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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

This book is printed on acid-free paper. Copyright ß 2004, Elsevier Inc. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the Publisher. The appearance of the code at the bottom of the first page of a chapter in this book indicates the Publisher’s consent that copies of the chapter may be made for personal or internal use of specific clients. This consent is given on the condition, however, that the copier pay the stated per copy fee through the Copyright Clearance Center, Inc. (www.copyright.com), for copying beyond that permitted by Sections 107 or 108 of the U.S. Copyright Law. This consent does not extend to other kinds of copying, such as copying for general distribution, for advertising or promotional purposes, for creating new collective works, or for resale. Copy fees for pre-2004 chapters are as shown on the title pages. If no fee code appears on the title page, the copy fee is the same as for current chapters. 0065-3233/2004 $35.00 Permissions may be sought directly from Elsevier’s Science & Technology Right Department in Oxford, UK: phone: (+44) 1865 843830, fax: (+44) 1865 853333, E-mail: [email protected]. You may also complete your request on-line via the Elsevier Science homepage (http://elsevier.com), by selecting ‘‘Customer Support’’ and then ‘‘Obtaining Permissions.’’ For all information on all Academic Press publications visit our Web site at www.academicpress.com ISBN: 0-12-034267-7 PRINTED IN THE UNITED STATES OF AMERICA 04 05 06 07 08 9 8 7 6 5 4 3 2 1

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

CONTENTS

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. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .


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

Copyright 2004, Elsevier Inc. All rights reserved. 0065-3233/04 $35.00

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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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


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

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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).


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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.)


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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


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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


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


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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.

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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


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¨ RKLUND AND GUSTAFSSON BJO

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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THE MEDIATOR COMPLEX

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

46


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.

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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).


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.

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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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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.

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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.


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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

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