mRNA Formation and Function

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Edited by Joel D. Richter Department of Molecular Genetics and Microbiology University of Massachusetts Medical Center Worcester, Massachusetts

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Front cover photograph: Simultaneous detection of fushi tarazu protein by specific antibody staining (brown) and of wingless m R N A by in situ hybridization (blue) in a gastrulating Drosophila embryo. Courtesy of Dr. Armen Manoukian, Ontario Cancer Institute, Toronto, Canada.

This book is printed on acid-free paper. Copyright 9 1997 by ACADEMIC PRESS. 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. Academic Press 15 East 26th Street, 15th floor, New York, New York 10010 http://www.apnet.com Academic Press Limited 24-28 Oval Road, London NW1 7DX, UK http://www.hbuk.co.uk/ap/ Library of Congress Cataloging-in-Publication Data m R N A formation and function / [edited by] Joel D. Richter. p.

cm.

Includes bibliographical references and index. ISBN 0-12-587545-2 (alk. paper) 1. Messenger RNA--Research--Methodology. OP623.5.M47M75 1997 572.8'8--dc21

I. Richter, Joel D.

PRINTED IN THE UNITED STATES OF AMERICA 97 98 99 00 01 02 MM 9 8 7 6 5 4 3

97-33343 CIP

2

1

CONTENTS

Contributors Preface

xiii xvii

CHAPTER

1

The T a t - T A R R N P , a Master Switch That Regulates HIV-1 Gene Expression Carlos Sufl~, Paul R. Bohjanen, Yi Liu, and Mariano A. Garcia-Blanco

I. Introduction II. Methods References

1 4 21

CHAPTER

2

HeLa Nuclear Extract- A Modified Protocol Cameron g. Miller, Sharon F. Jamison, and Mariano A. Garcia-Blanco

I. Growth of the HeLa Cells II. Extract Preparation Reference

CHAPTER

25 26 30

3

Functional Analysis o f Splicing Factors and Regulators Juan Valcdrcel, Concepci6n Martlnez, and Michael R. Green

I. Introduction II. Functional Analysis of General Splicing Factors III. Analysis of Regulatory Mechanisms in P r e - m R N A Splicing References

31 32 43 49

vi

Contents

CHAPTER

4

Identification of P r e - m R N A Splicing Factors and Analysis of R N A - P r o t e i n Interaction James G. Patton, Billy T. Dye, Daron C. Barnard, and James G. McAfee

I. Introduction II. Pre-mRNA Splicing and the Role of the Polypyrimidine Tract III. Analysis of R N A Binding Proteins by UV Cross-Linking IV. Analysis of Protein-RNA Interaction by Fluorescence Spectroscopy V. Analysis of Pre-mRNA Splicing Complexes and Identification of Associated Factors by UV Cross-Linking VI. Identification of R N A Binding Proteins by a cDNA Expression Library Screen References

CHAPTER

55 56 58 62

68 72 75

5

In Vitro Analysis of Mammalian Cell m R N A

3' Processing Gregory M. Gilmartin

I. Introduction II. General Considerations for the Analysis of nflKNA 3' mRNA Processing in Vitro III. Pre-mRNA Cleavage and Polyadenylation IV. 3' Processing Complex Formation V. 3' Processing Complex Stability VI. Probing 3' Processing Factor-Pre-mRNA Interactions by UV Cross-Linking VII. Application of in Vitro Selection to the Analysis of Sequences That Direct 3' Processing VIII. Conclusions References

79 80 82 85 88 90 93 97 97

Contents CHAPTER

vii 6

Rapid Identification and Cloning of Sequence-Specific RNA Binding Proteins Jeffrey Wilusz

I. II. III. IV. V.

Introduction Preparation of Extracts Preparation of RNAs Identification of Protein-RNA Complexes Determination of R N A Sequence Requirements for Protein Binding VI. Cloning cDNAs for R N A Binding Proteins by Northwestern Screening References CHAPTER

99 100 101 102 103 105 108

7

Analysis of Polyadenylation Phenotypes in Saccharomyces cerevisiae J. Scott Butler, Michael W. Briggs, and Aaron Proweller

I. II. III. IV.

Introduction Voly(A) Tail Analysis Polyadenylation Site Mapping by R T - P C R Preparation of Yeast mRNA 3' End Processing Extracts References CHAPTER

111 112 117 121 125

8

Poly(A) Polymerase/Cap-Specific 2'-O-Methyltransferase from Vaccinia Virus: Expression, Purification, Uses, and Protein-Ligand Interaction Assays Paul D. Gershon

I. Background II. Expression and Purification of the Poly(A) Polymerase/ 2'- O-Methyltransferase III. Uses of the Poly(A) Polymerase/2'-O-Methyltransferase IV. Assays Developed to Examine Protein-Ligand Interactions References

127 132 135 139 145

viii

Contents

CHAPTER 9 A Genetic Approach to Mapping C o d i n g R e g i o n Determinants o f m R N A Instability in Yeast Aidan N. Hennigan and Allan Jacobson

I. II. III. IV.

Introduction Experimental Methodology Experimental Results Discussion and Conclusions References

149 150 153 157 160

C H A P T E R 10 Identification o f the Protein That Interacts with the 3' End o f Histone m R N A William F. Marzluff, Michael L. Whitfield, Zbigniew Dominski, and Zeng-Feng Wang

I. II. III. IV. V.

Introduction Detecting Specific Protein-RNA Interactions In Vitro Processing of Histone Pre-mRNA Isolation of Polyribosomes Isolation and Characterization of R N A Binding Proteins Using the Yeast Three-Hybrid System Appendix: Cell Culture References

163 165 174 181 183 188 192

C H A P T E R 11 Analysis o f m R N P C o m p l e x e s Assembled in Vitro Martin Holcik and Stephen A. Liebhaber

I. Complex Formation II. Detection of RNP Complexes III. Analysis of RNP Complexes Identified b y Electrophoretic Mobility Shift Analysis References

196 200 202 208

ix

Contents

CHAPTER

12

Analysis of RNA Binding Proteins Using in Vitro Genetics Ite A. Laird-Offringa

I. II. III. IV.

Introduction Successfully Making Display Phage Binding of Displayed RNA Binding Proteins to RNA Application of in Vitro Genetics to the Study of RNA Binding Proteins V. Protocols References

CHAPTER

212 215 221 224 227 234

13

Interactions of Proteins with Specific Sequences in R N A Lucy G. Andrews and Jack D. Keene

I. Introduction II. Selection of Specific Sequences from Random RNA Libraries by RNA Binding Proteins as a First Step in Identification of in Vivo Targets III. Binding of Protein to Sequences in Cellular RNAs IV. Determination of Minimal Binding Sequences within RNA Ligands V. Selection of RNAs from 3' U T R Libraries VI. Summary References

CHAPTER

237

240 247 250 257 259 259

14

A Rapid Genetic Method for the Study of RNA Binding Proteins Chaitanya Jain and Joel G. Belasco

I. Introduction II. Background III. A Quantitative Model for Translation Repression by Heterologous RNA-BPs IV. Methods V. Strategies and Applications

263 264 265 267 274

Contents

VI. Troubleshooting VII. Application of the Genetic Methods to Rev VIII. Conclusions References

CHAPTER

280 281 282 283

15

Footprinting RNA-Protein Complexes with Hydroxyl Radicals Patricia Ansel-McKinney and Lee Gehrke

I. Introduction II. Preparing an R N A Template for Hydroxyl Radical Footprinting III. Binding and Cleavage Reactions IV. Footprinting Data V. Concluding Remarks References

286 288 292 293 299 300

C H A P T E R 16 Analysis o f R i b o s o m e L o a d i n g o n t o m R N A Species: I m p l i c a t i o n s for T r a n s l a t i o n a l C o n t r o l Hang/un Ruan, Cheryl Y. Brown, and David R. Morris

I. II. III. IV.

Polysome Profiles and Translational Control Extraction and Display of Polysomes R N A Analysis by Northern Blots Analysis by RNase Protection References

CHAPTER

305 307 312 315 320

17

Analysis of Picornavirus Internal Ribosome Entry Site Function in Vivo Graham J. Belsham

I. Introduction II. Cap-Dependent and Cap-Independent Translation III. Characteristics of Picomavirus IRES Elements

323 324 325

Contents

xi

IV. Characteristics of the Vaccinia/T7 R N A Polymerase Expression System go Analysis of Picornavirus IRES Function in Vivo VI. Distinguishing between Cap-Dependent and IRESDirected Translation VII. Complementation of Defective IRES Elements by CoExpression of the wt IRES in Trans References

CHAPTER

327 328 334 335 337

18

RNA Traffic and Localization Reported by Fluorescent Molecular Cytochemistry in Living Cells Marty R. Jacobson and Thoru Pederson I. Introduction II. Fluorescent Labeling of R N A III. Cell Culture, Microinjection, and Digital Imaging Processing IV. Examples V. What Is the Biological Authenticity of Fluorescent RNA? VI. Technical Problems and Their Resolution VII. Integrating Fluorescent R N A Cytochemistry with Other Methods VIII. What Does Fluorescent R N A Cytochemistry Actually Measure? IX. New Frontier Methods X. Closing Remarks References

CHAPTER

341 342 344 350 351 353 354 355 356 356 357

19

Detection of m R N A in Situ: Techniques for Studying Gene Expression in Drosophila melanogaster Tissues Armen S. Manoukian

I. Introduction II. Whole-Mount in Situ Hybridization to m R N A with DIG-Labeled Probes

361 362

xii

Contents III. Uses of the Whole-Mount Procedure IV. Detection of Multiple Gene Products V. Subcellular Localization of wingless m R N A References

365 367 369 369

C H A P T E R 20 Differential Display P r o t o c o l That Preferentially I d e n t i f i e s m R N A s o f M o d e r a t e to L o w A b u n d a n c e in a Microscopic

System

Ognian C. Ikonomov and Michele H. Jacob

I. II. III. IV.

Introduction Experimental Procedures Confirmation of Differential Expression Concluding Remarks References

Index

371 373 379 381 382

385

CONTRIBUTORS

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

Lucy G. Andrews (237), Department of Microbiology, Duke University Medical Center, Durham, North Carolina 27710 Patricia Ansel-McKinney (285), Department of Microbiology and Molecular Genetics, Harvard Medical School, Boston, Massachusetts 02139; HarvardM.I.T. Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139 Daron C. Barnard (55), Department of Molecular Biology, Vanderbilt University, Nashville, Tennessee 37235 Joel G. Belasco (263), Skirball Institute of Biomolecular Medicine, Department of Microbiology, New York University Medical Center, New York, New York 10016 Graham J. Belsham (323), BBSRC Institute for Animal Health, Pirbright, Woking, Surrey, GU24 ONF United Kingdom

Paul R. Bohjanen (1), Department of Molecular Cancer Biology, Division of Infectious Diseases, Department of Medicine, Duke University Medical Center, Durham, North Carolina 27710 Michael W. Briggs (111), Department of Microbiology and Immunology, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642 Cheryl Y. Brown (305), Department of Biochemistry, University of Washington, Seattle, Washington 98195 J. Scott Buffer (111), Department of Microbiology and Immunology, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642 Zbigniew Dominski (163), Program in Molecular Biology and Biotechnology, University of North Carolina, Chapel Hill, North Carolina 27599

Billy T. Dye (55), Department of Molecular Biology, Vanderbilt University, Nashville, Tennessee 37235 Mariano A. Garcia-Blanco (1, 25), Department of Molecular Cancer Biology, Division of Infectious Diseases, Departments of Medicine and Microbiology, Duke University Medical Center, Durham, North Carolina 27710

xiii

xiv

Contributors

Lee Gehrke (285), Department of Microbiology and Molecular Genetics, Harvard

Medical School, Boston, Massachusetts 02115; Harvard-M.I.T. Division of Health Sciences and Technology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02139 Paul D. Gershon (127), Department of Biochemistry and Biophysics/Institute of Biosciences and Technology, Texas A&M University, Houston, Texas 77030 Gregory M. Gilmartin (79), Department of Microbiology and Molecular Genetics and the Markey Center for Molecular Genetics, University of Vermont, Burlington, Vermont 05405 Michael R. Green (31), Howard Hughes Medical Institute and Program in

Molecular Medicine, University of Massachusetts Medical Center, Worcester, Massachusetts 01605 Aidan N. Hennigan (149), Department of Molecular Genetics and Microbiology, University of Massachusetts Medical School, Worcester, Massachusetts 01655 Martin Holcik* (195), Howard Hughes Medical Institute, Departments of Genetics and Medicine, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 Ognian C. lkonomoval" (371), Neurobiology Group, Worcester Foundation for Biomedical Research, Shrewsbury, Massachusetts 01545 Michele H. Jacob:~ (371), Neurobiology Group, Worcester Foundation for Bio-

medical Research, Shrewsbury, Massachusetts 01545 Allan Jacobson (149), Department of Molecular Genetics and Microbiology, University of Massachusetts Medical School, Worcester, Massachusetts 01655 Marty R. Jacobson (341), Cell Biology Group, Worcester Foundation for Biomedical Research, University of Massachusetts Medical School, Shrewsbury, Massachusetts 01545 Chaitanya Jain (263), Skirball Institute of Biomolecular Medicine, Department of Microbiology, New York University Medical Center, New York, New York 10016 Sharon F. Jamison~ (25), Department of Molecular Cancer Biology, Duke University Medical Center, Durham, North Carolina 27710 Jack D. Keene (237), Department of Microbiology, Duke University Medical Center, Durham, North Carolina 27710 * Present address: Molecular Genetics Laboratory, University of Ottawa, Children's Hospital of Eastern Ontario, Ottawa, Ontario KIH 8L1 Canada. t Present address: Department of Psychiatry, Wayne State University, Detroit, Michigan 48201. Present address: Department of Neuroscience, Tufts University School of Medicine, Boston, Massachusetts 02111. Present address: INTRONN LLC, Durham, North Carolina 27702.

Contributors

xv

lte A. Laird-Offringa (211), Departments of Surgery and of Biochemistry and

Molecular Biology, University of Southern California School of Medicine, Los Angeles, California 90033 Stephen A. Liebhaber (195), Howard Hughes Medical Institute, Departments of Genetics and Medicine, University of Pennsylvania School of Medicine, Philadelphia, Pennsylvania 19104 Yi Liu (1), Departments of Molecular Cancer Biology and Microbiology, Duke University Medical Center, Durham, North Carolina 27710 James G. McAfee (55), Department of Molecular Biology, Vanderbilt University, Nashville, Tennessee 37235 Armen S. Manoukian (361), Division of Cell and Molecular Biology, Ontario Cancer Institute, Department of Medical Biophysics, University of Toronto, Toronto, Ontario M5G 2M9, Canada Concepci6n Martinez (31), Gene Expression Programme, European Molecular Biology Laboratory, D-69117 Heidelberg, Germany William F. Marzluff (163), Program in Molecular Biology and Biotechnology, University of North Carolina, Chapel Hill, North Carolina 27599 Cameron R. Miller (25), Department of Molecular Cancer Biology, Duke University Medical Center, Durham, North Carolina 27710 David R. Morris (305), Department of Biochemistry, University of Washington, Seattle, Washington 98195 James G. Patton (55), Department of Molecular Biology, Vanderbilt University, Nashville, Tennessee 37235 Thoru Pederson (341), Cell Biology Group, Worcester Foundation for Biomedical Research, University of Massachusetts Medical School, Shrewsbury, Massachusetts 01545 Aaron Proweller (111), Department of Microbiology and Immunology, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642 Hangjun Ruan (305), Department of Biochemistry, University of Washington, Seattle, Washington 98195 Carlos Sufi~ (1), Department of Molecular Cancer Biology, Duke University Medical Center, Durham, North Carolina 27710 Juan Valcfircel (31), Gene Expression Programme, European Molecular Biology Laboratory, D-69117 Heidelberg, Germany Zeng-Feng Wang (163), Program in Molecular Biology and Biotechnology, University of North Carolina, Chapel Hill, North Carolina 27599 Michael L. Whitfield (163), Program in Molecular Biology and Biotechnology, University of North Carolina, Chapel Hill, North Carolina 27599 Jeffrey Wilusz (99), Department of Microbiology and Molecular Genetics, UMDNJ-New Jersey Medical School, Newark, New Jersey 07103

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PREFACE

In the past decade or so, there has been a virtual explosion in the field of R N A science. While most investigations had been limited to studying either bulk populations ofmRNAs or, in a few cases, specific mRNAs that are highly enriched in certain cells, such as globin m R N A in reticulocytes, almost any laboratory can now examine even very rare messages in any type of cell. Huge quantities of specific mRNAs are synthesized in vitro with bacteriophage R N A polymerases, minute quantities of specific mRNAs in single cells are measured by reverse transcription and the polymerase chain reaction (RT-PCR), specific, localized mRNAs are detected by in situ hybridization, and specific m R N A expression is disrupted by ribozyme or antisense oligonucleotide-directed cleavage or by antisense RNA-mediated translational silencing. And this is only the beginning! A wonderful new array of techniques useful for the identification and cloning of cDNAs for specific R N A binding proteins, which, of course, control m R N A processing, transport, translation, and degradation, is available. This volume deals with new methods used in R N A research. Although a grouping of the chapters would be somewhat arbitrary, there is a common theme as to what each method can accomplish. For example, six chapters deal with the isolation and characterization of specific R N A binding proteins, seven chapters examine R N A metabolism and associated regulatory proteins, four chapters are aimed at m R N A detection and localization, and two chapters take a genetic approach to the study of R N A function. This compilation of methods is meant to go well beyond that presented in the primary literature. These chapters contain the procedural details that may not be fully elaborated in .journal articles because of space limitations, details that can often make the difference between success and failure. The volume will be useful not only to newcomers to the R N A field but also to established investigators whose research has led them to explore different facets of R N A biology. Joel D. Richter

xvii

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The T a t - T A R RNP, a Master Switch That Regulates HIV-1 Gene Expression Carlos Sufi6 Department of Molecular Cancer Biology Duke University Medical Center Durham, North Carolina 27710

Paul R. B o h j a n e n Department of Molecular Cancer Biology Division of Infectious Diseases Department of Medicine Duke University Medical Center Durham, North Carolina 27710

Yi Liu Departments of Molecular Cancer Biology and Microbiology Duke University Medical Center Durham, North Carolina 27710

Mariano A. Garcia-Blanco Department of Molecular Cancer Biology Division of Infectious Diseases Departments of Medicine and Microbiology Duke University Medical Center Durham, North Carolina 27710

I. I N T R O D U C T I O N The study of the regulation of human immunodeficiency virus type 1 (HIV1) gene expression has increased our understanding of the biology of complex retroviruses and provided useful insights into eukaryotic transcriptional control. The HIV-1 viral protein Tat increases HIV-1 transcription more than 100-fold and is absolutely required for viral replication (Dayton et al., 1986; Fisher et al., 1986). Tat is a protein of 86 amino acids that functions by binding to an R N A element designated TAR (for trans-activation response element) (reviewed in Gait and Karn, 1993; Cullen, 1995). TAR is a highly structured R N A element located at the 5' end of all HIV-1 transcripts and is composed of double-stranded stems, a four-nucleotide bulge, and a six-nucleotide loop (Fig. 1: TAP.). Tat binds specifically to the TAR bulge in vitro and on binding alters the structure of the TAP. R N A (Colvin and Garcia-Blanco, 1992; Puglisi et al., 1992). Although the structure of TAP. and the requirements for T a t - T A R binding have been thoroughly investigated, little is known about the mechanism by which Tat increases HIV-1 transcription. Current models suggest a role for Tat in the regulation of transcriptional initiation as well as elongation (reviewed in Cullen, 1993; Jones and Peterlin, 1994; Karn et at., 1996). 1 m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

Figure 1 Model for transcriptional control by HIV-1 Tat. (A) Basal transcription of the HIV-1 promoter. In the absence of the T a t - T A R RNP, basal transcription is carried out by core R N A polymerase (pol) II complexes that lack elongation-competent components. These complexes are elongation incompetent and synthesize short transcripts that are released from the HIV-1 DNA template. (B) Activation of the HIV-1 promoter by the T a t - T A R R3qP complex. Tat may trans-activate the HIV-1 promoter by modifying and/or recruiting transcription components that transform elongation-incompetent R N A pol II complexes to elongationcompetent complexes that are now able to synthesize long transcripts. Part of this model is based on data by Sufi6 et al. (1997). All factors involved in transcription initiation and elongation have not been drawn to simplify the model.

4

Carlos Sufi6 et al.

Experimental data suggest that the primary role of TAR is to anchor Tat to a location near the start site of transcription (Selby and Peterlin, 1990; Southgate et al., 1990). Because mutations in the TAR loop that abolish Tat-mediated transactivation in vivo minimally affect Tat binding in vitro, it was postulated that host proteins recognize the TAR loop (Feng and Holland, 1988; Roy et al., 1990). Considerable genetic and biochemical evidence supports the existence of an essential cellular factor that binds to the TAR loop (Marciniak et al., 1990; Madore and Cullen, 1993; Bohjanen et al., 1996). Although genetic studies suggest that Tat binds to TAR as a preformed complex in vivo (Madore and Cullen, 1993), it remains unclear whether the cellular loop-binding factor is a component of this Tat-containing complex or if it binds to TAR independently. Numerous cellular proteins that bind to Tat or TAR have been identified and certain ones have been characterized in some detail (reviewed in Gaynor, 1993), but the role of most of these proteins remains unknown. Recent in vitro experiments have identified proteins that bind to Tat and are essential for Tat-mediated trans-activation (Sufi& and Garcia-Blanco, 1995a; Zhou and Sharp, 1995). It appears that Tat and cellular factors form a complex on TAR RNA. This complex, referred to as the T a t - T A R ribonucleoprotein (RNP), constitutes the core of a master switch that likely regulates the Tat-dependent increase in HIV-1 transcription by communicating with an R N A polymerase II transcription complex (Fig. 1). Complete elucidation of the mechanism by which the T a t - T A R R N P activates HIV-1 gene transcription will require identification, purification, and characterization of all essential components in a reconstituted system. The establishment of a cell-free system that faithfully recapitulates Tat-dependent activation of HIV-1 transcription was an essential step toward this objective (Marciniak et at., 1990). The in vitro transcription system has been used to characterize the functions of Tat and TAR cofactors (Sheline et al., 1991; Wu et al., 1991; Sufi~ and GarciaBlanco, 1995a; Zhou and Sharp, 1995) and has provided important insights into the mechanism by which Tat activates the HIV-1 promoter (Marciniak and Sharp, 1991; Laspia et al., 1993; Bohan et al., 1992; Kato et al., 1992; Graeble et al., 1993). We will discuss the basis of this system and show how it can be used to study the role of the T a t - T A R R N P in HIV-1 gene regulation.

II. M E T H O D S A. SYNTHESIS AND PURIFICATION OF RECOMBINANT H I V - 1 TAT PROTEIN Large quantities of experiments described in recombinant methods or protocols of purification

pure Tat protein were needed to carry out the in vitro this chapter. Purified Tat protein has been obtained by chemical synthesis. Due to its poor solubility, complex using strong denaturing agents have been developed

1. Tat-TAP, P,.NP

5

(Green and Loewenstein, 1988; Chun et al., 1990; Waszak et al., 1992). Previously, we described an easy procedure for the purification of large amounts of biologically active recombinant HIV-1 Tat protein using the pGEX-2TK vector (Pharmacia, Piscataway, NJ) (Sulk& and Garcia-Blanco, 1995a). This vector allows expression of the gene products as fusions with the glutathione S-transferase (Gst) of Schistosoma japonicum (Smith and Johnson, 1988). This plasmid encodes a specific thrombin cleavage site to split the fusion and a high-aflfinity protein kinase phosphorylation site that allows the labeling of the protein with [T-32p]ATP. Since the Gst-Tat fusion protein was poorly soluble, we have used a modification of the procedure described by Grieco et al. (1992) to allow better solubilization. Briefly, 1 liter of transformed BL-21 cells was grown to an 0D600 o f 0.8, and expression of the fusion proteins was induced by adding 0.1 m M isopropyl-/3-thiogalactopyranoside (IPTG, Sigma, St. Louis, MO) for 3 hr. Cells were pelleted and resuspended in 10 ml of phosphate-buffered saline (PBS) containing 2 m M ethylenediaminetetraacetic acid (EDTA), 50/zg/ml phenylmethylsulfonyl fluoride (PMSF, Sigma), 1 /zg/ml leupeptin, and 1 /zg/ml pepstatin. The bacterial pellet was lysed on ice by mild sonication and centrifuged at 12,000 g (rmax) for 10 rain at 4~ The supernatant fraction was mixed with Triton X-100 to 1% (v/v) and kept on ice. The pellet fraction was resuspended with 8 ml of a solution containing 1.5% (v/v) sodium N-lauroylsarcosinate (Fluka, Ronkonkoma, NY), 25 m M triethanolamine, and 1 m M EDTA (pH 8.0), mixed and placed on ice for 30 rain. The suspension was then centrifuged at 18,900 g (rmax) for 20 rain at 4~ Triton X-100 and dithiothreitol (DTT) were added to the new supernatant fraction to final concentrations of 2% and 1 mM, respectively, and the solution was mixed with the supernatant fraction resulting from the first centrifugation. The new solution was then rocked with 0.5 nil of glutathione-agarose beads (Sigma) for 1 hr at 4~ and the beads were washed extensively with 1% Triton X-100, 1% Tween-20, 1 m M DTT, and 50/zg/ml PMSF in PBS. After the beads were washed with 50 m M Tris HC1 (pH 8.0), Gst-fusion proteins were released with 10 mM glutathione (Sigma). Cleavage with thrombin (Sigma) was done with the Gst-fusion on the beads or in solution. This reaction was carried out in cleavage buffer (150 mMNaC1, 50 mMTris pH 8.0, 2.5 mMCaC12, 1 m M D T T ) containing 10 NIH U per milliliter of thrombin for 20 rain at room temperature. The reaction was stopped by adding 100/zg/ml PMSF. As shown in Figure 2, Gst-Tat and Tat proteins were purified to near homogeneity. The yields of HIV-1 Tat were up to 300/zg per liter of bacterial culture.

B. In Vitro TRANSCRIPTION SYSTEM AND H I V - 1 TAT Trans-AcTIVATION The development of transcription-competent, cell-free systems has facilitated the study of the complex interactions that regulate gene expression and has been

6

Carlos Sufi6 et al.

Figure 2 SDS-polyacrylamidegel electrophoresis(SDS-PAGE) gel showingthe purification of recombinant Gst-Tat and Tat proteins. Left: lane 1, supernatant following the first centrifugation after lysing the cells; lane 2, pellet following the first centrifugation after lysing the cells; lane 3, eluate from glutathione-agarosebeads showingpure Gst-Tat protein. Right: lane 1, Gst eluted with free glutathione after thrombin digestion; lane 2, glutathione-agaroseeluate following thrombin cleavageshowing pure Tat protein. Gels were stained with Coomassie blue. Lane M shows prestained molecular weight standards (Amersham) expressedas size in kilodaltons. The positions of the purified proteins (Gst-Tat, Gst, and Tat) are indicated by arrows.

critical in the identification of general transcription factors. In particular, we and others have used a previously described in vitro transcription system (Marciniak et al., 1990) to faithfully recapitulate the Tat- and T A R - d e p e n d e n t transcriptional activation of HIV-1 genes. The in vitro transcription activity of the HIV-1 promoter is exceptionally high in nuclear extract alone, masking the activation effect of Tat. Marciniak et al. (1990) were the first to overcome this problem by introducing a 30-min incubation time prior to the addition of labeled uridine triphosphate (UTP) in the transcription reaction. Under these conditions, basal transcription was diminished and strong activation of HIV-1 transcription was detected on addition of Tat (Marciniak et al., 1990). Kato et al. (1992) developed a cell-free trans-activation system with no preincubation. They found that the addition of sodium citrate at 6 - 8 m M in the transcription reaction decreased the basal level of the transcription, which enabled them to observe trans-activation by Tat. W e have reproduced this effect and found that 4 m M of sodium citrate is the best concentration in our

1. Tat-TAIL RNP

7

experimental conditions. A modification of the protocol by Marciniak et al. (1990) was used by Karn and co-workers (Graeble et al., 1993). They added 10 /zM ZnSO4 to stimulate specifically HIV-1 transcription and at the same time decreased 10-fold the concentration of nucleotide triphosphates in the reaction. Addition of Tat to the reaction greatly stimulated HIV-1 transcription under these conditions. Finally, we and others (Graeble et al., 1993) have also found that increasing the concentration of KC1 in the transcription mix lowered basal transcription and thus enhanced Tat-dependent transcriptional activation of the HIV-1 promoter. In our laboratory, in vitro transcription reactions were carried out in the presence of transcription-competent HeLa cell nuclear extract (Dignam et al., 1983) and linearized HIV-1 and adenovirus major late promoter (AdML) DNA templates containing specific promoter sequences (Sufi6 and Garcia-Blanco, 1995a). The in vitro transcription reactions were carried out at 30~ for 30 min in 25/zl of a mix containing 10/zl (approximately 15 mg protein/M) of nuclear extract, 14 N-2-hydroxy-ethylpiperazine-N'-2-ethanesulfonic acid-potassium hydroxide (HEPES-KOH, pH 7.9), 14% (v/v) glycerol, 68 mMKC1, 15 m M NaC1, 7 m M MgC12, 4 m M sodium citrate (pH 6.7), 250 ng poly(I) 9poly(C) (Pharmacia), 300 ng poly(dI) 9poly(dC) (Pharmacia), 1 ~ DTT, 10 mM creatine phosphate, 0.1 m M EDTA, 4 units of RNasin (Promega, Madison, WI), 625 /zM each ATP, CTP, and GTP, and 40 /zM of UTP, as well as 10 /zCi of [a-32p]UTP (3000 Ci/mmol, ICN, Costa Mesa, CA). The HIV-1 and AdML templates were normally used at 100 and 250 ng per reaction, respectively, but dose-response curves are recommended. After the incubation time, 100/zl of a solution containing 200 ~ NaC1, 300 m214 NaOAc, 10 ~ EDTA, 0.75% sodium dodecyl sulfate (SDS), 25/zg/ml yeast tRNA, 100 m M Tris-HC1 pH 7.5, and 50/zl of a solution containing 7 M urea, 100 m214 LiC1, 50 mA4 EDTA, 0.5% SDS, and 100 m M Tris-HC1 (pH 7.5), were added. Transcription products were extracted once with phenol-chloroform-isoamyl alcohol (25:24:1) and once with chloroform, and then precipitated with 50/zl of 7.5 M ammonium acetate and cold ethanol. Pellets were resuspended in gel-loading buffer and resolved by electrophoresis on 6% (19: 1) polyacrylamide-8 M urea gels that were dried and exposed. Figure 3 shows the results of typical transcription reactions. In this experiment, the in vitro transcription system was used to test the biological activity of recombinant HIV-1 Tat. Addition of recombinant Tat to the transcription reactions resulted in a specific increase in the level of the HIV-1 m R N A (Fig. 3). Tat did not significatively affect the level of transcription directed by the AdML promoter (Fig. 3). Addition of a single point mutant of an inactive Tat (K41A, Lys to Ala at position 41) did not activate transcription from the HIV-1 promoter (Fig. 3). Figure 4 shows the results of titrating sodium citrate (Fig. 4A) and KC1 (Fig. 4B), allowing optimization of Tat trans-activation. As mentioned above, optimal transactivation was achieved when sodium citrate and KC1 were present in the reaction at 4 and 68 mM, respectively. The level of Tat-mediated trans-activation (TA)

8

Carlos Sufi~ et al.

Figure 3 In vitro trans-activation. The transcriptional activity of HIV-1 Tat and K41A Tat mutant were assayed in an in vitro transcription system. Transcription reactions were carried out in the absence (lanes 1 and 5) or presence of 100, 250, and 500 ng Tat (lanes 2, 3, and 4) and K41A Tat mutant (lanes 6, 7, and 8) proteins as described in the text. The specific transcripts from the HIV-1 (HIV) and the control AdML templates are indicated. The molecular size markers (M, base pairs) were the pBR322 D N A - M s p I digest (Biolabs) labeled with [tx-32p]dCTP.

1. Tat-TAR RNP

9

Figure 4 Effect of sodium citrate and KC1 on the in vitro Tat trans-activation system. Transcription reactions were carried out in the absence (-) or presence (+) of HIV-1 Tat, with the addition of sodium citrate (A) or KC1 (B) at the final concentrations indicated on top of the figures. Runoff transcripts were resolved on 6% polyacrylamide-urea gels. The specific transcripts from the HIV-1 (HIV) and control AdML templates and the molecular size markers (M, base pair) are indicated at the right and left of each figure, respectively.

is given by the following formula (Marciniak et al., 1990; Sufi~ and GarciaBlanco, 1995a): HIV(+)/AdML(+) TA = HIV(-) AdML(-) w h e r e H I V ( - ) and H I V ( + ) are the level o f transcripts from the H I V - 1 p r o m o t e r in the absence or presence o f Tat, respectively, and A d M L ( - ) and A d M L ( + ) are the levels o f the transcripts from the A d M L p r o m o t e r in the absence or presence of Tat, respectively. T h e levels o f the transcripts were d e t e r m i n e d by quantification

10

Carlos Sufi~ et al.

using a Phosphorlmager (445SI Molecular Dynamics, Sunnyvale, CA). Images were stored using IPLab and ImageQuant computer programs. Tat trans-activation levels in HeLa cell nuclear extracts were usually around 10-fold (Marciniak et al., 1990; Kato et al., 1992; Sufi6 and Garcia-Blanco, 1995a; Zhou and Sharp, 1995). This in vitro transcription system recapitulates the requirement for a functional T A R element since transcription directed by an HIV-1 promoter containing a mutated T A R element is unchanged on Tat addition (Marciniak et al., 1990). To further optimize the effects of recombinant Tat on HIV-1 transcription, we carried out a time course. In vitro transcription reactions were done in the presence and absence of Tat and incubated at 30~ for different times. Reactions were stopped and processed as described above, and newly synthesized transcripts were resolved on polyacrylamide gels. Basal transcription and trans-activation by Tat could be detected after only 5 min of incubation (Fig. 5). Both basal and Tatactivated transcription increased over time, but the level of the Tat-responsive HIV-1 transcripts showed the biggest accumulation (Fig. 5). Tat-dependent transcriptional activation of the HIV-1 promoter did not significantly change after 30 min of incubation (Fig. 5 and data not shown), perhaps due to limiting amounts of cellular factors.

C . T A T AS AN AFFINITY REAGENT TO PURIFY N O V E L

HIV-1 COFACTORS The observation that HeLa cell nuclear extracts are fully competent to mediate Tat- and TAR-dependent transcriptional activation of the HIV-1 promoter indicated that this system could be separated into complementing fractions capable of recapitulating Tat trans-activation. We have previously shown that the Gst-Tat fusion protein activated the HIV-1 promoter to the same extent as recombinant Tat (Sufi6 and Garcia-Blanco, 1995a,b). This result allowed the use of Gst-Tat proteins as affinity reagents to separate nuclear extracts by chromatography in order to purify functionally relevant Tat-binding proteins. Gst-Tat proteins were covalently attached to glutathione-agarose matrices with dimethylpimelimidate (DMP) (Pierce, Rockford, IL) (Sufi8 et al., 1997). To couple the Gst-fusion protein to the glutathione matrix, the beads were washed sequentially with 0.1 M borate buffer (pH 8.0) and 0.2 M triethanolamine (pH 8.2). Coupling was performed with 40 m M DMP in 0.2 M triethanolamine (pH 8.2) (Koff et al., 1992) at room temperature for 1 hr. Beads were then washed with 40 m M ethanolamine (pH 8.2) and with 0.1 M borate buffer (pH 8.0). Uncoupled protein was eluted by washing the beads with a solution containing 10 m M glutathione. Column chromatography was then used to generate fractions that were assayed for their ability to support basal and Tat-activated transcription of the HIV-1 as well as the AdML promoters. This approach was successfully used to identify an activity in

1. T a t - T A R R N P

11

Figure 5 Time-course analysis of Tat trans-activation. Transcriptions reactions were carried out in the absence ( - ) or presence (§ of 250 ng recombinant Tat. After the indicated times, reactions were stopped and transcription products resolved by electrophoresis on polyacrylamide-urea gels. Exposure times of 24 and 72 hr are shown in the left and right panels, respectively. The specific transcripts from the HIV-1 (HIV) and control AdML templates and the molecular size markers (M, base pairs) are indicated at the right and left of the figure, respectively.

12

Carlos Sul~ et al.

H e L a nuclear extract required for Tat trans-activation o f the H I V - 1 p r o m o t e r but was n o t necessary for basal H I V - 1 or A d M L transcription in vitro (Sufi& and GarciaBlanco, 1995a). H e L a nuclear extract passed t h r o u g h a G s t - T a t affinity c o l u m n s h o w e d a reduction in Tat trans-activation o f the H I V - 1 p r o m o t e r (Fig. 6). Basal transcription directed from the H I V - 1 or A d M L p r o m o t e r s r e m a i n e d u n c h a n g e d (Fig. 6). N u c l e a r extract passed t h r o u g h a Gst (Fig. 6) or G s t - K 4 1 A Tat m u t a n t c o l u m n (data not shown) supported trans-activation by Tat. Importantly, the addition o f unfractionated nuclear extract to the extract passed t h r o u g h the G s t - T a t c o l u m n was sufficient to recover Tat trans-activation (Fig. 6) (Sufi~ and Garcia-

Figure 6 Specific depletion of the trans-activation by Gst-Tat. Gst and Gst-Tat affinity columns were prepared as described in this chapter, and HeLa nuclear extract was loaded on the columns. Unfractionated nuclear extract (lanes 1 and 2), the flowthrough from the Gst (lanes 3 and 4), and Gst-Tat (lanes 5 to 7) columns were used in transcription reactions in the absence (-) and presence (+) of 250 ng HIV-1 Tat. A mixture of the flowthrough from the Gst-Tat column plus 2 ~1 of unfractionated nuclear extract (lane 7) is also shown. Runoff transcripts from the HIV-1 (HIV) and control AdML templates and the molecular size markers (M, base pairs) are indicated at the right and left of the figure, respectively.

1. Tat-TAR RNP

13

Blanco, 1995a). Further fractionation of HeLa nuclear extract sequentially through Gst-K41A mutant and wild-type Tat affinity matrices have led to the purification of the human Tat coactivator CA150 (Sufi6 et al., 1997).

D. TAR RNA

AS AN AFFINITY REAGENT FOR

CHARACTERIZING THE T A T - T A R

RNP

T A R R N A has been shown to function as a decoy to selectively inhibit Tat-activated HIV-1 transcription in vitro (Marciniak et al., 1990; Bohjanen et al., 1996). T A R bulge and loop structures were both required for this inhibitory activity (Bohjanen et al., 1996). Although Tat bound to the T A R bulge in vitro (Weeks et al., 1990; Roy et al., 1990; Calnan et al., 1991), inhibition of Tatactivated HIV-1 transcription by TAlL decoys was not reversed by the addition of excess Tat (Bohjanen et at., 1996), strongly suggesting that T A R decoys do not function simply by binding to Tat. In contrast, the inhibition of HIV-1 transcription by T A R decoys was reversed by the addition of fractionated nuclear proteins (Sheline et al., 1991), suggesting that T A R R N A acts by interacting with Tat as well as with at least one cellular factor. Here, biotinylated T A R R N A was used as an affinity reagent for purifying and characterizing the cellular components of the T a t - T A R RNP. Biotinylated T A R R N A or T A R R N A containing a mutation in the loop (31/34 RNA; see Fig. 7) was prepared using the method of Milligan and Uhlenbeck (1989) with minor modifications. DNA oligonucleotide templates were synthesized by a commercial vendor (Genosys Biotechnologies, Inc., The Woodlands, TX). T7 R N A polymerase was prepared as described by Grodberg and Dunn (1988).

TAR

31/34 TAR

uo

C

A

C

GA o

cU o U

G

C U

G--C A--U G--C G--C A (U)21

CU U A

A G- G ~ V C U

G--C A--U G--C G--C A (15)21

Figure 7 The sequences of TAR and 31/34 TAR RNA used as affinity reagents.

