 102 Schnapp, A., 128–131, 137, 138, 141, 146 Schnapp, G., 130, 131, 137, 146 Schnitzler, G. R., 162, 167, 172, 173 Schotta, G., 213 Schramke, V., 186 Schramm, L., 94, 97 Schreck, R., 130 Schreiber, S. L., 182, 195, 206 Schroeder, A. J., 50 Schultz, M. C., 97, 131, 133, 139 Schultz, P., 9, 15, 28, 69, 71, 73, 74, 96, 114, 139 Schurter, B. T., 218 Scott, J., 47 Scott, M. P., 160, 161 Scott, P. H., 95, 96, 97 Seelig, H. P., 163 Segall, J., 67 Seither, P., 129, 130, 138 Sekeri-Pataryas, K. E., 196 Sekine, S., 8, 12, 20, 22 Selleck, W., 73, 185 Sells, B. H., 124
AUTHOR INDEX
Sendra, R., 183 Sengupta, S. M., 165 Sentenac, A., 9, 15, 24, 28, 79, 94–96, 98, 100, 101, 105, 106, 109, 113–115, 129, 139 Sethy, I., 94, 96, 98, 102 Sethy-Coraci, I. K., 94, 95, 96, 102, 103, 104, 114 Seto, A. G., 84 Sevenov, M. V., 132 Severinov, K., 20, 28, 29 Sha, W. C., 76, 83 Shaaban, S., 96, 98, 100, 101, 105, 109 Shachar, I., 76 Shao, Z., 74 Sharp, P. A., 14, 67, 94 Shatkin, A. J., 26 Shaw, P. E., 18 Shaw, P. J., 125 Shearn, A., 162 Sheffer, A., 11 Shematorova, E. K., 9, 28 Shen, W. C., 52, 77, 80, 83, 86 Shen, X., 163, 167 Shen, Y., 96 Shi, J. H., 72, 83 Shiekhattar, R., 163 Shilatifard, A., 25 Shiloach, J., 163 Shimizu, Y., 50 Shodai, T., 99 Shpakovski, G., 9, 28, 129 Shu, F., 85 Siaut, M., 9, 15, 28, 98 Siddiqi, I. N., 139 Sidorenkov, I., 19 Sif, S., 172 Sigler, P. B., 14, 71 Sil, A., 159 Silkov, A., 76 Silver, P. A., 215, 217 Silvian, L. F., 30 Simon, G. M., 191 Singer, D. S., 78, 80 Singer, H. A., 141, 146 Singer, M. S., 213 Singh, N., 54, 55 Sinn, E., 67, 96 Sipiczki, M., 49 Sirri, V., 125, 143
Smale, S. T., 14, 77, 78, 126, 140 Smerdon, J. S., 99, 100, 109, 110 Smith, C. L., 166 Smith, E. R., 193 Smith, J. S., 99, 100, 109, 110 Smith, M. M., 196 Smith, S. D., 131, 133, 141, 146 Smith, V. A., 140 Sogo, J. M., 147, 148 Soh, J., 186 Soll, D., 94, 96, 102 Sollner-Webb, B., 126–128, 130, 133 Solomon, W., 45, 52, 53, 80 Solow, S., 80 Song, C. Z., 129, 130, 132 Song, W., 47, 49, 50, 56 Song, Y., 143, 145 Sonu, M. S., 52 Sosunov, V., 25 Sosunova, E., 25 Sousa, K., 183 Spahr, H., 46, 48, 51, 59 Spangler, L., 17 Speer, J. L., 193 Speirs, J., 125 Sperling, J., 218 Spradling, A. C., 54, 55 Spring, H., 132 Stallcup, M. R., 206, 215, 217 Stargell, L. A., 71, 84 Staub, A., 183 Stefanovsky, V. Y., 124, 126, 127, 133, 135, 143 Steffan, J. S., 139 Steger, D. J., 74, 86, 183, 185, 188 Stein, T., 95, 97 Steinmetz, E. J., 26 Steitz, T. A., 21, 30 Stelzer, G., 85 Stern, D., 54, 55 Stern, M. J., 159 Sterner, D. E., 183, 189, 191 Sternglanz, R., 139, 182, 186 Stevenin, J., 74 Stevens, J. L., 54 Stillman, B., 186 Stillman, D. J., 50, 60, 192, 193 Strahl, B. D., 86, 194, 203, 204 Strahle, U., 75 Strasser, K., 26
239
240
AUTHOR INDEX
Strohner, R., 148, 163 Strubin, M., 79 Struhl, K., 26, 78, 79, 83, 182, 183 Stubblefield, B., 54 Stunnenberg, H. G., 76, 141 Stutz, F., 26 Suja, J., 132 Suka, N., 186, 195 Suka, Y., 195 Suldan, Z., 52, 53 Sullivan, E. K., 169 Sullivan, G. J., 133 Sun, X., 52, 54, 58, 193 Sun, Y., 9, 28 Sun, Z. W., 205 Sutcliffe, J. E., 96 Sutton, A., 186 Suzuki, H., 27 Suzuki, Y., 49, 50 Svejstrup, J. Q., 45, 56, 192, 193 Swanson, M. J., 84, 183 Swanson, R. N., 96, 114 Sweder, K. S., 167 Sweetser, D., 29 Swilling, N. W., 72, 81, 86, 190 Syntichaki, P., 188 Szentirmay, M. N., 26 Szymanski, P., 83
T Tabor, S., 21 Tabtiang, R. K., 49 Tafrov, S. T., 186 Taggart, A. K., 84 Tahirov, T. H., 30 Takada, S., 75 Takagi, Y., 27 Tamkun, J. W., 160, 161, 162, 163 Tan, C., 143, 145 Tan, Q., 27 Tan, S., 73, 183, 185, 191 Tanaka, N., 126, 128 Tanese, N., 67, 68, 79, 134 Tang, H., 14, 195 Tao, Y., 52, 80 Taunton, J., 182 Taylor, L. J., 144, 145 Taylor, P., 100, 109
Teichmann, M., 75 Temiakov, D., 30 Tempst, H., 53, 162 Tempst, P., 44, 48, 49, 51–54, 58, 74, 189, 192 Teng, Y., 196 Tengstrom, C., 54 Teunissen, H., 191 Teyssier, C., 215, 217 Thanos, D., 170, 186, 187, 188 Thireos, G., 188 Thoma, F., 96 Thomas, M. J., 25 Thomm, M., 29 Thompson, C. M., 46, 47 Thompson, E. A., 130, 137, 140 Thompson, M. M., 141 Thompson, N. E., 1–3, 8, 11, 19 Thorner, J., 85 Thuault, S., 69, 73 Thuillier, V., 24 Thuriaux, P., 9, 15, 28, 29, 96, 106, 109, 113, 115, 129 Tian, Q., 136 Tiensuu, T., 54, 55 Timmers, H. T., 15, 85 Tini, M., 196 Tjernberg, A., 75 Tjian, R., 16, 45, 52, 53, 54, 67–69, 71, 75–83, 86, 126, 128, 129, 131–134, 136, 137, 140, 188 Tocchini-Valentini, G. P., 95 Todone, F., 1, 2, 4, 9 Todorov, G., 144, 145 Tokomori, K., 68, 78 Tolentino, E., 126 Tong, K. I., 84 Topalidou, I., 188, 191 Tora, L., 69, 71–76, 134, 183 Torchia, J., 196 Torii, H., 99 Tosh, K., 96 Toulokhonov, I., 20 Tower, J., 126 Tran, P. T., 9 Travers, A. A., 188 Treich, I., 47, 49, 56, 159, 166, 167 Treitel, M. A., 47, 159 Tremethick, D. J., 163, 169 Trendelenburg, M. F., 132
241
AUTHOR INDEX
Trere, D., 124, 125 Trievel, R. C., 211 Trinkle-Mulcahy, L., 130, 131, 137 Trouche, D., 186 Trumpower, B. L., 49 Tsai, F. T., 14 Tschiersch, B., 206 Tschochner, H., 9, 15, 28, 44, 139, 147 Tse, C., 183 Tseng, H., 136 Tsukihashi, Y., 80 Tsukiyama, T., 160, 163, 166, 167, 169 Tuan, J. A., 141 Tuan, J. C., 136, 137, 141 Tuck, S., 54, 55 Tudor, M., 54, 55 Tugendreich, S., 98 Turley, S., 110 Turner, B. M., 158, 183, 202, 206 Tzamarias, D., 191
Vasslyleva, M. N., 8, 12, 20, 22 Vassylyev, D. G., 8, 12, 20, 22, 30 Veenstra, G. J., 75 Vente, A., 142, 146 Vermaak, D., 163 Verrijzer, C. P., 67, 68, 77, 78, 80, 191 Vervish, A., 186 Veschambre, P., 52, 54 Vignali, M., 187 Vigneron, M., 129 Viljoen, M., 47 Vingron, M., 130 Virbasius, A., 79, 85 Virbasius, C. A., 79, 83 Vogelauer, M., 195 Voit, R., 132, 133, 141–146 Vollmer, F., 75 von Hippel, P. H., 14 Vu, L., 127, 139
W U Ujvari, A., 18 Um, S. J., 196 Umehara, T., 186 Upadhya, R., 96 Upegui-Gonzalez, L. C., 186 Uptain, S. M., 24 Urnov, F. D., 43 Usheva, A. A., 14 Utley, R. T., 183, 186, 188, 204
V Vallier, L. G., 50 Van, M. V., 113, 115 van der Knaap, J. A., 85 van der Vliet, P. C., 85 van der Zee, S., 14 van Holde, K. E., 175 VanKanegan, M., 165 van Leeuwen, F., 202, 203, 213, 214 Van Mullem, V., 28 van Vuuren, H. J., 47 Varga-Weisz, P. D., 163, 166, 167, 169 Varon, M., 11 Varshavsky, A., 205
Wade, P. A., 163, 169, 175 Wagner, G., 2 Wai, H., 127 Waksman, G., 22 Walker, A. K., 72, 83 Walker, A. P., 99, 100, 109, 110 Walker, P., 126 Walker, S. S., 68, 72, 83 Walkinshaw, M. D., 100, 109 Wallberg, A. E., 188 Wallisch, M., 148 Wang, A., 195 Wang, B., 2 Wang, C. K., 67 Wang, D., 24, 25 Wang, E. H., 78, 80, 82, 86 Wang, G., 54 Wang, H., 218 Wang, J., 18, 21, 67, 83, 136, 143 Wang, M. D., 24 Wang, W., 15, 162, 163, 164 Wang, X., 17, 191 Wang, Y., 182, 194, 195 Wang, Z., 75, 96, 97, 101, 105, 109, 113, 114 Ward, J., 53 Warner, J. R., 95
242
AUTHOR INDEX
Wassarman, D. A., 82, 83 Waters, R., 196 Waugh, D. S., 19, 20 Weber, J. A., 17 Weeks, D. L., 75 Weeks, J. R., 29 Weil, P. A., 67, 68, 71, 73, 83, 85, 183 Weilbaecher, R. G., 23 Weinstein, I., 186 Weinzierl, R. O., 1, 2, 4, 9, 28 Weisenberger D., 146 Weiss, M. A., 2 Weiss, V. H., 215, 216, 217 Weissman, J. D., 78, 80 Werner, F., 1, 2, 4, 9, 28 Werner, J., 57 Werner, M., 24, 28, 98, 113, 115 Werten, S., 69, 73 West, M. L., 18 White, M. F., 169 White, R. J., 95–97, 115 Whitehouse, I., 169 Widmer, R. M., 147 Widom, J., 171, 174, 182 Wieczorek, E., 74, 76 Wilkinson, J. K., 128 Willis, I. M., 94–111, 113, 114 Willy, P. J., 14, 85 Wilm, M., 163, 166 Wilson, C. J., 47 Wilson, J. R., 207, 209, 210, 211, 213 Wilson, S. H., 21, 30 Wind, M., 24 Windle, J. J., 128, 133 Winfield, S., 54 Winsor, B., 67 Winston, F., 68, 79, 83, 86, 159, 160, 183, 189 Winter, A. G., 95, 96, 97 Wintzerith, M., 129 Wittmeyer, J., 176 Wittschieben, B. O., 192, 193 Wolf, V. J., 96 Wolffe, A. P., 43, 75, 81, 86, 163, 169, 204 Wolstein, O., 76 Wong, J., 163 Wood, A., 205, 214 Wood, C., 76, 83 Wood, J. M., 183
Woodcock, C. L., 166 Woodcock, G. R., 24 Wooddell, C. I., 19 Woods, A., 47 Workman, J. L., 74, 81, 82, 86, 148, 161, 162, 163, 165, 166, 167, 169, 173, 183, 185–189, 191, 192, 194, 202, 203 Worman, H. J., 191 Woychik, N. A., 4, 9, 11, 17, 27, 28 Wriggers, W., 29 Wright, A. P., 188 Wu, C., 160, 163, 166, 167, 169 Wu, J., 195 Wu, S. Y., 80 Wu, W. H., 193 Wuite, G. J., 24 Wurtz, J. M., 72, 75, 86 Wyrick, J. J., 46, 47, 51, 52, 60
X Xenarios, I., 195 Xiang, F., 195 Xiao, Z., 50 Xie, J., 73, 79 Xie, W. Q., 131, 133, 141, 146 Xie, X., 73 Xue, Y., 162, 163, 164
Y Yaffe, M. B., 100 Yalamanchili, P., 96 Yamaguchi, K., 130 Yamaguchi, Y., 25 Yamamoto, K., 129, 130, 159, 183 Yamamoto, O., 128 Yamamoto, R. T., 139 Yamamoto, T., 67, 78 Yamamura, S., 53, 55 Yamashita, S., 84 Yamit-Hezi, A., 76 Yan, H., 25 Yang, D., 162 Yang, J. C., 188 Yang, X. J., 47, 74, 81, 82, 86, 193 Yaniv, M., 160 Yankulov, K., 27
AUTHOR INDEX
Yano, K., 129, 130 Yates, J. R., III, 74, 86, 183, 185, 186, 194 Yatherajam, G., 71 Yau, P. M., 195 Yie, J., 170 Yin, Y. W., 30 Ying, C., 187 Yokomori, K., 80 Yokoyama, S., 8, 12, 20, 22, 30 Yoshinaga, S. K., 159 Young, M. K., 139, 163 Young, R. A., 4, 9, 11, 14, 18, 27, 29, 44, 46–47, 51, 52, 54, 55, 56, 60, 80, 83, 84, 86 Yu, D. Y., 196 Yu, G., 50 Yu, Y., 50, 196 Yuan, C. C., 9, 28 Yuan, C. X., 51–56, 58, 59, 62 Yuan, L., 186 Yuan, X., 15 Yudkovsky, N., 27, 57
Z Zahradka, P., 124 Zantema, A., 191 Zaros, C., 9, 15 Zatsepina, O. V., 130, 132 Zaychikov, E., 5 Zehring, W. A., 29 Zeng, L., 86 Zenklusen, D., 26
243
Zentgraf, H., 15, 130 Zerby, D., 75 Zhai, W., 136, 137, 141, 142, 145 Zhang, D., 75 Zhang, G., 28, 29 Zhang, H., 55, 136 Zhang, J., 18, 47, 135 Zhang, K., 195 Zhang, L., 71 Zhang, M., 162, 172, 173 Zhang, W., 183 Zhang, X., 51, 52, 53, 55, 58, 74, 207, 209, 210, 211, 216, 217 Zhang, Y., 52, 54, 58, 163 Zhang, Z., 73, 83 Zhao, J., 15 Zheng, S., 76, 83 Zhou, H., 97 Zhou, J., 83 Zhou, L., 216, 217 Zhou, M. M., 86 Zhou, Q., 67 Zhou, S., 45, 52, 53, 54, 68, 75, 76, 80, 134, 162, 164 Zhou, Y., 204 Zhu, X., 79, 85 Zhu, Y., 54, 55, 56 Zilahi, E., 49 Zillig, W., 29 Zitomer, R. S., 49 Zomerdijk, J. C., 130, 131, 134, 137, 138 Zou, S., 78, 80 Zwicker, J., 83 Zwicker, S., 80
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SUBJECT INDEX
A A-block mutations, 95–96 promoter element, 114 ABC10, 115 Acetyl-CoA binding sites, 82 Acetyl-lysines, 190–191 Acetylation chromatin-modifying complexes and, 182 H4, 188 H3 K14, 195 histone, 203 histone lysine, 182 nucleosomal, 158, 190f Pol I regulation by, 143–144 TAFI63, 144 Acetyltransferase(s) histone, 184t lysine and, 143 MYST, 185–186 in TAF1, 86 TFIIF and, 81–82 Achantamoeba castellanii, 147 Activation, mediators of, 79–80 Activator bypass experiments, 79 Activator-recruited cofactor (ARC), 53 ADA, 183 Ada, 183 Ada2, 185 SANT and, 191 Ada3, 185 Adenovirus Major Late (AdML) promoter, 73 AdoHcy, 211 AdoMet, 209–210 AdoHcy and, 211
in binding pocket, 217 Dot1 and, 214 methionine group of, 216 PRMTs and, 215 ADP, chromatin remodeling and, 167 ADP-ribosylation, 158 AgNOR number, 124 -amanitin, 4 NTP binding and, 22–23 Amino acid identity, 134 Angiotensin II, 141 Antiapoptotic responses, 76 Apoptotic signaling, 204 apoTBP, 69 Apyrase, 166 Arabidopsis thaliana, 124 ARC. See Activator-recruited cofactor Archaea, 28–29 Arginine methyltransferases, 215–218. See also PRMTs Arginine target residues, 218 Arrest, 23–25 ASHI, 207 Aspartate loop, 5–8 residues, 5 ATP binding, 166–167 cofactor, 16 hydrolysis, 166–167, 168, 172 Pol I and, 137 ATPase(s) cofactors, 167 DNA-stimulated, 161 DNA-stimulating, 159 Autoinhibition, 101–102 Autophosphorylating serine/threonine kinase, 80 245