14

Carlos Sufi4 et al.

Transcription reactions containing 40 mM Tris-HC1 (pH 8.1), 16 mM MgC12, 1 mM spermidine, 5 mM DTT, 1.6 mM each ofATP, GTP, and CTP, 0.16 m M UTP, 0.16 mM biotinylated UTP (biotin-21-UTP, Clontech, Palo Alto, CA), 0.2 nM r UTP (3000 Ci/mmol, ICN), 20 /zg/ml DNA template, 10 /zg/ml T7 R N A polymerase, 0.01% (v/v) Triton X-100, and 0.04% (w/v) polyethylene glycol (average molecular weight 8000) were incubated for 2 hr at 37~ The reaction mixtures were then separated by electrophoresis on 12% urea polyacrylamide gels. Gel slices containing the R N A of interest were identified by ultraviolet (UV) shadowing and were cut out of the gel. This R N A was eluted from gel slices overnight in a buffer containing 0.1% SDS, 0.5 M ammonium acetate, and 10 mM magnesium acetate. The R N A was recovered by ethanol precipitation and quantified by measuring radioactivity with a scintillation counter (LKB, Gaithersburg, MD). The sequence of the biotinylated TAlK and 31/34 R N A transcripts is shown in Figure 7. The poly(U) tail was added to increase the efficiency of biotinylation and to minimize biotinylation of critical U residues within the TAR structure. As shown in Figure 8, biotinylated TAR R N A inhibited Tat-activated transcription from the HIV-1 promoter without inhibiting AdML transcription (lanes 3-5). In contrast, biotinylated 31/34 R N A had no effect (lanes 6-8). Because biotinylation of TAR R N A did not suppress its ability to specifically function as a decoy, biotinylated TAR R N A was used as an affinity reagent to deplete nuclear extracts of trans-activating activity. Two depletion approaches were used, with similar results. In the first approach, biotinylated TAR R N A or 31/34 R N A (10 pmol) was added to standard 25/zl in vitro transcription reactions containing Tat and the HIV-1 promoter, and the reaction mixtures were incubated at 4~ for 10 min. Streptavidin-coated magnetic beads (Dynal, Lake Success, NY) were washed three times with a buffer containing 20 mMHEPES (pH 7.9), 20% glycerol, 0.2 mMEDTA, 100 mMKC1, 0.5 mM DTT, and 1 mg/ml bovine serum albumin (fraction V, Boehringer Mannheim, Indianapolis, IN). Washed beads (150/zg) were added, and the reaction mixtures were rotated for 45 min at 4~ The beads were removed using a magnetic separator, an additional 10 pmol of biotinylated TAlK or 31/34 R N A was added to the supernatants, and the mixtures were incubated at 4~ for 10 min. Again, 150/zg of washed streptavidin-coated beads was added, and the reaction mixtures were rotated for 45 min at 4~ The beads were removed, and the supematants were incubated at 30~ for 30 min to allow transcription. Newly transcribed R N A was isolated and separated by electrophoresis on 6% polyacrylamide gels. The gels were dried and exposed to film. In the second depletion approach, streptavidin magnetic beads that were not treated or were precoated with an excess of biotinylated T A R or 31/34 lKNA were washed in a buffer containing 20 mM HEPES (pH 7.9), 20% glycerol, 0.2 mM EDTA, 100 mM KC1, 0.5 mM DTT, and 1 mg/ml bovine serum

1. T a t - T A R R N P

15

Figure 8 Specific inhibition of Tat-activated HIV-1 transcription by biotinylated TAR. RNA. In vitro transcription reactions containing HIV-1 and AdML templates were carried out using HeLa nuclear extracts in the presence (+) or absence ( - ) of 50 ng Tat protein. Titrated amounts of biotinylated T A R or 31/34 R N A were added to the indicated reactions. The reactions were incubated for 30 min at 30~ and radiolabeled transcripts were isolated and separated by electrophoresis on a denaturing polyacrylamide gel. The positions of migration of the HIV-1 and AdML runofftranscripts are indicated with arrows. The positions of migration of single-stranded D N A markers are shown to the left of the figure as the number of deoxynucleotides.

16

Carlos Sufi~ et al.

albumin. The washed beads were added to standard 25 /zl in vitro transcription reactions containing Tat and the HIV-1 promoter, and the reaction mixtures were rotated at 4~ for 45 rain. The beads were removed, and an additional 150/zg of RNA-coated beads was added to the supernatants. The reaction mixtures were rotated again for 45 min at 4~ The beads were removed, and the supernatants were incubated at 30~ for 30 rain to allow transcription. Newly transcribed R N A was isolated and separated by electrophoresis on 6% polyacrylamide gels. The gels were dried and exposed to film. The result of an experiment utilizing both T A R affinity depletion approaches is shown in Figure 9. Tat-dependent activation of the HIV-1 promoter was specifically depleted by T A R R N A whether the R N A was added to the reaction in solution (lane 2) or was added in the form of precoated beads (lane 4). Less depletion was seen with the use of 31/34 R N A (lanes 3 and 5) or beads alone (lane 6). Current experiments involve characterization of the proteins that bound to the T A R RNA-coated beads.

E. A SOLID-PHASE BASED SYSTEM TO STUDY TAT TI'an$-AcTIVATION The use of biotinylated templates in cell-free transcription reactions has allowed the purification of active transcription complexes (Arias and Dynan, 1989). Karn and colleagues (1996; Keen et al., 1996) have used biotinylated templates to investigate the role of HIV-1 Tat in transcription elongation. We have used a modification of this system to analyze the effect of Tat at different stages of transcriptional initiation and elongation. The HIV-1 template was biotinylated by enzymatic methods. A plasmid containing the HIV-1 long terminal repeat (LTR) (Sufi8 and Garcia-Blanco, 1995a) was used as the template for the polymerase chain reaction (PCR). The P C R product extended from - 4 5 5 to +510 relative to the transcription start site. An EcoRI restriction site was inserted at the 5' end of the P C R product by using a primer that contained that site. The P C R product was digested with EcoRI, and the recessed terminus was filled in with biotin-14-deoxyadenosine triphosphate (dATP) (Gibco, Grand Island, NY) using the Klenow fragment ofDNA polymerase I (New England Biolabs, Beverly, MA). The incorporation of biotin-dATP into the P C R fragment was determined by an electrophoretic mobility shift assay as described by Diamandis and Christopoulos (1991). The biotinylated templates were then attached to 96-well streptavidin-coated microplates (Spectronics, Rochester, NY) and tested for their ability to support basal and Tat-activated transcription. Biotinylated templates (100 to 500 ng) were added to each well of the streptavidin-coated microplates in 25/,1 of binding buffer (PBS containing 0.1% bovine serum albumin and 0.1% Tween 20). The binding was carried out at room

1. T a t - T A R R N P

17

Figure 9 Depletion of Tat-activated HIV-1 transcription using a T A R affinity reagent. In vitro transcription reactions containing HIV-1 and AdML templates, HeLa nuclear extract, and 50 ng Tat protein were submitted to two rounds of depletion with biotinylated affinity reagents, as described in the text. Reactions were submitted to mock depletion (lane 1), depletion with soluble biotinylated T A R R N A (Soluble, lane 2) or 31/34 R N A (Soluble, lane 3), depletion with magnetic beads precoated with T A R R N A (Solid Phase, lane 4) or 31/34 R N A (Solid Phase, lane 5), or depletion with uncoated beads alone (lane 6). The reactions were incubated for 30 min at 30~ and radiolabeled transcripts were isolated and separated by electrophoresis on a denaturing polyacrylamide gel. The positions of migration of the HIV-1 and AdML runoff transcripts are indicated with arrows. The positions of migration of single-stranded D N A markers are shown to the left of the figure as the number of deoxynucleotides.

18

Carlos Sufi~ et al.

temperature for 1 hr, and the wells were then washed extensively with 20 m M H E P E S - K O H (pH 7.9), 20% (v/v) glycerol, 0.1 M KC1, 0.2 mM EDTA, and 0.5 mM DTT. In vitro transcription reactions were carried out in the wells as described above (see Section II,B). As shown in Figure 10, the results recapitulated the results from reactions using free templates. Addition of recombinant Tat induced a specific increase in the level of HIV-1 mRNA. As expected, the K41A Tat mutant did not activate transcription from the HIV-1 promoter.

1. Preinitiation Complex Formation Immobilized HIV-1 templates were incubated with 10 ~1 (10-20 mg/ml) of nuclear extract, 14 mA4 H E P E S - K O H (pH 7.9), 14% (v/v) glycerol, 68 m M KC1, 0.1 ~ EDTA, 7 ~ MgC12, and 1 nad~ DTT in a reaction volume of 25 /.L1at 30~ for 30 min. The wells of the microplate were then washed three times with 50 /~1 of a washing solution containing 20 mM H E P E S - K O H (pH 7.9), 20% (v/v) glycerol, 0.1 M KC1, 0.2 m21//EDTA, 0.5 m_M DTT, 100/zg/rnl PMSF, and 0.05% Nonidet P-40 (Sigma). At this stage, only complexes formed on the immobilized template would be expected to remain in the wells. To initiate tLNA synthesis, 25/zl of a solution containing 625/zM each of ATP, CTP, and GTP, 4 0 / z M of UTP, 10/zCi of [o~-32p]UTP (3000 Ci/mmol, ICN), 14 m M H E P E S - K O H (pH 7.9), 14% (v/v) glycerol, 68 mM KC1, 0.1 mM EDTA, 7 m21//MgC12, 15 mN/NaC1, 1 mM DTT, 250 ng poly(I) 9poly(C), 300 ng of poly(dI) 9poly(dC), 10 mdV/creatine phosphate, 4 m2~ sodium citrate (pH 6.7), and 4 units of tLNasin were added. The reactions were incubated at 30~ for 30 min, and transcription products were extracted and resolved by electrophoresis as described above. This procedure resulted in the synthesis of full-length HIV-1 tLNA (data not shown), which indicated that all factors required for basal transcription were present in the preinitiation complex that was retained on the immobilized templates prior to the initiation of transcription.

2. Elongation Complex Formation It has been shown that the Escherichia coli lactose repressor-operator complex can block the ongoing elongation of R N A polymerase II (Deuschle et al., 1990). Two consecutive lactose repressor (LacP,.) binding sites (LacO) were inserted into the HIV-1 template at the H i n d I I I site (nt +79). From 100 to 500 ng of the biotinylated templates was immobilized to the streptavidin-coated plates as described above. The templates were incubated with 1 big LacR at 30~ for 20 min in a 25-~1 solution containing 20 mM H E P E S - K O H (pH 7.9), 20% (v/v) glycerol, 0.1 M KC1, 0.2 mM EDTA, and 0.5 mM DTT. The solution in the wells was removed, and a solution containing 10/,1 (10-20 mg/ml) nuclear extract, 4 m M sodium citrate (pH 6.7), 4 units of RNasin, 14 mM H E P E S - K O H (pH 7.9), 14%

Figure 10 In vitro transcription and trans-activation on a solid-phase-based system. The transcriptional activity of HIV-1 Tat and K41A Tat mutant proteins was assayed in vitro with both free templates (lanes 1 to 4) and templates immobilized on a solid-phase system (lanes 5 to 8). Transcription reactions were carried out using HeLa nuclear extract in the absence (lanes 1, 3, 5, and 7) or presence of 50 ng Tat (lanes 2 and 6) and K41A Tat mutant (lanes 4 and 8) proteins. The specific transcripts from the HIV-1 and AdML promoters and the molecular size markers (in base pairs) are indicated at the right and left of the figure, respectively. Lanes 1 to 4 (free templates) and lanes 5 to 8 (immobilized templates) are from different exposures.

20

Carlos Sufi~ et al.

Figure 11 Pausingof elongation-competent RNA pol II complexes at the LacR binding sites (LacO). Transcription reactions were carried out as described in the text, and radiolabeled transcripts were isolated and separated by electrophoresis on polyacrylamide-urea gels. Lane 1: pausing of RNA pol complexes by the LacR at the LacO. Short transcripts attached to the pausing polymerase complexes are indicated by an asterisk. Lane 2: elongation of the short transcripts by the paused polymerase complexes on IPTG addition. Full-length transcripts are indicated with an arrow.

(v/v) glycerol, 68 m M KC1, 0.1 m M E D T A , 7 m M MgC12, and 1 m M D T T was added. Preinitiation complexes w e r e then f o r m e d on the template by incubating at 30~ for 10 min. R N A synthesis was initiated by adding 625 /xM each

1. Tat-TAR RNP

21

ATP, CTP, and GTP, 4 0 / z M UTP, 10/xCi [a-32p]UTP (3000 Ci/mmol, ICN), 7 m M MgC12, 250 ng poly(I) 9poly(C), 300 ng poly(dI) 9poly(dC), and 10 m M creatine phosphate. After incubation at 30~ for 1 rain, the reaction buffer was removed and the plates were washed three times with 50/xl of washing solution. At this stage, the R N A polymerase complex on the immobilized templates was expected to be stalled by the L a c k at the LacO sites at position + 79. To release the R N A transcripts from these complexes, we added 100/zl of a solution containing 200 m M NaC1, 300 m M NaOAc, 10 m M EDTA, 0.75% SDS, 25 /xg/ml yeast tRNA, 100 m M Tris-HC1 (pH 7.5), and 50/zl of a solution containing 7 M urea, 100 m M LiC1, 50 m M EDTA, 0.5% SDS, 100 m M Tris-HC1 (pH 7.5) to the plates. The released transcripts were extracted, precipitated, and resolved by electrophoresis, as described above. A set of transcripts about 80 nt in length was observed (Fig. 11, lane 1). This indicated that the elongating R N A polymerase was blocked by the L a c k at the LacO sites. The R N A polymerase did not fall off the template when it met the block since it remained in the plates after extensive washing. To test whether the paused polymerase was able to resume elongation on removal of the block, 5 0 0 / x M of lPTG was added to the plates after the wash, together with 625 /zM ATP, CTP, GTP, and UTP, 14 m M H E P E S - K O H (pH 7.9), 14% glycerol, 68 mMKC1, 0.1 m M E D T A , 7 m M MgC12, 15 m M NaC1, 1 m M D T T , 250 ng poly(I) 9poly(C), and 300 ng poly(dI) 9poly(dC), and 10 m M creatine phosphate. The reaction was carried out at 30~ for 10 rain, and the products were extracted and resolved as described. Since no radiolabeled ribonucleotides were added in the last part of the reaction, the transcript signals obtained by autoradiography should come from those of the stalled polymerase. As a result, full-length HIV-1 transcripts (576 nt) were observed in the presence of IPTG (Fig. 11, lane 2), which indicated that the stalled polymerase II complex remained fully active and was able to resume transcription elongation. Current experiments are focusing on the effect of Tat at each of the transcriptional stages using this system.

ACKNOWLEDGMENTS We thank Zvi Pasman for preparation ofT7 RNA polymerase. Work described in this chapter has been supported by the NIH and the Veterans Administration (to M.A.G.-B.) and the Ministry of Education and Science of Spain (to C.S.). P.R.B. was supported by a Howard Hughes Medical Institute Postdoctoral Fellowship for Physicians. M.A.G.-B. is an Established Investigator of the American Heart Association. The authors acknowledge the Keck Foundation for support to the Levine Science Research Center.

REFERENCES Arias, J. A., and Dynan, W. S. (1989). Promoter-dependent transcription by P,NA polymerase II using immobilized enzyme complexes.J. Biol. Chem. 264, 3223-3229.

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Bohan, C. A., Kashanchi, F., Ensoli, B., Buonaguro, L., Boris-Lawrie, K. A., and Brady, J. N. (1992). Analysis of Tat transactivation of human immunodeficiency virus transcription in vitro. Gene Expression 2, 391-407. Bohjanen, P. R., Colvin, R. A., Puttaraju, M., Been, M. D., and Garcia-Blanco, M. A. (1996). A small circular TAP, RNA decoy specifically inhibits Tat-activated HIV-1 transcription. Nucleic Acids Res. 24, 3733-3738. Calnan, B. J., Tidoe, B., Biancalana, S., Hudson, D., and Frankel, A. D. (1991). Arginine-mediated R N A recognition: The arginine fork. Science 252, 1167-1171. Chun, R., Glabe, C. G., and Fan, H. (1990). Chemical synthesis of biologically active tat transactivating protein of human immunodeficiency virus type 1. J. Virol. 64, 3074-3077. Colvin, R. A., and Garcia-Blanco, M. A. (1992). Unusual structure of the human immunodeficiency virus type 1 trans-activation response element. J. Virol. 66, 930-935. Cullen, B. R. (1993). Does HIV-1 Tat induce a change in viral initiation rights? Cell (Cambridge, Mass.) 73, 417-420. Cullen, B. R. (1995). Regulation of HIV gene expression. A I D S 9, $19-$32. Dayton, A. I., Sodroski, J. G., Rosen, C. A., Goh, W. C., and Haseltine, W. A. (1986). The transactivator gene of the human T cell lymphotropic virus type III is required for replication. Cell (Cambridge, Mass.) 44, 941-947. Deuschle, U., Hipskind, R. A., and Bujard, H. (1990). RNA polymerase II transcription blocked by Escherichia coli lac repressor. Science 248, 480-483. Diamandis, E.'P., and Christopoulos, T. K. (1991). The biotin-(strept)avidin system: Principles and applications in biotechnology. Clin. Chem. (Winston-Salem, N.C.) 37, 625-636. Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983). Accurate transcription initiation by R N A polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11, 1475-1489. Feng, S., and Holland, E. C. (1988). HIV-1 tat trans-activation requires the loop sequence within tar. Nature (London) 334, 165-167. Fisher, A. G., Feinberg, M. B., Josephs, S. F., Harper, M. E., Marselle, L. M., Reyes, G., Gonda, M. A., Aldovini, A., Debouk, C., Gallo, R. C., and Wong-Staal, F. (1986). The trans-activator gene of HTLV-III is essential for virus replication. Nature (London) 320, 367-371. Gait, M. J., and Karn, J. (1993). RNA recognition by the human immunodeficiency virus Tat and Rev proteins. Trends Biochem. Sci. 18, 255-259. Gaynor, R. B. (1993). Regulation of HIV-1 gene expression by the transactivator protein Tat. Curt. Top. Microbiol. Immunol. 193, 51-77. Graeble, M. A., Churcher, M. J., Lowe, A. D., Gait, M. J., and Karn, J. (1993). Human immunodeficiency virus type 1 transactivator protein, tat, stimulates transcriptional read-through of distal terminator sequences in vitro. Pro& Natl. Acad. Sci. U.S.A. 90, 6184-6188. Green, M., and Loewenstein, P. M. (1988). Autonomous functional domains of chemically synthesized human immunodeficiency virus Tat trans-activator protein. Cell (Cambridge, Mass.) 55, 1179-1188. Grieco, F., Hay, J. M., and Hull, R. (1992). An improved procedure for the purification of protein fused with glutathione-S-transferase. BioTechniques 13, 856-857. Grodberg, J., and Dunn, J. j. (1988). omp T encodes the Escherichia coli outer membrane protease that cleaves T7 RNA polymerase during purification. J. Bacteriol. 170, 1245-1253. Jones, K. A., and Peterlin, M. B. (1994). Control of RNA initiation and elongation at the HIV-1 promoter. Annu. Rev. Biochem. 63, 717-743. Karn, J., Churcher, M. J., Rittner, K., Keen, N.J., and Gaite, M. J. (1996). Control of transcriptional elongation by the human immunodeficiency virus Tat protein. In "Eukaryotic Gene Transcription" (S. Goodbourn, ed.), pp. 254-286. IRL Press, Oxford. Kato, H., Sumimoto, H., Pognonec, P., Chen, C.-H., Rosen, C. A., and Roeder, R. G. (1992). HIV-1 tat acts as a processivity factor in vitro in conjunction with cellular elongation factors. Genes Dev. 6, 655-666.

1. Tat-TAR RNP

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Keen, N. J., Gait, M. J., and Kam, J. (1996). Human immunodeficiency virus type-1 Tat is an integral component of the activated transcription-elongation complex. Proc. Natl. Acad. Sci. U.S.A. 93, 2505-2510. Koff, A., Giordano, A., Desai, D., Yamashita, K., Harper, J. W., Elledge, S., Nishimoto, T., Morgan, D. O., Franza, B. R., and Roberts, J. M. (1992). Formation and activation of a cyclin E-cdk2 complex during the G1 phase of the human cell cycle. Science 257, 1689-1694. Laspia, M. F., Wendel, P., and Mathews, M. B. (1993). HIV-1 Tat overcomes inefficient transcriptional elongation in vitro. J. Mol. Biol. 232, 732-746. Madore, S. J., and Cullen, B. R. (1993). Genetic analysis of the cofactor requirement for human immunodeficiency virus type 1 Tat function. J. Virol. 67, 3703-3711. Marciniak, R. A., and Sharp, P. A. (1991). HIV-1 Tat protein promotes formation ofmore-processive elongation complexes. EMBOJ. 10, 4189-4196. Marciniak, R. A., Calnan, B. J., Frankel, A. D., and Sharp, P. A. (1990). HIV-1 Tat protein transactivates transcription in vitro. Cell (Cambridge, Mass.) 63, 791-802. Milligan, J. F., and Uhlenbeck, O. C. (1989). Synthesis of small RNAs using T7 RNA polymerase. In "Methods in Enzymology" (J. E. Dahlberg and J. N. Abelson, eds.), Vol. 180, pp. 51-62. Academic Press, Orlando, FL. Puglisi, J. D., Tan, R., Calnan, B. J., Frankel, A. D., and Williamson, J. R. (1992). Conformation of the TAR RNA-arginine complex by N M R spectroscopy. Science 257, 76-80. Roy, S., Delling, U., Chen, C., Rosen, C. A., and Sonenberg, N. (1990). A bulge structure in HIV-1 TAR RNA is required for Tat binding and Tat-mediated trans-activation. Genes Dev. 4, 1365-1373. Selby, M. J., and Peterlin, B. M. (1990). Tram-activation by HIV-1 Tat via a heterologous RNA binding protein. Cell (Cambridge, Mass.) 62, 769-776. Sheline, C. T., Milocco, L. H., and Jones, K. A. (1991). Two distinct nuclear transcription factors recognize loop and bulge residues of the HIV-1 TAR RNA hairpin. Genes Dev. 5, 2508-2520. Smith, D. B., and Johnson, K. S. (1988). Single-step purification ofpolypeptides expressed in Escherichia coli as fusions with glutathione-S-transferase. Gene 67, 31-40. Southgate, C., Zapp, M. L., and Green, M. R. (1990). Activation of transcription by HIV-1 Tat protein tethered to nascent RNA through another protein. Nature (London) 345, 640-642. SufiS, C., and Garcia-Blanco, M. A. (1995a). Transcriptional trans activation by HIV-1 Tat requires specific coactivators that are not basal factors. J. Virol. 69, 3098-3107. Sufi6, C., and Garcia-Blanco, M. A. (1995b). Spl transcription factor is required for in vitro basal and Tat-activated transcription from the human immunodeficiency virus type 1 long terminal repeat. J. Virol. 69, 6572-6576. SufiS, C., Hayashi, T., Liu, Y., Lane, W. S., Young, R. A., and Garcia-Blanco, M. A. (1997). CA150, a nuclear protein associated with the RNA polymerase II holoenzyme, is involved in Tat-activated HIV-1 transcription. Mol. Cell. Biol. In press. Waszak, G. A., Hasler, J. M., McQuade, T. J., Tarpley, W. G., and Deibel, M. R., Jr. (1992). Purification of an active monomeric recombinant HIV-1 trans-activator. Protein Express. Purif 3, 126-133. Weeks, K. M., Ampe, C., Schultz, S. C., Steitz, T. A., and Crothers, D. M. (1990). Fragments of the HIV-1 Tat protein specifically bind TAR RNA. Science 249, 1281-1285. Wu, F., Garcia, J., Sigman, D., and Gaynor, R. (1991). Tat regulates binding of the human immunodeficiency virus trans-activating region RNA loop-binding protein TRP-185. Genes Dev. 5, 2128-2140. Zhou, Q., and Sharp, P. A. (1995). Novel mechanism and factor for regulation by HIV-1 Tat. EMBO d. 14, 321-328.

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HeLa Nuclear Extract" A Modified Protocol Cameron R. Miller Department of Molecular Cancer Biology Duke University Medical Center Durham, North Carolina 27710

Sharon F. Jamison* Department of Molecular Cancer Biology Duke University Medical Center Durham, North Carolina 27710

Mariano A. Garcia-Blanco Departments of Molecular Cancer Biology, Microbiology, and Medicine Duke University Medical Center Durham, North Carolina 27710

I. G R O W T H

OF THE

HeLa

CELLS

A. MATERIALS

100-ml glass spinner flask with paddle 250-ml glass spinner flask with paddle Two l-liter glass spinner flasks with paddles Two 8-liter glass spinner flasks with stir bars in place of paddles (all flasks are from BeHco Glass, Inc., Vineland, NJ)

B. METHODS

HeLa $3 cells (human epithelioid carcinoma, cervix) were obtained from ATCC stocks. The cells have been adapted to grow in Joklik's MEM medium (GIBCO-BRL, Gaithersburg, MD) containing 5% defined Bovine Calf Serum (Hyclone Lab., Logan UT), supplemented with penicillin-streptomycin (GIBCO) and NaHCO3 (MaUinckrodt, Phillipsburg, NJ) at 37~ The cells were grown in suspension and stirred using magnetic bars at medium speed. All flasks must be tightly sealed in order to keep the CO2 at appropriate levels. HeLa cells are grown at an ideal density of 5-8 X 105/ml (minimum density should be 3 X 10S/ml) in * Present address:

INTRONN LLC, West Main Street, Durham, North Carolina 27702. 25

m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

26

Cameron P,,. Miller et al.

20 liters of medium. HeLa cells double approximately every 24 hr, and in order to keep a density of 5-8 • 10S/ml, half of the volume of the cell suspension should be discarded daily and the same amount of fresh medium should be added until expansion is required (Table I).

II. E X T R A C T

PREPARATION

A. MATERIALS For a 20-liter batch:

Buffer A (0.3 liter): 10 m M N-2-hydroxyethylpiperazine-N'-2ethanesulfonic acid (HEPES)* (pH 7.9), 1.5 m54 MgC12, 10 m M KC1, 0.5 m M dithiothreitol (DTT) Buffer C (0.05 liter): 20 m M HEPES* (pH 7.9), 25% (v/v) glycerol, 0.42 M NaC1, 1.5 ~ MgC12, 0.2 ~ ethylenediaminetetraacetic acid (EDTA), 0.5 m M DDT, 0.5 m M phenylmethylsulfonyl fluoride (PMSF)'I"

Table I A 20-Liter B a t c h G r o w t h S c h e d u l e

Volume a (liters)

Volume discarded (cell suspension) (liters)

Volume added (fresh medium) (liters)

1 2 3 4 5 6 7 8 9

0.40 0.40 0.70 1.00 1.50 2.00 3.00 5.00 10.0

0.20 0.05 0.20 0.25 0.50 0.50 0.50 0 0

0.20 0.35 0.50 0.75 1.00 1.50 2.50 5.00 10.0

10

20.0

Day

Size flask b (liters) 1 1 1 1 2 2 2 2 2

X X X • • • • • •

1 1 1 1 1 1 8 8 8

a This is an example if the cell density is at 5 - 8 • 10S/ml each day before fresh medium is added. b Cells grow better at high volumes in the flasks. Keep these volumes as high as possible without increasing the risk of contamination before transferring the cells to larger flasks.

* Must be Calbiochem (La Jolla, CA) Free Acid Ultrol Grade brand, Cat. No. 391338. t PMSF used for B cell extracts only.

2. HeLa Nuclear Extract: A Modified Protocol

27

Buffer D (3.0 liter): 20 mM HEPES* (pH 7.9), 20% (v/v) glycerol, 0.2 mM EDTA, 0.1 M KC1, 0.5 mM DTT, 0.5 mM PMSFt

B. ~ T H O D S 1. Day Before Preparation a. Prepare buffers A, C, and D and store at 4~ (stir buffer D overnight). Prepare 1 M DTT solution and store at -20~ Prepare 500 ml sterile phosphate-buffered saline (PBS) and 500 ml sterile dH20 and store them at 4~ Buffer B is prepared only if $100 extract is made (Dignam et al., 1983). b. Bake one glass homogenizer (one tight and one loose pestle) in a dry oven, wrapped in aluminum foil, for 2 hr at a temperature greater than 100~ Cool to room temperature and store at 4~ c. Place two JA-10 rotors (Beckman, Fullerton, CA) and one JA-20 rotor (Beckman) at 4~

2. Steps Before Starting Preparation of Extract a. Thaw DTT; place it at 4~ (to be added to all buffers during third spin). b. Gather 12 Beckman II 500-ml bottles, 8 Beckman polycarbonate tubes (for use with the Beckman JA-20 rotor), 7 50-rnl Coming polypropylene tubes, 4 15-ml Corning polypropylene tubes, and 2 ice buckets. c. Place JA-10 rotors in centrifuges (the use of two centrifuges is preferable to decrease the time required for Step 3,a).

3. Nuclear Extraction The extraction procedure described here is a modification of the procedure ofDignam et al. (1983). The extraction takes approximately 11 hr from first spin to freezing of the aliquots a. Take out a 3-ml aliquot of the HeLa cell suspension and count in the hemocytometer (VW1K, NO. 15170-173). Pour the cell suspension into the 500-ml bottles to just below the upper curve of the bottles; all bottles should have approximately the same volume for all four spins (10 liters per spin). Spin cells in a Beckman JA-10 rotor for 10 rain at 1500 rpm (398 g) at 4~ Discard the supernatant fraction by pouting it into a beaker with bleach. Pour fresh cell suspension

28

Cameron R.. Miller et al. into the bottles, which have the cell pellets of the prior spin; you will note that the pellets will disperse. Repeat this process until all of the cells are pelleted. You should see the pellets increasing in volume with each subsequent spin. During the spins, do the following:

(1) First spinmcount the cells. (2) Second spin--calculate the volume of buffer C to be used: 3 ml per 10 9 cells. (3) Third spinmadd D T T to all the buffers that call for it. (4) Fourth spinmput six 50-ml Coming tubes on ice.

b. The following steps must be carried out in a cold room or on ice (it may be advisable to use an ice-water bath for some steps if they are done outside the cold room). Pour off most of the supernatant fraction until the level is just below the top of the cell pellet in one bottle and pour the supernatant fraction of a second bottle completely off. Using a quick swirling motion, release the pellet from the first bottle and quickly transfer the cell suspension to the bottle without medium. Check to be sure that all of the suspension was transferred. Quickly transfer it to the 50-ml polypropylene tube (Coming) and repeat this step until all of the cell pellet has been resuspended. Spin for 10 rain at 1500 rpm in an IEC centrifuge (model HN-SII), using the swinging bucket rotor, at 4~ c. Place the tubes on ice. Record the cell pellet volume on the side of each tube. Accurate measurement of the volume of the cell pellets is important. Decant the supernatant and gently tap the tubes to loosen the pellets. P, esuspend the pellets in chilled PBSmfive volumes per original cell pellet volume (e.g., for a 5-ml pellet, add 25 ml PBS). Cap the tubes and manually loosen the pellets by gently flicking the bottom of the tubes. Do not vortex or shake vigorously! R.esuspend the cells using a 10-ml pipette. Pipette up and down four times from the bottom for each tube or as necessary to ensure that the cell suspension is homogeneous. d. Spin for 10 rain at 1500 rpm in an IEC centrifuge in the swinging bucket rotor at 4~ Decant the supernatant and place the cells back on ice. e. Loosen the pellets and resuspend the cells in five volumes of chilled buffer A per original pellet volume, following the same procedure used in Step 3,c. This is a critical step; be as accurate as possible with volume measurements.

2. HeLa Nuclear Extract: A Modified Protocol

29

f. Let the resuspended cells stand for 10 min on ice to swell. g. Spin for 10 min at 2500 rpm in the IEC centrifuge in the swinging bucket rotor at 4~ Using a 25-ml pipette, remove as much of the supernatant as possible without disturbing the pellets. Cell pellets should have doubled in volume. h. Resuspend the swollen cell pellets in two volumes of buffer A. Note: Use the original cell pellet volume from Step 3,c. Do not use pipettes to resuspend the swollen cells; just loosen them enough to get all the pellets off the sides of the tubes because they may stick. i. Pour the suspension into a Dounce homogenizer. Lyse the cells with 10 strokes of the tight pestle (pestle A) at medium speed with a continuous and regular motion, holding the Dounce homogenizer vertically in an ice bucket. Optional: compare the swollen and lysed cells by microscopic inspection. j. Pour the lysed cells into Beckman polycarbonate clear-capped tubes for the JA-20 rotor. Spin for 15 min at 3000 rpm at 4~ (1090 g). Rinse the Dounce homogenizer with sterile deionized water after use, as it is needed in Step 3,o. k. The pellets will be soft and faint at this step; carefully remove the turbid, pink supernatant with a 10-ml pipette. (It can be saved for a pre-S100 extract in 50-ml Coming tubes.) 1. Spin the pellets again in the same Beckman tubes for 20 min at 14,500 rpm (25,400 g) at 4~ m. Pellets are the crude nuclei. (Discard the supernatant fraction or pool it with previous supernatant as pre-S100 if desired.) n. Divide and distribute a calculated amount of buffer C to each tube [see Step 3,a(2)] and resuspend just enough to release the pellets from the bottom of the tubes. This can be achieved by swirling; the pellets should be broken up in the following step. o. Pour the cell suspension into the previously rinsed Dounce homogenizer. Lyse the nuclei with 10 strokes of the loose pestle (pestle B) at medium speed, using a continuous and regular motion; hold the Dounce homogenizer vertically in an ice bucket. p. Pour the lysed suspension into the 15-ml Coming tubes and rotate them in a cold room for 30 min. The suspension should be thick and stringy in appearance. q. Pour the suspension into Beckman polycarbonate tubes and spin in the JA-20 rotor for 30 min at 14,500 rpm (25,400 g) at 4~ Five minutes before the end of the spin, hydrate the dialysis tubing (The Spectrum Companies, Gardena, CA, Cat. No. 132-650, MWCO: 6000-8000, diameter: 14.6 mm). Place in a l-liter beaker with a stir bar.

30

Cameron P.. Miller et

al.

r. Pipette the cloudy supematant fraction, the nuclear extract, into the dialysis membrane. Dialyze against buffer D as follows: 1 liter for 1 hr 1 liter for 2 hr 1 liter for 2 hr s. Mix the dialyzed nuclear extract in a 50-ml Coming tube; place it on ice and aliquot as desired. t. Freeze immediately in dry ice; leave it for at least 10 min. Snap freezing in liquid nitrogen is also recommended, although not necessary. Store in a - 8 0 ~ freezer or in liquid nitrogen. u. The protein concentration of the nuclear extract should be quantified after a quick centrifugation to remove large aggregates using the Bradford Dye Assay (Bio-Rad, Hercules, CA). The concentration should be approximately 15 mg/ml.

REFERENCE Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983). Accurate transcriptioninitiation by RNA polymeraseII in a soluble extractfrom isolatedmammaliannuclei. Nucleic Acids Res. 11, 1475-1489.

Functional Analysis of Splicing Factors and Regulators Juan Valc~ircel Gene Expression Programme European Molecular Biology Laboratory D-69117 Heidelberg, Germany

Concepci6n Martlnez Gene Expression Programme European Molecular Biology Laboratory D-69117 Heidelberg, Germany

Michael R. Green Howard Hughes Medical Institute and Program in Molecular Medicine University of Massachusetts Medical Center Worcester, Massachusetts 01605

Biochemical identification and analysis of components of the splicing machinery requires the development of functional assays to test their associated activities. Chromatographic and immunological procedures have been utilized to generate splicing-deficient nuclear extracts whose activity depends on the addition of purified factors. These procedures have also proved valuable in analyzing mechanisms of alternative splicing regulation.

I. I N T R O D U C T I O N Eukaryotic messenger RNAs are synthesized as precursors (pre-mRNAs) containing intervening sequences (introns), which are removed in the cell nucleus before the R N A is exported to the cytoplasm and translated. Intron removal and splicing together of the flanking sequences (exons) are essential steps in the expression of eukaryotic genes, and are often subject to regulation (for reviews, see Green, 1991; Moore et al., 1993; Kr~imer, 1995; Adams et al., 1996). Intron removal occurs by means of two transesterification reactions. In the first reaction, the 5' exon is excised and the 5' end of the intron forms a phosphodiester bond with the ribose 2 ' - O H group of an adenosine (termed the branchpoint), usually located close to the 3' end of the intron. In the second step of the reaction, the two exons are ligated and the intron is released in a lariat configuration. 31 m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

32

Juan Valcfircel et

al.

Pre-mRNA splicing takes place in macromolecular complexesmspliceo somesmcomposed of five small nuclear ribonucleoprotein particles (snRNPs) and a larger number of polypeptides that are not tightly associated with snRNPs (for reviews, see Madhani and Guthrie, 1994; Kr~imer, 1996). The snRNPs consist of one small nuclear R N A (snRNA) bound by two classes of polypeptides (reviewed by Ltihrmann et al., 1990): (1) proteins that are common to most snRNPs and (2) proteins that are specific for particular snRNPs. Non-snRNP proteins are diverse in structure and function, although families of factors with conserved motifs and domain organization have been identified (reviewed by Lamm and Lamond, 1993; Kr~imer, 1996). For example, two types of modules are often present in mammalian splicing factors and regulators: the R N P consensus motif (RNP-CS) and arginine-serine-rich (RS) domains (reviewed by Birney et al., 1993). Thus, an entire family ofpolypeptides involved in p r e - m R N A splicing~the SR proteins~have a similar modular structure consisting of one or two R N P - C S domains and a carboxy-terminal RS domain of variable length (for reviews, see Fu, 1995; Manley and Tacke, 1996; Valcfircel and Green, 1996). The first part of this chapter reviews biochemical methods used for the depletion ofnon-snRNP proteins from HeLa nuclear extracts and provides detailed protocols for the preparation of extracts in which the essential splicing factor U2AF has been depleted. The second part reviews biochemical assays that have been utilized to study mechanisms of alternative splicing in vitro.

II. F U N C T I O N A L

ANALYSIS OF GENERAL SPLICING FACTORS

The formal proof that a gene product is a component of the splicing machinery requires the demonstration that its absence compromises the splicing activity of an extract or a cell, as well as a demonstration that this "depleted" system can be complemented either by gene transfer in vivo or by adding back the purified or, optimally, the purified recombinant factor in vitro.

A. In Vitro DEPLETION SYSTEMS 1. Biochemical Depletion Early studies on pre-mRNA splicing were designed to biochemically ffactionate nuclear extracts and then reconstitute the reaction with a minimum number of purified components (Krainer and Maniatis, 1985; Furneaux et al., 1985; Kr~imer et al., 1987). Yeast genetics and biochemical analysis of mammalian spliceosomes, however, have estimated that the number of gene products required for pre-

3. Analysisof Splicing Factors

33

m R N A splicing is around 50-100, making the full reconstitution of splicing reactions from purified components a staggering project. Given this complexity, it is not surprising that standard procedures of biochemical fractionation are usually too unspecific to allow the depletion of single factors. A few rather simple biochemical procedures, however, have proved very useful in the characterization of some factors. Below we review some of these procedures.

a. Depletion of U 2 A F by Oligo(dT)-Cellulose

Chromatography

U2AF is a splicing factor that binds to the polypyrimidine tract that usually precedes the 3' splice sites of higher eukaryotic pre-mRNAs (Zamore et al., 1992; Singh et al., 1995). The factor is required for the association of U2 snRNP with the pre-mtLNA branchpoint region (Ruskin et al., 1988). U2 auxiliary factor (U2AF) purified from HeLa nuclear extracts consists of two subunits of 65 and 35 kDa, (Zamore and Green, 1989). Only U2AF 65 binds to the polypyrimidine tract, U2AF 35 being tethered to the pre-mtLNA through interaction with U2AF 65 (Zhang et al., 1992). One of the steps utilized in the purification protocol suggested a simple procedure to prepare extracts depleted of this factor (Zamore and Green, 1991) (Fig. 1). Because U2AF can bind poly(U) at high salt concentrations (1 M KC1), it was possible to optimize conditions that allow U2AF binding but not the binding of other poly(U)-binding factors. Subsequent studies revealed that U2AF 65 can

NE, 1MKC1

Depleted NE, 1M KC1 ]Dialysis ] Depleted NE Figure 1 PreparationofU2AF-depleted HeLa nuclear extracts by oligo(dT)-cellulose chromatography. Extracts are adjusted to 1 M KC1 and loaded on an oligo(dT)-cellulosecolumn. The flowthrough fraction corresponds to the depleted extract.