246
SUBJECT INDEX
B B-block promoter element, 95 B-lymphocyte(s) development, 76 TFIID in, 76 Backtracking, 23–25 Bacteria cleavage factors, 29 RNA polymerase of, 28–29 BAF57, 164–165 Basal transcription machinery, 79 Basic helix-loop-helix (bHLH) protein, 136 Basonuclein, 136 Bdf1. See Bromodomain Factor 1 Bdp1 binding, 109 Brf1 and, 101 recruitment, 105, 114 Tfc4 and, 106 TFC4 and, 106 Tfc4 interactions, 111 TPR1-TPR9 and, 113 Bdp1-Tfc4 interactions, 106–107 bHLH. See Basic helix-loop-helix protein Biochemical fractionation, 54 Brahma gene, 160 BRE, 14 Bre1 histone methylation and, 214 Rad6 and, 205–206 RING domain of, 191 Brf1, 15, 94 autoinhibition in, 109 binding, 109, 113 binding-site transition, 113–114 binding to Nt-TPR9, 102 binding to Tfc4, 101–102 conservation in, 97 as a ligand, 105 photo-crosslinking, 114 recruitment of, 102 structural domains of, 97–98 tetratricopeptid repeat domains and, 102–104 Tfc4 and, 98, 105, 111 Tfc4 fragments for, 102t Tfc4 mutant fragments for, 103t
TFIIB-core region of, 98 TPR1-TPR9 and, 113 Brf1-Tfc4 interaction, 97–105, 109 Brf1-TFIIIC, 95–97 BRG1, 160 multisubunit complexes of, 161 Bridge helix during backtracking, 23 bending, 22 movement, 23 BRM, 161 Bromo domain(s), 72, 188–189 containing proteins, 188–189 Gcn5, 190f substrate linking, 191 TAF1 and, 86 Bromodomain Factor 1 (Bdf1), 72, 86 kinase domain of, 190
C c-Jun, 84 C-terminal repeat domain (CTD) coupling and, 26 hyperphosphorylation, 58 Mediator and, 44 modifications, 18 phosphorylation, 13, 14, 18, 44–46 role of, 58 of Rpb1, 8 termination and, 26 truncations, 44 Caenorhabditis elegans Med proteins, 48 Mediator subunits in, 54 RNAi experiments in, 55 RNAi in, 75 Sur-2 protein, 53 TAFs of, 72 TFIID in, 83 Camptothecin, 136 Cancer cells AgNOR score of, 124 Pol II elongation and, 25 Cannonball gene product, 76 Capping, 26 Carboxy-terminal kinase I (CTDK-I), 192
SUBJECT INDEX
CARM1, 215 H3 and, 218 Casein kinase II (CK2), 97, 141 Catalysis, 21–23 CBP. See CREB-binding protein CBP HAT, 191 CBP/p300, 196 cdk. See Cyclin dependent kinase Cdk7/Kin28, 18 cDNA, Pol I and, 129 Cell cycle, G1 phase of, 57 Cell differentiation lymphoid, 143 Pol I during, 143 Cell viability, Srb7 and, 55 CF. See Core factor ChIP. See Chromatin immunoprecipitation assay Chromatin DNA in, 157 modifying complexes, 43 nuclease sensitivity in, 168 Pol I and, 147–149 regulators, 85 SNF and, 159 structure, 168 SWI and, 159 Chromatin immunoprecipitation (ChIP) assay ordered binding and, 57 TAF/TBP ratio and, 83 of TBP occupancy, 79 Chromatin-modifying complexes, 182 Chromatin remodeling apyrase and, 166 complexes, 43 initiation of, 174 mechanisms of, 168–174 Chromatin-remodeling complexes ATP-dependent, 164, 165f ATP hydrolysis and, 166 cofactor requirements, 167 in elongation, 192f enzymes and, 168 HAT complexes and, 187–191 helicase activity in, 167 recruitment of, 188 Chromatin structure changes in, 170f H3 phosphorylation and, 205
247
Chromo domain-containing proteins, 188–189 Chromosomal condensation, 204 Chromosome segregation defects, 50 Circular dichroism, of CTD, 8 cis-acting sequences, 136 CK2. See Casein kinase II Clamp domain, 5 carboxylate, 109 structure of, 8 Cleft domain, 13 Clr4 þ, 206 COMPASS, 207 methylation of, 212 Rad6 and, 214 Set1 subunit of, 211 Coordinate regulation, 95 Core factor (CF), 126, 138 UBF and, 131 in yeast, 139 Core promoter-binding factor (CPBF), 135 Core promoters, 77 Core subunits, 4 Cotransfection assays, USF and, 136 Coupling, to RNA processing, 26 Covalent modifications, nucleosomal, 158 CPBF. See Core promoter-binding factor CREB-binding protein (CBP), 186 CRSP, 54 Cryo-electron microscopy, Pol II core surface and, 5 Crystal shrinkage, 3 Crystallographic backbone models, of Pol II, 9–10 Crystallographic studies, of TBP, 69 Cse1, 50–51 Cse2, 50–51 CSE2 characteristics of, 51 mutations, 50 CSE1 mutations, 50 CTD. See C-terminal repeat domain CTD-phosphatase, 57 CTDK-I. See Carboxy-terminal kinase I CUP1, 52 CYB2 gene, 49 CYC7 gene, 49
248
SUBJECT INDEX
Cyclin C, 47 Cyclin dependent kinase (cdk), 44 Cyst formation, 147
D dATP, 167 Death cap mushroom, 22 Deoxynucleotide triphosphates, 167 Dephosphorylation of Pol II CTD, 13 reinitiation and, 26–27 termination and, 26 Dimer formation, 173 Dimerization, 132 Dinucleosome-like species, 172–173 Dinucleotides, 24 dmTAF2, 68 D468N mutation, 115 DNA affinity columns, 163 conformational changes, 172 in eukaryotic chromatin, 157 melting, 13, 15–16 in nucleosomes, 182 phosphodiester backbone of, 202 promoter, ribosomal, 132 radiolabeled, 173 separation, 20 shuttles, 30 template strand, 16 unwinding, 16 utilization, 158 DNA binding nonspecific, 14 TBP and, 84 DNA/Chromatin, 163 DNA duplex, 19 DNA microarray analysis, 51 DNA polymerase Klenow, 30 single-subunit, 21, 29–30 DNA-protein crosslinking, site-specific, 111 photocrosslinking, 94 DNA repair, 195–196, 204 Rad6 and, 205 DNA-RNA hybrid, 13 backtracking and, 24
clamp in, 8 destabilization of, 24, 25 elongation and, 19 maintaining, 20 DNA templates bases, 30 chromatin-assembled, 53 DNAse I digestion patterns, 171 footprinting, 78, 114 hypersensitivity, 160 TFIID and, 73 DNAse sensitivity, 172 Dock domain, 15 Domain-containing proteins, 189t Dot1, 213 deletion, 202–203 lysine target residues, 214 structure of, 214 Downstream promoter element (DPE), 77 DRIP complex. See Vitamin D3 receptor interacting proteins complex Drosophila hsp 70, 160 Drosophila melanogaster Brahma gene, 160 HAT proteins in, 193 Med78 and, 55 P insertions in, 54 polycomb group proteins in, 74 polythene chromosomes, 57 TFIID and, 83 TRF1 in, 75