34

Juan Valcfircel et al.

bind DNA as well as 1KNA (Kanaar et al., 1995; 1K. Singh and M. 1K. Green, unpublished observations), allowing the use of oligo(dT)-cellulose rather than poly(U)-Sepharose in the depletion procedure. Oligo(dT)-cellulose does not retain some of the other polypyrimidine-tract binding proteins and has fewer batch-tobatch variations. Below we describe a detailed, simple procedure that has consistently provided depleted extracts with uniform properties, and whose splicing activity can be rescued by addition of purified U2AF (Zamore and Green, 1991) or of the recombinant 65 subunit, which is apparently sufficient to rescue the splicing activity of the extract (Zamore and Green, 1991; Zamore et al., 1992). 1. Weigh 500 mg oligo(dT)-cellulose (Boehringer Mannheim, Mannheim, Germany, Cat. No. 808 229), and swell the beads in 10 ml ice-cold 1 M KC1 buffer D [20 mM N'-2-hydroxyethylpiperazineN'-2-ethanesulfonic acid (HEPES) pH 8.0, 0.5 mM ethylenediaminetetraacetic acid (EDTA), 20% glycerol, 1 mM dithiothreitol (DTT), 0.05% NP40]. Keep on ice for 30 min. Wash the beads twice with 10 ml of the same buffer: sediment the beads by centrifugation at 1000 rpm for 5 min and resuspend them in new buffer. The volume of the swollen beads can vary from batch to batch of oligo(dT)-cellulose. 2. Load the suspension of oligo(dT)-cellulose beads on a 10-ml column and pack the column by gravity. Wash the column with 5 ml of 1 M KC1 buffer D. 3. Dissolve 67 mg KC1 in 1 ml of HeLa nuclear extracts, prepared as described by Dignam et al. (1983), by repeated inversion in an Eppendorf tube. Measure the conductivity to confirm that the salt concentration is 1 M. If the salt concentration differs by more than 5% from 1 M, readjust the concentration either by diluting with more of the original extract or by dissolving more KC1 crystals. 4. Load 1 ml of the nuclear extract adjusted to 1 M KC1 in two steps of 0.5 ml each. Collect the flowthrough in aliquots of 0.5 ml. 5. Add 1 M KC1 buffer D to the column and collect the flowthrough in five or six aliquots of 0.5 ml. 6. Check the protein concentration (using, for example, the Bradford assay). Pool the two aliquots with the highest protein concentration. 7. Dialyze inmediately against 1 liter of 0.1 MKC1 buffer D for 5 hr at 4~ 8. To remove the aggregates formed during dialysis, centrifuge the extract for 5 min at 13,000 rpm in a microcentrifuge at 4~ Distribute the supernatant in 20-/xl aliquots and freeze in liquid nitrogen. This is considered the depleted extract. Keep at -70~ The protein concentration of the depleted extracts usually varies between 70% and 80% that of the original extract.

3. Analysisof Splicing Factors

35

It is generally useful to elute the material bound to oligo(dT), both to use it as a source of partially purified U2AF and to regenerate and reutilize the column. 9. Wash the column with 10 rnl of 2.5 M KC1 buffer D. 10. Elute with 2 M guanidine-HC1 (USB) buffer D. Add 250/,1 of solution and recover the flowthrough of each fraction separately. Test the protein concentration of each fraction and pool the fractions with the highest protein concentration. This eluate contains U2AF and other proteins that bind tightly to oligo(dT). Dialyze against 0.1 M KC1 buffer D for 10-18 h, changing the dialysis buffer once. 11. Once washed with guanidine-HC1 and extensively washed with 1 M KC1 buffer D, the column can be reutilized for further depletion experiments. The packed column can be stored at 4~ in the presence of 0.1% sodium azide or at - 7 0 ~ without any buffer.

b. Splicing Complementation Assays Below we describe a protocol designed to quantify U2AF activity sensitively in chromatographic fractions or purified protein preparations. Changes in the concentration of Mg 2+ and of the nuclear extract may be required for optimal splicing of different pre-mRNAs or other types of nuclear extracts. 1. Add, in this order: 1 /,1 of premix: [for five samples: 1 /,1 pre-mRNA (100,000 cpm, 29 fmol, 6 fmol/reaction) 0.5/,1 of 270 mM MgC12 0.5/,1 of 0.1 M ATP 2/,1 of 0.5 M creatine phosphate 1 /,1 of RNasin 40 U//,I] 2.5/,1 of 0.1 M KC1 buffer D containing the chromatographic fraction or purified protein. 3.5/,1 nuclear extract or depleted extract (adjust this volume for each type of extract according to its protein concentration). 2/,1 15% polyvinyl alcohol (PVA) Final volume = 9/,1. Final concentration: pre-mRNA: 6-10 fmol/9 /,1--- <1 nM; MgC12: 3 raM; KCI: 66.6 mM; ATP: 1.1 raM; creatine phosphate: 22.2 raM; RNasin: 0.9 U//,I; HEPES pH 8.0:13.3 raM; EDTA:

36

Juan Valcfircel et al.

0.13 m/l//; glycerol: 13.3%; PVA: 3.3%; NP40: 0.03%; DTT: 0.66 mM; nuclear extract: 39% ('--3/zg//zl final protein concentration). The nuclear extract is in buffer D 0.1 M KC1. The contributions of the nuclear extract to the final concentrations of HEPES, KC1, EDTA, and DTT have been taken into account in the final concentrations listed above. 2. Incubate at 30~ for 60-180 rain. 3. Add 94/zl of proteinase K solution: 1 /zl of 8/zg//zl tRNA 24/zl proteinase K, 10 mg/ml 69/zl proteinase K buffer 2• [200 mM Tris 7.5, 300 mad NaC1, 2% sodium dodecyl sulfate (SDS), 25 ~ EDTA] 4. Incubate at 65~ for 20 man. 5. Extract twice with 95/zl phenol/chloroform and once with 95/zl chloroform. 6. Precipitate with 300/zl ethanol. 7. Centrifuge at 13,000 rpm for 15 min at 4~ in a microcentrifuge, wash the pellet with 80% ethanol, and resuspend the dried pellet in 6/zl formamide-containing sequencing loading dye. 8. Boil 5 min and load on a polyacrylamide denaturing gel. Perform electrophoresis and autoradiography. Thirteen percent polyacrylamide gels are often employed because lariat intermediates and products usually migrate above the R N A precursor. This is particularly useful to quantify both steps of the reaction when the splicing efficiency is low and there is substantial degradation of the pre-mRNA in the extract. This assay has been utilized to analyze the functional activities of the different protein domains of U2AF 6s (Zamore et al., 1992; Valc~ircel et al., 1996). Figure 2 shows an example of this type of analysis. The amino-terminal 94 amino acids of U2AF 6s are essential for function and contain an RS domain. To understand the structural requirements of the RS domain, an adenovirus model substrate was spliced in vitro either in nuclear extracts or in oligo(dT)-depleted extracts that were complemented with purified recombinant U2AF 6s or a U2AF 6s mutant protein in which the 94 amino-terminal amino acids were substituted for by 7 arginineglycine dipeptides. This result supports the view that only arginine residues (and, more specifically, positively charged amino acids) are important for the function of the U2AF 6s RS domain (Valc~lrcel et al., 1996). c. Troubleshooting

This protocol must be optimized for each new batch of nuclear extract. The typical problems are either (1) that the extracts are not completely depleted, having

3. Analysis of Splicing Factors

37

Figure 2 Reconstitution of the splicing activity of U2AF-depleted nuclear extracts by complementation with recombinant U2AF 6s or a mutant derivative in the RS domain. For discussion see text. The concentrations of U2AF 6s and the mutant that replaces the RS domain by seven R G dipeptides used in the complementation assay were: 5, 15, 30, and 50 ng//.tl.

38

Juan Valcfircel et al.

an unacceptable background of splicing activity in the absence of exogenously added U2AF or (2) that the extracts are depleted of additional splicing components, and thus it is not possible to complement them by addition of recombinant U2AF 65 alone. These problems can usually be solved by adjusting the volume of nuclear extract applied to the column: a smaller volume if the extract is not well depleted or a larger volume if difficulties in complementation arise. Changes in the salt concentration at which the chromatography is performed are usually not useful: salt concentrations below 1 M KC1 result in extracts depleted in additional polypyrimidine-tract binding splicing factors like the polypyrimidine-tract binding protein (PTB)-associated splicing factor (PSF) (Patton et al., 1991, 1993; and see below); salt concentrations above 1 M KC1 usually result in extracts that are only marginally depleted in U2AF. A limited degree of optimization can also be accomplished by small variations in the amounts of nuclear extract utilized in the splicing assays: lower amounts if the extracts show a slightly high background of splicing activity; higher amounts if the extracts are not readily complemented by addition of U2AF. It is important to perform a titration curve to determine the concentration of U2AF that gives optimal complementation (Fig. 2). Concentrations above this optimal value result in lower levels of complementation, presumably due to the squelching of splicing factors that interact with U2AF. d. Assay for PSF Activity

Patton and colleagues observed that a variation of the depletion protocol described above allowed the detection of a splicing activity distinct from that of U2AF (Patton et al., 1991). When nuclear extracts were adjusted to 0.5 M instead of 1 M KC1 and chromatography in poly(U)-Sepharose was performed as indicated in the previous section, the splicing activity of the flowthrough fraction could not be restored by addition of purified U2AF, but it could be restored by the combination of U2AF and a fraction eluted from the column at 1 M KC1. Further work characterized a splicing activity present in the latter fraction as a 100-kDa polypeptide with affinity for polypyrimidine tracts, which is essential for the second step of the splicing reaction (Patton et at., 1993; Gozani et al., 1994).

e. Cytoplasmic $100 Extracts Are Depleted of SR Proteins Conventional protocols for preparation of nuclear extracts (Dignam et al., 1983; Kr~imer and Keller, 1990) involve the separation of nuclei from cytoplasm and membranes by centrifugation at low speed after cell lysis. The supernatant is then used to obtain cytoplasmic extracts by centrifugation at 100,000 g, which sediments membranes, organelles, and large aggregates. The supernatant of this second centrifugation is termed an $100 extract.

3. Analysisof Splicing Factors

39

It has been observed that $100 extracts contain significant amounts ofspliceosomal snRNPs and many other splicing factors, but they lack SR proteins. These extracts are not able to support splicing reactions in the absence of exogenously added SR proteins and are therefore useful depleted systems to study these factors. $100 complementation has been used as a criterion to define new members of the SR family (Krainer et al., 1990a; Ge and Manley, 1990; Fu et al., 1992; Zahler et al., 1993; Cavaloc et al., 1994) and for the functional characterization of their functional domains (Cficeres and Krainer, 1993; Zuo and Manley, 1993). Remarkably, any one of the SR proteins is able to stimulate splicing in $100 extracts, an observation that suggested redundant functions for these factors. Additional experimentation, however, identified differences in the in vitro activities of individual members of the family (Kim et al., 1992; Zahler et al., 1993; Fu, 1993; Zahler and Roth, 1995) and established the requirement for individual members in vivo (Ring and Lis, 1994; Peng and Mount, 1995). It is therefore unclear which is the specific rate-limiting step for splicing in $100 extracts (if there is only one) that can be stimulated any SR protein. f. Depletion of snRNPs with Antisense Oligonucleotides

A highly specific method for the depletion of particular snRNPs from mammalian extracts was developed by Lamond, Sproat, and co-workers (reviewed by Lamond and Sproat, 1993), based on hybridization of 2'-O-alkyl oligoribonucleotides to the snRNA component of the snRNPs. These oligonucleotides have three properties that make them particularly useful for this purpose: they are chemically stable in the extracts, they efficiently hybridize to complementary RNA, and the hybrids are resistant to cleavage by RNase H. Particular snRNPs bound by biotinylated 2'-O-alkyl oligo,ribonucleotides can then be removed by affinity chromatography on matrices containing avidin or anti-biotin antibodies. Detailed protocols for the antisense affinity depletion of R N P particles have been described by Blencowe and Barabino (1994). 2'-O-Methyl phosphonate-modified oligoribonucleotides have also been used for this purpose (Temsamani et al., 1991). Important insights into the function of mammalian snRNPs have been obtained using this depletion technique, including the involvement of U1 snRNP in U2 snRNP binding to the 3' splice site (Barabino et al., 1990) and the dispensability of U1 snRNP for splicing under conditions of excess SR proteins (Crispino et al., 1994).

g. Other Depletion Procedures (1) Heat Inactivation It has been observed that nuclear extracts heated to 43~ for 20 rain are able to support the first but not the second step of splicing (Krainer and Maniatis, 1985; Reed and Maniatis, 1988). This assay was used to

40

Juan Valc&cel et al.

obtain a chromatographic fraction able to complement those extracts that, in its most purified form, contained two 70-kDa proteins and a minor 100-kDa polypeptide (Delanoy and Caruthers, 1991). The assay was also crucial in studying a later event in the splicing process: the release of the spliced product from splicing complexes (Sawa and Shimura, 1991). Heat treatment has also been employed to inactivate specific components of extracts obtained from temperature-sensitive (ts) mutant yeast strains. For example, Company et al. (1991) observed that extracts prepared from a prp22 mutant strain pretreated at 30~ for 60 min showed a splicing defect similar to that observed in vivo when the strain was grown at a restrictive temperature. The defect could be suppressed both in vivo by a wild-type prp22 gene and in vitro by complementation with purified prp22 wild-type protein.

(2) Study of Partial Reactions Kfiimer et al. fractionated nuclear extracts in seven chromatographic fractions that were all required for in vitro pre-mR.NA splicing (Kr~imer et al., 1987). Three of these extracts were sufficient, however, to promote U2 sntkNP binding to the 3' splice site region (Kfiimer and Utans, 1991) and were purified and characterized further using this assay. These authors identified one of the fractions as U2AF and established the protein composition of the other two. The activity associated with one of these remaining fractions corresponded to a single heat-stable 75-kDa protein (Kr~imer, 1992), while the activity associated with the other fraction corresponded to seven polypeptides (Brosi et al., 1993a,b). 2. Immunodepletion Antibodies have been extensively used in the analysis of splicing factors. Immunoinhibition and immunodepletion experiments performed with antibodies generated against characteristic structural components of the snRNPs, for example, were important tools in the establishment of their essential roles in the splicing process (Padgett et al., 1983; Kfiimer et al., 1984; Krainer, 1988; Winkelmann et al., 1989). The development of specific antibodies against some non-snlLNP proteins has also been decisive in validating their function as splicing factors in the absence of other depletion procedures. Table I compares several immunodepletion protocols that have been successfully utilized. The examples have been selected to illustrate the variety of procedures and conditions employed. Although some of these differences can be attributed to the different affinities, specificities, or concentrations of the antibodies utilized, other small changes in the experimental procedure have a significant effect on the final result. While the antibody is usually immobilized on a resin, allowing minimal dilution of the extract, formation of antigen-antibody complexes before they are immobilized may be required in particular cases. While extracts are often depleted under conditions

Table I Immunodepletion

Factor

Antibody (AB)

Protocols ~

Conditions

Procedure Splicing reactions (100 ~,1) containing 30/,1 of NE are preincubated with ATP at 30~ for 30 min and then applied to a centrifuge filter unit containing the immobilized Ab. Incubation at 4~ for 30 rain. Depleted extract is collected by centrifugation through the filter unit. Incubation of NE and immobilized antibody in 0.42 M NaC1 at 0-4~ for 1 hr. Depleted extract and beads are separated by centrifugation. NE is passed through a column containing the immobilized Ab 5 times at room temperature. NE is circulated over the column for 2.5 hr at a flow rate of 0.5 ml/min in 0.5 M KC1.

SC35 (human SR protein)

Monoclonal (MAb)

3 mg of purified MAb immobilized in 100/,1 of Affigel 10 resin (Bio-Rad, Richmond, CA) for 30/,1 of HeLa NE

9G8 (human SR protein)

MAb

U2AF 35 (human RS-containing protein) CBP 80 (human cap-binding protein) prp-16 (yeast DEAH protein)

Polyclonal Ab

10 mg ammonium sulfate precipitate from ascites fluid immobilized in 270/zl of CNBr-activated Sepharose 4B CL (Pharmacia, Uppsala, Sweden) for 300/zl of HeLa NE 0.5 ml of antiserum immobilized in 250/,1 of proteinA-Trisacryl (Pierce, Rockford, IL) for 500/,1,1 of HeLa NE 5 ml of antiserum immobilized in 800/,1 of protein A-Sepharose 4 Fast Flow (Pharmacia, Uppsala, Sweden) for 4 ml of HeLa NE 125/,g of IgG for 60/,1 of yeast NE, then immobilized in protein A-Sepharose (Pharmacia, Piscataway, NJ)

a

Polyclonal Ab

Polyclonal Ab

NE, nuclear extract; CNBr, cyanogen bromide.

Reference Fu and Maniatis (1990)

Cavaloc et al. (1994)

Zuo and Maniatis (1996) Izaurralde et al. (1994)

NE is incubated with the antiserum at 0~ for Schwer and Guthrie 1 h, then added to the protein A beads and (1991) incubated at 4~ for 1 hr. Depleted extracts and beads are separated by centrifugation.

42

Juan Valdircel et al.

that support splicing, efficient depletion may require high salt concentrations. Preincubation in the presence of ATP has been found to improve the efficiency of depletion of particular factors, presumably because this treatment releases them from preformed complexes. It is always advisable to test how dilution of the extract before depletion, repeated exposure of the extract to the antibody (e.g., recircularization over the same column or chromatography on a different column), or different types of solid matrices affect the efficiency of a depletion procedure. Although most depletion protocols require considerable amounts of antibody, the columns can frequently be regenerated if the antibody has been covalently linked to the resin, for example, by treatment with 0.1 M triethylamine, pH 11.5.

B. In Vivo DEPLETION PROCEDURES Yeast genetics offers paradigmatic examples of in vivo depletion procedures. Techniques for the isolation of conditional mutants, synthetic-lethal mutants, and the targeted disruption of genes are basic tools for the analysis of all aspects of yeast biology, including 1KNA processing. The description of these techniques is beyond the scope of this review; detailed protocols, as well as examples of the use of these techniques, can be found in the comprehensive and excellent reviews by Guthrie and Fink (1991) and by Rymond and Rosbash (1992). The identification of splicing factors in another genetically tractable organism, the fruit fly Drosophila melanogaster, however, has followed a singular turn. Rather than exploiting Drosophila genetics to identify splicing factors, workers identified some of the genes encoding them by homology to their mammalian counterparts. Once the chromosomal locations of these genes were established, imprecise excision of P elements integrated nearby was used to knock out the genes (Kanaar et al., 1993; Ring and Lis, 1994; Rudner et al., 1996). Rescue of such mutants with complementary DNAs (cDNAs) then confirmed their essential nature and opened the way to their functional analysis in vivo. Surprisingly, splicing analysis performed with some of these mutants showed no obvious defects in the splicing of several constitutive and regulated pre-mRNAs (Ring and Lis, 1994; Rudner et al., 1996). A possible explanation for these results is that the splicing factors analyzed are only essential for splicing of a particular set of pre-mRNAs. Very recently, Wang et al. (1996) used the highly recombinogenic chicken B-cell line DT40 to generate cells in which both alleles of the gene encoding the SR protein ASF/SF2 were disrupted. Double inactivation of the gene was lethal, and expression of an exogenous cDNA encoding ASF/SF2 under a tetracyclineregulated promoter rescued the cells' viability, thereby allowing a functional analysis of the factor in tissue culture. This technique is very likely to become of widespread use in the analysis of mammalian genes in vivo.

3. Analysisof Splicing Factors

III. A N A L Y S I S OF R E G U L A T O R Y M E C H A N I S M S PRE-mRNA SPLICING

43

IN

Pre-mRNA splicing is often regulated, thereby generating multiple spliced mRNAs from a single primary transcript. The different mature mRNAs can encode distinct protein products with different properties (for reviews, see Smith et al., 1989; McKeown, 1992; Valcfircel et al., 1995; Adams et al., 1996; Chabot, 1996). Both in vitro and in vivo studies have revealed two mechanisms involved in prem R N A splicing regulation: (1) tissue- or stage-specific regulatory factors interact with the splicing machinery to promote or repress the use of particular splice sites, and (2) changes in the relative concentrations of general splicing factors affect splice site choice. In both mechanisms, regulation seems to occur at early stages of spliceosome assembly.

A. TISSUE- OR STAGE-SPECIFIC SPLICING REGULATORS Genetic studies in Drosophila and yeast have revealed the existence of regulatory factors that control specific splice site choices at particular developmental stages or cell types. Two examples are found in a cascade of regulated splicing occurring during sex determination in Drosophila (reviewed by Baker, 1989; Norton, 1994; MacDougall et al., 1995). A female-specific regulatory protein, Sexlethal (Sxl), controls splicing of the gene transformer (tra) such that in the presence of this factor, a non-sex-specific 3' splice site used in male flies is repressed and a female-specific 3' splice site is activated (Boggs et al., 1987; Sosnowski et al., 1989). Female-specific splicing of tra pre-mRNA allows translation of the protein transformer (TRA), which cooperates with another regulatory factor, the protein TRA-2, to stimulate the use of a female-specific 3' splice site in the gene doublesex (Nagoshi et al., 1988). The recapitulation of some of these regulatory phenomena in vitro has given insights into the molecular mechanisms involved. Sxl has been proposed to antagonize the splicing factor U2AF to regulate tra splicing (Valcfircel et al., 1993). TRA and TRA-2 have been proposed to stimulate the formation of a nucleoprotein complex containing SR proteins that is located in the exon downstream from the doublesex female-specific site (Tian and Maniatis, 1993). This complex can promote U2AF binding to the weak polypyrimidine tract of the female-specific site (Zuo and Maniatis, 1996).

1. Analysis of Spliced Products in Complex RNA Mixtures by Primer Extension Using Splice Junction Oligos A limitation to the in vitro analysis of long, alternatively spliced transcripts is the presence of multiple spliced products and intermediates, as well as degradation

44

Juan Valdtrcel

et al.

products of the different RNAs present in the mixture. All these R N A molecules may obscure the unambiguous detection of the spliced products when a uniformly labeled R N A substrate is utilized. Oligodeoxynucleotides complementary to the splice junctions have been used as sensitive probes for detecting specific spliced products without the interference of other R N A species. The spliced mRNAs are detected as primer extension products of the length of the first exon plus a few nucleotides of the second exon. Figure 3 is a schematic representation of the procedure. The primers should be designed such that the oligodeoxynucleotide cannot stably hybridize, particularly at its 3' end, with either of the exons but can form a stable duplex across the exons once they are spliced together. Eighteen-nucleotide primers complementary to the first 13 nucleotides of the downstream exon and the last 5 nucleotides of the upstream exon are usually adequate for this purpose. These numbers may need to be adjusted for specific RNAs if the primer generates unacceptably high levels of primer extension product on the pre-mRNA alone.

32p Oligo for Splice Junction 1 ~t

I Reverse Transcription I

_.m

32p

32p Oligo for Splice Junction 2

1

_.m

32p

Figure 3 Analysisof alternatively spliced products by primer extension using oligodeoxynucleotides complementary to the splice junctions. The primers are designed such that they cannot hybridize to each of the exons separately, but they can hybridize to a particular spliced product. Separate hybridization reactions containing spliced ILNAs and each of the labeled primers are set up. Primer extension on each of the reactions will therefore quantify the levels of each of the alternatively spliced products.

3. Analysisof Splicing Factors

45

a. Assay

Splicing assays and isolation of the R N A after splicing are performed as described before, except that the splicing reactions should be scaled up two or three times for convenience of analysis. Modifications of the ionic concentrations and/or the percentage of nuclear extract may be required for particular premRNAs. In the experiment described in Figure 4, the magnesium concentration was 2.4 m M and the percentage of nuclear extract was 29%. 1. After splicing reactions are completed and the R N A products isolated, resuspend the pellet of purified R N A in 24/,1 water. 2. Set up hybridization reactions for each of the splice junction oligodeoxynucleotides. Each reaction should contain: 12/xl corresponding to isolated R N A 4/xl 5• hybridization buffer [0.2 M piperazine-N,N'-bis(2ethanesulfonic acid) (PIPES) pH 6.4, 5 m M EDTA, 2 M NaC1] 2/,1 2% SDS 2/xl 5'-32p-labeled oligodeoxynucleotide (120 fmol: label 6 pmol with polynucleotide kinase and [~/-32p]ATP, bring the volume to 100/xl with water, and remove the enzyme by phenol extraction and the unincorporated radioactive nucleotide by gel filtration on Sephadex G-25) 3. Heat at 90~ for 1 n-fin. 4. Keep at 37~ for 6-12 hr in an oven or warm room (avoid thermal blocks to prevent condensation at the top of the tube). 5. Add 180/,l 0.5 M ammonium acetate and 200/,l isopropanol. Immediately centrifuge at room temperature for 5 min at 13,000 rpm on a tabletop microcentrifuge. Remove the supernatant and wash the pellet with 80% ethanol. This quick precipitation step is relatively selective for the R N A and R N A - D N A hybrids, leaving most of the nonhybridized oligodeoxynucleotide in the supernatant. 6. Dry the washed pellet and resuspend it in 9.5/,1 R T mix containing: 2/zl 5 • R T buffer (250 m M Tris pH 8.3, 370 m M KC1, 30 m M MgC12) 1 /,1 of 10 m M each dATP, dCTP, dGTP, dTTP 0.5 ~1 RNasin 0.5 ~1 0.2 M DTT 5.5/~1 water 7. Add 0.5/~1 of avian myeloblastosis virus (AMV) reverse transcriptase (10 U//zl) and incubate at 42~ for 60 min.

46

Juan Valcfircel et al.

Figure 4 Alternative splicing of tra p r e - n ~ N A in vitro. The structure of the p r e - m k N A and the alternative 3' splice sites are shown at the top. Splicing reactions were performed and analyzed as described in the text and in Valcfircel et al. (1993). The bands corresponding to the products of primer extension using splice junction oligos are shown in the lower part of the figure. Although the primer

3. Analysis of Splicing Factors

47

8. Add 10/zl formamide-containing loading dye, heat at 90~176 for 5 min, and load on a denaturing polyacrylamide gel of a percentage appropriate to the length of the primer extension product. 9. Perform electrophoresis and autoradiography. Figure 4 illustrates one example of the use o f in vitro systems and the detection method described above to recapitulate the regulation of alternative splicing of tra pre-mRNA by the Sxl protein. The figure also shows that while Sxl is able to repress one particular splice site and produce a switch in splice site usage, a U2AF 6s derivative lacking the KS domain acts as a general splicing inhibitor and represses the use of both 3' splice sites. Thus, two polypyrimidine-tract binding proteins that are unable to recruit U2 snRNP act as splicing inhibitors. The higher specificity of Sxl for particular 3' splice sites (Singh et al., 1995), however, explains its specific regulatory function during Drosophila development.

B.

CHANGES

IN THE LEVELS

OF GENERAL

SPLICING

FACTORS

The SP,. protein ASF/SF2 was purified as an activity promoting the use of proximal 5' splice sites (Ge and Manley, 1990; Krainer et al., 1990b). Therefore, a general splicing factor can act to modulate the relative use of alternative splice sites, an observation that has been extended to other members of the SP,. family as well as to heterogeneous nuclear R N P (hnP, NP), proteins, both in vitro (Fu et al., 1992; Mayeda and Krainer, 1992; Zahler et al., 1993; Zahler and Roth, 1995) and in vivo (Cficeres et al., 1994; Wang and Manley, 1995). The techniques described in the previous sections can also be applied to this type of analysis. Figure 5 shows that changes in the concentration of U2AF, accomplished by addition of recombinant U2AF 6s to in vitro reactions, result in changes in the relative use of two alternative 3' splice sites. An increase in the use of the proximal low-affinity splice site is observed when the protein concentration

extension products obtained with each splice junction primer have the same length, the bands corresponding to the two products are pasted one above the other for convenience of comparison. In the absence of the Sex-lethal (Sxl) protein (central lane), splicing occurs only to the non-sex-specific site. The presence of Sxl represses the use of the non-sex-specific site and activates the female-specific site (left lane). The concentration of recombinant Sxl [expressed as a fusion protein with glutathione Stransferase (GST)] necessary to promote a 50% switch in the use of the alternative 3' splice sites of tra pre-mRNA was 75 ng//zl. At the same concentration, addition of a U2AF 6s derivative lacking the KS domain inhibits the use of both splice sites (fight lane). Sxl is a therefore a splicing repressor with high specificity for particular 3' splice sites. U2AF 6s, on the other hand, is a general splicing factor required for recognition of most (or all) polypyrimidine tracts. The KS domain is essential for U2AF function but not for polypyrimidine-tract recognition.

48

Juan Valcfircel et al.

Figure 5 Changes in the concentration of U2AF 6s modulate the use of alternative 3' splice sites. Splicing reactions were performed and analyzed as in Figure 4, except that the relative positions of the two alternative 3' splice sites of tra were inverted in the p r e - m R N A substrate utilized (F/N). As a result, the female-specific 3' splice site, for which U2AF has lower affinity, is located proximal to the 5' splice site, and the non-sex-specific 3' splice site, to which U2AF binds with higher affinity, is located distal to the 5' splice site. With this configuration of splice sites, the female-specific site is utilized to some extent (42%) even in the absence of Sxl regulation. The high-affinity, non-sex-specific site, however, is still preferentially used (58%) (first row). Addition of recombinant U2AF 65 to the splicing reaction results in an increase in the use of the proximal, low-affinity site at the expense of the distal, high-affinity site (third row). Thus, under conditions of higher U2AF availability, the strength of the sites becomes less relevant than their position relative to the 5' splice site. Sxl is still able to promote the use of the female-specific site at the expense of the non-sex-specific site (second row), and the effects of Sxl and U2AF 65 are synergistic in this substrate (fourth row).

is increased. This result indicates that, under conditions of equal occupancy of 3' splice sites, the distance to the 5' splice site becomes a more decisive factor in the choice between alternative 3' splice sites. The result also suggests that physiological variations in U2AF concentration have the potential to affect the relative use of competing alternative 3' splice sites in vivo.

3. Analysis of Splicing Factors

49

Perspectives Substantial progress has been made in understanding the basic pathway of the splicing reaction and the sequential interactions of snRNPs and non-snRNP polypeptides with the splice sites. Detailed information is now available on where and when specific RNAs and proteins contact the pre-mRNA. Much less is known, however, about the role of their associated activities in spliceosome function. Increasingly sophisticated assays need to be designed to explore those functions with native and recombinant factors. Also urgently needed are in vivo assays in which more reductionistic in vitro approaches can be validated or contrasted. While yeast offers a unique opportunity to perform both types of experiments, recent developments in tissue culture techniques promise to extend some of these possibilities to mammalian cells as well (Wang et al., 1996). The study of mechanisms of splicing regulation is still quite incomplete. Very few in vitro systems have been established that are able to recapitulate splicing decisions that occur during cell differentiation and development. It seems likely that key players in these decisions remain to be identified, particularly in mammalian cells.

ACKNOWLEDGMENTS We thank Young-Sun Lin for technical advice, Ravinder Singh for communicating results before publication, and members of the EMBL Gene Expression Programme for comments on the manuscript. J.V. and C.M. are supported by EMBO. M.R.G. is a Howard Hughes Investigator, and work in M.R.G.'s lab is supported by a grant from NIH.

REFERENCES Adams, M., Rudner, D., and Rio, D. (1996). Biochemistry and regulation of pre-mRNA splicing. Curt. Opin. Cell Biol. 8, 331-339. Baker, B. (1989). Sex in flies: The splice of life. Nature (London) 340, 521-524. Barabino, S., Blencowe, B., Ryder, U., Sproat, B., and Lamond, A. (1990). Targeted snRNP depletion reveals an additional role for mammalian U1 snRNP in spliceosome assembly. Cell (Cambridge, Mass.) 63, 293-302. Birney, E., Kumar, S., and Krainer, A. R. (1993). Analysis of the RNA-recognition motif and RS and RGG domains: Conservation in metazoan pre-mRNA splicing factors. Nucleic Acids Res. 21, 5803-5816. Blencowe, B., and Barabino, S. (1994). Antisense aflfinity depletion of RNP particles. Methods Mol. Biol. 37, 67-76. Boggs, R., Gregor, P., Idriss, S., Belote, J., and McKeown, M. (1987). Regulation of sexual differentiation in D. melanogastervia alternative splicing of RNA from the transformer gene. Cell (Cambridge, Mass.) 50, 739-747.

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Brosi, R., Gr6ning, K., Behrens, S.-E., Ltihrmann, R., and Kr~imer, A. (1993a). Interaction of mammalian splicing factor SF3a with U2 snRNP and relation of its 60-kD subunit to yeast PRP9. Sdence 262, 102-105. Brosi, R., Hauri, H.-P., and Kr~imer, A. R. (1993b). Separation of splicing factor SF3 into two components and purification of SF3a activity. J. Biol. Chem. 268, 17640-17646. C~ceres, J., and Krainer, A. (1993). Functional analysis ofpre-mRNA splicing factor SF2/ASF structural domains. E M B O J . 12, 4715-4726. Cficeres, J., Stature, S., Helfman, D., and Krainer, A. R. (1994). Regulation of alternative splicing in vivo by overexpression of antagonistic splicing factors. Science 265, 1706-1709. Cavaloc, Y., Popielarz, M., Fuchs, J.-p., Gattoni, R., and Stevenin, J. (1994). Characterization and cloning of the human splicing factor 9G8: A novel 35 kDa factor of the serine/arginine protein family. E M B O J . 13, 2639-2649. Chabot, B. (1996). Directing alternative splicing: Cast and scenarios. Trends Genet. 12, 472-478. Company, M., Arenas, J., and Abelson, J. (1991). Requirement of the RNA helicase-like protein PRP22 for release of messenger RNA from spliceosomes. Nature (London) 349, 487-493. Crispino, J., Blencowe, B., and Sharp, P. (1994). Complementation by SR proteins of pre-mRNA splicing reactions depleted of U1 snRNP. Science 265, 1866-1869. Delanoy, P., and Caruthers, M. (1991). Detection and characterization of a factor which rescues spliceosome assembly from a heat-inactivated HeLa cell nuclear extract. Mol. Cell. Biol. 11, 34253431. Dignam, J., Lebovitz, R., and Roeder, R. (1983). Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11, 1475-1489. Fu, X.-D. (1993). Specific commitment of different pre-mRNAs to splicing by single SR proteins. Nature (London) 365, 82-85. Fu, X.-D. (1995). The superfamily of arginine/serine-rich splicing factors. R N A 1, 663-680. Fu, X.-D., and Maniatis, T. (1990). Factor required for mammalian spliceosome assembly is localized to discrete regions in the nucleus. Nature (London) 343, 437-441. Fu, X.-D., Mayeda, A., Maniatis, T., and Krainer, A. R. (1992). General splicing factors SF2 and SC35 have equivalent activities in vitro and both affect alternative 5' and 3' splice site selection. Proc. Natl. Acad. Sci. U.S.A. 89, 11224-11228. Furneaux, H., Perkins, K., Freyer, G., Arenas, J., and Hurwitz, J. (1985). Isolation and characterization of two fractions from HeLa cells required for mRNA splicing in vitro. Proc. Natl. Acad. Sci. U.S.A. 82, 4351-4355. Ge, H., and Manley, J. (1990). A protein factor, ASF, controls cell-specific alternative splicing of SV40 early pre-mRNA in vitro. Cell (Cambridge, Mass.) 62, 25-34. Gozani, O., Patton, J., and Green, M. (1994). A novel set of spliceosome-associated proteins and the essential splicing factor PSF bind stably to pre-mRNA prior to catalytic step II of the splicing reaction. EMBO.]. 13, 3356-3367. Green, M. (1991). Biochemical mechanisms of constitutive and regulated pre-mRNA splicing. Annu. Rev. Cell Biol. 7, 559-599. Guthrie, C., and Fink, G. R. (1991). Guide to yeast genetics and molecular biology. In "Methods in Enzymology" (C. Guthrie and G. R. Fink, eds.), Vol. 194. Academic Press, San Diego, CA. Izaurralde, E., Lewis, J., McGuigan, C., Jankowska, M., Darzynkiewicz, E., and Mattaj, I. W. (1994). A nuclear cap binding protein complex involved in pre-mRNA splicing. Cell (Cambridge, Mass.) 78, 657-668. Kanaar, R., Roche, S., Beall, E., Green, M., and Rio, D. (1993). The conserved pre-mRNA splicing factor U2AF from Drosophila: Requirement for viability. Science 262, 569-573. Kanaar, R., Lee, A., Rudner, D., Wemmer, D., and Rio, D. (1995). Interactions of the Sex-lethal RNA binding domains with RNA. E M B O J . 14, 4530-4539.

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Kim, Y.-J., Zuo, P., Manley, J., and Baker, B. (1992). The Drosophila RNA-binding protein RBP1 is localized to transcriptionally active sites of chromosomes and shows a functional similarity to human splicing factor ASF/SF2. Genes Dev. 6, 2569-2579. Krainer, A. R. (1988). Pre-mRNA splicing by complementation with purified human U1, U2, U4/ U6 and U5 snRNPs. Nucleic Acids Res. 16, 9415-9431. Krainer, A. R., and Maniatis, T. (1985). Multiple factors including the small nuclear ribonucleoproteins U1 and U2 are necessary for pre-mRNA splicing in vitro. Cell (Cambridge, Mass.) 42, 725-736. Krainer, A. R., Conway, G., and Kozak, D. (1990a). Purification and characterization of pre-mRNA splicing factor SF2 from HeLa cells. Genes Dev. 4, 1158-1171. Krainer, A. R., Conway, G., and Kozak, D. (1990b). The essential pre-mRNA splicing factor SF2 influences 5' splice site selection by activating proximal sites. Cell (Cambridge, Mass.) 62, 35-42. Kr~imer, A. (1992). Purification of splicing factor SF1, a heat-stable protein that functions in the assembly of a presplicing complex. Mol. Cell. Biol. 12, 4545-4552. Kr~imer, A. (1995). The biochemistry of pre-mRNA processing. In "Pre-mRNA Processing" (A. I. Lamond, ed.), pp. 35-64. R. G. Landes Co., Austin, TX. Kr~imer, A. (1996). The structure and function of proteins involved in mammalian pre-mRNA splicing. Annu. Rev. Biochem. 65, 367-409. Kr~imer, A., and Keller, W. (1990). Preparation and fractionation of mammalian extracts active in premRNA splicing. In "Methods in Enzymology" (J. E. Dahlberg and J. N. Abelson, eds.), Vol. 181, pp. 3-19. Academic Press, San Diego, CA. Kr~imer, A., and Utans, U. (1991). Three protein factors (SF1, SF3 and U2AF) function in pre-splicing complex formation in addition to snRNPs. EMBOJ. 10, 1503-1509. Kr~imer, A., Keller, W., Appel, B., and Ltihrmann, R. (1984). The 5' terminus of the RNA moiety of U1 small nuclear ribonucleoprotein particles is required for the splicing of messenger RNA precursors. Cell (Cambridge, Mass.) 38, 299-307. Kr~imer, A., Frick, M., and Keller, W. (1987). Separation of multiple components of HeLa cell nuclear extracts required for pre-messenger RNA splicing. J. Biol. Chem. 262, 17126-17137. Larrim, G., and Lamond, A. (1993). Non-snRNP protein splicing factors. Biochim. Biophys. Acta 1173, 247-265. Lamond, A., and Sproat, B. (1993). Antisense oligonucleotides made of2'-O-alkylRNA: Their properties and applications in RNA biochemistry. FEBS Lett. 325, 123-127. Ltihrmann, R., Kastner, B., and Bach, M. (1990). Structure of spliceosomal snRNPs and their role in pre-mRNA splicing. Biochim. Biophys. Acta 1087, 265-292. MacDougall, C., Harbison, D., and Bownes, M. (1995). The developmental consequences of alternate splicing in sex determination and differentiation in Drosophila. Dev. Biol. 172, 353-376. Madhani, H., and Guthrie, C. (1994). Dynamic R N A - R N A interactions in the spliceosome. Annu. Rev. Genet. 28, 1-26. Manley, J., and Tacke, R. (1996). SR proteins and splicing control. Genes Dev. 10, 1569-1579. Mayeda, A., and Krainer, A. R. (1992). Regulation of alternative pre-mRNA splicing by hnRNP A1 and splicing factor SF2. Cell (Cambridge, Mass.) 68, 365-375. McKeown, M. (1992). Alternative mRNA splicing. Annu. Rev. Cell Biol. 8, 133-155. Moore, M., Query, C., and Sharp, P. (1993). Splicing of precursors to messenger RNAs by the spliceosome. In "The RNA World" (R. F. Gesteland and J. F. Atkins, eds.), pp. 303-357. Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. Nagoshi, R., McKeown, M., Burtis, K., Belote, J., and Baker, B. (1988). The control of alternative splicing of genes regulating sexual differentiation in D. melanogaster. Cell (Cambridge, Mass.) 53, 229-236. Norton, P. (1994). Alternative pre-mRNA splicing: Factors involved in splice site selection. J. Cell Sci. 107, 1-7.

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Padgett, R., Mount, S., Steitz, J., and Sharp, P. (1983). Splicing of messenger RNA precursors is inhibited by antisera to small nuclear ribonucleoprotein. Cell (Cambridge, Mass.) 35, 101-107. Patton, J., Mayer, S., Tempts, P., and Nadal-Ginard, B. (1991). Characterization and molecular cloning of polypyrimidine-tract binding protein: A component of a complex necessary for pre-mRNA splicing. Genes Dev. 5, 1237-1251. Patton, J., Porro, E., Galcer~n, J., Tempts, P., and Nadal-Ginard, B. (1993). Cloning and characterization of PSF, a novel pre-mRNA splicing factor. Genes Dev. 7, 393-406. Peng, X., and Mount, S. M. (1995). Genetic enhancement of RNA-processing defects by a dominant mutation in B52, the Drosophila gene for an SR protein splicing factor. Mol. Cell. Biol. 15, 6273-6282. Reed, R., and Maniatis, T. (1988). The role of mammalian branchpoint sequence in pre-mRNA splicing. Genes Dev. 2, 1268-1276. Ring, H., and Lis, J. (1994). The SR protein B52/SRp55 is essential for Drosophila development. Mot. Cell. Biol. 14, 7499-7506. Rudner, D., Kanaar, R., Breger, K., and Rio, D. (1996). Mutations in the small subunit of the Drosophila U2AF splicing factor cause lethality and developmental defects. Proc. Natl. Acad. Sci. U.S.A. 93, 10333-10337. Ruskin, B., Zamore, P., and Green, M. (1988). A factor, U2AF, is required for U2 snRNP binding and splicing complex assembly. Cell (Cambridge, Mass.) 52, 207-219. Rymond, B., and Rosbash, M. (1992). Yeast pre-mRNA splicing. In "The Molecular and Cellular Biology of the Yeast Saccharomyces" O. Broach, J. Pringle, and E. Jones, eds.), Vol. 2, pp. 143-192. Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. Sawa, H., and Shimura, Y. (1991). Alteration ofRNase H sensitivity of the 3' splice site region during the in vitro splicing reaction. Nucleic Acids Res. 19, 3953-3958. Schwer, B., and Guthrie, C. (1991). PRP16 is an RNA-dependent ATPase that interacts transiently with the spliceosome. Nature (London) 349, 494-499. Singh, R., Valc~rcel, J., and Green, M. R. (1995). Distinct binding specificities and functions of higher eukaryotic polypyrimidine tract-binding proteins. Science 268, 1173-1176. Smith, C., Patton, J., and Nadal-Ginard, B. (1989). Alternative splicing in the control ofgene expression. Annu. Rev. Genet. 23, 527-577. Sosnowski, B. A., Belote, J. M., and McKeown, M. (1989). Sex-specific alternative splicing of RNA from the transformer gene results from sequence-dependent splice site blockage. Cell (Cambridge, Mass.) 58, 449-459. Temsamani, J., Agrawal, S., and Pederson, T. (1991). Biotinylated antisense methylphosphonate oligodeoxynucleotides. J. Biol. Chem. 266, 468-472. Tian, M., and Maniatis, T. (1993). A splicing enhancer complex controls alternative splicing ofdoublesex pre-mRNA. Cell (Cambridge, Mass.) 74, 105-114. Valc~rcel, J., and Green, M. (1996). The SR protein family: Pleiotropic effects in pre-mRNA splicing. Trends Biochem. Sci. 21, 296-301. Valc~rcel, J., Singh, R., Zamore, P., and Green, M. (1993). The protein Sex-lethal antagonizes the splicing factor U2AF to regulate alternative splicing of transformer pre-nflkNA. Nature (London) 362, 171-175. Valcfircel, J., Singh, R., and Green, M. (1995). Mechanisms of pre-mRNA splicing regulation. In "Pre-mRNA Processing" (A. I. Lamond, ed.), pp. 97-112. R. G. Landes Co., Austin, TX. Valcfircel, J., Gaur, R., Singh, R., and Green, M. (1996). Interaction ofU2AF 6s RS region with premRNA branch point and promotion of base pairing with U2 snRNA. Science 273, 1706-1709. Wang, J., and Manley, J. (1995). Overexpression of the SR proteins ASF/SF2 and SC35 influences alternative splicing in vivo in diverse ways. RNA 1, 335-346. Wang, J., Takagaki, Y., and Manley, J. (1996). Targeted disruption of an essential vertebrate gene: ASF/SF2 is required for cell viability. Genes Dev. 10, 2588-2599.

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Winkelmann, G., Bach, M., and Ltihrmann, R. (1989). Evidence from complementation assays in vitro that U5 snRNP is required for both steps of mRNA splicing. E M B O J . 8, 3105-3112. Zahler, A., and Roth, M. (1995). Distinct functions of SR proteins in recruitment of U1 small nuclear ribonucleoprotein to alternative 5' splice sites. Proc. Natl. Acad. Sci. U.S.A. 92, 2642-2646. Zahler, A., Stolk, J., Lane, W., and Roth, M. (1992). SR proteins: A conserved family of pre-mRNA splicing factors. Genes Dev. 6, 837-847. Zahler, A., Neugebauer, K., Stolk, J., and Roth, M. (1993). Distinct functions of SR proteins in alternative pre-mRNA splicing. Science 260, 219-222. Zamore, P., and Green, M. (1989). Identification, purification and biochemical characterization of U2 small nuclear ribonucleoprotein auxiliary factor. Proc. Natl. Acad. Sci. U.S.A. 86, 9243-9247. Zamore, P., and Green, M. (1991). Biochemical characterization of U2 snRNP auxiliary factor: An essential pre-mRNA splicing factor with a novel intranuclear distribution. EMBOJ. 10, 207-214. Zamore, P., Patton, J., and Green, M. (1992). Cloning and domain structure of the mammalian splicing factor U2AF. Nature (London) 355, 609-614. Zhang, M., Zamore, P., Carmo-Fonseca, M., Lamond, A., and Green, M. (1992). Cloning and intracellular localization of the U2 small nuclear ribonucleoprotein auxiliary factor small subunit. Proc. Natl. Acad. Sci. U.S.A. 89, 8769-8773. Zuo, P., and Maniatis, T. (1996). The splicing factor U2AF 35mediates critical protein-protein interactions in constitutive and enhancer-dependent splicing. Genes Dev. 10, 1356-1368. Zuo, P., and Manley, J. (1993). Functional domains of the human splicing factor ASF/SF2. EMBO J. 12, 4727-4737.

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Identification of pre-mlKNA Splicing Factors and Analysis of RNA-Protein Interaction James G. Patton Department of Molecular Biology Vanderbilt University Nashville, Tennessee 37235

Billy T. Dye Department of Molecular Biology Vanderbilt University Nashville, Tennessee 37235

D a r o n C. B a r n a r d

James G. McAfee Department of Molecular Biology Vanderbilt University Nashville, Tennessee 37235

Department of Molecular Biology Vanderbilt University Nashville, Tennessee 37235

I. I N T R O D U C T I O N Pre-mRNA splicing occurs in a large particle termed the spliceosome, composed of multiple protein and protein-lKNA complexes (reviewed in 2, 22, 31). Unlike the situation of self-splicing introns, it seems likely that metazoan intron excision and exon ligation are 1KNA-catalyzed events (7, 28, 34, 52), probably involving the U family of small nuclear ribonucleoproteins (snRNPs; 26, 49). However, protein factors are clearly required for metazoan pre-mRNA splicing (3, 23, 24). Indeed, the size of the spliceosome (40-60S) underscores the multiplicity of its components and the complexity of the splicing process. Genetic systems have allowed for the identification of many proteins involved in pre-mRNA splicing (3). For mammalian splicing, cell-free systems have dramatically increased our understanding of the splicing reaction and allowed for biochemical purification of some of these factors. This chapter will deal with relevant methods that can be used to identify 1KNA binding proteins and analyze RNA-protein interactions. Most of the accompanying illustrative experiments will focus on proteins involved in mammalian pre-mRNA splicing and are presented primarily as examples of the various techniques that can be applied to many 1KNA-protein interactions. 55 rrtRNA Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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II. P R E - m R N A S P L I C I N G A N D T H E R O L E O F T H E POLYPYRIMIDINE TRACT A. MATERIALS AND METHODS 1. In Vitro Transcription

Template DNAs (47) were linearized with appropriate enzymes, and 1 /xg of DNA was transcribed in the presence of 250 /xM 7mG(5')ppp(5')G cap structure (New England Biolabs, Beverly, MA) in 1• Transcription Buffer (Promega, Madison, WI) containing 10 m M dithiothreitol (DTT), 500 /xM ATP, 500 /xM UTP, 125 /xM GTP, 125 /xM CTP, 40 U RNasin (Promega), 20 /xCi [c~-32p]CTP, and 20 U SP6 ILNA polymerase. The total volume was 20/xl, and the reactions were incubated at 37~ for 1 hr. Following transcription, the volume of each reaction was brought up to 200/xl with water for extraction with phenol, phenol/chloroform, chloroform/isoamyl alcohol (24: 1), and ethanol precipitation. Prior to phenol extraction, 1 /xl of each diluted reaction was removed and spotted onto a glass fiber filter, and 1 /xl of each reaction was precipitated with 10% trichloroacetic acid (TCA) in the presence of 100 /xg yeast tRNA on ice for 20 min. TCA-precipitated tLNAs were then trapped on glass fiber filters, and the incorporated 32p was analyzed by scintillation counting to determine the R N A yield.

2. In Vitro Splicing In v i t w splicing was carried out as described previously (35). Briefly, 5 ng of 32p-labeled pre-mtLNA was incubated with 15/xl of nuclear extract in 500/xM ATP, 20 m M phosphocreatine, 500/xM MgC12, 500/xM DTT, and 40 U RNasin in a 25-/xl reaction volume at 30~ for either 0 or 2 hr. Nuclear extracts were prepared as described (1) and dialyzed against Buffer E [20 m M Tris pH 8.0, 20% glycerol, 0.1 m KC1, 0.2 m M ethylenediaminetetraacetic acid (EDTA), and 0.5 m M DTT]. Reactions were stopped by the addition of 50/xg proteinase K and incubation at 65~ for 5 min in 100 m M Tris (pH 7.8), 12.5 m M EDTA, 150 m M NaC1, and 1% sodium dodecyl sulfate (SDS). RNAs were extracted as described above, and the resulting pellets were washed for 5 min in 70% ethyl alcohol (EtOH). The pellets were then dried and resuspended in 8/xl formamide containing 1 • Tris-borate-EDTA (TBE), 10% glycerol, and 0.01% Bromophenol Blue and xylene cyanol, followed by heating at 65~ for 10 min before electrophoresis. Denaturing polyacrylamide gels (8%) were prepared with a 19:1 ratio of acrylamide to bisacrylamide in the presence of 8 M urea. The gels were

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fixed in 10% TCA, 10% acetic acid, and 30% methanol, dried, and visualized using a PhosphorImager.

B. RESULTS AND SUMMARY The development of cell-free systems has allowed tremendous progress in understanding the mechanisms of mammalian splicing and in identifying the factors involved (23, 31). Pre-mR.NA splicing occurs via two transesterification reactions and can be carried out in HeLa cell nuclear extracts using in vitro prepared pren ~ N A substrates. As an example of how crucial cis-acting elements within the pre-mR.NA can be identified with the use of such assays, we prepared a series of pre-mR.NAs containing variable-length polypyrimidine tracts. The parental substrate, p G 3 + D X , is derived from exons 2 and 3 of oe-tropomyosin (oe-TM; 45) and contains an unusually long polypyrimidine tract adjacent to an atypical upstream branch point sequence. Normally, exons 2 and 3 of c~-TM are mutually exclusive due to the fact that the branch point between them is too close to the 5' splice site of exon 2, precluding effective interaction between U1 snRNP at the 5' splice site and U2 snP, NP at the branch point (45). However, the two exons can be spliced to one another provided that spacer sequences are added between the 5' splice site and the branch point. In Figure 1, two different spacer sequences (DX and DS) were utilized to show that the length of the resulting lariat structure dramatically changes the mobility of the lariat-exon intermediate and the lariat product on denaturing polyacrylamide gels. The effect of the polypyrimidine tract on splicing is also clearly shown in Figure 1 by comparing the splicing efficiency of a series of substrates containing decreasing polypyrimidine tract lengths. Decreasing the tract from its original 51 nucleotides to 39, 23, or 17 nucleotides has relatively little effect on splicing efficiency. In contrast, decreasing the tract to nine pyrimidines inhibits splicing, while a tract containing a single pyrimidine is unable to stimulate recognition of the adjacent branch point, leading to complete abolition of splicing (Fig. 1). These results demonstrate the importance of the pyrimidine tract in pre-mRNA splicing. Several P,.NA binding proteins have been identified that interact with the pyrimidine tract (10, 11, 40, 50, 51). The ability to use in vitro splicing reactions and mutated pre-mR.NA substrates allows identification of important cis-acting P,.NA sequences, which in turn allows identification of proteins that bind to these sequences and mediate or regulate pre-mR.NA splicing. Such analyses have enabled identification of important cis-acting sequences at the 5' and 3' splice sites, branch point sequences, exon sequences, and other intron sequences (15, 23, 31, 46). Such mutations generally affect the binding of trans-acting factors that can then be identified by various biochemical approaches. One technique that is commonly used to identify R N A binding proteins is ultraviolet (UV) cross-linking.

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III. A N A L Y S I S O F R N A B I N D I N G P R O T E I N S BY UV CROSS-LINKING A. MATERIALS AND METHODS A 135-nucleotide R N A encompassing the branch point/polypyrimidine tract from intron 2 of c~-TM (B3P3) was generated by in vitro transcription (as described above) of a subclone from pGC + DX. The 32p-labeled R N A (10 ng) was incubated with proteins from an HeLa cell nuclear extract or with recombinant proteins at 4~ for 15 min in a buffer containing 2 m M MgC12, 0.5 m M DTT, 5 U RNasin, and 60% protein solutions (v/v) of variable concentrations (dialyzed as described above). Following the binding reaction, the mixtures were transferred to microtiter dishes and placed in a foil-lined box under a UV light source (7-cm distance) emitting at 254 nm. Typically, a standard transilluminator was used as the UV light source by inversion over the foil-lined box. Irradiation was performed for 5-10 min at room temperature, after which the reactions were transferred back to their original tubes and the RNAs were digested by addition of 20 U micrococcal nuclease, 0.125 mg/ml RNase A, and a final concentration of 1.3 m M CaC12. RNase digestion was allowed to proceed for 1 hr at 37~ Proteins containing covalently linked 32p-RNA (primarily single nucleotides) were analyzed on 9% SDS Laemmli gels, fixed in 10% TCA, 10% HOAc, and 30% methyl alcohol (MeOH), dried, and exposed to film or Phosphorlmager screens. Recombinant polypyrimidine tract binding protein (PTB) and U2 auxiliary factor (U2AF) were prepared as follows. An EcoRI fragment containing the fulllength PTB complementary D N A (cDNA) was cloned into pET28A (Novagen, Madison, WI), resulting in the fusion of 34 amino acids at the amino terminus of PTB, including a hexahistidine tag that allows purification of the resultant fusion protein, pETPSF was constructed as described (36). Overnight cultures were diluted 1:10 in LB medium and grown at 37~ until a n OD600 of 0.5 was attained. Isopropyl-/3-thiogalactopyranoside (IPTG) was then added to a final concentration

Figure 1 The effect of the polypyrimidine tract on the efficiency of in vitro pre-mRNA splicing. In vitro splicing reactions were performed using HeLa nuclear extract for the indicated time. Two different pre-mRNA substrates were used that contained introns of different length due to the insertion of either the DS fragment or the DX fragment into the intron between exons 2 and 3 of cz-TM. The length of the intron affects the mobility of the lariat-exon intermediate and the lariat product on denaturing polyacrylamide gels, whereas the size of the m R N A between the two substrates remains the same. To demonstrate the effect of decreasing polypyrimidine tract strength on pre-mRNA splicing, successive deletions (as depicted below) of the tract in the DX substrate were spliced for 2 hr and analyzed as shown. Precursors, intermediates, and the products of splicing are as indicated.

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of i raM, and growth was continued for an additional 2 hr at 37~ for PTB and 7 hr at 37~ for PSF. The proteins were purified under nondenaturing conditions with Ni-NTA resin (Qiagen, Chatsworth, CA). Following growth, bacterial cells were resuspended in three volumes of Sonication Buffer [SB; 50 mM NaPO3 pH 8.0, 300 mM NaC12, 5% glycerol, 1 m M phenylmethylsulfonyl fluoride (PMSF)], digested with 5 mg/ml lysozyme, sonicated, and applied to a 2-ml Ni-NTA column. Following extensive washing with SB containing 20 mM imidazole, the proteins were eluted with a 0-0.5 M imidazole SB gradient, and peak fractions were dialyzed into Buffer A (20 mM Tris-C1 pH 8.0, 150 mM KC1, 0.5 m M DTT, 0.2 mM EDTA, 5% glycerol, 1 mM PMSF).

B. RESULTS AND SUMMARY UV cross-linking is a potentially valuable tool in analyzing p r o t e i n - R N A interaction. Figure 1 demonstrates the importance of the pyrimidine tract in prem R N A splicing. In order to identify trans-acting factors that recognize the pyrimidine tract, a short R N A was prepared that consists of just the region surrounding the wild-type branch point/polypyrimidine tract (B3P3) upstream of exon 3 in ot-TM. As shown in Figure 2, incubation of the B3P3 I:(NA probe in HeLa cell nuclear extract leads to label transfer from the tkNA to a single 57-kDa protein, PTB (10, 35). The efficiency and specificity of the PTB-pyrimidine tract interaction allowed purification and cloning of PTB, with the use of cross-linking to monitor the purification (12, 35). With the use of a biochemical complementation strategy, the essential splicing factor U2AF (40, 54) was identified and also found to bind specifically to the polypyrimidine tract. Indeed, U2AF binds B3P3 with higher affinity than does PTB (55). Subsequently, PTB and U2AF were found to compete for binding to pyrimidine tracts such that PTB could repress splicing by preventing the binding of U2AF (25, 37, 44). To illustrate the use of UV crosslinking to analyze protein-RNA interaction, the competition between U2AF and PTB was demonstrated by cross-linking a set amount of recombinant PTB with increasing amounts of recombinant U2AF 65, the large subunit of U2AF. As shown in Figure 2, PTB is efficiently cross-linked both in HeLa cell nuclear extracts and in recombinant form. In whole nuclear extract, the levels of U2AF are not sufficient to prevent PTB cross-linking or to allow detectable U2AF cross-linking. In contrast, recombinant U2AF 65 can be cross-linked to R N A under these conditions, but the efficiency of cross-linking is much lower than that of PTB, further accounting for the fact that U2AF 65 cross-linking in nuclear extract is not observed. When recombinant PTB and U2AF 65 are combined, U2AF 65 is able to prevent PTB cross-linking, even at slightly lower concentrations than those of PTB, suggesting that U2AF 6s binds to the polypyrimidine tract with higher affinity.

F i g u r e 2 Competitive R N A binding between U2AF and PTB. A 32p-labeled, 135-nt R N A encompassing the branch point/ polypyrimidine tract upstream of exon 3 of oe-TM was incubated with HeLa nuclear extract (right panel) or with recombinant U2AF 65 or recombinant PTB (center panel) and subjected to UV cross-linking. Following digestion of the R N A , 32p-labeled proteins were separated on 9% SDS gels. In nuclear extract, PTB is eflqciently cross-linked and no U2AF is detectable. However, crosslinking with recombinant proteins shows that PTB and U2AF compete for R N A binding. The input recombinant proteins used in the center panel are shown on the right after Coomassie Blue staining.

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Several caveats to UV cross-linking studies are in order. First, the lack of a UV cross-linking signal does not necessarily imply a lack of binding for a given protein. UV cross-linking is dependent on an appropriate residue located an optimal distance from the labeled nucleotide, and even remarkably high-affinity binding proteins may not bind in a manner that promotes cross-linking (16). Second, UV cross-linking is inherently nonequilibrium, and since the R N A - p r o t e i n interaction is covalent, the reaction is driven toward binding. Thus, nonspecific and lowaffinity binding is often detectable, necessitating the addition of cold competitor RNAs in order to determine binding specificity (33, 54). Last, the labeled nucleotide can dramatically alter cross-linking results such that independent experiments must often be performed with RNAs labeled with different nucleotides. Despite these caveats, UV cross-linking provides a rapid, easy way to identify p r o t e i n - R N A interactions. In the samples shown in Figure 2, cross-linking was performed with the use of unmodified proteins, radiolabeled RNAs, and irradiation at 254 nm. Cross-linking can also be performed with longer-wavelength light and with modified proteins or RNAs or in the presence of bifunctional cross-linking reagents (16). Depending on the conditions used, different proteins can potentially be identified. With recently described techniques allowing the ability to label RNAs at specific positions (32), identification of proteins targeted to distinct R N A sequences by UV cross-linking is an even more attractive technique. Thus, UV cross-linking, when cautiously interpreted, can be an effective technique to identify proteins involved in specific R N A - p r o t e i n interactions.

IV. A N A L Y S I S O F P R O T E I N - R N A INTERACTION FLUORESCENCE SPECTROSCOPY

BY

A. MATERIALS AND METHODS The fluorescent substrate polyethenoadenosine (poly[r(sA)]) was generated by treatment of poly(A) R N A with chloracetaldehyde (48). High-pressure liquid chromatography (HPLC)-purified oligoribonucleotides were purchased from Cruachem and used without further purification. The extinction coefficients (8260) for each oligoribonucleotide were calculated with the use of the nearest-neighbor approximation method (6). The names, sequences, and 8260 values for each of the oligoribonucleotides used in these studies are as follows: P3: PTB winner: U2AF winner:

rCUUUCUCUUUCUCUCUCCCU rUCUUCUCUUCUCUUCUCUUC rUUUUUUCCCCUUUUUUUUCC

(8260 = (8260 = (8260 - -

8340 M-1 cm-*) 8505 M -1 cm-1) 8895 M -i cm-1)

Fluorescence measurements were carried out using an SLM Aminco Bowman AB-2 Luminescence spectrometer. The excitation and emission slits were set at

4. RNA-Protein Interaction in Splicing

63

4 and 8 nm, respectively. Binding experiments with PTB were conducted in 10 m M Tris (pH 8.0), 1 m M EDTA, and 100 m M NaC1. Fluorescence titrations were conducted by exciting a solution of poly[r(sA)] at 310 nm and measuring the change in fluorescence emission as a function of increasing PTB concentration. The fluorescence emission was corrected for background levels and dilution due to the addition of protein sample. Dilution of the poly[r(eA)] solution with protein sample did not exceed 2.5% of the initial volume. The fluorescence enhancement (E) was calculated with the following equation:

E=

F - Fi Fi

where F is the fluorescence emission observed at each titration point and F~ is the initial fluorescence emission observed in the absence of protein. Quantitative binding parameters for the interaction of PTB with poly[r(eA)] (control titrations) were determined by simulating theoretical curves to fit the experimental data using the McGhee-von Hippel noncooperative binding equation (29). Competition titrations were used to determine the relative equilibrium constants (Kcomp)for the interaction of PTB with the above oligoribonucleotides. In these experiments, the fluorescence emission of a solution containing 1.6/zM of competitor oligoribonucleotide and 0 . 8 / z M of poly[r(sA)] was monitored as a function of increasing PTB concentration. An expression for Kcomp can be derived by considering the following equilibria: PTB + eA <--) P T B - e A and PTB + comp ~ PTB-comp where PTB, eA, and comp are the steady-state concentrations ofPTB, poly[r(eA)], and the competitor RNA, respectively. P T B - s A and PTB-comp are the equilibrium concentrations of the P T B - e A and PTB-competitor complexes, respectively. Based on the above equilibria, the mass action expression defining Kcompand KeA yields the following equations: [PTB-eA] K~gA = ([PTB]f~ee)([eA]f~ee) and

Kcomp

[PTB-comp] ([PTB]free) ([comp]free)

In the competition titrations, [PTB]free for both mass action expressions is the same. As a result, both expressions can be equated through [PTB]free. The solution for Kcompthen becomes

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et al.

( [ P T B - c o m p ] ) (KsA)([8A] free) Kcomp =

( [ P T B - e A ] ) ([comp]free)

T h e concentration o f P T B b o u n d to the c o m p e t i t o r ( P T B - c o m p ) can be calculated f r o m the difference in the c o n c e n t r a t i o n o f P T B required to p r o d u c e the same E observed in the presence and absence o f competitor. Since all extinction coefficients are reported in nucleotides, this value is multiplied by the binding site size to obtain the concentration o f c o m p e t i t o r nucleotides b o u n d . Assuming that a linear relationship exists b e t w e e n the fractional change in E and the fractional saturation o f poly[r(eA)], [cA]free can be d e t e r m i n e d as follows:

[sA]free = [sA]total- ['~A]bound or [~a]ffee = [~A]total- (E/Emax)([~A]total) w h e r e [•A]total and [eA]bound are the total and b o u n d concentrations o f poly[r(eA)] in nucleotides, respectively. These equations can be rearranged to calculate the c o n c e n t r a t i o n o f [CA]bound in nucleotides. Kcomp determinations for each c o m p e t i t o r oligoribonucleotide were calculated at five different e n h a n c e m e n t values. T h e average value o f Kcomp, along w i t h the standard deviation, is as s h o w n in Figure 3B. Fluorescence experiments w i t h U 2 A F w e r e c o n d u c t e d on the same fluorescence i n s t r u m e n t in buffer containing 20 m M Tris p H 7.9, 200 m M NaC1, and 0.2 m M E D T A . H o w e v e r , because U 2 A F contains a single tryptophan, titrations w e r e carried out by measuring the decrease in the intrinsic t r y p t o p h a n fluorescence o f U 2 A F as a function o f increasing c o n c e n t r a t i o n o f different oligonucleotides. Fluorescence measurements were corrected for b a c k g r o u n d and scatter due to the addition o f nucleic acid. T h e fluorescence emission (F) o f the protein at any p o i n t t h r o u g h o u t the titration was d e t e r m i n e d by integration o f the area o f an emission scan from 345 to 370 n m . Fluorescence q u e n c h i n g (Q) was calculated as follows:

Figure 3 Equilibrium analysis of PTB and U2AF binding to RNA. The relative and intrinsic equilibrium constants for the interaction of recombinant PTB and U2AF with RNAs containing either the in vitro selected PTB-RBS, the U2AF-RBS, or a portion of the polypyrimidine tract upstream of exon 3 in c~-TM (P3) were determined with the use of fluoresence spectroscopy. (A, B) The binding isotherms for the binding of recombinant PTB to poly[r(eA)] alone (solid line) or in the presence of each of the three RNA competitors (PTB-RBS, P3, and U2AF-RBS) are as shown. The relative association c o n s t a n t s (Kcomp)are shown in (B), along with standard deviations derived from calculations performed at multiple points along the binding isotherms. Panels (C) to (E) are double reciprocal plots of binding data for the interaction of recombinant U2AF65 with the same RNAs used in panels (A) and (B). The experimental data were fit by linear regression analysis with correlation coefficients greater than 0.97. The slope of each line defines the KD and is displayed as an inset within each plot.

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Q- F-F~ F The fractional saturation of binding sites on the protein by oligonucleotide is defined by the ratio of Q to the maximum quenching observed when all sites are saturated (Qm~x). Thus, the ratio Q/Qmax defines the binding density, v (fractional saturation of binding sites on the protein). The mass action expression for the interaction of U2AF with oligonucleotide, assuming the simplest model of a single binding site on the protein, then becomes

K~=

[oligo]~ee [W2AF]~ee [U2AF-oligo]

where KD is the dissociation constant. The numerator in the above equation is the product of the free concentration of oligoribonucleotide and U2AF, and the denominator is the concentration of bound complex. Since Q and Qmaxare empirically determined, an expression where these values are related to the equilibrium constant needs to be defined. The fractional saturation of sites can be calculated as follows: Q Qm~x

[W2AF-oligo] _ [U2AF]total

[U2AF-oligo] [W2AF-oligo] + [U2aF]~ee

By solving the mass action expression for [U2AF-oligo] and substituting this into the above equation, the relationship between v and the KD becomes [oligo]~ee KD + [oligo]free Taking the reciprocal of both sides of this equation results in a linear equation with a slope equal to the KD. KD values were evaluated from double reciprocal ("Benesi-Hildebrand") plots of 1 / v versus 1/[oligo] and determined by linear regression analysis of the data using Kaleidagraph software (v2.13).

B. RESULTS AND SUMMARY

RNA-protein interactions can be evaluated with the use of a number of methods, such as fluorescence, UV absorption, circular dichroism, and nuclear magnetic resonance (NMR) spectroscopy. The only requirement is that the spectroscopic signal, which may be either on the R N A or on the protein, is changed on formation of the RNA-protein complex. The advantage of these methods is their ability to indirectly monitor the steady-state concentrations of the free and bound ligand. In nonequilibrium methods such as filter binding, band shift assays, or UV cross-linking, free and bound reactants are measured after perturbation of

4. RNA-Protein Interaction in Splicing

67

equilibrium. The study of protein-nucleic acid interactions with fluorescence spectroscopy typically involves monitoring the intrinsic tryptophan fluorescence derived from the protein of interest. However, because PTB does not contain tryptophan, its interaction with the above oligoribonucleotides was determined by analysis of competition titration data with a fluorescently labeled substrate RNA. The binding of PTB to poly[r(sA)] results in enhanced fluorescence. The affinity of a given oligoribonucleotide was then determined based on its ability to inhibit the PTB-induced fluorescence enhancement of the poly[r(sA)]. Shown in Figure 3A are binding isotherms for the interaction of PTB with poly[r(sA)] alone, or in the presence of RNAs containing the optimal PTB binding site, P3 (the pyrimidine portion of B3P3; see above), or the optimal U2AF binding site. The optimal RNA-binding sequences for PTB and U2AF were determined with the use of iterative genetic selection techniques (37, 44) and will be referred to as the PTB R N A binding site (PTB-RBS) or the U2AF R N A binding site (U2AF-RBS). Visual inspection of the data shown in Figure 3A indicates that the PTB-RBS and P3 R N A inhibit the binding of PTB to poly[r(sA)] far better than does the U2AFRBS. This is more quantitatively represented by the bar graph in Figure 3B, where the average Kcomp value calculated from five different E values is shown. The relative affinity of PTB for its selected R N A sequence is approximately 2 times that for binding to P3 R N A and 12 times that for its interaction with the U2AFRBS. Though we have analyzed these data to determine relative affinities, it is important to note that the experiment can be designed to determine intrinsic affinities for the interaction of PTB with each oligo with the use of more rigorous analytical approaches. In contrast to the forward titrations involving the binding of PTB to poly[r(sA)], U2AF binding to R N A can be monitored directly as the protein binds R N A by measuring the change in fluorescence of a single tryptophan residue within U2AF. Shown in Figure 3 C - E are double reciprocal plots of binding data for the interaction of U2AF with RNAs containing the PTB-Rt3S, P3, or the U2AF-RBS. In these experiments, because the fluorescent signal is an intrinsic property of the protein, there is a direct correlation between the fractional change in the signal and the fractional saturation of binding sites on the protein by the oligoribonucleotide. If the number of oligoribonucleotide binding sites on the protein is known, intrinsic equilibrium constants can be determined. In our analysis of the data shown in Figure 3C-E, we have assumed a one-to-one interaction for the binding of U2AF to the oligoribonucleotides. Linear regression analysis of these data yielded correlation coefficients exceeding 0.98 for each of the data sets, indicating that the one-site model gives a good fit to the experimental data. An analysis of the data shown in Figure 3 C - E indicates that U2AF binds its selected R N A sequence and the PTB-RBS with essentially equal affinity. The interaction of U2AF with P3 R N A is only slightly lower than that observed for the specific U2AF- or PTB-selected sequences. It seems clear that the preferred binding sites

68

James G. Patton et

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for PTB and U2AF overlap. This finding agrees with published data derived from nonequilibrium techniques (25, 37, 44) and the UV cross-linking experiments shown in Figure 2. However, the cross-linking data (Fig. 2) suggest that U2AF binds with higher affinity to 1KNA than does PTB, whereas the fluorescence data suggest that U2AF and PTB bind with relatively similar affinities. Since the nuclear concentration of PTB is much higher than that of U2AF, it is suggested that if the two proteins bind with relatively similar affinity to overlapping sites, additional factors must play a role to enable U2AF binding and pre-mRNA splicing. Consistent with this idea, several such candidate factors have been proposed (17, 27, 53). The use of fluorescence spectroscopy to monitor the interaction of PTB with the oligoribonucleotides described here has allowed us to determine the relative affinities for the interaction of the protein with these substrates under equilibrium conditions. The relative difference in the affinities between oligoribonucleotide substrates directly reflects the difference in their intrinsic affinities. In contrast, the analysis of the U2AF binding data has not only yielded true equilibrium dissociation constants, but the ability to fit the data to a binding model has revealed the stoichiometry of the interaction. These observations demonstrate that the sensitivity of spectroscopic approaches like fluorescence can yield data with a high degree of precision for the determination of affinities and also for information about the stoichiometry of the interaction.

V. A N A L Y S I S O F P R E - m R N A S P L I C I N G C O M P L E X E S AND IDENTIFICATION OF ASSOCIATED FACTORS BY UV CROSS-LINKING A. MATERIALS AND METHODS 1. In Vitro Splicing Reactions and E Complex Assembly

For analysis of spliceosomes by gel filtration, large-scale splicing reactions (750 /.~1) were performed containing 0.5 m M ATP, 20 m M phosphocreatine, 2 m M MgC12, 0.5 m M DTT, 1 ng/~l capped, uniformly 32p-labeled p r e - m R N A substrate (transcribed as above), 40% HeLa nuclear extract in buffer E (20 rnM Tris pH 7.9, 100 m M KC1, 0.2 m M EDTA, 0.5 m M DTT, 20% glycerol), and additional buffer E to obtain a final concentration of 60 m M KC1. E complex assembly reactions were performed under identical conditions, except that ATP, phosphocreatine, and MgC12 were omitted and the nuclear extract was preincubated at 30~ for 30 man to deplete any endogenous ATP. Reactions were incubated at 30~ for either 45 man (splicing) or 40 min (E complex) immediately prior to gel filtration.

4. RNA-Protein Interaction in Splicing

69

2. Gel Filtration of Splicing Complexes A 500-/xl aliquot of either a splicing reaction or an E complex assembly reaction was loaded onto a 1.6 • 50-cm Sephacryl S-500 gel filtration column equilibrated and run in 20 mM Tris (pH 7.8), 0.1% Triton X-100, 60 ~ KC1, and 2.5 mM EDTA. Fractions (1 ml) were collected at a flow rate of 6 ml/hr, and 200-/xl aliquots from each fraction were quantitated by liquid scintillation counting. R N A from 200-/xl aliquots of the indicated fractions was isolated by phenol extraction and ethanol precipitation, separated by electrophoresis on denaturing 8% polyacrylamide:bisacrylamide gels, and visualized with a PhosphorImager.

3. UV Cross-Linking UV cross-linking was performed by irradiating aliquots (400 /xl) of the indicated gel filtration fractions for 10 min with 254 nm light at a distance of 7 cm from the source. To ensure adequate irradiation, the samples were divided into 40-/xl aliquots before cross-linking. The aliquots were then combined, 1KNase A digested (0.125 mg/ml for 1 hr at 37~ precipitated with 25% TCA, and resuspended in a volume suitable for SDS gel electrophoresis on 9% Laemmli gels.

B. RESULTS AND SUMMARY Spliceosomes form in a stepwise, ATP-dependent fashion (5, 38, 39, 42). Prespliceosomes and catalytic spliceosomes can be identified by nondenaturing gel electrophoresis or by gel filtration chromatography (20, 21, 30, 42). The advantage of chromatographic separation is twofold. First, the earliest prespliceosome complex (E complex) can only be identified by gel filtration, as it apparently dissociates upon electrophoresis (30). Second, the various complexes can be enriched or biochemically purified, allowing analysis of the associated components (4, 8, 14). To illustrate the utility of this method in identifying specific spliceosome components, we performed in vitro splicing reactions and then subjected these reactions to gel filtration chromatography. As shown in Figure 4A, pre-mRNAs incubated with nuclear extract in the absence of ATP form a predominant complex referred to as the H complex due to association with the pre-mRNA of abundant R N A binding proteins called hnRNP proteins (heterogeneous ribonucleoproteins; 9). However, a small shoulder that elutes slightly ahead of the H complex constitutes the E complex (Fig. 4A). Under appropriate conditions, the ratio of H to E complexes can be altered to generate increased amounts of E complex (30). The

70

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Figure 4 Chromatographic separation of splicing complexes and UV cross-linking. Separation of splicing complexes by gel filtration of in vitro splicing reactions performed in the presence (A) or absence (D) of ATP. Peaks containing the H, E, A, and B complexes are as indicated, along with the void volume (V0) and total volume (V~) of the column. RNAs recovered from each of the complexes were isolated by extraction of the appropriate fractions (as indicated) and separated on 9% denaturing gels (B, E). The pre-mRNA substrate, splicing intermediates, and spliced products are as indicated. UV cross-linking of proteins to the pre-mtkNA substrate was performed on the indicated fractions, and the 32p-labeled proteins were separated on 9% SDS gels (C, F). Molecular weight markers are as shown (M).