E E4 ligase, 205 E1A protein, 53 EDTA, 209 EIBF, 135 Electron microscopy, of Pol II, 1 Elongation, 13, 18–20 chromatin modifiers and, 194 deregulation of, 25 factors, 2 HAT proteins in, 192–194 inhibiting, 22–23 pause and, 24 phosphoproteins in, 18 in Pol I, 146–147
249
SUBJECT INDEX
TIF-IC and, 131 Topoisomerase I and, 136 transition to, 18 Elongator complex destabilization of, 25 Elp3 in, 192–193 functional model of, 18–19 HAT proteins in, 193 stability, 19 Elp3, 192–193 ELP3, 193 EM structures, 60 Embryogenesis, 55 Embryonic development, Srb7 and, 55 Enhancer sequences, 133 Enhancers, 126–127 Enzymatic activities, of TFIIF, 80–82 Epigenetic mechanisms, 128 Epigenetic regulation, 194–195 Epithelial cells, Pol I in, 145 ES cells, 55 Esa1, 196 Escape commitment, 16 Estrogen receptor , 56 Exonuclease III footprinting, 18 E(z) preSET regions of, 209 SET domains and, 207
F Factor C, 130 Fcp1, 27 Fidelity, 21–23 Finger proteins, 136 Foot domain, 13 Funnel, 5
VP16, 50 Gal11, 49–50 interactions with, 55 med2 and, 52 GAL11, 50 Gal4-E1A, 53 Gal4-VP16, 53, 57 Galactose metabolizing enzymes, 49 Mig1 and, 47 ROX3 and, 49 Snf1 and, 47 Gcn4, 55 Gcn5, 182 bromo domain, 190f H3 K14 and, 195 HAT complexes and, 183 roles of, 188–189 SAGA and, 189 Gen5 HAT activity, 184 Gene regulation, global, 51–52 General transcription factors (GTFs), 43–44 TBP and, 52 Genes GAL4-regulated, 48 GCN4-regulated, 48 MAT1-regulated, 48 Gluatamates, in PRMTs, 217 Glucocorticoid(s) receptor, 56 rRNA synthesis and, 140 Glucogenic growth, 47 Glucose repression, 50 Snf1 and, 47 GreA, 29 GreB, 29 GST-TBP, 68 GTFs. See General transcription factors
G G1 phase, 57 GAL1, 49 GAL4, 173 fusion proteins, 161 nucleosome complex, 162 Gal4 Gal11 and, 55 Srb4 and, 55–56
H H1 nucleosomes and, 157–158 Pol I and, 148 H3, 139 CARM1 and, 218 COMPASS and, 207
250
SUBJECT INDEX
H3 (cont.) Dot1 and, 214 Lys4 of, 213 lysine residue methylation of, 211 phosphorylation of, 204–205 R17 of, 218 R26 of, 218 H4, 139 acetylation of, 188 PRMT1 and, 218 R3 of, 218 Rmt1 and, 218 H3/H4 heterodimers, 201–202 H3 K14, 195 H3 K79, 214 H2A modification, 205 Hamster cells, TFIID and, 83 Hat1, 182 HATs. See Histone acetyltransferase(s) H2A.X, 195 H2B, 191 H2B/H2A heterodimers, 202 hBRG1, 190 hBRM, 160 Hda1, 195 HDACs. See Histone deacetylases Heat shock-induced pathways, 204 Heat shock promoter, 57 Heavy atom clusters, 3 Heavy-metal compounds, nonstandard, 13 HeLa cells, 53 Helicase activities ATP-dependent, 16 in chromatin-remodeling enzymes, 167 Heterochromatic gene silencing, 213 Hexokinase 2, 49 hGcn5, 74 HIS3, 188 Histone acetyltransferase (HAT) complex(es), 74, 86, 182–187 chromatin-remodeling complexes and, 187–191 CREB-binding protein and, 186 DNA repair and, 195–196 in elongation, 192f PHD and, 190–191 recruitment of, 188 regulation of, 186–187
RING and, 190–191 in S. cerevisiae Mediator, 44 SANT and, 190–191 transcription and, 195–196 transcription regulation and, 183–185 transcriptional elongation and, 192–194 type A, 182 Histone acetyltransferases (HATs), 81–82, 158, 184t catalytic domains, 190 Elp3, 192–193, 194 HBO1, 186 Histone code, 194–195, 203 hypothesis, 86 Histone deacetylases (HDACs), 158 histone deacetylation and, 204 identification of, 182 Histone fold, 201 domains, 72–73 Histone methyltransferases catalytic data of, 211f H3 K14 and, 195 Histone octamer peeling, 171–172 transfer, 169, 173–174 Histone proteins arginine methyltransferases and, 215 residues, 205f Histone tails, 202 covalent modifications, 203 modification sites, 203f phosphorylation of, 204 Histone(s) acetylation, 203 acetylation of, 81 core, 181–182 deacetylation, 204 H1, 157 lysine residue methylation of, 211 methylation, 206 modifications, 182 monoubiquitination of, 82 ubiquitination, 205–206 HMG boxes, 131–132 HMG proteins DNA binding of, 164–165 HAT proteins and, 187
251
SUBJECT INDEX
HMKTs methylation of, 212 methyltransferase activity of, 211 HMTase, 207 HO activation, 57 RGR1 and, 50 transcriptional regulation of, 188 HO endonuclease, 158–159 HO transcription, 49 Holoenzymes, 8 Pol I, 138, 149 Pol II, 44, 46, 51, 61f promoter recruitment of, 14 Srb proteins and, 46–47 Hop TPR-ligand complexes, 109 Hortologous factors, 134 HP2 recruitment, 213 hpr1, 51 Hrs1/Pgd1 Gcn 4 and, 55 med2 and, 52 hSRB7, 53 hSRB10, 53 hSRB11, 53 hsTAF4, 73 hsTAF4B, 76 hsTAF6, 76 Human origin recognition complex, 186 HXK2, 49 H190Y, 103 mutation, 108 hydroxy radical cleavage, 5 Hypertranslocation, 24
I IME1, 50 Inchworming, 23 Induced fit, 30 INF- promoter, 188 Initiation, 13 complex assembly, 14–15 TFIID and, 67–68 Initiation-elongation transition, 15–18, 17–18 Initiator (Inr), 77–78 INO1, 188
Inr-binding proteins, 78 Insulin, rRNA synthesis and, 140 Intergenic spacer region, of rDNA repeats, 125 Interspecific crosses, 127 Invertase, 49 ISWI nucleosome disruption and, 171 NURF and, 163 ISWI-chromatin, 148
J JBP1, 215 JBP1/PRMT5, 218
K K4, 213 Kin28, 44–46 Srb4 and, 52 KIN28 mutation, 50 Kinase activity, TAF1 and, 81 Kinase(s) cyclin-dependent, 96 domains, 81 Ku antigen, 135
L LEF-1, 131 Leucine zipper motif, 51 LexA-Med1 fusion, 48 Ligand binding identification of sites, 108–109 PTS1 peptide and, 111 in tetratricopeptide repeats, 104f, 107f in TPR arrays of Tfc4, 108–111 Ligand bonding, of TPRs, 100 L469K, 106 mutation, in TPR7, 109 Lobe, 5 Lower jaw, 5 Lymphocyte(s) rRNA activation and, 146 stimulation, 141
252
SUBJECT INDEX
Lys4, 211 methylation of, 213 Lys9, 211 Lys20, 211 Lys27, 211 Lys36, 211 Lysine acetylation of, 143, 203 residues, 212f target residues, 211–213, 214 Lysine methyltransferases, 204f, 206–214 non-SET domain containing, 213–214
M Male X-chromosome dosage compensation, 193 MAP Kinase, 55 MCM1 transcription, 49–50 2-MDa, 54 Med1, 48 MED1, 48 Med2, 48, 50 MED2, 48 deletion, 50 med2, 52 Med4, 50–51 Med6, 48, 52 embryogenesis and, 55 MED6, 48 Med7 Cse2 and, 50–51 embryogenesis and, 55 MED8, 49 Med8, 49 Med11, 49 Med78, 55 Med proteins, 48–49 Med230/sop-1 activity, 55 Med130/sur-2, 55 Med220/TRAP220, 55 Mediator complex(es), 27 analogues of, 53–54 CTD and, 58 function of, 43 head region, 60–62 in higher eukaryotes, 52–55 human, 52 identification of, 44
mammalian, 53–54 med2, 52 metazoan, 52 metazoan subunits, 54–55 metazoan v. yeast, 59 middle region, 60 mouse, 54 Rgr1 in, 60 RNA polymerase and, 44–46 S. cerevisiae subunits of, 45t S. cerevisiae v. human, 58 S. cerevisiae v. S. pombe, 59 Saccharomyces cerevisiae, 43–52 SBF and, 57 scaffolds, 57 Srb4 on, 60 structure-function, 58–62 subunit composition in yeast of, 46–51 tail region, 59–60 TFIID and, 80 transcriptional activation in, 55 Messenger RNA. See mRNA Metal A, 5 location of, 21 Metal B, 8 location of, 21 NTP and, 21 Methyl group transfer, 212f Methylation arginine, 215 chromatin-modifying complexes and, 182 histone, 206 histone residues and, 205f MF1 transcription, 49 Mig1 repressor, 47 Misincorporated nucleotides, 25 Modular functional domains, of SAGA HAT complex, 185 MOF, 193 Monoubiquitinating activity, 82 Monoubiquitination, of H2B, 191 Mot1, 85 Motifs, in TAFs, 72 MOZ, 185–186 mRNA, 124 processing, 26 proofreading, 25 synthesis, 1
SUBJECT INDEX
mtTF1, 131 Multicopy suppression, 109 Mushroom toxin, 4 elongation and, 22–23 Mutagenesis, site-directed, 104 Mutations, gain-of-function, 102 MYST acetyltransferases, 185–186
N N-CoR, 191 NAD, 144 Nascent transcript, 16 proofreading of, 25 NAT, 54 NC2, 51, 85 NF-kappaB, 53 NG-monomethylarginine, 215 NGN0 G-asymmetric dimethylarginine, 215 NGNG-symmetric dimethylarginine, 215 Nhp6 proteins, 96 NoRC. See Nucleolar remodeling complex NORs. See Nucleolar organizer regions Nt-TPR5, 113–114 Nt-TPR9 Brf1 binding to, 102 H190Y and, 103 inhibition isotherm of, 101 NTP. See Nucleotide triphosphate NuA3, 193 NuA4 HAT complex, 196 Nuclear extract, human, 128 Nuclear magnetic resonance of CTD, 8 Pol II and, 1 Nuclear receptor (NR), 56 Nuclease sensitivity, in chromatin, 168 Nucleic acids cleavage of, 30 polymerization of, 30 Nucleolar dominance, 127 Nucleolar organizer regions (NORs), 124 Nucleolar remodeling complex (NoRC), 148 Nucleolus, Pol I and, 125 Nucleophilic attack, 211
Nucleosomal acetylation, 190f Nucleosomal plasmid supercoiling, 169 Nucleosome-nucleosome interactions, 169 Nucleosome remodeling, 158, 203 as enzymatic process, 168 Nucleosome-remodeling complexes, 158–163 enzymes, 163–168 factor, 148 Nucleosome structure, 201–202, 203f bulging/looping model of, 174, 176 spooling/peeling model of, 174–175 twist model of, 174, 175–176 Nucleosome(s) chromatin in, 201 composition of, 157 crystal structure of, 175 displacement, 173 disruption, 171–172 DNA in, 182 entry/exit point of, 164 histone-DNA contacts of, 175 histone-induced supercoiling in, 202 multisubunit complexes of, 161–163 passive movement of, 169 sliding, 169–171 spacing, 171 transcriptional regulation and, 181 Nucleotide addition cycle, 21 Nucleotide triphosphate (NTP), 21, 167 20 -OH group of, 21 NURF, 163 ATPase activity and, 167 NUT1, 49 Nut1 Gal 11 and, 50 proteins, 49 NUT2, 49 Nut2 embryogenesis and, 55 proteins, 49
253
254
SUBJECT INDEX
O O-helix, 22 Oligonucleotides, in pause and arrest, 24 ORC1 subunit, 186 Oscillation. See Backtracking
P p53, 145 monitor cell proliferation, 144 tumor suppressor, 96–97 P/CAF HAT complex, 74 p38 MAPK pathways, 195 p42/p44 MAPK pathways, 195 p55 protein, 182 PAF53, 130 UBF and, 137 Pausing, 23–25 PC2, 53–54 PC4, 54 Pc-G proteins. See Polycomb group proteins PC3/topoisomerase I, 54 PCAF bromo domain of, 190 recruitment of, 143–144 in transcriptional elongation, 193 PCF1-1, 102–103 PCF1-2, 102–103 Perixisome proliferator-activated receptor , 56 PEX5, 111 PEX5/PTS-1 peptide cocrystal structure, 109 Pgd1, 50 PGD1 deletion, 50 PHD domain, 190–191 regulation through, 191 PHO5, 192 Phorbol esters, 140 Phosphate starvation, 174 Phosphodiester bonds, 22–23 Phosphoproteins, in elongation, 18 Phosphorylation chromatin-modifying complexes and, 182 of CTD, 14
of H3, 204–205 of histone tails, 204 initiation-elongation transition and, 18 nucleosome and, 158 of Pol II CTD, 13 serine, 142 serine 5, 18 of UBF, 140–143 Photocrosslinking to DNA, 114 DPE interactions, 78 TAF2 and, 78 Phytohemagglutinin, 141 Pol I elongation and, 146 PIC assembly, 71–72 Plant hybrids, interspecific, 128 Pleiotropic effects, 50 Pocket proteins p107, 96 p130, 96 Point mutations, on TAFs, 73 Pol I. See RNA polymerase I Pol II. See RNA polymerase II Pol II-dependent genes, 43 Pol II-Mediator complex, 27 Pol II-TFIIS complex lid, 20 rudder, 20 structure, 20, 25 zipper, 20 Pol III. See RNA polymerase III Poliovirus protease, 96 Polycomb group (Pc-G) proteins, 160 Polycomb repressive complex 1 (PRC1), 74 Polymerase recycling, 26–27 Polymerase-TFIIS complex, 11–13, 12f Pore domain, 5 elongation and, 19 metal B and, 8 Post-SET domains, 207–209 p67phox, 110 pRb. See Retinoblastoma protein PRC1. See Polycomb repressive complex 1 Pre-SET domains, 207–209 Preinitiation complex assembly, 95–97
255
SUBJECT INDEX