71

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E complex forms in the absence of ATP and is a stable complex representing the commitment of a given pre-mRNA to the splicing pathway (18, 19, 30, 42). As shown in Figure 4B, no splicing intermediates are detected for either the H or the E complex, as expected for prespliceosome complexes. To identify proteins that are part of either complex, we isolated both complexes and subjected such fractions to UV cross-linking using the same conditions as in Figure 2. For the purposes of this chapter, we have not attempted to specifically identify any of

72

James G. Patton et al.

the labeled proteins; rather, the objective is to illustrate the methodology. The experiment shown in Figure 4C used a continuously labeled pre-mRNA substrate, and distinct cross-linking species are detectable for both complexes, particularly the H complex. Such cross-linking analysis can also be performed with substrates labeled at individual nucleotides, affording analysis of protein-RNA interaction at a specific site rather than the global approach shown here (8). In addition, specifically labeled substrates can decrease the levels of the more efficiently crosslinked hnRNP proteins and increase the signal-to-noise ratio. It should be stressed that the number of proteins seen in Figure 4C is an underestimate due, in part, to some of the previously mentioned difficulties with UV cross-linking and also to degradation of certain proteins on irradiation with UV light, particularly 254-nm wavelengths (16). Nevertheless, distinct proteins can be identified within each complex. When splicing reactions are allowed to proceed in the presence of ATP, additional prespliceosome complexes (A and B complexes) and catalytic spliceosomes (C complexes) assemble and can be identified by gel filtration chromatography (Fig. 4D). The A and B complexes are prespliceosomes, whereas the C complex is short-lived and contains catalytic spliceosomes that have undergone the first step of splicing (14). When we subjected pre-mRNA substrates to such analysis, a dramatic decrease in H complex was observed as the A, B and C complexes assembled (Fig. 4D). Without the use of mutant substrates or of heat- or EDTAtreated extracts (14), it is virtually impossible to isolate C complexes, so the labeled peak contains mostly A and B complexes. Following catalysis, C complexes rapidly dissociate such that the amount of detectable C complex is quite low. Examination of the RNAs from these peaks shows that splicing products and intermediates accumulate in the peaks containing the A, B, and C complexes, whereas only unspliced pre-mRNA is detected in the region corresponding to H complexes (Fig. 4E). Cross-linking analysis of proteins in each of the various complexes is also shown (Fig. 4F). It seems clear that multiple proteins that are unique to a given complex can be identified with the use of a continuously labeled substrate. Such analysis can be extended if site-specific labeling is used. With this strategy, many proteins can be identified, which can then be further characterized with biochemical purification and cross-linking analysis.

VI. I D E N T I F I C A T I O N O F R N A B I N D I N G P R O T E I N S BY A cDNA EXPRESSION LIBRARY SCREEN

A. MATERIALS AND METHODS Appropriate plating cells were prepared and library titrations were initially performed following standard protocols (41) to allow plating of a complementary DNA (cDNA) expression library at relatively low density (i.e., plaques readily

4. RNA-Protein Interaction in Splicing

73

distinct from one another). This necessitated increased numbers of plates, but protein expression seems to be weaker on plates containing large numbers of plaques. Following initial incubation at 42~ for 4-6 hr, the lawns, containing barely visible plaques, were overlayed with nitrocellulose filters soaked in 10 mM IPTG (blot dry before overlaying) and incubated overnight at room temperature. The following morning, the filters were marked with india ink, and each filter was carefully removed and placed in binding buffer (50 mM Tris pH 8, 150 mM KC1, 1 mM EDTA, and 1 mM DTT). After all of the filters were lifted and the plates were returned to 4~ the filter-bound proteins were denatured/renatured by immersion in binding buffer plus 6 M urea. The filters were gently agitated at room temperature in the urea buffer for 10 min, after which the buffer was removed and replaced with binding buffer plus 3 M urea and agitated for 6 min. Again, the urea-containing buffer was removed and replaced with binding buffer plus 1.5 M urea and agitated for 5 min. The urea concentration was sequentially halved until a concentration of 0.093 M urea was reached, after which the filters were placed in binding buffer without urea for two 10-min incubations at room temperature. The R N A probe was prepared by transcription (as above) and resuspended in binding buffer plus 0.2 mg/ml yeast transfer R N A (tRNA). The probe (500,000 cpm/filter) was added to the filters, and the binding reaction was allowed to proceed with gentle agitation for 1 hr at room temperature. (The yeast tRNA concentration will vary with every situation. Too much t R N A may result in no signal, whereas too little may result in digestion of the probe R N A by abundant RNases associated with the filters.) Following the binding reaction, the filters were washed in binding buffer. After the last wash, the filters were dried extensively and exposed to film. (The affinity of the protein-RNA interaction will determine how long to wash the filters, but generally three or four washes for 10-15 min each is sufficient.)

Hints 1. The proper amount and type of nonspecific KNA to add to the binding reaction must be empirically determined for each probe. The more nonspecific KNA that can be included, the better, to avoid rapid degradation of the probe. 2. The binding reaction should never be allowed to proceed much longer than 1 hr to prevent KNAse digestion. 3. Do not use Blotto buffer or dehydrated milk solutions of any kind at any step, as they contain high levels of KNase.

B. RESULTS AND SUMMARY The use of cDNA expression library screens can sometimes obviate the need for extensive, difficult biochemical purification schemes to identify and characterize

74

James G. Patton et al.

Figure 5 Identification of R N A binding proteins by a cDNA expression library screen. A smooth muscle cDNA library (Stratagene, La Jolla, CA) in the Uni-ZAP Xt< vector was plated at low density and screened with a short R.NA derived from splicing regulatory elements flanking exon 3 in oe-TM. With this strategy, the clone was plaque purified, isolated, and sequenced.

4. RNA-Protein Interaction in Splicing

75

specific R N A or D N A binding proteins. However, such screens have generally been used to identify D N A binding proteins (43) and not R N A binding proteins. For R N A binding proteins, purification schemes can be devised using cross-linking as a screening strategy, as mentioned above. However, if specific R N A sequences have been identified that are necessary for a given R N A - p r o t e i n interaction, c D N A expression library screens offer an attractive, fast alternative to traditional approaches. As an example of this approach, an R N A sequence involved in smooth muscle regulation of oe-TM alternative splicing was recently identified by mutational analysis of splicing substrates (13). To identify candidate R N A binding proteins that interact with the newly identified regulatory sequence, we screened a smooth muscle c D N A expression library with an R N A containing a multimer of the regulatory sequence, similar to approaches used to identify specific transcription factors (43). The major drawback to this type of screen is the presence of abundant RNases associated with the plaque-bearing filters such that digestion of the R N A probe constantly decreases potentially positive signals. Using the modifications outlined above, we have overcome such obstacles and have cloned multiple R N A binding proteins with this approach (J. G. Patton, unpublished). As shown in Figure 5, specific clones can be identified and plaque purified based on their ability to bind specific R N A probes. Sequence analysis of this particular clone has shown that it is indeed an R N A binding protein, as demanded by the screening strategy. W e are now in the process of obtaining full-length clones corresponding to this c D N A and are attempting to determine whether the encoded protein regulates the splicing ofoe-TM. As with all such library screens, the difficulty lies in separating the false positives from the true positives. However, the ability to quickly identify candidate factors rather than using the laborious biochemical purification methods allows rapid progress toward more functional experiments.

ACKNOWLEDGMENTS This work was supported by grants from the NIH (RO1 GM50418) and the American Heart Association to J.G.P.B.T.D. is supported by an NIH training grant GM08554, and J.G.M. is a Ford Foundation Fellow. The Molecular Dynamics Phosphorlmager was purchased with funds from an NSF grant (BIR-9419667).

REFERENCES 1. Abmayr, S. M., Workman, J. L., and Roeder, R. G. (1988). The pseudorabies immediate early protein stimulates in vitro transcription by facilitating TFIID :promoter interactions. Genes Dev. 2, 542-553. 2. Adams, M. D., Rudner, D. Z., and Rio, D. R. (1996). Biochemistry and regulation of premRNA splicing. Curr. Opin. Cell Biol. 8, 331-339.

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3. Beggs, J. D. (1995). "Yeast Splicing Factors and Genetic Strategies for Their Analysis." R. G. Landes Co., Austin, TX. 4. Bennett, M., Michaud, S., Kingston, J., and Reed, R. (1992). Protein components specifically associated with prespliceosome and spliceosome complexes. Genes Dev. 6, 1986-2000. 5. Bindereif, A., and Green, M. R. (1987). An ordered pathway ofsnRNP binding during mammalian pre-mRNA splicing complex assembly. E M B O J . 6, 2415-2424. 6. Borer, P. N. (1975). In "Handbook of Biochemistry and Molecular Biology" (G. D. Fasmann, ed.), pp. 359-360. CRC Press, Cleveland, OH. 7. Cech, T. R. (1993). Structure and mechanism of the large catalytic RNAs: Group I and Group II introns and ribonuclease P. In "The RNA World" (R. F. Gesteland and J. F. Atkins, eds.), pp. 239-269. Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. 8. Chiara, M. D., Gozani, O., Bennett, M., Champion-Arnaud, P., Palandjian, L., and Reed, R. (1996). Identification of proteins that interact with exon sequences, splice sites, and the branchpoint sequence during each stage of spliceosome assembly. Mol. Cell. Biol. 16, 3317-3326. 9. Dreyfuss, G., Matunis, M. J., Pinol-Roma, S., and Burd, C. G. (1993). hnRNP proteins and the biogenesis of mRNA. Annu. Rev. Biochem. 62, 289-322. 10. Garcia-Blanco, M. A., Jamison, S. F., and Sharp, P. A. (1989). Identification and purification of a 62,000-dalton protein that binds specifically to the polypyrimidine tract of introns. Genes Dev. 3, 1874-1886. 11. Gerke, V., and Steitz, J. A. (1986). A protein associated with small nuclear ribonucleoprotein particles recognizes the 3' splice site of premessenger RNA. Cell (Cambridge, Mass.) 47, 973-984. 12. Gil, A., Sharp, P. A., Jamison, S. F., and Garcia-Blanco, M. A. (1991). Characterization ofcDNAs encoding the polypyrimidine tract binding protein. Genes Dev. 5, 1224-1236. 13. Gooding, C., Roberts, G. C., Moreau, G., Nadal-Ginard, B., and Smith, C. W.J. (1994). Smooth muscle specific switching ofc~-tropomyosin mutually exclusive exon selection by specific inhibition of the strong default exon. E M B O J . 13, 3861-3872. 14. Gozani, O., Patton, J. G., and Reed, R. (1994). A novel set of spliceosome-associated proteins (SAPs) and the essential splicing factor PSF bind stably to pre-mRNA prior to catalytic step II of the splicing reaction. E M B O J . 13, 3356-3367. 15. Green, M. R. (1986). Pre-mRNA splicing. Annu. Rev. Genet. 20, 671-708. 16. Hanna, M. M. (1989). Photoaffinity cross-linking methods for studying RNA-protein interactions. In "Methods in Enzymology" (J. E. Dahlberg, andJ. N. Abelson, eds.), Vol. 180, pp. 383-418. Academic Press, Orlando, FL. 17. Hoffman, B. E., and Grabowski, P.J. (1992). U1 snRNP targets an essential splicing factor, U2AF 65, to the 3' splice site by a network of interactions spanning the exon. Genes Dev. 6, 2554-2568. 18. Jamison, S. F., and Garcia-Blanco, M. A. (1992). An ATP-independent U2 small nuclear ribonucleoprotein particle/precursor mRNA complex requires both splice sites and the polypyrimidine tract. Proc. Natl. Acad. Sci. U.S.A. 89, 5482-5486. 19. Kohtz, J. D., Jamison, S. F., Will, C. L., Zuo, P., Ltihrmann, R., Garcia-Blanco, M. A., and Manley, J. (1994). Protein-protein interactions and 5' splice site recognition in mammalian m R N A precursors. Nature (London) 368, 119-124. 20. Konarska, M. M., and Sharp, P. A. (1986). Electrophoretic separation of complexes involved in the splicing of precursors to mRNAs. Cell (Cambridge, Mass.) 46, 845-855. 21. Konarska, M. M., and Sharp, P. A. (1987). Interactions between small nuclear ribonucleoprotein particles in formation of spliceosomes. Cell (Cambridge, Mass.) 49, 763-774. 22. Kr~imer, A. (1995). The biochemistry ofpre-mRNA splicing. In "Pre-mRNA Processing" (A. I. Lamond, ed.), pp. 35-64. R.G. Landes Co., Austin, TX. 23. Kr~imer, A. (1996). The structure and function of proteins involved in mammalian pre-mRNA splicing. Annu. Rev. Biochem. 65, 367-409.

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24. Lamm, G. M., and Lamond, A. I. (1993). Non-snRNP protein splicing factors. Biochim. Biophys. Acta 1173, 247-265. 25. Lin, C.-H., and Patton, J. G. (1995). Regulation of alternative 3' splice site selection by constitutive splicing factors. R N A 1, 234-245. 26. Lfihrmann, R. (1988). snRNP proteins. In "Structure and Function of Major and Minor Small Nuclear Ribonucleoprotein Particles" (M. L. Birnstiel, ed.), pp. 71-99. Springer-Verlag, Berlin. 27. Lynch, K. W., and Maniatis, T. (1996). Assembly of specific SR protein complexes on distinct regulatory elements of the Drosophila doublesex splicing enhancer. Genes Dev. 10, 2089-2101. 28. Madhani, H. D., and Guthrie, C. (1994). Dynamic R N A - R N A interactions in the spliceosome. Annu. Rev. Genet. 28, 1-26. 29. McGhee, J. D., and von Hippel, P. H. (1974). Theoretical aspects of DNA-protein interactions: cooperative and non-cooperative binding of large ligands to a one-dimensional homogeneous lattice. J. Mol. Biol. 86, 469-489. 30. Michaud, S., and Reed, R. (1991). An ATP-independent complex commits pre-mRNA to the mammalian spliceosome assembly pathway. Genes Dev. 5, 2534-2546. 31. Moore, M. J., Query, C. C., and Sharp, P. A. (1993). Splicing of precursors to mRNA by the spliceosome. In "The RNA World" (R. F. Gesteland and J. F. Atkins, eds.), pp. 303-358. Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. 32. Moore, M. J., and Sharp, P. A. (1992). Site-specific modification ofpre-lnRNA: the 2'-hydroxyl groups at the splice sites. Science 256, 992-997. 33. Mullen, M. P., Smith, C. W. J., Patton, J. G., and Nadal-Ginard, B. (1991). a-Tropomyosin mutually exclusive exon selection: Competition between branch point/polypyrimidine tracts determines exon choice. Genes Dev. 5, 642-655. 34. Nilsen, T. W. (1994). R N A - R N A interactions in the spliceosome: Unraveling the ties that bind. Cell (Cambridge, Mass.) 78, 1-4. 35. Patton, J. G., Mayer, S. A., Tempst, P., and Nadal-Ginard, B. (1991). Characterization and molecular cloning of polypyrimidine tract binding protein: A component of a complex necessary for pre-mRNA splicing. Genes Dev. 5, 1237-1251. 36. Patton, J. G., Porto, E. B., Galceran, J., Tempst, P., and Nadal-Ginard, B. (1993). Cloning and characterization of PSF, a novel pre-mRNA splicing factor. Genes Dev. 7, 393-406. 37. P6rez, I., Lin, C-H., McAfee, J. G., and Patton, J. G. (1997). Mutation of PTB binding sites causes misregulation of alternative 3' splice site selection in vivo. R N A 3, 764-778. 38. Pikielny, C. W., Rymond, B. C., and Rosbash, M. (1986). Electrophoresis ofribonucleoproteins reveals an ordered assembly pathway of yeast splicing complexes. Nature (London) 324, 341-345. 39. Ruby, S. W., and Abelson, J. (1988). An early hierarchic role of U1 small nuclear ribonucleoprotein in spliceosome assembly. Science 242, 1028-1035. 40. Ruskin, B., Zamore, P. D., and Green, M. R. (1988)./~ factor, U2AF, is required for U2 snRNP binding and splicing complex assembly. Celt (Cambridge, Mass.) 52, 207-219. 41. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). "Molecular Cloning," 2nd ed. Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. 42. Seraphin, B., and Rosbash, M. (1989). Identification of functional U1 snRNA-pre-mRNA complexes committed to spliceosome assembly and splicing. Cell (Cambridge, Mass.) 59, 349-358. 43. Singh, H., LeBowitz, J. H., Baldwin, A. S.J., and Sharp, P. A. (1988). Molecular cloning of an enhancer binding protein: Isolation by screening of an expression library with a recognition site DNA. Cell (Cambridge, Mass.) 52, 415-423. 44. Singh, R., Valcarcel, J., and Green, M. R. (1995). Distinct binding specificities and functions of higher eukaryotic polypyrimidine tract binding proteins. Science 268, 1173-1176. 45. Smith, C. W. J., and Nadal-Ginard, B. (1989). Mutually exclusive splicing of a-tropomyosin exons enforced by an unusual lariat branch point location; implications for constitutive splicing. Cell (Cambridge, Mass.) 56, 749-758.

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46. Smith, C. W. J., Patton, J. G., and Nadal-Ginard, B. (1989a). Alternative splicing in the control of gene expression. Annu. Rev. Genet. 23, 527-577. 47. Smith, C. W. J., Porto, E. B., Patton, J. G., and Nadal-Ginard, B. (1989b). Scanning from an independently specified branch point defines the 3' splice site of mammalian introns. Nature (London) 342, 243-247. 48. Steiner, R. F., Kinner, W., Lunasin, A., and Delac, J. (1973). Fluorescent derivatives ofpolyribonucleotides containing 8-adenosine. Biochim. Biophys. Acta 294, 24-37. 49. Steitz, J. A., Black, D. L., Gerke, V., Parker, K. A., Kramer, A., Frendeway, D., and Keller, W. (1988). Functions of the Abundant U-snRNPs. In "Structure and Function of Major and Minor Small Nuclear Ribonucleoprotein Particles" (M. L. Birnstiel, ed.), pp. 115-154. SpringerVerlag, Berlin. 50. Swanson, M. S., and Dreyfuss, G. (1988). RNA binding specificity of hnRNP proteins: A subset bind to the 3' end ofintrons. EMBOJ. 7, 3519-3529. 51. Tazi, J., Alibert, C., Temsamani, J., Reveillaud, I., Cathala, G., Brunel, C., and Jeanteur, P. (1986). A protein that specifically recognizes the 3' splice site of mammalian pre-mRNA introns is associated with a small nuclear ribonucleoprotein. Cell (Cambridge, Mass.) 47, 755-766. 52. Weiner, A. M. (1993). mRNA Splicing and autocatalytic introns: Distant cousins or the products of chemical determinism? Cell (Cambridge, Mass.) 72, 161-164. 53. Wu, J. Y., and Maniatis, T. (1994). Specific interactions between proteins implicated in splice site selection and regulated alternative splicing. Cell (Cambridge, Mass.) 75, 1061-1070. 54. Zamore, P. D., and Green, M. R. (1989). Identification, purification, and biochemical characterization of U2 small nuclear ribonucleoprotein auxiliary factor. Proc. Natl. Acad. Sci. U.S.A. 86, 92439247. 55. Zamore, P. D., Patton, J. G., and Green, M. R. (1992). Cloning and domain structure of the mammalian splicing factor U2AF. Nature (London) 355, 609-614.

In Vitw Analysis of Mammalian Cell mRNA

3' Processing Gregory M. Gilmartin Department of Microbiology and Molecular Genetics and the Markey Center for Molecular Genetics University of Vermont Burlington, Vermont 05405

I. I N T R O D U C T I O N The biosynthesis of a functional eukaryotic messenger RNA (mRNA) requires the posttranscriptional modification of both the 5' and 3' ends of the newly synthesized transcript. While the 5' end acquires a 7-methyl diguanosine triphosphate (mYGpppG) cap, the 3' end of the pre-mRNA is subject to endonucleolytic cleavage followed by the addition ofa poly(A) tail. The only known exceptions are those mRNAs of higher eukaryotes encoding histone proteins. Although their 5' ends are capped and their 3' ends are generated by endonucleolytic cleavage, these mRNAs do not receive a poly(A) tail. The machinery responsible for histone pre-mRNA 3' processing is clearly distinct from that which directs polyadenylation. This chapter will focus solely on the 3' processing ofpolyadenylated mRNAs; histone 3' processing is discussed in detail in Chapter 10. The poly(A) tail appears to play a variety of roles in the life of an mRNA. In addition to its role in promoting mRNA stability, it appears to participate in nuclear export, as well as in the subsequent recruitment of the message into ribosomes. The 3' processing of the pre-mRNA may also contribute directly to the regulation of gene expression through the modulation of both the level and the coding capacity of the mature message. Modulation ofmRNA protein-coding capacity is exemplified by the regulation oflgM heavy chain mRNA synthesis, in which poly(A) site choice contributes to the production of the secreted versus the membrane form of the/z heavy chain protein during the course of B-cell development. Despite the ubiquity of mRNA polyadenylation, the sequence conservation among the cis-acting elements responsible for directing 3' processing is surpris79 m R N A F o r m a t i o n and F u n c t i o n Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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ingly limited (reviewed by WaNe, 1995). In mammalian cells, only the hexamer AAUAAA, which resides 10-30 nucleotides upstream of the cleavage site, exhibits a high degree of sequence conservation among poly(A) sites. Immediately downstream of the cleavage site is an amorphous U- or GU-rich element that possesses neither a conserved sequence nor apparent secondary structure. These two elements, which together comprise the "core" poly(A) site, appear to be sufficient, at least in some cases, to direct 3' processing in vivo. Sequences that flank the core poly(A) site have been shown to influence the efficiency of 3' processing. Sequences that enhance 3' processing have been found upstream of the human immunodeficiency virus type 1 (HIV-1), SV40 late, adenoviruses L1, L3, and L4, and hepatitis B virus core poly(A) sites. These elements function in an orientation- and positiondependent manner. Although generally U-rich, they exhibit little sequence or apparent structural similarity. The development of an in vitro 3' processing system by Moore and Sharp (1985), capable of efficient 3' cleavage and polyadenylation of exogenously added pre-mRNA, has permitted the detailed investigation of the biochemistry of 3' processing. Although tightly coupled in vivo, the endonucleolytic cleavage and poly(A) addition reactions can be carried out independently in vitro. Fractionation of mammalian cell nuclear extracts has demonstrated the requirement for at least six factors for the in vitro reconstitution of 3' processing (reviewed by WaNe, 1995). Cleavage and polyadenylation specificity factor (CPSF) binds the premRNA on recognition of the AAUAAA hexamer. The interaction of CPSF with cleavage stimulation factor (CstF), which binds the downstream element, results in the formation of a stable ternary complex. Endonucleolytic cleavage of the premRNA requires the subsequent recruitment of cleavage factors Im and IIm, and in most cases of poly(A) polymerase (PAP). Following cleavage, CPSF directs the addition of adenylate residues to the 5' cleavage product by PAP, while poly(A) binding protein II enhances the processivity of PAP and confers poly(A) tail length control. In vitro, CPSF and PAP are sufficient for specific poly(A) addition. This chapter will focus on in vitro methods for the analysis of m R N A 3' processing factors and the dissection of the cis-acting elements of the poly(A) site. Functional assays for the evaluation of both poly(A) site cleavage and poly(A) addition are presented. The analysis of RNA-protein and protein-protein interactions involved in 3' processing are addressed by protocols for gel mobility shift analysis and ultraviolet light (UV) cross-linking. Finally, an in vitro selection protocol is discussed for the examination of the contributions of specific pre-mRNA sequences to poly(A) site recognition and processing.

II. G E N E R A L C O N S I D E R A T I O N S F O R T H E A N A L Y S I S OF m R N A 3' P R O C E S S I N G in Vitro The initial work demonstrating the in vitro 3' processing of exogenous premRNA (Moore and Sharp, 1985) was carried out in a HeLa cell nuclear extract

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originally developed by Dignam et al. (1985) for the analysis of R N A polymerase II transcription initiation. Subsequent modifications of the Dignam protocol, developed in the Keller lab (Ruegsegger et al., 1996), have yielded superior extracts active for pre-mILNA 3' processing. The Keller nuclear extract may be used directly for 3' processing assays or may be dialyzed into an appropriate buffer prior to use. Dialysis, however, consistently results in a loss of processing activity on the order of approximately twofold. The protocols presented below address the use of both dialyzed and undialyzed nuclear extracts. The extracts are quick frozen in liquid N2 and stored at -80~ Freeze-thawing does not appear to have a large impact on processing activity, but in principle, it should be avoided whenever possible. Two functional 3' processing assays may be performed in a HeLa cell nuclear extract: (1) endonucleolytic cleavage and (2) poly(A) addition. As discussed below, the use of ATP analogs allows for the independent analysis of these two reactions. Although it is possible to both cleave and polyadenylate a pre-mRNA substrate in vitw, such coupled reactions are generally avoided due to the difficulty encountered in the interpretation of the results. Poly(A) addition is not obligatorily coupled to cleavage of the t
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are usually at least partially degraded in nuclear extract, 3' end-labeling allows for their positive identifcation.

III. P R E - m R N A C L E A V A G E AND POLYADENYLATION Our current understanding of the biochemistry of 3' processing stems, in large part, from the ability to faithfully process exogenously added pre-mRNAs in a reaction dependent on the known cis-acting sequence elements (Hart et al., 1985; Moore and Sharp, 1985; Moore et al., 1986; Sperry and Berget, 1986; Zarkower et al., 1986). An additional advantage of the in vitro system is the ability to uncouple the endonucleolytic cleavage reaction from the poly(A) addition reaction and independently examine the reaction parameters, the sequence requirements, and the participation of trans-acting factors for each step of 3' processing. Two parameters allow for the independent analysis of endonucleolytic cleavage and poly(A) addition. The first is the differential requirement for Mg2+: poly(A) addition requires Mg 2+, while the endonucleolytic cleavage reaction can proceed in its absence. Ethylenediaminetetraacetic acid (EDTA) may therefore be used to examine cleavage in the absence of poly(A) addition. The second parameter is the requirement for ATP. While both reactions require the presence of ATP, only poly(A) addition requires ATP hydrolysis. An ATe analog such as c~,/3-methylene ATP will therefore support cleavage but not poly(A) addition. Furthermore, since poly(A) addition requires a 3'-OH, the use of the cordycepin (3'dATP) allows for cleavage but prevents the addition of more than a single adenylate residue. Cordycepin is generally the preferred analog for the analysis ofpoly(A) site cleavage in nuclear extract. Metal ions can be further exploited to examine two modes of poly(A) addition: (1) specific (AAUAAA-dependent) and (2) nonspecific (AAUAAA-independent). In the presence of Mg 2+, poly(A) addition requires both an AAUAAAcontaining ILNA substrate and CPSF (the factor responsible for AAUAAA recognition). The replacement of Mg 2+ with Mn 2+, however, allows for the polyadenylation of exogenously added 1KNA in the absence of CPSF in a sequence-independent manner. Such an assay may be particularly useful in the evaluation of poly(A) polymerase activity during protein purification.

A. PROTOCOL: POLY(A) SITE CLEAVAGE IN HELA CELL NUCLEAR EXTRACT 1. Reagents HeLa cell nuclear extract (prepared by the method of 1Kuegsegger et al., 1996): dialyzed (---24 mg protein/ml dialyzed into buffer B) or undialyzed (---29 mg protein/ml)

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3' Processing

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oe-32p-Labeled pre-mRNA substrate, ---25 fmol//xl ('-'4 X 103 Cerenkov cpm/fmol), resuspended in H20 Buffer A [20 mM N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES) (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM phenylmethylsulfonyl fluoride (PMSF), 0.5 mM dithiothreitol (DDT)], required if undialyzed extract is used Buffer B [100 mM KC1, 20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if dialyzed extract is used 3' dATP (12.5 mM) eolyvinyl alcohol (PVA) (10%) tRNA (0.5 mg/ml) tRNA (10 mg/ml) ETS buffer [10 mM EDTA, 10 mM Tris (pH 7.8), 0.5% SDS] Proteinase K (20 mg/ml) Sodium acetate (pH 6.1) (3 M) Phenol, phenol/chloroform/isoamyl alcohol (25:24: 1) 95% Ethanol, 75% ethanol 7 M Urea polyacrylamide gel, 90% formamide loading dye

2. M e t h o d s

1. Reactions (25/a,1) are set up on ice by the addition of individual components in the following order: 5/xl PVA 1 /a,1 3' dATP 1 /zl tRNA (0.5 mg/ml) 12/,1 Buffer B and 5/,1 dialyzed nuclear extract or 14.5/,1 buffer A and 2.5/,1 undialyzed nuclear extract 1/,1 32p-Labeled pre-mRNA substrate 2. Incubate the reactions at 30~ for 30 to 90 mins. 3. Add 175/a,1 ETS buffer and 5/,1 proteinase K. Incubate the reactions for 10 min at 37~ 4. Extract the reaction with 200/a,1 phenol. Remove the aqueous phase and add 25/xl of 3 M sodium acetate and 1/xl tRNA (10 mg/ml). Reextract the aqueous phase with 200/,1 phenol/chloroform/isoamyl alcohol. 5. Precipitate the RNA with 2.5 volumes of 95% ethanol, wash the pellet with 75% ethanol, and dry the pellet. Resuspend the pellet in 5/,1 loading dye, boil for 1 min, and quickly chill in an ice-water bath. Load onto a denaturing polyacrylamide gel (prerun for 30 rain).

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Gregory M. Gilmartin B. PROTOCOL: POLY(A) ADDITION IN H E L A CELL NUCLEAR EXTRACT 1. Reagents

HeLa cell nuclear extract (prepared by the method of Ruegsegger et al., 1996): dialyzed (---24 mg protein/ml dialyzed into buffer B) or undialyzed (---29 mg protein/M) oe-32p-Labeled precleaved P,.NA substrate, "--25 fmol/~l ("-4 • 103 Cerenkov cpm/fmol), resuspended in H20 Buffer A [20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if undialyzed extract is used Buffer B [100 mM KC1, 20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if dialyzed extract is used 10 • Poly(A) mix containing 10 mM ATP, 50 mM creatine phosphate, and 7 mM MgC12 PVA (10%) tP,.NA (0.5 mg/ml) ETS buffer [10 mM EDTA, 10 mM Tris (pH 7.8), 0.5% sodium dodecyl sulfate (SDS)] Proteinase K (20 mg/ml) Sodium acetate (pH 6.1) (3 M) tRNA (10 mg/ml) Phenol, phenol/chloroform/isoamyl alcohol (25:24: 1) 95% ethanol, 75% ethanol 7 M Urea polyacrylamide gel, 90% formamide loading dye 2. Methods

Reactions (25/~1) are set up on ice by the addition of individual components in the following order: 5 ~1 PVA 2.5/~1 Poly(A) mix 1 /.tl tP,.NA (0.5) mg/ml) 10.5/.tl Buffer B and 5/.tl dialyzed nuclear extract or 13/,1 buffer A and 2.5/,1 undialyzed nuclear extract 1/zl 32p-Labeled pre-mR.NA substrate Polyadenylation reactions are camed out in the same manner as for the poly(A) site cleavage reactions discussed above, although the incubation period may be shortened to 15 min or less at 30~ The specificity of cleavage and poly(A)

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addition may be addressed through the use of an RNA containing an AAUAAA hexamer point mutation. Single-point mutations, such as AACAAA, have been shown to block processing both in vitro and in vivo.

IV. 3' P R O C E S S I N G

COMPLEX

FORMATION

Gel mobility shift analysis has been used extensively for the investigation of protein-nucleic acid interactions. It has been particularly useful for the dissection of the RNA-binding properties of specific 3' processing factors, the analysis of the order of processing complex assembly, and the evaluation of the contributions of protein-protein interactions to complex assembly. Although this assay may be performed in a nuclear extract, the interpretation of such assays may be limited by the complex milieu of RNA-binding proteins. The assay has been extremely useful, however, in the analysis of purified or partially purified processing factors. The protocol presented below uses nuclear extract, but it may be easily adapted for the use of fractionated extracts. Figure 1 illustrates the sequence requirements for 3' processing complex assembly on an HIV-1 pre-mRNA in a HeLa cell nuclear extract. Two specific complexes are observed. Previous work has shown that the faster-migrating "A" complex represents the interaction of CPSF with the pre-mRNA and, as demonstrated in Figure 1, requires the presence of the AAUAAA hexamer. The more slowly migrating "B" complex represents the interaction of CPSF and CstF, and its formation requires both the AAUAAA hexamer and the downstream element. Assembly of the CPSF-CstF-RNA complex does not require ATP, so complexes may be examined in the absence of processing. Heparin, an acid mucopolysaccharide, is added following processing complex assembly to reduce the interaction of nonspecific RNA-binding proteins with the pre-mRNA. The processing complexes that assemble at the poly(A) site are surprisingly large, given the simplicity of the 3' processing reaction, and their resolution therefore requires a very-low-percentage polyacrylamide gel [3%, 80:1 or 100:1 (acrylamide : bisacrylamide)].

A. PROTOCOL: ANALYSIS OF 3' PROCESSING COMPLEX FORMATION IN H E L A CELL NUCLEAR EXTRACT 1. Reagents HeLa cell nuclear extract (prepared by the method of Ruegsegger et al., 1996): dialyzed (~24 mg protein/ml dialyzed into buffer B) or undialyzed (---29 mg protein/ml)

Figure 1 Sequence requirements for 3' processing complex formation in a HeLa cell nuclear extract. 32p-Labeled wild-type (use/CPS) and mutant HIV-1 pre-mRNAs were incubated in a HeLa cell nuclear extract for 10 min at 30~ Following treatment with heparin at 0~ the RNA-protein complexes were resolved on a 3% nondenaturing polyacrylamide gel. u s e / C P S / A A C A A A contains a single-point mutation (AAUAAA to AACAAA); use/CPS/Adse contains a substitution of vector sequences for the poly(A) site downstream element: Ause/CPS contains a linker substitution within the 3' processing enhancer; CPS contains a deletion of upstream sequences encompassing the 3' processing enhancer; CPS/AACAAA and CPS/Adse are analogous to CPS but contain mutations in the hexamer and downstream element, respectively. Reproduced from Gilmartin et al. (1992).

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c~-32p-Labeled R N A substrate, ---25 fmol//zl ('--4 X 103 Cerenkov cpm/ fmol), resuspended in H20 Buffer A [20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 01. mM PMSF, 0.5 mM DTT], required if undialyzed extract is used Buffer B [100 mM KC1, 20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if dialyzed extract is used PVA (10%) tRNA (0.5 mg/ml) Heparin (100 mg/ml) 3% Nondenaturing polyacrylamide gel [80:1 or 100:1 (acrylamide : bisacrylamide)] 0.5X Tris-borate-EDTA (TBE)[25 mM Tris-borate (pH 8.3), 0.5 mM EDTA], gel electrophoresis equipment: 20 X 20-cm glass plates, 1.5-mm spacers, 15-well (9-mm-wide teeth) comb; the notched plate is treated with dimethyldichlorosilane (5% in chloroform); excess silane is removed with ethanol; do not treat the plates with RainX 2. Methods

1. Gel preparation: nondenaturing gel mix (80 ml) contains the following: 6 ml acrylamide [40%, 80:1 or 100:1 (acrylamide : bisacrylamide)] 8 m15• TBE 66 ml H20 Add 560/xl ammonium persulfate (10%) and 48/xl N , N , N ' , N ' tetramethylethylenediamine (TEMED). Pour gel, clamp the comb and three sides of the plates, and allow to polymerize for at least 1 hr at room temperature. Gel, running buffer (0.5 • TBE), and electrophoresis apparatus must be equilibrated to 4~ prior to use, preferably by storage at 4~ overnight. 2. Gel mobility shift reactions (20/xl) are set up on ice by the addition of individual components in the following order: 2 ~1 PVA 1/xl tRNA (0.5 mg/ml) 14/xl Buffer B and 2/xl dialyzed nuclear extract or 14/xl buffer A and 2/xl undialyzed nuclear extract. 1/xl 32p-Labeled pre-mRNA substrate 3. Incubate the reactions at 30~ for 10 min. 4. Return the reactions to ice, add 1/xl heparin, and incubate the reactions for 10 min on ice.

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Gregory M. Gilmartin 5. Load the reactions directly onto a nondenaturing polyacrylamide gel (no loading dye is used) that has been prerun for 30 min at 300 V at 4~ Use gel-loading pipette tips to layer the reactions directly onto the bottom of the well. 6. Run gel at 300 V for 90 to 120 min at 4~ Transfer gel to 3MM paper and dry.

V. 3' P R O C E S S I N G

COMPLEX

STABILITY

The gel mobility shift assay may be extended to examine the stability of specific R N A : protein complexes. As detailed below, 3' processing complexes are assembled on a 32p-labeled pre-mRNA and subsequently challenged with a large excess of unlabeled poly(A) site containing RNA. The stability of the initial complex may therefore be monitored over time, as any processing factors released from the 32p-labeled pre-mRNA will subsequently be sequestered by the excess unlabeled pre-mRNA. The ability of a poly(A) site to direct the assembly of a stable processing complex has been shown to correlate closely with the processing efficiency of the pre-mRNA. Figure 2 illustrates the use of the gel mobility shift assay to examine the stability of complexes formed in a HeLa cell nuclear extract on a set of HIV-1 pre-m_R.NAs. The wild-type pre-m_R.NA (use/CPS) directed the formation of a stable complex, whereas disruption of the TAR. R N A stem-loop structure (use/ stem1 and use~stem2) dramatically reduced complex stability (Fig. 2; compares lanes 1-3 to lanes 4-9). Compensatory changes that restored the R N A stem-loop structure (use~stem1~2) restored the ability of the R N A to form a stable processing complex (Fig. 2, lanes 10-13).

A. PROTOCOL: ANALYSIS OF 3' PROCESSING COMPLEX STABILITY IN H E L A CELL NUCLEAR EXTRACT 1. Reagents HeLa cell nuclear extract (prepared by the method of Ruegsegger et al., 1996): dialyzed (---24 mg protein/ml dialyzed into buffer B) or undialyzed (---29 mg protein/ml) ~x-32p-Labeled R N A substrate, ---25 fmol//zl ( " 4 • 103 Cerenkov cpm/ fmol), resuspended in H20 Unlabeled pre-mRNA substrate, ---2.5 pmoles//zl Buffer A [20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if undialyzed extract is used

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Figure 2 Stability of 3' processing complexes formed on HIV-1 pre-mRNAs. 32p-Labeled wildtype (use/CPS) and mutant HIV-1 pre-mRNAs were incubated in a HeLa cell nuclear extract for 10 min at 30~ An aliquot was removed at time 0 and treated with heparin at 0~ An ---100-fold molar excess of unlabeled wild-type pre-rnRNA was added to the remaining reaction, and aliquots were removed after additional incubation for 10 and 20 rain at 30~ Each aliquot was treated with heparin at 0~ The RNA-protein complexes were resolved on a 3% nondenaturing polyacrylamide gel. Reproduced from Gilmartin et at. (1992).

Buffer B [100 m M KC1, 20 m M H E P E S (pH 7.9), 0.2 m M E D T A , 10% glycerol, 0.1 m M P M S F , 0.5 m M D T T ] , r e q u i r e d if dialyzed extract is used e V A (10%) tRNA

(0.5 m g / m l )

H e p a r i n (30 m g / m l ) 3% N o n d e n a t u r i n g p o l y a c r y l a m i d e gel [ 8 0 : 1 or 1 0 0 : 1 (acrylamide :bisacrylamide)], 0 . 5 X T B E [25 m M T r i s - b o r a t e (pH 8.3),

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Gregory M. Gilmartin 0.5 m M EDTA], gel electrophoresis equipment: 20 • 20-cm glass plates, 1.5-mm spacers, 15-well (9-mm-wide teeth) comb; the notched plate is treated with dimethyldichlorosilane (5% in chloroform); excess silane is removed with ethanol; do not treat the plates with RainX

2. Methods 1. Nondenaturing gel preparation: as discussed above. 2. Gel mobility shift react.3ns (30/xl) are set up on ice by the addition of individual components in the following order: 3/~1 PVA 1.5/xl tRNA (0.5 mg/ml) 21.5/M Buffer B and 3/~1 dialyzed nuclear extract or 21.5/~1 buffer A and 3/~1 undialyzed nuclear extract. 1/M 32p-Labeled pre-mRNA substrate 3. Incubate the reactions at 30~ for 10 rain. 4. Remove a 5-/~1 aliquot and add to a cold microfuge tube containing 1/~1 of heparin on ice. 5. To the remaining reaction add 1 ~1 unlabeled pre-mRNA and continue the incubation at 30~ Remove 5.2-/,1 aliquots at 5, 10, 20, and 30 min and add to cold microfuge tubes containing 1/xl heparin on ice. 6. Load the reactions directly onto a nondenaturing polyacrylamide gel (no loading dye is used) that has been prerun for 30 min at 300 volts at 4~ Use gel-loading pipette tips to layer the reactions directly onto the bottom of the well. 7. Run gel at 300 V for 90 to 120 min at 4~ Transfer gel to 3MM paper and dry.