formation of, 56 TBP and, 170 PRMT1, 215, 218 PRMTs AdoMet and, 215 arginine residues and, 215 arginine target residues in, 218 catalytic domain of, 215–216 catalytic mechanism of, 217–218 properties of, 208f sequence alignment of, 216f structure of, 215–217 type I, 215 Processivity, 18–20 Proliferator-activated receptor , 56 Prometaphase TAFI110 during, 142 TBP during, 142 Promoter clearance, 16 escape, 13, 18 proximal, 133 recognition, sequence-specific, 14 structures, 79 swapping experiments, 80 type II, 93 type III, 93 Promoter DNA binding, 13, 14 duplex, 15 loading, 14–15 melting, 14 Promyelocytic leukemic cells, 143 Proofreading, 23–25 Protein-DNA crosslinking elongation and, 18 site specific, 14 Protein kinase AMP-activated, 47 in CTD phosphorylation, 44–46 cyclin C-dependent, 47 UBF and, 141 Protein-protein interactions, on TFIID, 69 Proteins polycomb group, 74 viral-transforming, 97 Proteolytic degradation, 205
Protrusion, 5 Proximal promoter, 126 PTRF, 148 PTS1 peptide, 111 Putative null mutations, 55
R R17, 218 R26, 218 Rad6 Bre1 and, 205–206 histone methylation and, 214 ubiquitin in, 205 RAR, 56 Ras, 55 RB. See Retinoblastoma protein Rb pocket, 145 rDNA in Arabidopsis thaliana, 124 chromatin structure of, 147 gene, 125 gene, eukaryotic, 125f growth-regulated transcription of, 130 promoter, 127 promoter region of, 137 repeats, 125 space-promoters, 127 synthetic template, 128 terminators, 127 transcription, 129 in Xenopus, 124 rDNA transcription cis-acting sequences in, 136 Ku antigen and, 135 protein factors in, 136 SL1 in, 136–137 UBF in, 136–137 Regulation, 26–27 by tumor suppressor proteins, 144–146 Reinitiation, 26–27 of Pol I transcription, 130 Repression, ROX3 and, 49 Restriction enzymes, in nucleosomes, 168 Retinoblastoma protein (pRb), 144–145
256 Retinoblastoma protein (RB), 96 Retinoic X receptor , 56 Rgr1, 49–50 in Mediator complex, 60 N-terminal domain, 50 RGR1 HO and, 50 SUC2 and, 50 rgr1, 50 Rgr1/TRAP170 subunit, 59 Ribonucleoprotein (RNP), 9 Ribosomal RNA. See rRNA Ribosome production, 124 RING domain, 190–191 regulation through, 191 RMR1, 49 Rmt1, 218 RNA 30 -end, 24 30 -end processing, 26 chain elongation, 21 export, 26 extrusion of, 24 interactions, 17 maturation events, 13 polymerization activity, 129–130 processing, 26 processing factors, 13 ribosomal, 27–28 surveillance, 26 transcripts, 13 transfer, 27–28 50 -RNA capping, 18 RNA interference (RNAi), 54–55 TLF and, 75 RNA-Pol II contacts, 16–17 RNA polymerase archaeal, 28–29 bacterial, 28–29 holoenzyme, 8 IIB, 46 Mediator interactions with, 44–46 nuclear, 77 single-subunit, 29–30 spacer-promoters, 127 structure, 29 subunits, 5t ten-subunit core, 4–8 in transcriptional arrest, 24
SUBJECT INDEX
RNA polymerase I (Pol I), 3 A34.5 subunit, 28 A49 subunit, 28 acetylation and, 143–144 assembly, 136–138 associated factors, 129–131 auxiliary factors, 131–136 during cell differentiation, 143 chromatin and, 147–149 core enzyme, 129 DNA melting in, 16 elongation in, 146–147 factors in, 135–136 glucocorticoids and, 140 H1 and, 148 holoenzyme, 138, 149 insulin and, 140 mitotic deactivation of, 142 nucleolus and, 125 PAF53 and, 130 phorbol esters and, 140 posttranslational modification of, 140 pRb and, 145 proximal promoter, 126 regulation of, 147 regulatory mechanisms, 140–147 reinitiation, 130 role of, 123 serum deprivation and, 140 SL1 and, 133 subunits, 129 termination, 127 TIF-IC and, 131 transcription, 124, 126 transcription system, 129 UBF and, 131 viral infection and, 140 vs. Pol II, 27–28 in yeast, 138–139 yeast v. mammalian, 129 RNA polymerase II (Pol II), 1 12-subunit, 9–11 10-subunit core, 1 atomic core structures of, 3–4 backtracking, 23–24 bridge helix of, 22 complete yeast, 10f core elongation complex, 12f core surface, 4
SUBJECT INDEX
crystallographic backbone models, 9–10 CTD, 8, 13, 58 dissociation of, 13 elongation complex, 19 Elp3 in, 192 fidelity, 21 function of, 13–27 holoenzyme, 46, 61f human, 52 hyperphosphorylated, 56 molecular weight of, 3 movement, 23 NTP binding site, 21 preinitiation complex, 43–44 recruitment, 56 recycling, 13 RNA chain elongation by, 21 Rpb4/7 interaction, 10 saddle domain, 17 Soh2 and, 51 specificity of, 21–22 Srb4 on, 51 structural conservation of, 2–3 structural elements of, 5, 6–7t structural studies of, 2t structure determinations of, 3–4 structure of, 3–13 subunits, 4 TAFs, 134 TFIIH and, 16 TFIIS in, 11 transcribing complexes, 16 transcribing genes, 77 transcription factors, 14 translocation, 22 upstream face of, 15 v. Pol I, 124 vs. other polymerases, 27–30 RNA polymerase III, 3 C17/C25 complex, 28 C11 subunit, 28 C31 subunit, 28 C34 subunit, 28 C82 subunit, 28 effectors, 115 in eukaryotes, 93 factors, 114–115 gene transcription, 97 genes, 95 reporter gene transcription, 105
257
Tfc4 interactions with, 112f type II genes, 94 v. Pol I, 124 vs. Pol II, 27–28 RNA synthesis, 13 Pol II specificity and, 21 RNAi. See RNA interference RNAPII, 192. See also RNA polymerase II 5S RNA, 93 RNase P RNA, 93 7SL RNA, 93 RNP. See Ribonucleoprotein Rossman fold, 216 ROX3, 49 ROX3, 47 Nut proteins and, 49 Rox3 protein, 49 RPA49, 130 RPABC10 Brf1 and, 101 Rpb1, 4 aspartate loop, 8 C-terminal region of, 8 trigger loop, 12 Rpb2, 4 acidic residues in, 8 zinc ions in, 8 Rpb4, 9 Fcp1 and, 27 Rpb5, 4 Rpb6, 4 Rpb7, 9 N-terminal RNP-like domain of, 11 Rpb8, 4 Rpb9, 4 transcribing complex stability and, 17 Rpb10, 4 Rpb12, 4 Rpb4/7 complex, 1, 4, 9 C17/C25 and, 28 counterparts of, 28 function of, 15 Pol II interaction, 10 in recycling, 27 rpb4 deletion strain, 3 RPC53, 115 Rpd3, 195 RRN3, 147 dissociation of, 138 RRN5, 139
258
SUBJECT INDEX