VI. P R O B I N G 3' P R O C E S S I N G F A C T O R - P R E - m R N A INTERACTIONS BY UV CROSS-LINKING UV light-induced RNA-protein cross-linking has been used to identify several of the RNA-binding components of the pre-mRNA 3' processing complex. Exposure of processing complexes assembled in a HeLa cell nuclear extract to 254-nm light is sufficient to induce specific covalent cross-linking of the 160-kDa and 30-kDa subunits of CPSF and the 64-kDa subunit of CstF to the pre-mRNA. Recent analysis of purified cleavage factor Im (CFI) demonstrated that all three subunits of the heterotrimeric complex (68, 59, and 25 kDa) could be UV crosslinked to the pre-mRNA (Ruegsegger et at., 1996).

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Mthough this is a potentially useful assay, the interpretation of UV crosslinking data may be fraught with difficulties. The assay relies on the formation of a covalent RNA-protein bond followed by the removal of most, if not all, of the non-cross-linked R N A with RNase. Consequently, the outcome of the assay is dependent on the nucleotide content of the protein contact site and the choice of [a-32p]NTP used for synthesis of the pre-mRNA. Therefore, while a positive signal is a strong indication of protein-RNA contact, the absence of a signal cannot be taken as evidence for the lack of RNA-protein contact. Likewise, UV cross-linking cannot be used for the quantitative comparison of protein binding to RNAs of different sequence. The use of an AAUAAA hexamer mutant is a necessary control for the identification of poly(A) sitespecific UV cross-linked products amid the myriad nonspecific RNA-binding proteins present in a nuclear extract. The use of site-specific internal 32p-labeling of the pre-mRNA (Moore and Sharp, 1992) may be useful for the localization of protein contact sites on RNA. This technique has been used to demonstrate an interaction of the 160-kDa subunit of CPSF with the upstream 3' processing enhancer of the HIV-1 poly(A) site (Gilmartin et al., 1995). Figure 3 illustrates the UV cross-linking pattern of CPSF to both uniformly labeled and site-specifically labeled HIV-1 pre-mRNAs.

A. PROTOCOL: O V CROSS-LINKING OF 3' PROCESSING FACTORS IN HELA CELL NUCLEAR EXTRACT 1. Reagents HeLa cell nuclear extract (prepared by the method of 1Kuegsegger et al., 1996): dialyzed (---24 mg protein/ml dialyzed into buffer B) or undialyzed ('--29 mg protein/ml) a-32p-Labeled 1KNA substrate, ---25 fi'nol//xl (~4 • 103 Cerenkov cpm/ fmol), resuspended in H20 Buffer A [20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if undialyzed extract is used Buffer B [100 mM KC1, 20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT], required if dialyzed extract is used tlKNA (0.5 mg/ml) Heparin (100 mg/ml) RNase A (1 mg/ml) 72-Well microtiter plate (5.5 X 8 cm, Nunc, Naperville, IL) SDS-polyacrylamide gel, loading dye

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Figure 3 UV cross-linking of CPSF and CstF to HIV-1 p r e - n ~ N A s . 32p-Labeled HIV-1 premRNAs were incubated with CPSF or CstF, and the reactions were treated with heparin at 0~ irradiated with UV light, and then digested with RNase A. The resulting labeled proteins were then analyzed on a 10% polyacrylamide-SDS gel. Lanes 1, 3, 5, and 7 contain the wild-type HIV-1 prem R N A substrate, lanes 2, 4, 6, and 8 contain the HIV-1 p r e - m R N A substrate with the single-point mutation (U to C) in the hexamer. (A) UV cross-linking of partially purified CPSF or CstF to uniformly [c~-32p]UTP-labeled HIV-1 pre-mRNAs. (B) UV cross-linking of partially purified CPSF to RNAs specifically 32p-labeled within the upstream element (sequences - 7 3 to - 145 upstream of the AAUAAA hexamer). (C) UV cross-linking of glycerol gradient-purified CPSF to RNAs specifically labeled within the upstream element. Reproduced from Gilmartin et al. (1995).

2. M e t h o d s 1. Reactions (20 hal) are set up on ice by the addition of individual components in the following order (note that PVA is omitted from

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these reactions since it is not compatible with SDS-polyacrylamide gel electrophoresis): 1/,1 t R N A (0.5 mg/ml) 16 ~1 Buffer B and 2/~1 dialyzed nuclear extract or 16 ~1 buffer A and 2/,1 undialyzed nuclear extract 1/,1 32p-Labeled pre-mRNA substrate (~-25 fmol) 2. Incubate the reactions at 30~ for 5 mins. 3. Return the samples to ice and add 1/zl of heparin. Incubate for 5 min on ice. 4. Transfer the samples to the microtiter plate. 5. Irradiate the samples (uncovered) with 254-nm wavelength light [Ultraviolet Light Products (UVP, San Gabriel, CA) model UVG-54] at a distance of 5 cm for 10 min on ice in a 4~ room. 6. Return the samples to the original microfuge tubes, add RNase A to 50 mg/ml, and incubate for 15 min at 37~ 7. Add protein gel-loading buffer, heat the samples at 100~ for 2 min, and load onto an SDS-polyacrylamide gel.

VII. A P P L I C A T I O N OF In Vitro S E L E C T I O N T O T H E A N A L Y S I S OF S E Q U E N C E S T H A T D I R E C T 3' P R O C E S S I N G The cis-acting sequence elements of the mammalian poly(A) site have been extensively probed with site-directed mutagenesis and base-modification techniques. The data derived from these studies, coupled with sequence comparisons of a wide range of m R N A 3' ends, have revealed a very limited set of sequence requirements for 3' processing. The only conserved sequence, AAUAAA, is also the only sequence found to be sensitive to both single-base substitution and base modification (Conway and Wickens, 1987). In vitro selection has therefore been used to examine the contribution of sequences flanking the AAUAAA hexamer to poly(A) site recognition and processing (Graveley et al., 1996, 1997). The analysis of poly(A) site cis-acting elements by in vitro selection may be based on either a functional assay [cleavage or poly(A) addition] or an RNA-protein binding assay. The protocol presented below and diagrammed in Figure 4 uses a poly(A) addition assay to examine the impact of sequences upstream of the AAUAAA hexamer. This in vitro selection strategy relies on the ability to enrich for efficient poly(A) addition substrates from a pool of RNAs randomized at specific positions. RNAs are polyadenylated in the presence of purified CPSF and recombinant poly(A) polymerase, and the polyadenylated products are selected by binding to oligo(dT) beads. The selected RNAs are then subject to reverse

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Figure 4 Schematic diagram of the in vitro selection strategy for the isolation of efficient HIV-1 premRNA poly(A) addition substrates. Reproduced from Graveley et al. (1996).

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transcriptase-PCR (RT-PCR), and the products are transcribed to yield a new pool of polyadenylation substrates, enriched for those RNAs that are efficiently polyadenylated.

A. PROTOCOL: In Vitro SELECTION FOR EFFICIENT POLYADENYLATION SUBSTRATES 1. Reagents Purified CPSF (Gilmartin et al., 1995) and recombinant poly(A) polymerase (Wahle et al., 1991) Randomized pool of RNA substrates, 75 fmol, resuspended in H20 eVA (10%) Buffer B [100 mM KC1, 20 mM HEPES (pH 7.9), 0.2 mM EDTA, 10% glycerol, 0.1 mM PMSF, 0.5 mM DTT] MgC12/ATP solution (17.5 mM MgC12, 25 mM ATP) tRNA (0.5 mg/ml) ETS buffer [10 mM EDTA, 10 mM Tris (pH 7.8), 0.5% SDS] Proteinase K (20 mg/ml) Sodium acetate (pH 6.1) (3 M) Glycogen (20 mg/ml) Phenol, phenol/chloroform/isoamyl alcohol (25:24: 1) 95% Ethanol, 75% ethanol Dynabeads Oligo(dT)2s (Dynal, Great Neck, NY) resuspended in 2X binding buffer [20 mM Tris-HC1 (pH 7.5), 1.0 M LiC1, 2 mM EDTA] (5/,g Dynabeads//,1) Dynabead wash buffer [10 mM Tris-HC1 (pH 7.5), 0.15 M LiC1, 1 mM EDTA] EDTA (pH 7.5) (2 mM) Reagents and primers for RT-PCR; the 5' primer contains the SP6 RNA polymerase promoter to allow for transcription of the PCR product; both 5' and 3' primers contain appropriate restriction sites to facilitate the cloning of the PCR products Reagents for SP6 RNA polymerase in vitro transcription 2. Methods 1. Reactions (25 p,1) are set up on ice by the addition of individual components in the following order: 5/xl PVA 12/xl Buffer B

96

Gregory M. Gilmartin 1 /zl tRNA (0.5 mg/ml) 5/xl (10 fmol) CPSF 1 /xl (300 fmol) PAP 1 /xl Randomized pool of R N A substrates (75 fmol) 2. Incubate the reactions at 30~ for 10 min to allow for the assembly of polyadenylation complexes. 3. Add 1 /xl MgCl2/ATP solution and incubate for 2 min at 30~ 4. Add 175/xl ETS buffer and 5/xl proteinase K. Incubate the reactions for 10 min at 37~ 5. Extract the reaction with 200/xl phenol. Remove the aqueous phase and add 25/xl of 3 M sodium acetate and 1/xl glycogen. Reextract the reaction with 200/xl phenol/chloroform/isoamyl alcohol. 6. Ethanol precipitate the RNA, wash the pellet with 75% ethanol, and dry the pellet. Resuspend the pellet in 20/xl RiO. 7. Heat the R N A sample to 65~ for 2 min and add to 100/xg Dynabeads Oligo(dT)25 in 20/zl 2• binding buffer. Incubate at 25~ for 5 min. 8. Wash the beads twice with 50/xl of Dynabead wash buffer. 9. Elute the bound R N A by the addition of 10/xl of 2 mM EDTA followed by incubation at 65~ for 2 min. 10. Anneal the selected R N A with 20 pmoles of 3' primer in a reaction mixture containing 100 mM Tris-HC1 (pH 8.3), 10 mM mgC12, 10 mM DTT, and 500/xM deoxynucleotide triphosphates (dNTPs) by heating to 70~ for 10 min followed by a 5-min incubation on ice. 11. Add 2.5 U of avian myleoblastosis virus (AMV) reverse transcriptase (USB, Cleveland, OH) to the reaction and incubate at 37~ for 30 min. 12. Extract the reaction with phenol/chloroform/isoamyl alcohol, ethanol precipitate, and resuspend the cDNA in 20/xl RiO. 13. Add the cDNA to a 0.5-ml microfuge tube containing 50 pmol of 5' P C R primer and 50 pmol of 3' P C R primer, 200/xM dNTPs, Amplitaq buffer, and 1 U of Taq DNA polymerase. Amplify for 40 cycles. 14. Transcribe the P C R product with SP6 R N A polymerase and gel purify the R N A product. This new pool, enriched for RNAs that were efficiently polyadenylated, may then be used in the next round of selection.

The stringency of selection may be increased during subsequent rounds by reducing the amount of CPSF in the reaction or by increasing the KC1 concentration of the polyadenylation reaction. Since the actual selection is for RNAs that are retained on oligo(dT)beads, RNAs will be enriched that bind the beads by base-pairing as well as non-base-pairing interactions. To eliminate those RNAs

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from the pool that bind to the beads by non-base-pairing interactions, polyadenylated RNAs are separated from the nonpolyadenylated species by denaturing gel electrophoresis prior to the cloning of the selected RNAs. The polyadenylated RNAs are eluted from the gel and subjected to RT-PCR. The PCR products are then cloned and sequenced.

VIII.

CONCLUSIONS

The ability to carry out 3' cleavage and polyadenylation in vitro has provided the basis for the investigation of the biochemistry of 3' processing. Although the basic components are now reasonably well established, we are only beginning to understand the mechanism by which poly(A) sites are selected within the context of a complex pre-mRNA. It has become increasingly clear, however, that 3' processing is influenced by other R N A processing events, particularly splicing. A system for the coupling of splicing and polyadenylation has been developed by Niwa et al. (1990) to examine this interaction. Future progress in the biochemistry and regulation of 3' processing will certainly require an understanding of the interactions of the polyadenylation complex with other components of the mRNA processing machinery. These studies, along with the use of in vitro selection, may provide an insight into the enigma posed by a processing reaction that utilizes a very limited set of R N A sequences to direct the assembly of a very large protein complex to execute an apparently simple endonucleolytic cleavage and poly(A) addition reaction.

REFERENCES Conway, L., and Wickens, M. (1987). Analysis o f m R N A 3' end formation by modification interference: The only modifications which prevent processing lie in AAUAAA and the poly(A) site. E M B O J. 6, 4177-4184. Cooke, C., and Alwine, J. C. (1996). The cap and 3' splice site similarly affect polyadenylation efficiency. Mol. Cell. Biol. 16, 2579-2584. Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983). Accurate transcription by R N A polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11, 1475-1489. Gilmartin, G. M., McDevitt, M. A., and Nevins, J. R. (1988). Multiple factors are required for specific R N A cleavage at a poly(A) addition site. Genes Dev. 2, 578-587. Gilmartin, G. M., Fleming, E. S., and Oetjen, J. (1992). Activation ofHIV-1 pre-mRNA 3' processing in vitro requires both an upstream element and TAR. E M B O J . 11, 4419-4428. Gilmartin, G. M., Fleming, E. S., Oetjen, J., and Gravely, B. R. (1995). CPSF recognition of an HIV1 m R N A 3'-processing enhancer: Multiple sequence contacts involved in poly(A) site definition. Genes Dev. 9, 72-83. Graveley, B. R., Fleming, E. S., and Gilmartin, G. M. (1996). R N A structure is a critical determinant of poly(A) site recognition by cleavage and polyadenylation specificity factor. Mol. Cell. Biol. 16, 4942-4951.

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Graveley, B. IL., Fleming, E. S., and Gilmartin, G. M. (1997). Restoration of both structure and function to a defective poly(A) site by in vitro selection. J. Biol. Chem. 271, 33654-33663. Hart, R. P., McDevitt, M. A., and Nevins, J. Ik. (1985). Poly(A) site cleavage in a HeLa nuclear extract is dependent on downstream sequences. Cell (Cambridge, Mass.) 43, 677-683. Huang, Z., and Szostak, J. W. (1996). A simple method for 3'-end labeling of RNA. Nucleic Acids Res. 24, 4360-4361. Lingner, J., and Keller, W. (1993). 3'-End labeling of ILNA with recombinant poly(A) polymerase. Nucleic Acids Res. 21, 2917-2920. Manley, J. L., Yu, H., and Ryner, L. (1985). R N A sequence containing hexanucleotide AAUAAA directs efficient mlLNA polyadenylation in vitro. Mol. Cell. Biol. 5, 373-379. Moore, C. L., and Sharp, P. A. (1985). Accurate cleavage and polyadenylation of exogenous R N A substrate. Cell (Cambridge, Mass.) 41, 845-855. Moore, M. J., and Sharp, P. A. (1992). Site-specific modification of pre-mtkNA: The 2'-hydroxyl groups at the splice sites. Science 256, 992-997. Moore, C. L., Skolnik-David, H., and Sharp, P. A. (1986). Analysis of R N A cleavage at the adenovirus 2 L3 polyadenylation site. EMBOJ. 5, 1929-1938. Niwa, M., Rose, S. D., and Berget, S. M. (1990). In vitro polyadenylation is stimulated by the presence of an upstream intron. Genes Deu. 4, 1552-1559. Ikuegsegger, U., Beyer, K., and Keller, W. (1996). Purification and characterization of human cleavage factor I-m, involved in the 3' processing of messenger R N A precursors.J. Biol. Chem. 271, 61076113. Sperry, A. O., and Berget, S. M. (1986). In vitro cleavage of the simian virus early polyadenylation site adjacent to a required downstream TG sequence. Mol. Cell Biol. 6, 4734-4741. Wahle, E. (1995). 3'-end cleavage and polyadenylation of mlLNA precursors. Biochim. Biophys. Acta 1261, 183-194. WaNe, E., Martin, G., Schlitz, E., and Keller, W. (1991). Isolation and expression of cDNA clones encoding mammalian poly(A) polymerase. EMBOJ. 10, 4251-4257. Zarkower, D., Stephenson, Ik., Sheets, M., and Wickens, M. (1986). The AAUAAA sequence is required both for cleavage and polyadenylation of simian virus 40 per-mlkNA in vitro. EMBOJ. 6, 177-186.

Rapid Identification and Cloning of Sequence-Specific RNA Binding Proteins Jeffrey Wilusz Department of Microbiology and Molecular Genetics UMDNJ-New Jersey Medical School Newark, New Jersey 07103

The identification and characterization of sequence-specific R N A binding proteins can be a key step toward elucidating mechanisms of regulation ofpostranscriptional processes. In this chapter, a UV cross-linking approach to identify sequence-specific R N A binding proteins in in vitro reconstitution assays is presented in detail, followed by an overview of approaches used to identify the protein binding site on the transcript. Finally, a Northwestern screening approach for the rapid identification ofcDNA clones expressing the R N A binding protein of interest is discussed. In summary, these methodologies provide a reasonably rapid and efficient strategy for the identification and molecular characterization of R N A binding proteins.

I. I N T R O D U C T I O N A typical result in the early stages in studies of m R N A biology is the identification of a region of a transcript that plays a pivotal role in the control of a posttranscriptional process. The next key focus of the study invariably becomes determining the mechanism by which this R N A signal functions. It is likely that many R N A signals work through the sequence- or structure-specific assembly of protein-RNA complexes (i.e., Dreyfuss et al., 1996; Nagai, 1996). There are several general approaches to identifying these complexes. A genetic approach is 99 m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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excellent but is not always applicable if the biological system of interest is not easily amenable to genetic selection. Biochemical purification and reconstitution is another strategy. Aside from being relatively labor intensive, this approach requires the development or availability of an in vitro system that faithfully reproduces all aspects of the basal or regulated process of interest. Such systems are not available for all 1KNA processing events, and available systems often do not reproduce regulatory aspects of the process. A third approach, and the one that will be the focus of this chapter, is to identify proteins which specifically interact with the 1KNA domain of interest and to obtain complementary DNA (cDNA) clones which encode these proteins. The information and reagents generated by this strategy can prove to be key to the development of testable hypotheses concerning the mechanistic and regulatory aspects of 1KNA processing. The experimental strategy of identifying and cloning sequence-specific 1KNA binding proteins has several advantages. First, experimental progress is often rapid and inexpensive relative to more time-consuming biochemical fractionation and reconstitution efforts. Second, 1KNA binding assays are generally easier and much less complex than reconstitution assays based on activity measurements. Third, the identification of 1KNA binding proteins by UV cross-linking provides an approximate molecular weight for the protein. Finally, the strategy does not require a priori the development of an in vitro system that faithfully reproduces a regulated 1KNA-processing event. The experimental strategy also has its disadvantages. First, not all protein1KNA interactions are stable enough to be detected by standard methodologies. Second, this approach has the inherent risk that the identified 1KNA-protein interaction may not be functionally significant. The need to eventually determine the functional signficance of the identified protein cannot be overemphasized. Finally, the probability of success in the identification and cloning oflKNA binding proteins is very much dependent on the quality of the reagents used (the extracts, cDNA library, etc.). The methodologies presented below have been used successfully in our laboratory to identify and characterize proteins that interact with polyadenylation substrate 1KNAs in HeLa nuclear extracts (Wilusz and Shenk, 1988, 1990; Qian and Wilusz, 1991, 1994). Keep in mind that, in principle, these methods can be adjusted to accommodate different cell types, subcellular fractions, 1KNAs, and so on.

II. P R E P A R A T I O N

OF EXTRACTS

Prepare nuclear and/or cytoplasmic extracts following the detailed protocol ofDignam et al. (1983), with the following modifications/considerations:

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1. We have obtained more consistent results with extracts prepared from actively growing HeLa $3 cells rather than frozen cell pellets. 2. Reducing dialysis times to 1 hr also results in more consistent extracts. 3. The addition of protease inhibitors, especially in all buffers used at and beyond the nuclear salt wash extraction step, can be beneficial. Nuclear extracts prepared by this procedure should be dispersed into small aliquots and can be stored for long periods of time (>2 years) at -80~

III. P R E P A R A T I O N

OF R N A s

RNAs are prepared by runoff transcription using phage polymerases as described (Melton et al., 1984), with the following modifications/considerations: 1. Adjust the relative ratios of hot to cold nucleotides in the transcription reaction so that 1 out of every 10 uridine triphosphates (UTP) [or another nucleotide triphosphate (NTP) of choice] is contributed by the radiolabeled reagent. This gives RNAs of sufficient specific activity to see the results of the experiments described below in overnight exposures to X-ray film. 2. It is recommended that all RNAs be gel purified prior to use to allow the purification of the desired transcript away from the smaller products and larger "runaround" transcription products commonly generated by phage polymerases. Ensuring that the R N A probes used in these assays are of sufficient quality is important to generate consistent results and makes troubleshooting much easier in the long run. 3. After localization of the R N A on a 5% acrylamide gel containing 7 M urea with the use of a brief (30-sec) exposure to film, KNA bands are excised with a clean razor blade and eluted overnight at room temperature in 500/xl of a buffer containing 25 mM Tris (pH 7.6), 400 mM NaC1, and 0.1% sodium dodecyl sulfate (SDS). Proteinase K may be added to this mixture (2/zl of a 10-mg/ml solution) to remove any low levels of ribonucleases that may be present. Approximately 50-75% of the R N A will elute from the gel during this incubation. The R N A is then further purified by phenolchloroform-isoamyl alcohol extraction and concentration with ethanol. 4. The molar amounts of R N A probes can be determined by scintillation counting of a small portion, taking into account the specific activity of the radiolabel, the total number of uridylates (or other bases, depending on the radiolabel used), and the relative ratio of radiolabeled to cold nucleotide.

Jeffrey Wilusz

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IV.

IDENTIFICATION

PROTEIN-RNA

OF

COMPLEXES

The following reconstitution methodology is based on identifying proteins associated with polyadenylation substrate vitro 3' end processing system originally described by Moore The principles of the approach described below, however, are types of extracts and RNAs.

our experience in RNAs using an in and Sharp (1985). applicable to many

A. P R O T E I N - R N A RECONSTITUTION In a microcentrifuge tube, add 3.25 ~1 of 10% polyvinyl alcohol [Sigma (St. Louis, MO) cold water soluble], 1/zl of a 12.5 m M A T P / 2 0 0 mMphosphocreatine solution, 1 /~1 of radiolabeled R N A (200,000 cpm; approximately 10-50 fmol), and 7.5 /~1 of nuclear salt wash extract prepared as described above. Reagents should be gently mixed with the pipettor. Incubate the mixture at 30~ for 5 to 10 rain. Notes: The use of ATP is optional but may be required in the assembly of some protein-RNA complexes. One/~1 of a 10 m M solution of ethylenediaminetetraacetic acid (EDTA) may improve the stability of unstable RNAs in the extracts due to the presence of magnesium-dependent exonucleases (Ford et al., 1997). Incubation of RNAs in nuclear extracts at higher temperatures (37~ often results in unacceptable levels of nonspecific nuclease activity.

B. ~

CROSS-LINKING R N A - P R O T E I N COMPLEXES

Transfer the reaction mixture from the microfuge tube to a single well of a standard 96-well microtiter dish. Place the dish on ice and position it directly under a 15-W germicidal light (we routinely use a Sylvania G18T8 bulb in a standard fluorescent fixture available from many hardware stores). The distance between the sample and the light should be approximately 4 cm. Irradiate the sample for 10 min on ice. Notes: During irradiation, the R N A bases absorb UV light and form very short-lived flee radical intermediates (Smith, 1976). Only amino acid residues that are very closely associated with the R N A bases will therefore become cross-linked. In the few cases in which the site of cross-linking on the protein has been assayed, phenylalanine residues located in R N A binding domains appear to be favorite crosslinking targets (Merrill et al., 1988). Due to this requirement for close proximity to the R N A bases, only a subset of R N A binding proteins can be efficiently crosslinked to the transcript. The use of alternative binding assays, such as gel shifts,

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should be explored if UV cross-linking fails to give satisfactory results. The 10minute dose of irradiation was determined in time course studies which assessed the approximate time of maximal cross-linking at the peak of linear kinetics. It is recommended that a similar time course of sample irradiation be used for all studies to ascertain that protein-P.NA cross-linking is occurring with linear kinetics and no significant lag period.

C. ANALYSIS OF U V CROSS-LINKED PROTEIN-RNA COMPLEXES Transfer the irradiated sample from the well to a microfuge tube. Add 1/a,1 IkNase A (10 mg/ml) and incubate at 37~ for 15 min. Add 13.5/xl of a standard 2 • protein sample loading dye (containing SDS and fl-mercaptoethanol) and place the reaction mixture at 100~ for 3 rain. Precipitated proteins in the sample are then removed by a brief (1- to 2-min) spin in a microfuge. Samples are then analyzed on a standard 10% acrylamide/SDS gel and visualized by autoradiography following a 12- to 18-hr exposure to X-ray film.

Notes: The above procedure will generate a protein containing a short piece of radiolabeled tLNA which will normally migrate within 1-2 kda of its apparent molecular weight. RNase T1 or a mixture of enzymes can be substituted for IkNase A in the above protocol if the ILNA binding sequence is relatively rich in purines. The amount and specific activity of ILNA in the reaction mixture have been optimized to allow analysis of the data following a convenient overnight exposure. Film exposure times may be adjusted to take into account any changes in these parameters. This protocol will allow the identification oflKNA binding proteins, as well as the determination of their approximate molecular weight. UV cross-linking analysis is also sufficiently quantitative to be used in competition studies to determine the specificity of the interaction you have identified. In addition, the requirement for monovalent and divalent cations, ATP, and other cofactors for binding can be readily assessed with this methodology.

V. D E T E R M I N A T I O N OF R N A S E Q U E N C E REQUIREMENTS FOR PROTEIN BINDING There are numerous strategies to identify the sequence requirements for specific interaction of proteins with RNAs. These include (1) deletion and substitution mutagenesis, (2) RNase H mapping, and (3) site-specific radiolabeling/UV

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cross-linking. The combined use of two or more of these strategies is often helpful. A summary of these strategies is presented below, and the reader is referred to specific references for more information and specific methodologies.

A. DELETION AND SUBSTITUTION MUTAGENESIS The construction of site-directed mutations using recombinant technology is a powerful tool to determine the influence of sequence elements on R N A - p r o t e i n interactions. One should first consider delimiting the approximate 3' limit of the R N A element by the relatively easy generation of a series of 3' deletion derivatives with the use of different restriction enzymes to prepare templates for runoff transcription reactions. Additional internal mutations can then be constructed through traditional cloning approaches (Sambrook et al., 1989) or polymerase chain reaction (PCR) approaches (Innis et al., 1995). Internal mutations, for example, can be readily generated using a megaprimer P C R approach, as described previously (Aiyar and Leis, 1993).

B. RNASE H MAPPING The site of protein cross-linking on the input R N A molecule can be approximated by this approach which was first applied by MacDonald et al. (1994). One limitation of this technique, however, is that it requires an available antibody to immunoprecipitate the cross-linked protein-RNA complex. Following isolation of the protein-RNA complex, complexes are incubated with DNA oligonucleotides specific for various regions of the RNA. RNase H is then added to cleave the 1LNA specifically at the sites of oligo hybridization. The 5' or 3' RNase H cleavage products are then analyzed in an acrylamide gel to determine which will migrate aberrantly due to the presence of cross-linked protein. This allows the location of the protein cross-linking site to be placed 5' or 3' to the site of oligo hybridization.

C. SITE-SPECIFIC R N A LABELING R N A binding proteins are visualized in UV cross-linking analysis due to the transfer of radiolabel from the transcript to the protein by covalent attachment of small fragments of RNA. The placement of a radiolabel at specific sites on the transcript, therefore, will allow the localization of that residue to the proximity of the R N A binding element. The bridge ligation method of Moore and Sharp (1992) allows the site-specific labeling of RNAs at selected internal positions.

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VI. CLONING cDNAs FOR RNA BINDING PROTEINS BY NORTHWESTERN SCREENING The molecular characterization of R N A binding proteins is a pivotal next step in their characterization and in the development of future reagents for investigations into their mechanism of action. The goal of this section is to describe an approach for the rapid cloning of cDNAs for R N A binding proteins without the need for labor-intensive biochemical purification. Many R N A binding proteins interact with their substrates while immobilized on solid support matrices such as nitrocellulose. The detection of immobilized R N A binding proteins with the use of an R N A probe is called Northwestern screening (Qian and Wilusz, 1993). For successful Northwestern analysis, several factors must be taken into account. First, the equilibrium and kinetic binding constants of the RNA-protein complex must be relatively high (10 9 M -1) to retain a stable association during the wash protocol. Second, a minimal R N A binding element should be used in the probe to reduce the number of false positives obtained during screening. R.NA probes containing appropriately spaced multimers of the minimal binding element can be especially useful in this strategy. Third, the conformation of the R N A binding domain must be maintained in the immobilized protein. Specific renaturation protocols have been developed to address this issue (Vinson et al., 1988). Fourth, the integrity of the R N A probe must be maintained throughout the procedure. It is recommended that probes be gel purified, as described above, to ensure their integrity prior to screening, as well as assayed following their incubation with the colony lifts to ensure that probe integrity is generally maintained throughout the procedure. Fifth, if any selectable properties o f R N A - p r o t e i n interaction are noted in preliminary studies (e.g., salt resistance), they should be incorporated into the wash protocol to increase the selectivity of the screen. Finally, the cDNA library use should be of sufficient quality to allow the detection of low-abundance clones.

A. PREPARATION OF FILTER LIFTS FROM LAMBDA eDNA LIBRAR~S

1. Preparation of plating bacteria. Inoculate a single colony of the plating bacteria appropriate for the cDNA library to be used in 50 ml of LB broth (10 g tryptone, 10 g NaC1, and 5 g yeast extract per liter) containing 50/zg/ml ampicillin and 0.2% maltose. Allow the culture to grow to saturation overnight at 37~ Collect the cells by pelleting at 3000 g for 10 min and resuspend the cells in one half of their original volume with 10 m M MgSO4.

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JeffreyWilusz 2. Plating the library. Infect 1 ml of bacterial suspension with an aliquot of 50,000 phage from the library. Incubate the infection at 37~ for 20 rain to allow the phage to efficiently absorb to the cells. Add 9 ml of top agarose (LB broth plus 10 m M MgSO4 and 0.6% agarose) prewarmed to 45~ to the suspension. Mix the sample by inverting the tube several times and quickly pour the sample onto LB-agar plates (150 X 15 ram). Let the top agarose solidify at room temperature for 5-10 rain. Invert the plates and incubate them at 37~ for 3.5-4 hr until pinpoint plaques appear. 3. Induction of recombinant proteins. Soak the necessary number of nitrocellulose filters (137 mm BA-S85 from Schleicher & Schuell, Keene, NH) in a 10 m M solution of isopropyl-/~-thiogalactopyranoside (IPTG) for 30 min in order to impregnate them with the inducer. Allow the filters to air dry, and number them with a ballpoint pen to facilitate identification following screening. Carefully overlay the filters onto the plates containing visible pinpoint plaques. Incubate the plates in an upright position for approximately 4 hr at 37~ to allow induction of recombinant proteins and their transfer to the nitrocellulose filter following phage-induced cell lysis. Prior to removing the filters from the plates, mark their relative positions on the lawn with several asymmetrical sticks with an 18-gauge needle. Allow the filter lifts to air dry for 10 min.

B. NORTHWESTERN SCREENING OF FILTER LIFTS

1. Blocking unoccupied regions of the filter. Incubate the filters in blocking buffer [10 m M Tris-C1 (pH 7.8), 150 m M NaC1, 5 mg/ml Torula yeast core R N A (Sigma, St. Louis, MO)] at 37~ for 1 hr with mild shaking. Use approximately 100 ml of blocking buffer per dozen filters in a 125 • 65-ram crystallizing dish (Fisher Scientific, Pittsburgh, PA). 2. Optional protein renaturation step. An increase in the binding of nucleic acids to immobilized proteins has been noted to occur in some instances if the transferred proteins are first denatured in 6 M guanidine-HC1 and then allowed to slowly renature. Immobilized proteins are treated sequentially with 10-rain washes of 6 M, 3 M, 1.5 M, 0.75 M, 0.375 M, 0.187 M, and 0 M guanidine-HC1 in a buffer containing 20 m M N-2-hydroxyethylpiperazine-N'-2ethanesulfonic acid-potassium hydroxide (HEPES-KOH pH 7.5), 25 m M NaC1, and 1 m M dithiothreitol (DTT). 3. Washing. Wash the filters three times for 5 rain each in 10 m M TrisC1 (pH 7.8) to remove all traces of blocking buffer. Use approximately

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5.

6.

7.

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100 ml of wash buffer per dozen filters in a crystallizing dish or Pyrex baking dish. Binding of R N A probe to immobilized proteins. Incubate the filters for 1 hr at room temperature with the 1LNA probe at a concentration of 5 X 10 s cpm per milliliter in Northwestern (NW) buffer containing 10 m M Tris-C1 (pH 7.8), 1 mm EDTA, 50 m M NaC1, 0.02% Ficoll, 0.02% polyvinyl pyrrolidone, and 0.02% bovine serum albumin (Boehringer Mannheim, Indianapolis, IN, ultrapure grade). Use approximately 70 ml of solution per dozen filters in a 125 X 65-mm crystallizing dish. It is highly recommended that 400/xl of the probe solution be saved following the incubation in order to check the integrity of the probe on a denaturing gel to aid in troubleshooting the assay. Removal of unbound RNA. Excess 1KNA probe is removed from the filter by four consecutive 10-min washes at room temperature with 100 ml of N W buffer. If conditions are found (e.g., high salt) that can be used to increase the selectivity of the assay for the protein-lKNA interaction of interest, they should be incorporated into the wash buffer at this step. Identification of candidate-positive plaques. Allow the filters to air dry, tape them to Whatman 3MM paper, cover the filters with plastic wrap, and expose them to X-ray film overnight with an intensifying screen. The filters may be arranged asymmetrically or marks may be made on the Whatman paper to facilitate alignment of the film to the filter for the localization of positive clones. A 16-hr exposure is generally sufficient to detect positive signals. Longer times may be necessary if 1LNA probes have been significantly degraded during the incubation procedure. If more than 50% of the probe has been degraded during incubation, a poor signal-to-noise ratio may be observed. Using the asymmetrical needle marks and the numbers previously marked on the filters, align candidate-positive signals with the corresponding regions of the phage lawn. Core candidate-positive regions of the lawn with the large end of a Pasteur pipette. Place the agar plug in 1 ml of SM buffer [5.8 g NaC1, 2 g MgSO4, 50 ml 1 M Tris-C1 (pH 7.8), and 0.1% gelatin per liter] and allow the phage to elute overnight at 4~ One drop of chloroform should be added to samples to inhibit bacterial growth. Following elution, 1 /xl of buffer should contain 5000-10,000 phage particles. Verification of candidate-positive phage. Repeat the Northwestern screening assay described above using the candidate-positive phage. Approximately 1000 phage should be mixed with 200/xl of bacteria and plated onto 100 X 15-mm LB agar plates. Following the appearance of plaques, overlay the lawns with 87-mm nitrocellulose

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filters and perform the screening procedure as described above. The amount of buffers used in incubations and washes should, of course, be adjusted to appropriate volumes for the number of filters used in this secondary screening. A true-positive result should now give 20-100 positive signals per filter and is therefore readily differentiated from spurious spots. 8. Plaque purification of positive phage. Select positive phage samples and repeat the screening process a third time using 50-300 phage per 100 X 15-mm plate. Following screening, select a well-isolated positive plaque for coting. Elute the phage as described above. The recombinant phage should now be pure and ready for further analysis with the use of standard methodologies (Sambrook et al., 1989).

ACKNOWLEDGMENTS I wish to thank all members of the laboratory, especially Drs. Zuwei Qian and Paramjeet S. Bagga, for their input. This work was supported by grants from the NIH and the New Jersey Cancer Commission.

REFERENCES Aiyar, A., and Leis, J. (1993). Modification of the megaprimer method o f P C R mutagenesis: Improved amplification of final product. BioTechniques 14, 366-369. Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983). Accurate transcription initiation by R N A polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res. 11, 1475-1489. Dreyfus, G., Hentze, M., and Lamond, A. I. (1996). From transcript to protein. Cell (Cambridge, Mass.) 85, 963-972. Ford, L. F., Bagga, P. S., and Wilusz, J. (1997). The poly(A) tail inhibits the assembly of a 3' to 5' exonuclease in an in vitro R N A stability system. Mol. Cell. Biol. 17, 398-406. Innis, M. A., Gelfard, D. H., and Sninsky, J. (1995). " P C R Strategies." Academic Press, San Diego, CA. MacDonald, C. C., Wilusz, J., and Shenk, T. (1994). The 64 kilodalton subunit ofCstF polyadenylation factor binds to pre-mRNAs downstream of the cleavage site and influences cleavage site location. Mol. Cell. Biol. 14, 6647-6654. Melton, D. A., Krieg, P. A., Rebagliati, M. R., Maniatis, T., Zinn, K., and Green, M. R. (1984). Efficient in vitro synthesis of biologically active R N A and R N A hybridization probes from plasmids containing a bacteriophage SP6 promoter. Nucleic Acids Res. 12, 7035-7056. Merrill, B. M., Stone, K. L., Cobianchi, F., Wilson, S. H., and Williams, K. R. (1988). Phenylalanines that are conserved among several RNA-binding proteins from part of a nucleic acid-binding pocket in the A1 heterogeneous nuclear ribonucleoprotein..J. Biol. Chem. 263, 3307-3313. Moore, C. L., and Sharp, P. A. (1985). Accurate cleavage and polyadenylation of exogenous R N A substrates. Cell (Cambridge, Mass.) 36, 581-591. Moore, M. M., and Sharp, P. A. (1992). Site-specific modification of pre-mRNA: The 2' hydroxyl groups at splice sites. Science 256, 992-997. Nagai, K. (1996). RNA-protein interactions. Curt. Biol. 6, 53-61.

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Qian, z., and Wilusz, J. (1991). An RNA-binding protein specifically interacts with a functionally important domain of the downstream element of the SV40 late polyadenylation signal. Mol. Cell. Biol. 11, 5312-5320. Qian, Z., and Wilusz, J. (1993). Cloning of a cDNA encoding an RNA binding protein by screening expression libraries using a northwestern strategy. Anal. Biochem. 212, 547-554. Qian, Z., and Wilusz, J. (1994). GRSF-I: A poly(A) + mRNA binding protein which interacts with a conserved G-rich element. Nudeic Acids Res. 22, 2334-2343. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). "Molecular Cloning: A Laboratory Manual," 2nd ed. Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. Smith, K. C. (1976). The radiation-induced addition of proteins and other molecules to nucleic acids. In "Photochemistry and Photobiology of Nucleic Acids" (S. Y. Wang, ed.), Vol. 2, pp. 187-218. Academic Press, New York. Vinson, C. R., LaMarco, K. L., Johnson, P. F., Landschulz, W. H., and McKnight, S. L. (1988). In situ detection of sequence-specific DNA binding activity specified by a recombinant bacteriophage. Genes Dev. 2, 801-806. Wilusz, J., and Shenk, T. (1988). A 64 kd nuclear protein binds to RNA segments that include the AAUAAA polyadenylation motif. Cell (Cambridge, Mass.) 52, 221-228. Wilusz, J., and Shenk, T. (1990). A uridylate tract mediates efficient hntLNP C protein-RNA cross linking and functionally substitutes for the downstream element of the polyadenylation signal. Mol. Cell. Biol. 10, 6397-6407.