RRN6, 139 RRN7, 139 RRN9, 139 RRN10, 139 RRN11, 139 RRN3/Pol I complex, 138 RRN3 protein, 130–131 RRN3/TIF-IA, 15 in yeast, 139 rRNA gene structure, 124–127 lymphocytes, 146 mammalian promoter, 126 mitogenic stimulation of, 141 p53 and, 145 Pol I-transcribed, 95 precursor, 123 production, 124 species-specific transcription, 127–128 subunits, 123 synthesis, 141 transcription, 128–136 RSC, 173 bromo domains of, 189 nucleosome particles and, 165 structural analysis of, 166 Rsc1, 189 Rsc2, 189 Rsk-2 kinase, 195 RYE genes, 47
S S-adenosyl-L-methionine, 207 Saccharomyces cerevisiae assembly in, 94 CTD of, 44 histone ubiquitination in, 205 Med proteins of, 48 Mediator complex, 43–52 Mediator complex identification, 44 Mediator subunits of, 45t methylation in, 213 Pol II of, 3 rpb4 deletion strain of, 3 Rpb4 in, 9 Srb4 ts strain and, 51 transcriptional activation in, 55
Saccharomyces pombe histone methyltransferase in, 206 Med8 in, 49 mediator complex of, 59 Saddle domain, 15 Pol II, 17 SAGA, 74 bromo domains of, 188 Gcn5 and, 183–184, 189 transcriptional regulation and, 188 SAGA HAT complex, 57, 185 SAM binding site Dot1 and, 214 on PRMTs, 215–216 SANT, 190–191 regulation through, 191 Sas2, 185–186 Sas3, 195 SBF, 57 Scaffold proteins, UBF as, 132 Schizosaccharomyces pombe, TFIIIC in, 95 scMediator, 59 head region of, 60 ScTfc4, 100 Nt-TPR9 region of, 101 Selectivity factor 1 (SL1), 133–135 phosphorylation status of, 142 in rDNA transcription, 136–137 species specificity of, 128 TBP/TAF and, 134 SEP15, 49 Sequence markers, in Pol II structure analysis, 3 Serine 5, 18 484, 142 phosphorylation, 142 in UBF, 140 Serum deprivation, 140 SET domains, 206–213 catalytic core of, 210 catalytic mechanism of, 210–211 properties of, 208f structure, 207–210 tyrosine residue in, 210 Set2 histone methyltransferase, 194 SET2-RNAPII, 194 SII. See TFIIS Sin4, 49–50 Gal 11 and, 50
SUBJECT INDEX
SIN4, 47 GAL1 and, 50 Nut proteins and, 49 sin4, 50 ScMediator and, 59 Sin5, 50 Sir2, 214 Sir4, 214 SL1. See Selectivity factor 1 SLIK/SALSA Gcn5 and, 183 SMCC. See SRB/MED-containing cofactor complex SNF, 159, 161 SNF genes characterization of, 158–160 isolation of, 158–160 Snf1 kinase, 47 snf1/mig1 cells, 47 snf1 suppression, 48 SNF2h, 148 soh1, 51 Soh2, 51 soh2, 51 Soh4, 51 soh4, 51 Soh1 protein, 51 sop-1, 55 Space-promoters, rDNA, 127 Specificity, 21–23 Sporulation deficiency, 50 SRB2, 46 Srb4 Gal4 and, 55–56 on Mediator complex, 60 ts strain, 51 SRB4, 46 Srb5, 46 SRB5, 46 Srb7, 55 Cse2 and, 50–51 human homologue of, 52 Srb8, 47 SRB8, 47 Nut proteins and, 49 Srb9, 47 SRB9, 47 Nut proteins and, 49 Srb10, 47 SRB10, 47
259
Nut proteins and, 49 Srb11, 47 SRB11, 47 Nut proteins and, 49 Srb8-11 module, 47 SRB genes, 46 SRB/MED-containing cofactor complex (SMCC), 51, 53 Srb proteins, 46–47 Srb10/Srb11 CDK-cyclin pair, 54 Srb10-Srb11 cyclin kinase complex, 48 SREBP-la, 53 SRY, 131 SSN2, 52 SSN7, 49 Ssn6-Tup1 corepressor complex, 47 STAGA complex, 74 Gcn5 and, 183 Sth1, 189 SUA7, 51 SUC2, 49 RGR1 and, 50 transcriptional regulation of, 188 SUC2 gene, 159 Sucrose Mig1 and, 47 Snf1 and, 47 utilization, 159 sup9e-A19-supS1 gene, 102 sur-2, 55 Su(var)3-9 histone methyltransferase in, 206 preSET regions of, 209 SET domains and, 207 SUV39H1, 206 SUV39H2, 206 SWI, 159, 161 Swi3, 191 SWI genes characterization of, 158–160 isolation of, 158–160 SWI/SNF, 161 DNA affinity columns and, 163 DNAse I digestion and, 171 DNAse I footprinting and, 162 HAT complexes and, 187 nucleosome particles and, 165 promoter activation and, 170
260
SUBJECT INDEX
SWI/SNF (cont.) structural analysis of, 166 transcriptional regulation and, 188 SWI2/SNF2 ATPase(s), 164f helicase activity of, 167 subunits, 166 Swi/Snf chromatin-remodeling complex, 57 Swi4p, 49 Swi6p, 49 Synthetic reporter gene, 49
T T7, 30 T-lymphocyte development, 76 T1671 mutation, 108 T3-response elements, 53 TAF1 acetyltransferase activity in, 86 C-terminal kinase domains, 81 HAT activity in, 81–82 human, 80 monoubiquitinating and, 82 promoter DNA and, 78 TAF2, 78 TAF6, 78 TAF9, 78 TAF12, 73 TAF250 kinase domain of, 190 recruitment and, 188 TAF9-TAF6 heterotetramer, 73 TAF-TAF interactions, 69 TAFI63 acetylation, 144 human v. mouse, 144 TAFI110, 142 TAFs. See TBP-associated factors Tailed template, 19 TATA-binding protein (TBP), 14, 52, 67–68, 69–72, 134f archeal homologues of, 29 C-terminus of, 69, 72 crystallographic studies of, 69 GAL4 and, 161 N-terminus of, 69, 72 nucleosome structure and, 175
occupancy of, 79 Pol II and, 15 preinitiation complex and, 170 during prometaphase, 142 SL1 and, 134 subcomplexes, 69 TFIIA and, 84 v. TRF1, 75 in yeast, 139 TATA box, 14, 77–78 TFIID and, 67 TATA sequences, 175 TBP. See TATA-binding protein TBP-associated factors (TAFs), 52, 70t, 72–74 basal transcription machinery and, 79 C-terminal acidic domain of, 72 conservation in, 72 D. melanogaster, 72 histone fold domains and, 72–73 histone-like, 73 lobes, 73–74 mapping, 73 motifs in, 72 point mutations on, 73 SL1 and, 134 subcomplexes, 69 variant, 76 yeast, 68 yeast viability and, 82 TBP-containing complexes, 68 TBP-DNA complex, 71 NC2 and, 85 TAF and, 84 TBP-free TAF complex (TFTC), 76 EM structure of, 74 Gcn5 and, 183 TBP-like factor (TLF), 75 TBP-sans-TAFs complex, 76 TBP/TAF complexes, 135 TBP-TAF interactions, 69 TCF-1, 131 Telomeres COMPASS and, 207 Gal11 and, 50 Termination, 13, 26–27 Terminators, rDNA, 127 12-O-tetradecanoylphorbol-13-acetate (TPA), 144
SUBJECT INDEX
Tetrahymena thermophila, 182 Tetratricopeptide repeats (TPRs), 94 arrays, 101 crystal structures, 108 groove, 99 ligand-binding in, 107f ligand-binding sites in, 104f models for, 109–111 motifs, 101 mutations of, 102–104 organization of multiple, 110f phylogenetic analysis of, 108 structural model of, 99f structure of, 98–99, 109 superhelical array, 109 superhelix, 99, 108 in Tfc4, 98–101 Tfc1, 114 Tfc4, 94, 114–115 A-block promoter element and, 114 ABC10 and, 115 Bdp1 and, 106 Bdp1 interactions, 111 Brf1 and, 105 Brf1 binding to, 101–102 Brf1 interactions, 98, 111 fragments for Brf1, 102t gain-of function mutations in, 96 interactions with Pol III factors, 112f ligand binding, 108–109 modeling, 108 mutant fragments, 103t photo-crosslinking, 114 tetratricopeptide repeats in, 98–101 in TFIIIB, 103–104 TPR arrays of, 101 TPR models for, 109–111 TFC4, 106 Tfc7, 114 TFIC, 130 TFIIA, 14 GAL4 and, 161 interactions, 84 phosphorylation of, 80 reinitiation and, 27 TFIIB, 14
archeal homologues of, 29 formation of, 43–44 Pol II and, 15 SANT and, 191 Soh4 and, 51 TFIIB-DNA complex, 94 TFIIB-related factor 1. See Brf1 TFIID, 14 alternative, 75–76 in B-lymphocytes, 76 components, 68–76 DNA binding of, 81 DNase I and, 73 DNase I footprinting, 78 footprint, 68 formation of, 43–44 functions, 76–82 initiation and, 84 Mediator complex and, 80 necessity of, 83 promoter recognition, 77–79 purification of, 68 recruitment of, 188 regulation of, 84–85 reinitiation and, 27 requirement for, 82–84 SL1 and, 134 structure, 68–76 subcomplexes, 80 subunit structure of, 71f subunits, 68–69 TAF250 and, 188 TATA box and, 67 TBP/TAF and, 134 transcription initiation and, 67–68 TFIID-Chromatin, 85–86 TFIIE, 14 archeal homologues of, 29 formation of, 43–44 Gal11 and, 50 reinitiation and, 27 TFIIF, 14 enzymatic activities of, 80–82 formation of, 43–44 Pol II and, 15 TFIIH, 14 DNA melting and, 15–16 formation of, 43–44 Gal11 and, 50
261
262 TFIIH (cont.) Kin28 and, 45 kinase activity, 52 Pol II and, 16 Pol II transcription and, 16 reinitiation and, 27 TFIIIA, 93 TFIIIA-5S gene complex, 94 TFIIIB, 94 assembly, 94, 95 mitotic repression and, 96 SL1 and, 134 TATA-directed assembly of, 97–98 TBP/TAF and, 134 Tfc4 fragments in, 103–104 TFIIIC-dependent assembly of, 111 TFIIIC-DNA and, 111–114 TFIIIB-DNA complex, 95 TFIIIC, 93 B-block binding subunit, 106 human, 96 poliovirus protease and, 96 subunits, 94 TFIIIB interactions, 96 TFIIIC2, 96 TFIIIC-DNA Brf1 recruitment by, 101 TFIIIB and, 111–114 TFIIS, 2, 11–13 archeal homologues of, 29 binding, 22 cleavage modes and, 30 contraction, 24 Pol II and, 4 Pol II elongation and, 25 proofreading and, 25 TFIIS-mediated rescue, 25 TFS, 29 TFTC. See TBP-free TAF complex Thermus aquaticus, 28 Thymine DNA glycosylase, 196 Thyroid hormone, 53 Thyroid hormone receptor coactivator complex (TRAP), 51, 52 purification of, 53 TIF-IA, 130 RRN3 and, 130–131 TIF-IA/RRN3, 130, 137
SUBJECT INDEX
TIF-IC, 131 Pol I elongation and, 146 TIP5, 148 Tip60, 185–186 TLF knockout mice, 75 Topoisomerase I, 136 TPA. See 12-O-tetradecanoylphorbol-13acetate TPR2, 102 TPR7 function of, 106 L469K mutation in, 109 TPR9, 109 TPR arrays, 106 TPR1-TPR5, 113 TPR1-TPR9 Bdp1 and, 113 Brf1 and, 113 H190Y and, 103 TPR6-TPR9 ligand-binding channel of, 106 ligand-binding grooves of, 113 non-TPR consensus residues, 108 TPRs. See Tetratricopeptide repeats TR, 56 Tra1, 183 trans, histone octamer in, 173–174 Transcribing complexes, 17 Transcript cleavage activity, 24 Transcription abortive, 16 activators, 43–44 arrest, 23–24 basal, 49 bubble, 16 cell-free, 128 cycle, 13–14 elongation, 13 epigenetic regulation of, 194–195 fidelity of, 21 HAT proteins and, 183–185 HO, 49 inhibitors, 84–85 initiation, 13, 67–68, 84 MCM1, 49–50 mechanism, 2 MF1, 49 mRNA processing and, 26 pause, 23–24
263
SUBJECT INDEX
by Pol III, 93 regulation, 26–27 regulation, in eukaryotes, 181 reinitiation, 26–27 ribosomal gene, 124 species-specific, 127–128 start site, 15 TBP functions in, 77 termination, 13, 26–27 Transcription bubble, 13 maintenance, 18–20 Transcription complex assembly, 137f Transcriptional activity, 55–62 Transcriptional silencing, 205 Translational repositioning, 169 Translocation, 21–23 TRAP220, 62 TRAP230, 62 TRAP240, 62 TRAP. See Thyroid hormone receptor coactivator complex TRF1, 75 Trigger loop, Rpb1, 12 Trithorax, 206 tRNA, 93 TRX N-terminal of, 209 SET domains and, 207 ts13 cell line, 82 ts lethality, 50 TTF-1, 144 TTF-I/Reb1, 148 Tumor suppressor proteins, 144–146 Ty1 elements, 50 Tyrosine, in SET domain, 210
U U937, 143 U6 snRNA, 93 UAF, 138 UBF2, 133 UBF. See Upstream binding factor Ubiquitin activity, 190 in Rad6, 205 Ubiquitination
chromatin-modifying complexes and, 182 histone, 205–206 nucleosome and, 158 UCE, 126 UBF and, 131 Upper jaw, 5 Upstream binding factor (UBF), 131–133, 137f acetylation of, 143 activation, 133 carboxy-terminal acidic domain of, 136 enhancers and, 133 functional domains of, 132f phosphorylation of, 140–143 posttranslational modification of, 140 in rDNA transcription, 136–137 reactivation of, 142 serine phosphorylation and, 142 SL1 and, 133 Upstream stimulating factor (USF), 136 URS2 region, 49 USA, 53–54 USF. See Upstream stimulating factor
V Van der Walls interactions, 216 Vascular smooth muscle stimulation, 141 VDR. See Vitamin D3 receptor Viral infection, rRNA synthesis and, 140 Vitamin D3 receptor interacting proteins (DRIP) complex, 53 Vitamin D3 receptor (VDR), 53, 56 VP16 ARC and, 53 Gal11 and, 55
W Watson-Crick base pairing, 21 Wnt signaling, 55 Worms putative null mutations and, 55 sop-1(RNAi), 55
264
SUBJECT INDEX
Y
X X-ray analysis, of Pol II, 1 X-ray crystallography elongation and, 18 of Pol II, 1 single-subunit DNA polymerase, 29–30 single-subunit RNA polymerase, 29–30 of T. aquaticus, 28 Xenopus enhancers in, 126–127 interspecific crosses with, 127 rDNA genes in, 124
Ybf2/Sas3, 185–186 YGP1, 47
Z Zeste, 206 Zinc cluster, 209 Zinc ions, 4 in Rpb2, 8 Zipper protein, 136