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Analysis of Polyadenylation Phenotypes in Saccharornyces cerevisiae Michael W. Briggs Department of Microbiology and Immunology University of Rochester School of Medicine and Dentistry Rochester, New York 14642

j. Scott Butler Department of Microbiology and Immunology University of Rochester School of Medicine and Dentistry Rochester, New York 14642

Aaron Proweller Department of Microbiology and Immunology University of Rochester School of Medicine and Dentistry Rochester, New York 14642

I. I N T R O D U C T I O N The past 10 years have witnessed a vast expansion of our knowledge of how m R N A 3' untranslated regions (UTRs) function to signal the position and efficiency of the posttranscriptional cleavage and polyadenylation reactions that form most mature messenger R N A (mRNA) 3' ends in eukaryotes. Experiments directed at understanding structure-function relationships in mRNAs have also begun to shed light on the roles that 3' UTRs and poly(A) tails play in influencing m R N A stability, translation efficiency, and perhaps rates of nuclear export. Many of these experiments rely on genetic approaches in Saccharomyces cerevisiae that have as their goal the identification of gene products that influence the role that m R N A 3' ends play in each of these processes. The identification of genes by classical genetic selections and screens, as well as with screens for synthetic lethality and protein-protein interactions, has provided evidence indicating direct roles for gene products in m R N A 3' end processing, m R N A stability, translation, and nuclear export. However, the interactions between all of these pathways often make it difficult to determine whether a specific mutation affects a particular pathway directly or indirectly. Thus, when trying to assign a specific biological role to a newly identified gene product, it is necessary to employ a battery of techniques to determine where in a pathway the new 111 m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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protein acts. This is particularly important in the study of mutations suspected to affect m R N A 3' end processing (cleavage and polyadenylation) in S. cerevisiae. Several cautionary tales underscore how difficult it can be to assign a direct role for a gene product in the 3' end processing pathway. In the cases of the R N A 1 and P R P 2 0 genes, strains with conditional mutations yielded mR.NAs with abnormally long m R N A 3' ends, unusually long poly(A) tails, and defects in nuclear export of n ~ N A s (1,2). These mutant strains, however, yielded mlLNA 3' end processing extracts with slightly diminished but otherwise normal processing activities, indicating that the phenotypes in vivo do not result from direct defects in the processing enzymes (2). In the case of mutations in R N A 14 and R N A 1 5 , initial experiments showed that A C T 1 n ~ N A and total poly(A) decayed with half-lives shorter than those seen in normal cells, which led to the conclusion that these genes encode components of the m_R.NA turnover pathway (3). However, synthetic lethality experiments and measurements of n ~ N A 3' end processing in vitro clearly showed that these enzymes are central components of the cleavage and polyadenylation pathway (4). Finally, an in vitro screen of a library of temperaturesensitive mutants that identified the poly(A) polymerase mutation pap 1-1 encountered additional strains with defective processing activities in vitro but with no apparent polyadenylation defects in vivo (5). In each of these cases, the true nature of these mutations became clear only after several techniques focusing on 3' end formation in vivo and in vitro were applied to the problem. Below we describe methods useful for analyzing m R N A 3' end formation in vivo and in vitro. More standard procedures for isolating yeast tLNA, poly(A) + ILNA fractionation and Northern blot hybridization are described elsewhere (5).

II. P O L Y ( A )

TAIL

ANALYSIS

This protocol describes how to label the 3' ends of 1LNA with cytidine 3',5'-[5'-32p]bisphosphate and ILNA ligase, followed by hydrolysis of the nonpoly(A) portions of the RNA. The remaining labeled poly(A) stretches are separated by electrophoresis and visualized by autoradiography. The method was first developed by Ahlquist and Kaesberg (6) and was later used by Sachs and Davis (7) to demonstrate changes in poly(A) tail lengths in yeast strains deficient in poly(A)binding protein activity. In yeast, poly(A) tails are rapidly synthesized to a length of 60-80 nucleotides and are trimmed one nucleotide at a time to a length of about 10 nucleotides before the mR.NA is degraded. The cytidine 3',5'-bisphosphate (pCp) labeling method allows the display of all of the intermediates in this process, thus facilitating an assessment of which step may be affected in any given mutant. Moreover, if care is exercised during the labeling, the assay will yield a quantitative picture of how much poly(A) exists in the cell at any given time.

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A. PREPARATION OF CYTIDINE 3',5'-[5'-32pIBISPHOSPHATE Cytidine 3',5'-[5'-32p]bisphosphate is comercially available (Amersham); however, it is quite expensive. Fortunately, it can be prepared quickly and reliably, without any special reagents or workup of the product. The reaction features the transfer of32p from 5' [T-32p]ATP to cytidine 3'-monophosphate, catalyzed by T4 polynucleotide kinase. Since this enzyme has a 3' phosphatase activity, it is necessary to use a phosphatase-free preparation available from Boehringer Mannheim to ensure that the product retains the 3' phosphate required by 1KNA ligase.

1. Reagents 5• T4 Kinase Buffer 250 mM Tris-HC1, pH 7.5 50 mM MgC12 25 mM Dithiothreitol (DTT) 10/xM Cytidine 3'-Monophosphate Sigma Chemicals (St. Louis, MO)

T4 Polynucleotide Kinase (3' Phosphatase Free) Boehringer Mannheim (Indianapolis, IN) 10 U//xl 5' [T-32p]ATP Dupont NEN, Wilmington, DE; 6000 Ci/mmol; 10 mCi/ml

1. Transfer 20/xl (200/xCi) of 5'D,-32p]ATP to a clean, sterile microfuge tube and spin in a Speed-Vac under a vacuum until dry. 2. Add to the dry 5'[T-32p]ATP the following: 10/xl sterile, distilled water 3/xl 5• T4 polynucleotide kinase buffer 1/xl cytidine 3'-monophosphate (3'-CMP) (10/xM) 1 /xl T4 polynucleotide kinase (10 U//xl) 3. Incubate at 37~ for 4 hr. 4. Heat at 90~ for 5 min to inactivate the polynucleotide kinase. 5. Dilute to 20/xl with sterile, distilled water. The phosphorylation of 3'-CMP with polynucleotide kinase typically results in the transfer of more than 80% of 32p from ATP to 3'-CMP. The yield of this reaction can be determined by thin layer chromatography if knowledge of the specific activity of the product is desired.

B. POLY(A) TAIL LABELING AND ANALYSIS In this step cytidine 3',5'-[5'-32p]bisphosphate is ligated to the 3' ends of all of the 1KNAs in a sample of total 1KNA. Since only about 1-2% of total yeast

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R N A is mRNA, there is a large excess of carrier ribosomal R N A (rRNA) and transfer R N A (tRNA) in every reaction. This means that even large differences in the amount of mRNA from sample to sample will have little effect on the overall reaction rate. Thus, one can compare the effects of conditions predicted to change mRNA levels without worrying about their effects on labeling efficiency. The sacrifice in specific activity made by labeling total R N A rather than poly(A)+RNA is, in practice, of little consequence, as the poly(A) tail ladders are visible within hours after separation on thin polyacrylamide gels.

1. Reagents 2 • Ligase Buffer 100 mM HEPES, pH 8.3 20 mM MgC12 25% Glycerol

T4 R N A Ligase New England Biolabs (Beverly, MA); 10 U//,1 Ligase Reaction Mix (per reaction) 0.5/.d R N A ligase (10 U//.tl) 0.5/xl RNasin (40 U//.d) 0.5/.d Cytidine 3',5'-[5'32p]bisphosphate (10/xCi//2,1) 1.2/xl Ligase stock Proteinase K Stop Solution (per reaction) 6/xl 0.2 M Na2EDTA 2/xl 10% SDS 2/xl Proteinase K (20 mg/ml)

Ligase Stock (0.1 ml) 50/2,1 2• Ligase buffer 1/.d 2.5 mM rATP 12.5/.d Glycerol 12.5/,1 DMSO 24/,1 H20 RNasin Promega (Madison, WI); 10 U//xl Stop Solution 0.3 M NaC1 0.01 M Tris-HC1, pH 8 0.01 M Na2EDTA

12% Acrylamide Gel (80 ml) 33.6 g Urea 9.6 g Acrylamide 0.5 g Bisacrylamide 4 ml 10X TBE (500 mM Tris-borate, pH 8.3; 10 mM EDTA) 250/zl 10% Ammonium persulfate 15/zl TEMED

1. Heat a sample of total R N A (1 btg//-d) at 65~ for 10 min.

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2. Add 1/zl (1 /zg//zl) of the R N A to 3.0/zl of Ligase Reaction Mix by drawing the solution into and out of the pipette tip. 3. Incubate the mixture at 4~ for 16 hr. 4. Stop the reaction by adding 21/zl Stop Solution. 5. Determine the efficiency of ligation for each sample by spotting 1/zl on a 2-cm Whatman DE81 filter, washing at room temperature twice for 5 rain in 0.5 M Na2HPO4 (5 M/filter), once in water for 1 min, and once in ethanol for 1 rain. The filters are then allowed to dry, and radioactivity is determined by scintillation counting. If care is taken to add equal amounts of R N A to each sample, the specific activities of the labeled RNAs usually vary by less than 20% and are typically in the range of 1-5 X 10 6 cpm//zg.

2. RNase Digestion and Gel Electrophoretic Analysis 1. To 20/~1 of labeled R N A add: 40/zl 2X RNase Solution (1 M NaC1; 40 m M Tris-HC1, pH 8; 2 mM MgC12; 1 mg/ml tRNA) 1 /zl 10 mg/ml RNase A 0.2/zl RNase T1 (USB, Cleveland, OH, 475 U//zl) 39/zl H20 2. 3. 4. 5. 6. 7. 8. 9.

Incubate for 3 hr at 37~ Add 10/zl Proteinase K Stop Solution. Incubate at 37~ for 15 rains. Extract the mixture once with phenol: chloroform: isoamyl alcohol (48:1:1). Add 200/zl ethanol, incubate at - 2 0 ~ for at least 30 min, and collect the pellet by microcentrifugation at 13,000 rpm for 10 min. Wash the wet pellet with 200/zl 70% ethanol and dry in a Speed Vac. Resuspend the pellet in 10/zl FBX tracking dye (80% formamide, 0.1% xylene cyanol, 0.1% bromophenol blue). Heat for 3 rain at 100~ Load 4/.~1 on a 12% acrylamide, 0.5 X TBE gel. Run the gel at 2000 V until the bromophenol blue dye has migrated 30 cm. Expose the gel to Kodak X-Omat A R film for 4 48 hr at - 7 0 ~ or to a Molecular Dynamics storage phosphor screen for 1-5 hr.

Figure 1 shows a typical poly(A) tail gel from a normal strain and one carrying the temperature-sensitive pap1-1 mutation before and after shift of the cells to 35~ Although the specific activities of the two pap1-1 samples were each 5 X 10 6 cpm//~g, it is clear that there is far less poly(A) in pap1-1 cells 30 min after inactivation of poly(A) polymerase. More detailed analysis using this method

F i g u r e 1 Poly(A) tail distribution in a PAP1 strain and a pap 1-1 strain after shift of the cells to 35~ for 30 mins. The strains A364A (PAP1) and UR3148-1B (pap1-1) were grown in YPD at 25~ to an A600 of 0.2-0.5, cells were collected from half of each culture, and the remaining cells were incubated at 35~ for 30 min before being collected. Total R N A was prepared from each sample, and poly(A) tails were labeled as described in the text. The values to the right of the figure indicate the lengths of radiolabeled D N A molecular length markers.

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showed that poly(A) disappears from the cell with a half-life of about 20 rain (8). Since this is the average half-life of m R N A in yeast, the results supported the conclusion that the defect is in the synthesis of poly(A) rather than in poly(A) trimming. This technique is also an excellent way to determine if different conditions or mutations alter global poly(A) tail lengths (1,2). Since deadenylation occurs predominantly in the cytoplasm, mutations that slow the export of m K N A out of the nucleus may increase the proportion of mRNAs with the longest poly(A) tails, thereby giving the impression that poly(A) metabolism is altered (1,2). When interpreting changes in average steady-state poly(A) tail lengths, it is important to remember that they are influenced by the rate of m R N A decay as well as the rate of deadenylation. Thus, slow deadenylation or accelerated turnover of the cytoplasmic pool of m K N A could each result in an increase in the proportion of mRNAs with the longest poly(A) tails. Questions aimed at determining whether a mutation affects poly(A) tail synthesis or deadenylation can be approached with the use of chase experiments to measure the half-life of poly(A) tails (9). Decker and Parker (10) developed a pulse chase technique that allows one to accurately measure deadenylation rates of specific mRNAs. This technique clearly demonstrated that real4 and rna15 mutations do not alter the deadenylation rate ofPGK1 mRNAs (11).

III. P O L Y A D E N Y L A T I O N

SITE M A P P I N G B Y R T - P C R

Polymerase chain reaction (PCR) amplification of complementary DNAs (cDNAs) synthesized by reverse transcriptase (RT) primed with oligo(dT) provides a rapid and accurate method for the determination of polyadenylation sites in S. cerevisiae (12). Because of its ease and accuracy, this method has largely supplanted $1 nuclease mapping as the preferred technique for poly(A) site mapping in many laboratories (8,13,14). Moreover, comparison of results obtained with the R T P C R method with those from S1 nuclease mapping indicated that the former technique provides more reliable information regarding the bona fide polyadenylation site (8,12). The R T - P C R method is not without potential pitfalls. Yeast 3' untranslated regions are highly A and U rich and sometimes contain poly(A) stretches that act as priming sites for oligo(dT) (12). In other cases, a gene may utilize numerous poly(A) sites, resulting in R T - P C R patterns that are initially quite confusing (14). Finally, in the case of multiple poly(A) sites, the P C R may favor the amplification of some cDNAs over others if they constitute the majority of the transcripts or if they are significantly shorter than others. Thus, as with any other technique, R T - P C R determination of a poly(A) site should be supported by other methods such as RNase protection or determination of the polyadenylation site in vitw (8,12).

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A critical decision in preparing for an R T - P C R experiment is the choice of the primers. For cDNA synthesis and subsequent P C R we have found 5' TsGGATCCTI7 3' to work well (8,12). The BamH1 restriction site incorporated within oligo(dT) facilitates subsequent cloning of the P C R products and enhances the stability of the primer template hybridization during PCR. A second strategy employs an oligo(dT) primer redundant for A, C, and G at the 3' terminus (13). This guarantees that the primer will initiate synthesis at the poly(A) site rather than within the poly(A) tail itself. However, since poly(A) tails in yeast are only 60-90 nucleotides in length, random hybridization of the oligo(dT) primer within the tail will reduce the poly(A) stretch in the cDNA to the size of the primer over 30 cycles of amplification (12). The second or 5' primer should be chosen to hybridize as close to the termination codon as possible to ensure amplification of all the potential poly(A) sites. The primer should be 16 nucleotides or longer, 45-55% G + C, and may contain a restriction enzyme site at its 5' end to facilitate subsequent cloning and sequencing of the product.

1. Reagents Oligo(dT) BamH1

10 • P C R Buffer 500 m M KC1 25 m M MgC12 100 m M Tris-HC1, pH 8.4

5' TsGGATCCT17 M-MLV Reverse Transcriptase GIBCO-BRL (Gaithersburg, MD); 200 U//xl Taq Polymerase

2 mM dNTP Mixture of all four dNTPs in water, Final concentration 2 mM, pH 7

GIBCO-BRL; 2.5 U//xl

2. Reverse Transcription 1. To a sterile 0.5 ml microfuge tube add: 2/~1 10 • P C R buffer 2/~1 2 m M dNTP 1 IM Oligo(dT)-BamH1 (25 pmol//xl) 1 /~1 Total yeast R N A (1 /~g//~l) 13/xl Sterile distilled water 2. Incubate at 65~ for 5 min to denature the RNA. 3. Allow to cool to 25~

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4. Add 1/xl M-MLV reverse transcriptase (200 U//xl; GIBCO-BtLL) 5. Incubate at 37~ for 60 min.

3. P C R

1. To the tLT reaction add: 60/xl Sterile, distilled water 8 ~1 10 • PCR. buffer 8/xl 2 m M dNTP 1 Ixl Oligo(dT)-BamH1 (25 pmol//xl) 2/xl 5' Oligo primer (25 pmol//xl) 1 Ixl Taq DNA polymerase (2.5 U//xl; GIBCO-BR.L) 2. Overlay the reaction mixture with 2 drops (---20/xl) of mineral oil. 3. Carry out the P C R in a thermocycler using 30 cycles of the following steps: Step 1:1 min, 30 sec Step 2 : 2 min Step 3 : 2 min

at 94~ at 55~ at 72~

4. The reaction products can be analyzed by electrophoresis in a standard 1% agarose gel using Tris-borate buffer. Typically 10/xl of the PCR. reaction is sufficient to see the products using standard ethidium staining protocols. Although the R T - P C R method can provide an estimate of the position of a yeast polyadenylation site, it is often desirable to determine the exact position in the mR.NA. Moreover, a single band on a gel can sometimes harbor a collection of P C R products derived from cDNAs with multiple polyadenylation sites (8). If restriction enzyme cleavage sites are included in the PCI< primers, the DNA can be hydrolyzed with the appropriate enzymes, gel purified, and cloned into a bacterial plasmid for further amplification and DNA sequencing. Alternatively, PCt< products can be cloned directly with the use of the T-A cloning kit available from Stratgene (La Jolla, CA). Sequence analysis of a collection of independent clones is required to determine whether the clones were derived from a population of mRNAs with a single or multiple poly(A) sites. If a quantitative assessment of the relative usage of several polyadenylation sites is required, RNase protection analysis can be camed out with the use ofantisense probes prepared by transcription of the cDNA clones (8). Figure 2 shows the result of P,.T-PCR mapping of the TCM1 and PAB1 transcripts from a pap 1-1 strain before and after inactivation ofpoly(A) polymerase. Although no effort was made to make the assay quantitative, the results suggest

Figure 2 R T - P C R synthesis of cDNAs spanning the poly(A) sites of PAB1 and T C M 1 mRNAs from R N A prepared from UR3148-1B (pap1-1) before (0) and 60 rain after (60') shift to 35~ Reverse transcription/PCR using a downstream oligo(dT) BamH1 primer 3'-TTTTTTTTTTTTTTTTTCCTAGGTTTTT-5' and an upstream primer containing an EcoRI restriction sequence 5 ' - G G A A T T C C G G G T G G T T T C G T C C A C T A C G G T G - 3 ' for T C M 1 and 5 ' - G G A A T T C C G C C A C C T C A A G A G G T C T T C C C - 3 ' for PAB 1 was carried out as described in the text. The values on the left of the figure indicate the positions and lengths of 123-bp D N A molecular length markers.

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that there is little or no poly(A) + PAB1 m R N A but some poly(A) + T C M 1 m R N A after polyadenylation shutoff. Indeed, subsequent analysis of these transcripts by RNase protection confirmed these findings. (8). Moreover, the single band produced by R T - P C R for PAB1 m R N A was shown by DNA sequencing and RNase protection analysis to be derived from a population of PAB1 mRNAs with several adjacent polyadenylation sites, while that for T C M 1 is derived from a single site. (8).

IV. P R E P A R A T I O N OF Y E A S T m R N A PROCESSING EXTRACTS

3' E N D

A yeast strain that displays conditional loss of poly(A) + n ' ~ N A by either of the methods described above is certainly a candidate for a m R N A 3' end processing mutant. However, loss of steady-state poly(A) + may result from defects in transcription, splicing, or acceleration of poly(A) tail trimming as well. The simplest, most direct test of a 3' end processing defect is measurement of the activities in extracts derived from the strains in question. The protocol described here is a variation of the splicing preparation developed by Lin et al. (15-17) that was subsequently modified to stabilize polyadenylation activity. A miniature scale version of this procedure was also developed to allow the screening of a large number of strains from a collection of temperature-sensitive mutants (5).

1. Reagents Buffer S

Buffer A

1 M sorbitol 50 mM Tris-HC1, pH 7.8 10 mM MgC12 30 mM DTT

10 mM H E P E S - K +, pH 7.0 1.5 mM MgC12 10 mM KC1 0.5 mM DTT

Buffer B

Zymolyase

20 mM HEPES-K +, pH 7.0 0.2 mM EDTA 50 mM KCL 20% (v/v) Glycerol 0.5 mM DTT

Zymolyase 100T; ICN Biochemicals (Cleveland, OH)

2. Processing Extract Preparation 1. Grow yeast strain in 1 liter YPD at 30~ to an A600 of 4-6. 2. Pellet the cells by centrifugation (1000 g for 5 rain at 2500 rpm in a JA 10 rotor).

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9. 10.

11. 12. 13. 14. 15. 16. 17. 18. 19. 20.

Resuspend cells in 15 ml Buffer S at 25~ Incubate without shaking at room temperature for 15 min. Pellet cells as in Step 2 (3000 rpm, JA 20 rotor). Resuspend cells as in Step 3. Add 0.045 ml, 20 mg/ml Zymolyase 100T. Shake the suspension gently at 30~ for 40 man or until a 1/100 dilution of the suspension in 1% SDS gives an A550 1/10th of the starting value. Collect spheroplasts by centrifugation, as in Step 2. Gently resuspend spheroplasts in 8 ml buffer A. It is important to resuspend the spheroplasts gently, without drawing them into and out of a pipette, as this will cause them to lyse. They can be resuspended by gently stirring with the end of a 1-ml serological pipette and then poured into the homogenizer (Step 11). Lyse spheroplasts with five strokes in a tight-fitting Dounce homogenizer (Wheaton glass homogenizer, pestle A). Add sterile 2.0 M KC1 to a final concentration of 0.2 M. Stir gently on ice for 30 man. Clear by centrifugation (22,000 g for 30 min at 13,500 rpm in a JA 20 rotor). Collect the supernatant. Pellet polyribosomes by ultracentrifugation (145,000 g for 60 man at 45,500 rpm in a Ti 70.1 rotor). Collect the supernatant. Add ammonium sulfate (2.2 g/10 ml) and stir gently until dissolved. Then let the solution sit without stirring for 20 man at 4~ Collect the precipitate by centrifugation (15,000 g for 20 man at 11,500 rpm in a JA 21 rotor). P, esuspend the pellet in a small (i.e., 1-2 ml) volume of buffer B and dialyze for 3 hr against 1000 vol of the same buffer. Freeze the extract in small volumes in dry ice-ethanol and store at -70~ Determine the approximate protein concentration of the extract by measuring the A260 and the A280 of a 1/50 dilution in water. A good extract for m R N A 3' end formation will have an A280 above 6 units/ ml and an A260/A280 ratio of about 1. The extracts can be frozen and thawed repeatedly over many months, with little loss of activity.

There are several key factors to consider when preparing to carry out this protocol. First, the activity of the extract is dependent on the protein concentration, which, in turn, depends on the number of cells. Thus, one needs to harvest the cells at a high density without allowing them to enter stationary phase. Second, the strain chosen must be easily converted to spheroplasts so that they can be efficiently lysed. Third, an active yet relatively pure preparation of zymolyase must

7. PolyadenylationPhenotypes in S.

cerevisiae

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be used. Less expensive preparations than that recommended here have protease levels that make them unacceptable for preparation of active extracts. Fourth, the spheroplasts must be collected and resuspended gently so that they are not broken until the homogenizer is used. Rough treatment of the spheroplasts can ruin the preparation because apparently it breaks open vacuoles containing proteases. This procedure has been modified to allow the processing of large numbers of cells as a prelude to chromatographic ffactionation of the enzymatic activities (18). The large-scale procedure disrupts the cells with glass beads, thus avoiding the expense and protease problems associated with the use of zymolyase. Importantly, the modified procedure yields excellent processing extracts.

3. Polyadenylation Assay in vitro 1. The following reagents (per reaction) are added to a sterile microfuge tube on ice: 1/xl 1 /xl 1 /xl 4/xl 2/xl

750 mild KC1 20 mM Mg(Ac)2 20 mild ATP Sterile, distilled water Yeast processing extract

2. Start the reaction by adding 1/xl radiolabeled synthetic precursor IkNA (such as CYC1 or GAL716) to a final concentration of 1-100 nM (this is typically 2-200 • 103 cpm). 3. Incubate at the desired temperature (25~176 for the desired length of time. The reaction is usually over within 30 rain. 4. Stop the reaction by adding 2/xl stop solution (proteinase K, 2mg/ ml; EDTA, 130 m_M; SDS, 2.5%). 5. Incubate for 15 min at 37~ 6. Add 25/xl ethanol (-20~ and incubate at - 2 0 ~ for at least 15 min. 7. Collect the pellet by microcentrifugation (13,000 rpm) for 10 min. 8. Dry the pellet in a Speed Vac (Savant Instruments, Farmingdale, NY) for 5 min. 9. Resuspend the pellet in 10/xl FBX tracking dye (80% formamide, 0.1% xylene cyanol, 0.1% bromophenol blue). 10. Heat at 90~ for 3 min. 11. Separate the products by electrophoresis on a 4% acrylamide, 0.08% bisacrylamide, 8 M urea gel in 50 mM Tris-borate (pH 8.3). Expose the gel to Kodak X-Omat AP, film for 4-48 hr at - 7 0 ~ or to a Molecular Dynamics (Sunnyvale, CA) storage phosphor screen for 1-5 hr.

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It is quite easy to prepare extracts from yeast that have very active cleavage and polyadenylation activities. Unfortunately, it is also easy to obtain extracts with little or no activity. Thus, p o o r activity from a strain suspected o f having a m u t a t i o n directly affecting m R N A 3' end processing is not necessarily supportive evidence. O n l y after several attempts, including a demonstration o f proficiency in an extract from a normal strain, can one begin to seriously consider that p o o r processing activity reflects a defect in vivo. Because the level o f activity b e t w e e n two extracts is rarely the same (see Fig. 3), it is necessary that an extract from a m u t a n t strain demonstrate a conditional p h e n o t y p e such as thermosensitivity or sensitivity to depletion (4,5) that is similar to that observed in vivo before differences b e t w e e n m u t a n t and normal extracts can be considered as evidence o f a processing defect. This can be quite difficult in practice, as extracts from strains with thermosensitive processing defects often have little or no activity in vitro (4,5,19). H o w e v e r , care and persistence are often rewarded, as seen in Figure 3, which demonstrates the

Figure 3 Temperature dependence of mRNA 3' end processing in polyadenylation extracts derived from a PAP1 (EJ101) or a pap1-1 (UR3148-1B) strain. Extracts were prepared from cells grown at the permissive temperature (25~ for UR3148-lB. Processing fractions were carried out at the indicated temperatures for 30 min, as described in the text.

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t e m p e r a t u r e sensitivity o f p o l y a d e n y l a t i o n activity in an extract f r o m a strain w i t h the t e m p e r a t u r e - s e n s i t i v e p a p I - 1 m u t a t i o n in the gene for poly(A) polymerase. T h e v e r y l o w activity o f the m u t a n t e n z y m e is typical o f extracts f r o m t h e r m o s e n s i tive m u t a n t s and p r o b a b l y results f r o m instability o f the altered protein.

ACKNOWLEDGMENTS Research in the author's laboratory is supported by a grant from the National Science Foundation (MCB-931664). M.W.B. and A.P. were supported by a training grant from the U.S. Public Health Service (5-T32-AI070362).

REFERENCES 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.

Piper, P. W., and Aamand, J. L. (1989). J. Mol. Biol. 208, 697. Forrester, W., Stutz, F., Rosbash, M., and Wickens, M. (1992). Genes Dev. 6, 1914. Minvielle-Sebastia, L., Winsor, B., Bonneaud, N., and Lacroute, F. (1991). Mol. Cell. Biol. 11, 3075. Minvielle-Sebastia, L., Preker, P., and Keller, W. (1994). Science 266, 1702. Patel, D., and Butler, J. S. (1992). Mol. Cell. Biol. 12, 3297. Alhquist, P., and Kaesberg, P. (1979). Nucleic Acids Res. 7, 1195. Sachs, A. B., and Davis, R. W. (1989). Cell (Cambridge, Mass.) 58, 857. Proweller, A., and Buffer, S. (1994). Genes Dev. 8, 2629. Parker, R., Herrick, D., Peltz, S. W., andJacobson, A. (1991). In "Methods in Enzymology" (C. Guthrie and G. R. Fink, eds.), Vol. 194, p. 415. Academic Press, San Diego, CA. Decker, C. J., and Parker, R. (1993). Genes Dev. 7, 1632. Mandart, E., and Parker, R. (1995). Mol. Cell. Biol. 15, 6979. Sadhale, P. P., Sapolsky, R., Davis, R. W., Buffer, J. S., and Platt, T. (1991). Nucleic Acids Res. 19, 3683. Russo, P., Li, W.-Z., Hampsey, D. M., Zaret, K. S., and Sherman, F. (1991). E M B O J . 10, 563. Heidmann, S., Obermaier, B., Vogel, K., and Domdey, H. (1992). Mol. Cell. Biol. 12, 4215. Lin, R.-J., Newman, A. J., Cheng, S.-C., and Abelson, J. (1985). J. Biol. Chem. 260, 1880. Butler, J. S., and Platt, T. (1990). Science 242, 1270. Butler, J. S., Sadhale, P. P., and Platt, T. (1992). Mol. Cell. Biol. 10, 2599. Chen, J., and Moore, C. (1992). Mol. Cell. Biol. 12, 3470. Preker, P. J., Linger, J., Minvielle-Sebastia, L., and Keller, W. (1995). Cell (Cambridge, Mass.) 81, 379.

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Poly(A) Polymerase/Cap-Specific 2'- O-Methyltransferase from Vaccinia Virus" Expression, Purification, Uses, and Protein-Ligand Interaction Assays Paul D. Gershon Department of Biochemistry and Biophysics/Institute of Biosciences and Technology Texas A&M University Houston, Texas 77030

I. B A C K G R O U N D A. VACCINIA VIRUS GENE EXPRESSION AND m R N A MODIFICATION: GENERAL Viral gene-regulatory systems have contributed pivotally to the elucidation of messenger R N A (rnRNA) synthesis and modification mechanisms (Banerjee, 1980; Hercyk et al., 1988; Moss, 1996). Prominent among model viral systems have been those of vaccinia, the prototypic poxvirus. As a complex cytoplasmic animal virus, vaccinia displays a unique life cycle (Moss, 1996). This is centered upon three characterized stages of gene expression: an early stage, which precedes genome replication, and the intermediate and late stages, which follow this event. Stage-specific promoters, approximately 30 base pairs in length (Baldick et al., 1992; Davison and Moss, 1989a,b), and transcription factors (Kovacs and Moss, 1996; Moss, 1990, 1996; Moss et al., 1991) regulate vaccinia gene expression. A virus-encoded multisubunit DNA-dependent R N A polymerase transcribes all three gene classes. While a sequence-specific transcriptional termination mechanism has been demonstrated for vaccinia early genes (Moss et al., 1991), termination mechanisms for intermediate and late genes are currently ill defined (Moss, 1996). The early gene transcription apparatus, including the DNA-dependent R N A polymerase and rnRNA modification enzymes, is packaged within the virus particles during their assembly and is thereby carried into the cytoplasm of the newly infected cell (Kates and McAuslan, 1967; Munyon et al., 1967). Immediately upon 127 m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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or after infection, vaccinia early genes are transcribed and modified within the infecting virion, after which nascent early m R N A is extruded into the cytoplasmic milieu. Unlike most eukaryotic mRNAs, vaccinia transcripts are not subject to splicing. Nonetheless, mature vaccinia mRNAs exhibit eukaryotic features, including a 5' cap structure and a 3'-poly(A) tail. Studies ofvaccinia m R N A and associated RNA-modifying enzymes have contributed to the elucidation of the m R N A 5' cap structure (Wei and Moss, 1975), the demonstration of mRNA-associated poly(A) (Kates and Beeson, 1970), the characterization of enzymatic activities contributing to cap formation (Martin et al., 1975), and the isolation and characterization of a poly(A)-polymerizing enzyme (Moss et al., 1975). The structure of the vaccinia m R N A 5' cap is identical to that of its cellular counterpart, comprising a 7-methylguanosine (mVG) residue linked via a 5 ' - 5 ' triphosphate bridge to the first transcribed ribonucleotide (N1) of the transcript, which is always a purine. The resulting cap 0 structure (mVG(5')pppG/A) always becomes methylated at the 2' position, leading to a "cap I" structure (mVGpppGm/Am). A fraction of the total m R N A population is also 2'-O-methylated at the second transcribed nucleotide (N2), leading to a "cap II" structure (Moss et al., 1979). If N2 contains an adenine base, this is frequently methylated at position 6 of its ring, leading to a doubly methylated nucleotide (m6Am; Boone and Moss, 1977). Vaccinia m R N A 3' ends possess poly(A) tails with a median length of around 100 nt (Nevins and Joklik, 1975). This is a little shorter than the tails typically added to cellular transcripts. Vaccinia-encoded RNA-modification functions can be recapitulated in vitro by two heterodimers, namely, the multifunctional capping enzyme and the poly(A) polymerase (PAP)/cap-specific 2'-O-methyltransferase. Genes for each of the four polypeptide chains have been identified using biochemical procedures. This has been facilitated by (1) the presence of the enzymes within the virion, from which they can be purified relatively easily and in relatively large amounts, and (2) the genes being encoded within a relatively small, compact, double-stranded DNA genome.

B. m R N A

CAP FORMATION IN VACCINIA-INFECTED CELLS

Purified vaccinia virus particles and extracts thereof contain catalytic activities for the addition of m R N A 5' cap 0 and cap I structures. Cap formation proceeds according to a well-defined reaction sequence comprising (1) removal of the m R N A 5'-terminal phosphate; (2) guanylylation of the 5' end via a 5 ' - 5 ' triphosphate linkage; (3) 7-methylation of the added (terminal) guanine base; and (4) 2'O-methylation of the ribose of the first transcribed nucleotide. The first three steps (i.e., formation of the cap 0 structure) are catalyzed by the vaccinia capping enzyme (Martin et al., 1975; Moss et al., 1979), whose properties and uses have

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been reviewed (Shuman, 1995; Shuman and Moss, 1990). The fourth step (cap 0 to cap I conversion) is carried out by a separate enzyme, the vaccinia-encoded, capspecific S-adenosyl-t-methionine : m R N A (nucleoside-2'-O-)-methyltransferase. Although this enzyme was originally purified and characterized as a monomeric 38-kDa protein (Barbosa and Moss, 1978a,b), it was later shown to be genetically identical to the smaller (VP39) subunit of the vaccinia PAP heterodimer (Schnierle et al., 1992; Section I,C). The two findings are not incompatible, since VP39 is present in severalfold molar excess over its dimerization partner, VP55, in both the virion and infected-cell cytoplasm (Gershon et al., 1991; P. D. Gershon, unpublished data), consistent with the presence of both monomeric and VP55associated VP39 in virion and cell extracts. In vaccinia-infected cells, the formation of cap II and its m 6 A m counterpart proceeds via the action of cellular (cytoplasmic) methyltransferases (Banerjee, 1980; Boone and Moss, 1977; Langberg and Moss, 1981). A cap II-forming 2 ' - 0 methyltransferase from HeLa cells has been partially purified and characterized (Langberg and Moss, 1981) and has been shown to be located in the cytoplasm. HeLa cell m6A methyltransferase has been partially purified (1Kottman et al., 1994). Genes for the cellular cap II- and m6A-forming methyltransferases do not appear to have been reported at this time.

C. m R N A

P O L Y ( A ) TAIL FORMATION IN

VACCINIA-INFECTED CELLS Mthough 5' cap formation likely occurs through a similar series of enzymatic steps for both vaccinia virus and cellular organisms, mlKNA 3' end formation appears to occur by a fundamentally different mechanism. Whereas the polyadenylylation of cellular m R N A 3' ends is preceded by sequence-specific lnRNA recognition and cleavage (see other chapters in this volume), polyadenylylation of vaccinia early mRNAs follows sequence-specific transcriptional termination (Moss et al., 1991). The mechanism of 3' end formation for the majority of vaccinia intermediate and late mlKNAs is ill defined but likely also involves some form of transcriptional termination followed by poly(A) tail addition. A single documented exception (Antczak et al., 1992) suggests a sequence-specific 3' end cleavage event for one abundant vaccinia late mRNA. Poly(A) tails are added to vaccinia transcripts by the heterodimeric viral PAP. All characterized PAPs catalyze the following reaction (Edmonds, 1990): Polyribonucleotide + nATP

Mg2+ or M n 2+

polyribonucleotide (A)n + PPi

Despite their commonality of function, PAP enzymes isolated from different sources appear to be only minimally conserved at the amino acid sequence level.

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Indeed, despite zealous efforts to find sequence similarity (Martin and Keller, 1996), an inability to align the vaccinia PAP with a broad nucleotidyltransferase superfamily containing all other known PAPs, as well as CCA-adding enzymes (Holm and Sander, 1995), has been specifically noted (Yue et al., 1996). Interestingly, despite the presence of PAP genes within the eukaryotic, eubacterial, and viral kingdoms (Manley, 1995), the recently reported complete genome sequence of an archaeon (Bult et al., 1996) did not note an isolog of any known PAP. A PAP was purified from vaccinia virion extracts more than 24 years ago (Moss and Rosenblum, 1974; Moss et al., 1973, 1975). This PAP was shown to be a heterodimer of approximately 80,000 molecular weight, with subunit molecular weights of---51,000 and ---35,000 (Moss et al., 1975). The PAP was also isolated from vaccinia-infected HeLa cell cytoplasm (Brakel and Kates, 1974) and shown to be heterodimeric (Nevins andJoklik, 1977). Later, the larger and smaller subunits of the vaccinia PAP were shown to be encoded by open reading frames ElL and J3P,. of the vaccinia virus genome, respectively, and were named VP55 and VP39, respectively (Gershon et al., 1991). Enzymatic properties of the vaccinia PAP heterodimer have been described (Moss et al., 1975; Nevins and Joklik, 1977; Shuman and Moss, 1988; Thomson and Gershon, 1995) and have recently been reviewed in detail (Gershon and Moss, 1996). Structural and functional studies have shed considerable light on th~ mechanism of action of the vaccinia PAP. Thus, individually overexpressed and purified VP55 and VP39 have catalytic and stimulatory roles, respectively, in poly(A) tail addition (Gershon et al., 1991). Purified VP55 polyadenylates an mP,.NA 3' end bimodally: 30 to 35 adenylates are added in a rapid, highly processive initial burst before an abrupt transition to slow, nonprocessive adenylate addition (Gershon and Moss, 1992). By polyadenylylation of primers containing short, preformed 3'oligo(A) extensions, this transition was shown to be regulated by the net length of the oligo(A) tail rather than by the number of adenylate additions catalyzed (Gershon and Moss, 1992). RNA primers composed only of oligo(A) are polyadenylylated in an exclusively nonprocessive mode, indicating that some property of the nonpolyadenylylated I"nP,NA 3' end signals the initial rapid polyadenylylation burst. This signal was found not to reside in specific RNA sequences. Instead, VP55 has an affinity for multiple uridylate residues within the 3'-terminal 31 to 40 nt of a nonpolyadenylylated RNA primer (Gershon and Moss, 1993b). Even DNA primers in which the deoxythymidylates are replaced with ribouridylates can be polyadenylylated with the processive burst. The abrupt loss of processivity after addition of 30-35 adenylates appears to result from the translocation of VP55 to the nascent poly(A) tail, which lacks uridylates for stable interaction (Gershon and Moss, 1993b). VP55 interacts with R N A via two major uridylate-recognition sites, which might have roles in polymerase translocation (Deng and Gershon, 1997). The VP39 subunit has no independent PAP activity. However, this subunit dramatically stimulates the rate ofpoly(A) tail elongation for those RNA molecules

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that are extended only slowly and nonprocessively by the VP55 subunit alone, i.e., those that either possess 30- to 35-nt oligo(A) tails or are composed solely of poly(A) (Gershon and Moss, 1993a). In the presence of VP39, elongation of these primers is converted to a rapid, semiprocessive reaction. The apparent predilection of VP39 for 1KNA composed of polypurine (Gershon et at., 1991), along with other data (Gershon and Moss, 1993a), suggest a mechanism whereby VP39 anchors VP55 to the growing tail.

D.

T H R E E - D I M E N S I O N A L STRUCTURE AND

S T R U C T U R E - F U N C T I O N RELATIONSHIPS WITHIN THE VACCINIA

P A P V P 3 9 SUBUNIT Analysis of broad panels of VP39 point mutants has identified regions of the VP39 primary structure required for methyltransferase function (Schnierle et al., 1994; Shiet al., 1996). Mutants with methyltransferase-specific defects were analyzed in novel assays for S-Adenosyl methionine (AdoMet)-protein and capprotein interaction (Section IV,C), leading to the identification of regions of the VP39 primary structure important for interaction with the AdoMet cofactor and the terminal (m7G) nucleotide of the mP, NA cap. These studies also established that VP39's two cysteine residues play no role in methyltransferase activity, contrasting VP39's mechanism of action with that of the nucleic acid pyrimidine-specific methyltransferases, within which a cysteine side chain appears to play an integral (covalent) role in the methyl transfer mechanism (Chen et al., 1991; Kealey et al., 1994; Verdine, 1994). Solution of the 1.85-A crystal structure of a VP39 point mutant complexed with its AdoMet cofactor (Hodel et al., 1996) provided the first three-dimensional structure of an P,.NA methyltransferase. The structural data (an outline of which is currently provided at http://www-ibt.tamu.edu/gershon/pg.html) showed VP39 to be a compact, single-domain protein whose core, which is based upon the P,.ossmann fold (Rossmann et al., 1974), appears to be structurally similar to a consensus fold for the catalytic domains of four other methyltransferases (Schluckebier et al., 1995). The N- and C-terminal regions of VP39 are appended to the core and occlude one of its faces. The AdoMet cofactor molecule appears to be held by a network of 13 hydrogen bonds plus several potential van der Waals contacts. A single clear salt link is apparent between the oe-ammonium group of the AdoMet methionine moiety and a neighboring aspartate side chain carboxyl. Strikingly, this single carboxyl was absent from the only VP39 point mutant that was defective in AdoMet binding (Shi et al., 1996). The surface of VP39 is composed almost entirely of loops and helices. This contrasts with the architecture of almost all other known P,.NA binding proteins, within which a solvent-exposed platform of/3 strands typically packs against a pair

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of o~ helices. Topologically, the VP39 surface resembles a severely oblate sphere, one face of which is bisected by a cleft. One end of the cleft deepens into a hydrophobic pocket lined with aromatic side chains. The methyl group of the AdoMet molecule protrudes into the central part of the cleft; the remainder of the molecule is positioned within an adjacent shallow pocket. The arrangement of the protruding AdoMet-methyl group and cleft suggests that the latter is an R N A binding site for the methyl transfer reaction. The section of cleft between the donor methyl and the hydrophobic pocket is of sufficient length to bind four or five transcribed nucleotides of RNA. The function of the face of VP39 opposite that containing the cleft is currently unclear. In addition, sites of contact between VP39 and its dimerization partner, VP55, are currently ill defined.

II. E X P R E S S I O N A N D P U R I F I C A T I O N OF T H E POLY(A) POLYMERASE/2'-O-METHYLTRANSFERASE The vaccinia PAP subunits are not available commercially but have been expressed and purified on a noncommercial basis. This is relatively straightforward, and a number of options are available, as outlined in the following sections.

A. PURIFICATION FROM VACCINIA VIRIONS The vaccinia PAP (VP55-VP39 heterodimer) and cap-specific 2'-O-methyltransferase (VP39 monomer), like many other vaccinia enzymes, are present within the virion. Until recently, most studies with these enzymes employed material isolated from vaccinia virions that had been purified using large-scale procedures. Protocols for the maintenance of HeLa S-3 cell suspension cultures, preparation of virus (strain WR) particles from infected HeLa cells, and purification of virus by sucrose density gradient centrifugation have been described in detail (Earl and Moss, 1991; Shuman and Moss, 1990). Details of PAP and 2'-O-methyltransferase extraction and purification from whole virus particles have also been described and reviewed in detail (Barbosa and Moss, 1978a; Gershon and Moss, 1996; Moss et al., 1975). Briefly, twice gradient-purified virions are treated with dithiothreitol (DTT) and sodium deoxycholate, insoluble proteinaceous material is removed by centrifugation, and the supernatant is forced through a hypodermic needle to shear the released DNA. After the addition of glycerol, Triton X-100 detergent and ethylenediamine tetraacetic acid (EDTA), DNA is removed using a diethylaminoethyl (DEAE)-cellulose column equilibrated in buffer containing 0.25 M NaC1. Under these conditions, the flowthrough fractions contain all of the known vaccinia enzymes. These fractions are diluted to reduce the NaC1 concentration to 5080 mM, and this material is typically applied to a second DEAE-cellulose column

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equilibrated at the lower salt concentration. The unbound (flowthrough) fractions from this column, containing the PAP (VP55-VP39 heterodimer) and 2 ' - 0 methyltransferase (VP39 monomer), can then be applied to a single-stranded DNAagarose column. NaCl-gradient fractions from this column contain, in ascending order, capping enzyme, VP39 monomer, and VP55-VP39 heterodimer. Due to difficulties in separating VP39 monomer from closely eluting proteins, purification schemes for the 2'-O-methyltransferase have typically employed an alternative procedure (outlined below). Highly purified PAP can, however, be obtained from the single-stranded DNA-agarose column fractions. After the fractions are assayed for polyadenylylation activity (Section III,A), pooled active fractions are applied to a heparin-agarose column, which is eluted with an NaC1 gradient (Gershon et al., 1991; Shuman and Moss, 1988). PAP heterodimer in the resulting gradient fractions should be essentially free of other proteins, as judged by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). As an alternative to heparin-agarose, a Mono S HR5/5 column (Pharmacia, Piscataway, NJ) has been used for the final purification step (Gershon et al., 1991). Although the two subunits of the PAP have been found to separate slightly with the use of heparin-agarose (P. D. Gershon, unpublished data), they do not separate detectably during Mono S column chromatography. The purification of cap-specific 2'-O-methyltransferase from flowthrough fractions of the initial DEAE-cellulose column has been described (Barbosa and Moss, 1978a). Briefly, the material is dialyzed against a sodium acetate-based buffer (pH 6.0), containing 0.2 M NaC1, and applied to a CM-Sephadex column that has been equilibrated in the same buffer. After a linear gradient of 0.2-0.6 M NaC1 is run in the column loading buffer and column fractions are assayed for cap-specific 2'-O-methyltransferase activity (Section III,E), VP39 can be obtained in sufficiently pure form for peptide sequencing (Schnierle et al., 1992). The further removal of minor contaminants can be achieved by dialysis of pooled peak fractions back into a Tris-based buffer containing 0.05 M NaC1 and application to an ADPagarose column. After a linear 0.05-0.5 M NaC1 gradient is run in the loading buffer, SDS-PAGE of gradient fractions containing 2'-O-methyltransferase activity typically shows a single 39-kDa protein band. Although the original purification protocol suggested the occasional use of a poly(U)-Sepharose column as a final purification step (Barbosa and Moss, 1978a), this was subsequently found to be unnecessary.

B. VACCINIA VIRUS OVER.EXPRESSION OF V P 5 5 AND V P 3 9 : PURIFICATION Recombinant vaccinia viruses have been constructed for individual overexpression of the PAP VP55 and VP39 subunits in HeLa cells. The recombinant

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viruses, named vPG172a and vPG175, contain additional copies of the VP55 and VP39 genes, respectively, under control of a phage T7 promoter (Gershon and Moss, 1992, 1993a). HeLa cells can be co-infected with a T7 KNA polymeraseexpressing recombinant vaccinia virus and either vPG172a or vPG175 (Fuerst et al., 1986; Moss et al., 1990). Elevated levels of VP55 and VP39 expression are obtained after 18 hr of infection. The extraction and purification procedures employed for the overexpressed VP55 and VP39 have recently been described in detail (Gershon and Moss, 1996).

C. Escherichia coli OVEREXPRESSION OF G S T - V P 3 9 : PURIFICATION Attempts have been made to express both 6His and GST (glutathione-Stransferase) fusions of VP55 in E. coli. However, only very low levels of expressed protein are obtained. Furthermore, under a variety of expression and extraction conditions, less than 10% of the total recombinant protein is typically recovered in the soluble fraction. Thus, kilogram amounts of E. coli paste have provided only ---1-3 mg of purified VP55 protein (P. D. Gershon, unpublished data). VP55 recovered from the soluble protein extract is, however, active. Due to these difficulties, E. coli expression of VP55 will not be described here. VP39 has been successfully expressed to high levels in E. coli as a GST fusion protein and purified to apparent homogeneity (Hodel et al., 1996; Schnierle et al., 1994; Shi et al., 1996). The GST-VP39 fusion was constructed by cloning the VP39 coding region between the NcoI and SalI sites of plasmid pGEX-KG (Guan and Dixon, 1991; Schnierle et al., 1994). GST-VP39 can be expressed from the resulting plasmid (pPG177), as described (Gershon and Moss, 1996; Shi et al., 1996). Briefly, extract is made by sonication, and the cleared supernatant is adsorbed to GSH-agarose beads. After the beads are washed, fusion proteins can be released by either GSH elution or thrombin treatment, to specifically release VP39 from the bead-bound GST moiety (Shi et al., 1996), as recently described in detail (Gershon and Moss, 1996). The resulting VP39 can be purified to apparent homogeneity by heparin-agarose column chromatography, eluting with a linear NaC1 gradient (0-500 mM). Although the procedure has been described in detail elsewhere (Gershon and Moss, 1996), some minor modifications have been made. First, truncations of the recombinant protein have been found to correlate with extended cleavage times, perhaps in combination with protein freeze-thawing. To circumvent this, small amounts of phenylmethylsulfonyl fluoride (PMSF) can be added at the end of the thrombin cleavage step. Second, it has been found that some mutants of VP39 express to higher levels in E. coli at 27~ than at 37~ Third, no deleterious effects were observed upon reducing the NaC1 concentration

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in the extraction buffer from 250 m M to 100 mM. The latter concentration permits higher concentrations of recombinant protein to be employed in assays. In one recent study (Hodel et al., 1996), the standard extraction and purification procedure permitted 160 mg of a point mutant of VP39 to be obtained in highly purified form. This large amount of purified protein was derived from 14 liters of fermentation culture after sonication of the resulting 80 g (wet weight) ofE. coli cell pellet into 300 ml of extraction buffer. Recombinant protein was adsorbed to 20 ml of glutathione-agarose beads, and the VP39 moiety was released by incubation with 30 U thrombin protease (Novagen Inc., Milwaukee, WI) at 25~ for 4 hr in a total extraction buffer volume of 40 ml. Eluted protein was adsorbed to a 5-ml heparinagarose column (HiTrap, Pharmacia), and the highly purified recombinant VP39 mutant was eluted as described above. Peak fractions contained the highly purified protein at a concentration of several milligrams per milliliter. Highly purified, concentrated, wild-type VP39 has also been obtained with the above large-scale procedures (A. E. Hodel, P. D. Gershon, and F. A. Quiocho, unpublished).

D . BACULOVIRUS OVEREXPRESSION OF V P 3 9 VP39 has been overexpressed from recombinant baculovirus. For this purpose, the G S T - V P 3 9 fusion gene was amplified as a PstI-HindIII fragment from the plasmid pPG177 (Section II,C), and cloned between the PstI and HindIII sites of plasmid pBluebac III (Invitrogen Inc., San Diego, CA). After the generation, purification and amplification of recombinant baculovirus, High Five insect cells (Invitrogen) were infected at multiplicities of 5-10, and protein was expressed over a 2.5-day period. Recombinant VP39 can be purified as described in Section II,C. Baculovirus has no known advantages over E. coli for VP39 expression.

III. U S E S O F T H E P O L Y ( A ) P O L Y M E R A S E / 2 ' - O - M E T H Y L T R A N S F E R A SE

A. POLY(A) TAIL ADDITION The vaccinia PAP is unusual in being able to rapidly and processively add long poly(A) tails to R N A substrates without requiring the presence of specific R N A sequences. Conditions for doing this vary, depending upon the importance of complete substrate utilization and regulated tail length. Reaction variables include (1) whether the VP55 subunit alone or both subunits are present in the reaction; (2) the time and temperature of the reaction; (3) whether Mn 2+ is present as well as/in place of Mg2+; (4) concentrations of monovalent cation; (5) relative concentrations of PAP, RNA, and ATP; and (6) the presence/absence of ATP

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analogs. Instead of exhaustively discussing these parameters, some appropriate PAP assays will be described and pointers provided to the primary literature. An electrophoretic time-course PAP assayhas been described and subsequently refined (Gershon and Moss, 1992, 1993a,b; Shuman and Moss, 1988). This assay employs a discrete 5' end-labeled 1KNA primer, and polyadenylylation products are analyzed by gel electrophoresis. As originally reported (Shuman and Moss, 1988), reaction conditions were similar to those described for [c~-32p]AMP incorporation into trichloroacetic acid (TCA)-precipitable 1KNA (Gershon and Moss, 1996; Moss et al., 1975), except for a reduction of the MnC12 concentration to 0.6 mM, the omission oflabeled ATP, and the use of a small (10-/xl) reaction volume and the 1KNA primer U10 after 5'-end labeling of the latter using [T-32p]ATP and T4 polynucleotide kinase. After a "time zero" sample is removed, reactions are initiated by adding enzyme to a mixture of the other components. Aliquots of the reaction mixture are then withdrawn at various times and mixed with equal volumes ofdeionized formamide. Reaction products are analyzed by electrophoresis in polyacrylamide gels containing 7 Murea and Tris-borate-EDTA (TBE) buffer followed by autoradiography. Modifications have been made to this assay (Gershon and Moss, 1992, 1993a,b). Specifically, chemically or enzymatically synthesized primers have been used, which are generally more than 25 nt in length. These are normally gel-purified after synthesis but prior to labeling. Labeling is usually terminated by incubating labeling reactions at 65~ for 15 n'fin, obviating a second gel-purification step. In general, primer is used in more than threefold molar excess over enzyme. For the majority ofpolyadenylylation reactions, MnC12 is replaced by MgC12. In reactions where MnC12 is still used, MnC12 94H20 is dissolved in water immediately prior to use, and the MgC12, Tris-HC1, and DTT in the assay mixtures, and in the polynucleotide kinase labeling reactions, are substituted for by MnC12, N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid-sodium hydroxide (HEPES-NaOH), and 2-mercaptoethanol, respectively (Thomson and Gershon, 1995). The above polyadenylylation assay is effective when used with either monomeric VP55, the reconstituted PAP subunits, or the purified heterodimer.

B. R N A 3'-END LABELING IKNA can be 3'-end labeled with a radioactive chain-terminating ATP analog, cordycepin monophosphate (3'-deoxyadenosine-5'-monophosphate or 3'dAMP). This analog can be transferred to 1KNA 3' ends using either a recombinant form of the PAP encoded by Saccharomyces cerevisiae (Lingner and Keller, 1993) or the heterodimeric vaccinia PAP (Thomson and Gershon, 1995). The vaccinia PAP may provide advantages in terms of its low 1KNA substrate specificity and ready inactivation. A typical 1KNA 3'-end labeling reaction contains 50 /xM MnC12 94H20, 1KNA primer (typically 0.6 /xM in 3' ends), [o~-32p]cordycepin

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5'-triphosphate (0.2 la,M, 5000 Ci/mMol, DuPont NEN, Wilmington, DE), 50 m M Tris-HC1 (pH 9.0), 50 m M NaC1, 5 m M 2-mercaptoethanol, "--25 nM VP55, -~300 nMVP39, and diethyl pyrocarbonate (DEPC)-treated H 2 0 in a total volume of 10/xl. Reactions are incubated at 30~ for 60 min. As a termination step, reactions can be heated to 70~ for ~15 min. Poly(A) + or hybrid-selected R N A labeled in this manner can be used directly for poly(A) tail length determination by nuclease digestion (Section Ill,D). The importance of using solid MnC12 that has been unsealed less than 6 months previously and dissolved in RNase-free H20 immediately prior to use is emphasized. An artifact has been observed after R N A 3'-end labeling in the above manner, using either yeast or vaccinia PAP as the labeling enzyme (Deng and Gershon, 1997). Thus, R N A 3'-end labeling followed by partial hydrolysis (i.e., either RNase A or alkaline treatment) always leads to a strong band migrating with the ---8-nt hydrolysis product, accompanied by hypohydrolysis of ~ 1 - to ---8-nt oligonucleotide fragments. This unusual effect is observed whether or not the R N A has been gel-purified after 3'-end labeling, and cannot be recapitulated by incubating 5'-end labeled R N A with all of the 3'-end labeling reagents except 3'-end labeling enzyme, indicating that the 3 '-end labeling reagents are not modifying the R N A directly. The artifact is not observed when 5'-end labeled, chemically synthesized U34-3'dA is hydrolyzed, indicating that it cannot be attributed to the presence of the 3'dA residue per se. It is also R N A sequence-independent. Thus, by process of elimination, it is apparently attributable to the presence of a 32p_ labeled 3'dA (CoMP) nucleotide attached to the R N A 3' end. Perhaps the 32p 3'dA moiety leads to an R N A modification involving the nucleotide at position " - 8 , such as an intramolecular cross-link.

C. P O L Y ( N ) T R A C T A D D I T I O N Occasionally, it may be desirable to tag or 3'-end label an R N A with a short 3' homopolymer tract other than oligo(A). Oligo(C), oligo(G), and poly(U) tracts can be added to R N A 3' ends (Gershon and Moss, 1992; Shuman and Moss, 1988; Thomson and Gershon, 1995) due to the donor nucleotide promiscuity of vaccinia PAP in the presence of Mn 2+ (Gershon and Moss, 1992). Reaction conditions are as described for poly(A) tail addition in the presence of MnC12 (Section III,A), except for the replacement of Mn 9ATP in the reaction with M n " CTP, M n " GTP, or M n " UTP (Gershon and Moss, 1993b). Under these conditions, oligo(C), oligo(G), and poly(U) tails, respectively, are added. Whereas the poly(A) tails that are rapidly and processively synthesized by monomeric VP55 are ~35 nt in length, VP55 processively synthesizes poly(U) tails several hundred nucleotides in length (Gershon and Moss, 1993b). Oligo(C) tails are added very slowly, so that only a few nucleotides are typically added during the lifetime of a

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reaction, and oligo(G) tail synthesis always ceases abruptly after the addition of 14-15 nt. Internally labeled oligo(N) tails can also be synthesized. Conditions for this reaction have recently been described in detail (Gershon and Moss, 1996; Thomson and Gershon, 1995).

D. MICRO-ASSAY FOR POLY(A) TAIL LENGTH DETERMINATION BY RNASE A DIGESTION The convenient 1KNA 3'-end labeling reaction described in Section III,B is highly compatible with poly(A) tail length determination after digestion of the body of the m R N A using ribonuclease A. Thus, poly(A) + or hybrid-selected 1KNA samples are 3'-end labeled and the labeling reactions terminated, as described in Section III,B. The resulting samples are made 0.5 M in NaC1 and 0.6 mg/ml in yeast tRNA (GIBCO-BRL, Gaithersburg, MD) that has been extracted twice with phenol-chloroform and once with chloroform, precipitated with ethanol, and redissolved in DEPC-treated H20 to a final concentration of 5 mg/ml. In 96-well format, 2-/xl aliquots of the terminated, supplemented, labeling reactions are mixed with 2.8-/xl aliquots of"poly(A)-binding suspension." The latter comprises streptavidin-paramagnetic particles (SA-PMPs; Promega, Madison, WI), collected from the manufacturer's storage buffer using a permanent magnet and resuspended in one-fifth volume of 0.5 M NaC1 containing 2 /xM biotinylated oligo(dT) probe (Promega). After incubation of the mixtures for 5 min at room temperature, SA-PMPs are collected using a permanent magnet, supernatants are removed, and the isolated PMPs mixed with 4-/xl aliquots of 50% formamide in DEPC-treated H20. Additional 2-/,1 aliquots of the terminated, supplemented, labeling reactions are mixed with 0.5-/,1 aliquots of i or 10/xg/ml Ribonuclease A (affinity purified; Ambion, Austin, TX) and incubated at 37~ for 10 min. The resulting digests are each incubated with 2.8-/xl aliquots of poly(A)-binding suspension, after which the SA-PMPs are collected using a permanent magnet and resuspended in 50% formamide. Finally, the supernatants from all SA-PMP suspensions in 50% formamide are loaded on preelectrophoresed polyacrylamide gels (10% T, 3% C) containing urea (7 M), buffered with Tris (100 mM), borate (90 mM), and Na2EDTA (1 mM), pH 8.3, after again isolating SA-PMPs using a permanent magnet. After electrophoresis, the incorporated radioactivity in dried gels is imaged by autoradiography.

E. CAP-SPECIFIC 2 ' - O - M E T H Y L A T I O N It may be necessary to convert the cap 0 structure to cap I, e.g., for m R N A expression studies. This conversion can be effected by the cap-specific

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(nucleoside-2'-O-)-methyltransferase activity of VP39 in the presence of AdoMet. In previously described assays for VP39's 2'-O-methyltransferase activity, brome mosaic virus (BMV) R N A was employed as a substrate (Barbosa and Moss, 1978b; Schnierle et al., 1992, 1994). Although commercially available packages (Promega) are too small and expensive for use in assays, the R N A can be prepared in large quantities given appropriate greenhouse facilities. Specific n ~ N A species requiring a cap I structure may be initially synthesized with a type 0 cap by in vitro transcription of a phage R N A polymerase promoterdriven template in the presence of dinucleotide cap analog. Alternatively, specific diphosphate/triphosphate-terminated transcripts may be treated with the vacciniacapping enzyme (Shuman and Moss, 1990). Cap-specific 2'-O-methyltransferase reaction conditions have been described elsewhere (Barbosa and Moss, 1978b; Gershon and Moss, 1996; Schnierle et al., 1992). Briefly, reactions contain 25 mM H E P E S - N a O H (pH 7.5), 1 mM DTT, and VP39. For assaying VP39 activity, reactions are supplemented with 1 p , M S-[methyl-3H]Adenosyl-L-methio nine (---6.9 Ci/mmol, Dupont NEN) and 5 /.tg BMV RNA. After incubation at 37~ for 30 min, enzyme can be inactivated by incubating samples at 100~ for 2 min. Radiolabeled R N A is quantitated using DEAE filters, which are subsequently washed and quantitated by scintillation counting. To subject chosen R N A species to the cap 0 to cap I conversion reaction, this R N A should be included in the reaction in place of BMV R N A and 3H-AdoMet replaced with unlabeled AdoMet at a concentration of 2-500 /.tM. To maximize conversion levels, VP39 can be included in reactions in either equimolar amounts with the R N A or stoichiometric excess. This is most easily accomplished after E. coli expression of VP39 using the large-scale protocol (Section II,C), since highly purified VP39 can be obtained at concentrations of several milligrams per milliliter directly from peak fractions of the final column gradient. VP39 can effect cap 0 to cap I conversion in capping enzyme buffer, which is similar to the above HEPES/DTT buffer except for the presence of MgCI2 (P. D. Gershon, unpublished data). Thus, it should be possible to synthesize cap Iterminated R N A directly from the triphosphate-terminated counterpart, in a single tube, by employing mixtures of VP39 with the vaccinia-capping enzyme.

IV. A S S A Y S D E V E L O P E D T O E X A M I N E PROTEIN-LIGAND INTERACTIONS During mechanistic studies of the VP55 and VP39 subunits of the vaccinia PAP/2'-O-methyltransferase heterodimer, assays have been developed to study protein interactions with R N A and AdoMet substrates. Some of these are described in the following sections.

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A. R N A BINDING ASSAYS Several R N A binding assays have been applied to VP55 and VP39. An effective nitrocellulose filter tLNA binding assay has been described (Gershon et al., 1991). Application of the electrophoretic mobility shift assay (EMSA) to the analysis of VP55-ILNA interactions has also been described (Deng and Gershon, 1997; Gershon and Moss, 1993b) and comprises one component of the selection procedure described in Section IV,D. Surface plasmon resonance-based techniques have been developed to assay the 1KNA binding properties of VP39 (Shiet al., 1996). One of these is described in the next section.

B. SURFACE PLASMON RESONANCE-BASED ASSAY FOR CAP-PROTEIN INTERACTION Since VP39 does not methylate the dinucleotide cap-analog (Barbosa and Moss, 1978b), direct cap binding by VP39 cannot be easily assayed. However, a surface plasmon resonance-based assay has been developed for analysis of cap 0-VP39 interactions based on the differential affinity of VP39 for m7GpppG- and pppG-terminated RNAs in the context of the "BIAcore" surface plasmon resonance instrument (Biacore AB, Piscataway, NJ; Shi et al., 1996). The two RNAs are immobilized, via 3' biotin moieties, within adjacent flowcells of the BIAcore biosensor chip, after which VP39, or mutants thereof, are injected in the mobile phase. The instrument can detect association of the protein with the immobilized R N A in real time. Details of the assay, which might be useful for analysis of other cap-binding proteins, follow. 5'-Capped, 3'-biotinylated R N A species are synthesized using T7 R N A polymerase. Poly(A) R N A was chosen for the body of the transcripts because of the apparent efficacy of the cap 0-terminated form as a substrate for VP39's cap-specific methyltransferase activity (Barbosa and Moss, 1978b). Prior to transcription, oligodeoxynucleotides with the following sequences: 5'-A(T)s0CTATAGTGAGTCGTATTAGG-3' and 5'-CCTAATACGACTCACTATAGA-3' are mixed in DEPC-treated water and incubated at 70~ for 5 min and then at 37~ for 15 min. The resulting transcription template comprises a double-stranded T7 promoter followed by a single-stranded region templating the synthesis of R N A with the sequence G(As0)U. Standard T7 transcription reactions (Megashortscript, Ambion) can be performed according to the manufacturer's instructions, except for the use of altered nucleoside triphosphate compositions. Thus, for uncapped

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R N A synthesis, reactions should contain 15 m M GTP, 30 m M ATP, and 2 m M Biotin-21-UTP (Clontech Labs., Palo Alto, CA). For the synthesis of quantitatively 5'-cap 0-terminated RNA, GTP is replaced with the cap 0 analog m7GpppG (0.9 mM, Boehringer). The successful incorporation of biotinylated UMP at the R N A 3' end can be confirmed by electrophoresis of aliquots of the transcription reactions on denaturing polyacrylamide gels followed by electroblotting to Hybond N + membranes (Amersham Life Science, Arlington Heights, IL), incubation of membranes with streptavidin-horseradish peroxidase conjugate (GIBCO-BRL), and chemiluminescent detection. After transcription, reactions are incubated for 15 min with DNase I and the I:(NA products are purified by PAGE. Running buffer for the BIAcore instrument comprises 10 m M HEPESN a O H (pH 8.0), 0.005% Tween 20, and, unless otherwise stated, 60 m M NaC1. After each ---48-hr period of use, fresh running buffer is made using DEPC-treated water. Before initial use, the BIAcore should be subjected to two cycles of the "desorb" routine (as programmed in the instrument software) in order to eliminate possible ribonuclease contamination. In the original study (Shi et al., 1996), streptavidin [---2000 resonance units (RU)] was immobilized within each of two flowcells of the BIAcore sensor chip by amine coupling (O'Shannessy et al., 1992) using the Amine Coupling Kit (Biacore Inc., Piscataway, NJ). Equally effective are sensor chips precoated with streptavidin (P. D. Gershon, unpublished data), which can be purchased from the manufacturer. The capped and uncapped RNAs ('--750 R U of each) are separately immobilized to the individual streptavidin-coated surfaces. The R N A is introduced by autoinjections that are manually interrupted after achieving a AInU of 750. Analytical cycles are initiated by autoinjection to the immobilized R N A surfaces of 40-/xl samples of GST-tagged VP39, or mutants thereof, diluted to a final concentration of 42 nM in running buffer containing 0.5 /xl of ribonuclease inhibitor (5' ~ 3') at flow rates of 5 /zl/min. Since the primary kinetic effect of the 5' cap structure relates to the protein-RNA association rate, protein injections are usually not followed by washout of the flowcell with running buffer, as would be required for analysis of dissociation kinetics. Instead, analytical cycles are usually terminated with three 40-/xl injections of high-salt regeneration buffer (10 m M HEPES-NaOH, 0.5 M NaC1, 0.005% Tween 20), each at a flow rate of 40/xl/min, to remove all traces of protein. Each protein sample is injected consecutively to the two R N A surfaces before continuing to the next sample. For each experiment, negative and positive controls comprise the GST moiety alone and wild-type GST-VP39, respectively. GST-VP39 is injected to each of the two R N A surfaces at both the start and end of each experiment. RNA-derivatized flowcells are not used for more than 36 hr or if the association parameters for the final GST-VP39 injection of an experiment differ from those for the initial injection by more than ---25%. There is little difference in rates of wild-type VP39 association with the capped and uncapped R N A species when running buffer contains 20 or 60 m M

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NaC1 (Shi et al., 1996). However, there is a distinct difference in running buffer containing 100 m M NaC1. This is mainly because the higher salt concentration specifically suppresses association of VP39 with the uncapped R.NA (Shi et al., 1996). For several point mutants of VP39, the association rate difference is clearly abolished, leading to the identification of surface residues with apparent roles in VP39 binding to the terminal cap nucleotide (Shi et al., 1996).

C. SOLID-PHASE ASSAY FOR PROTEIN-ADoMET INTERACTION Although association/dissociation constants for methyltransferase-AdoMet interaction can be inferred from activity data (Barbosa and Moss, 1978b), a direct binding assay might be useful for mutant proteins possessing reduced catalytic activity. Although such assays exist in the literature (Kossykh et al., 1993; Segal and Eichler, 1989), nonequilibrium assay conditions were considered inappropriate for VP39 due to the suspicion of a weak interaction, and the traditional equilibrium binding assay was considered inappropriate due to the tendency of VP39 to bind nonspecifically to dialysis membranes and the possible requirement for large amounts of protein. An assay was therefore developed to circumvent these problems and is described below. For each replicate assay of each VP39 mutant, a single colony of freshly transformed E. coli (strains DH12S or HB101) containing the plasmid encoding the (mutant) protein is expanded into i liter of superbroth. Following this, cultures are induced and harvested, then extracted and centrifuged, as described in Section II,C. Typically, 25 ml of soluble protein extract is obtained from a l-liter culture. Extracts can be stored at -70~ For each assay, an adequate volume of GSHagarose beads is washed twice with an excess volume of extraction buffer (containing 0.25 M NaC1) and is then made to a 50% slurry with this buffer. Four aliquots of protein extract (typically 0.4, 1.3, 4, and 12 ml) are mixed with four 50-/2,1 aliquots of the 50% GSH-agarose bead slurry and subjected to 60 min of rotation at ambient temperature. For the most abundantly expressed proteins, 12 ml of protein extract is typically sufficient to almost fully saturate a 25-/,1 aliquot of packed beads with GST-VP39 fusion protein. The beads are then sedimented by centrifugation, the supernatants carefully removed, and the beads washed three times with 1-ml volumes of extraction buffer (containing 0.25 M NaC1), followed by two washes with 0.3-ml volumes of AdoMet binding buffer (40 m M HEPES-NaOH, pH 7.5; 1 m M DTT, referred to as buffer AB). Prior to the centrifugation step of the second wash with buffer AB, bead suspensions are divided into three 0.1-ml aliquots in 0.6-ml tubes. One aliquot is mixed with 4X SDS-PAGE loading buffer and stored at -70~ The two remaining aliquots are centrifuged and the supernatants replaced with 0.5-ml volumes of buffer AB containing 5 nM S-[methfl-3H]Adenosyl-L-methionine

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(10.6 Ci/mMol, DuPont NEN). S-[methyl-3H]AdoMetis stored as recommended by the manufacturer and diluted with buffer AB immediately before use. Bead suspensions are rotated for 30 rain at ambient temperature and then centrifuged briefly. After careful removal of supematants with the use of an ultra-fine pipette tip to fully drain the beads without bead loss, 0.4-ml aliquots of scintillation fluid (Ultima gold, Packard Instruments, Meriden, CT) are added to the drained beads, which are then suspended by vigorous agitation. The 0.6-ml assay tubes are placed inside miniature pony vials (Packard Instruments), and duplicate counts obtained for each vial in the tritium channel of a scintillation counter. Mean tritium cpm values are then calculated from the duplicate values. For protein quantitation, the stored ahquots of beads in SDS-PAGE loading buffer (above) are heated and centrifuged briefly, and 20-/.,1 aliquots of the supernatants subjected to SDS-PAGE. For use as staining standards, four different amounts of bovine serum albumin (BSA) (1, 3, 10, 30 /zg, always taken from the same stocks frozen in SDS-PAGE loading buffer) are loaded on each SDS-PAGE gel. After electrophoresis, the gels are stained with Coomassie Brilliant Blue tL250 and then scanned using a computing densitometer containing a scanning laser (Molecular Dynamics, Sunnyvale, CA). The relevant protein bands are quantitated by volume integration with background subtraction to yield "protein amount" values. For correction of g.l-to-gel staining differences, protein amount values for the BSA standards loaded on a specific gel are plotted against the amounts of BSA loaded (/zg), and the slope of the line is obtained by linear regression employed as the "staining correction factor" for that gel. Protein amount values for sample lanes can then be divided by the staining correction factor for that gel to yield "staining-corrected protein amount" values for gel-to-gel comparison. These values are plotted against the corresponding mean tritium cpm values (above). Since each lane of the stained gel corresponds to two S-[methyl-3H]-AdoMetbinding reactions, two mean tritium cpm values are plotted against each staining-corrected protein amount value. Linear regression of the resulting plot yields a slope that is employed as the final AdoMet binding coefficient. Normally, each mutant is assayed two to five times and mean binding coefficient values are obtained, along with error bars indicating the average deviations of individual values from the mean.

D . R N A BINDING SITE CHARACTERIZATION BY DOUBLE SELECTION FROM THE OLIGONUCLEOTIDE POOL VP55 interacts with R N A through the specific recognition of uridylate residues (Deng and Gershon, 1997; Gershon and Moss, 1993b). VP55 completely fails to recognize cytidylates, despite the structural similarity of the cytosine and uridine rings. A procedure has been developed for characterizing the positions within an R N A at which uridylates are recognized against an oligodeoxycytidylate

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background (Deng and Gershon, 1997). Since the ribose sugar is used as a reporter to read out positions of selected uridylates, it is important to control for co-selection for the sugar itself (Deng and Gershon, 1997). Despite the necessity for this additional safeguard, the procedure may be generally applicable in the study of protein-RNA interactions. The procedure employs a chemically synthesized chimeric oligonucleotide pool, 34 nt in length (the minimum oligonucleotide size capable of stable complex formation with VP55 in the EMSA) in which a deoxyriboC background is spiked at a level of---10% with riboU. The chimera is synthesized by solid-phase 2'-O-Fpmp chemistry (Cruachem, Hemdon, VA) according to the manufacturer's instructions, using an ABI model 392 D N A / R N A synthesizer. The oligonucleotide is gel-purified after complete deprotection but before use, as described previously (Gershon and Moss, 1992). A single phosphoramidite bottle is used, containing a mixture of dC and rU phosphoramidites. Prior to pool synthesis, the following procedure was designed to calibrate coupling rates for 2'-O-Fpmp riboU versus deoxyriboC phosphoramidites, thereby ensuring predictable spiking levels in the resulting oligonucleotide: Several 9mer oligonucleotides of sequence dC4-N-dC4 were synthesized, in which N comprised either 100% dC, 100% rU, or a mixture ofdC and rU phosphoramidites that was distinct for each oligonucleotide (P. D. Gershon, unpublished data). The 9mers were 5'-end labeled using [T-32p]ATP and T4 polynucleotide kinase, subjected to complete hydrolysis by treatment with 50% trifluoroacetic acid at 70~ for 5 min, dried under vacuum, redissolved in 1 M Tris-HC1 (pH 9.0), and electrophoresed on a denaturing 20% polyacrylamide gel. For each oligonucleotide, the relative abundances of the labeled bands corresponding to the intact oligo and cleaved product were scored, enabling the rU coupling rate to be plotted against the proportion of rU phosphoramidite in the mixture. The clearest experimental data have been obtained using a double-selection procedure (Deng and Gershon, 1997). The first selection step comprises 3'-end labeling of the 34mer oligonucleotide pool using [ol-32p]CoTP in combination with VP55 (Section Ill,B). Reaction times are employed that would be expected to lead to only ~25% of maximal labeling levels. Oligonucleotides are then deproteinized by phenol/chloroform extraction and precipitated with a mixture of glycogen, sodium acetate, and ethanol. In the second selection step, the deproteinized oligonucleotide is incubated with a second aliquot of VP55 and subjected to the EMSA (Section IV,A). The resulting EMSA complex is excised from the nondenaturing gel, and the oligonucleotide is extracted from the crushed gel slice by overnight diffusion in 1% SDS/1 mM EDTA followed by phenol/chloroform extraction and precipitation with a mixture of glycogen, sodium acetate, and ethanol. The 32p-labeled, doubly selected oligonucleotide is then partially hydrolyzed by supplementing samples with a 1/20 volume of I 21//sodium bicarbonate and heating at 90~ for 10 or 20 rain (Whoriskey et al., 1995). The resulting samples

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are electrophoresed in 9% or 10% polyacrylamide/urea/TBE gels 0.4 m m or 0.8 m m in thickness, after supplementing the lower buffer tank with 1 M sodium acetate, in order to establish an electrolyte gradient (Ausubel et al., 1987). Gels are soaked extensively in 30% methanol/5% glacial acetic acid after electrophoresis and prior to drying. In control experiments, such soaking was shown to cause no detectable loss of small oligonucleotide fragments (data not shown).

REFERENCES Antczak, J. B., Patel, D. D., Ray, C. A., Ink, B. S., and Pickup, D.J. (1992). Site-specific RNA cleavage generates the 3' end of a poxvirus late mRNA. Proc. Natl. Acad. Sci. U.S.A. 89, 12033-12037. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K., eds. (1987). "Current Protocols in Molecular Biology." Wiley (Interscience), New York. Baldick, C.J., Keck, L. G., and Moss, B. (1992). Mutational analysis of the core, spacer, and initiator regions of vaccinia virus intermediate class promoters. J. Virol. 66, 4710-4719. Banerjee, A. K. (1980). 5'-Terminal cap structure in eucaryotic messenger ribonucleic acids. Microbiol. Rev. 44, 175-205. Barbosa, E., and Moss, B. (1978a). mRNA(nucleoside-2'-)-methyltransferase from vaccinia virus: Purification and properties. J. Biol. Chem. 253, 7692-7697. Barbosa, E., and Moss, B. (1978b). mRNA(nucleoside-2'-)-methyltransferase from vaccinia virus: Characteristics and substrate specificity. J. Biol. Chem. 253, 7698-7702. Boone, R. F., and Moss, B. (1977). Methylated 5' terminal sequences of vaccinia virus mRNA species made in vivo at early and late times after infection. J. Virol. 79, 67-80. Brakel, C., and Kates, J. R. (1974). Poly(A) polymerase from vaccinia virus-infected cells. I. Partial purification and characterization. J. Virol. 14, 715-723. Bult, C. J., White, O., Olsen, G. J., Zhou, L., Fleischmann, R. D., Sutton, G. G., Blake, J. A., FitzGerald, L. M., Clayton, R. A., Gocayne, J. D., Kerlavage, A. R., Dougherty, B. A., Tomb, J.-F., Adams, M. D., Reich, C. I., Overbeek, R., Kirkness, E. F., Weinstock, K. G., Merrick, J. M., Glodek, A., Scott, J. L., Geoghagen, N. S. M., Weldman, J. F., Fuhrmann, J. L., Nguyen, D., Utterback, T. R., Kelley, J. M., Peterson, J. D., Sadow, P. W., Hanna, M. C., Cotton, M. D., Roberts, K. M., Hurst, M. A., Kaine, B. P., Borodovsky, M., Klenk, H.-P., Fraser, C. M., Smith, H. O., Woese, C. R., and Venter, J. C. (1996). Complete genome sequence of the methanogenic archaeon Methanococcus jannaschii. Science 273, 1058-1073. Chen, L., MacMillan, A. M., Chang, W., Ezaz-Nikpay, E., Lane, W. S., and Verdine, G. L. (1991). Direct identification of the active-site nucleophile in a DNA (cytosine-5-)-methyltransferase. Biochemistry 30, 11018-11025. Davison, A. J., and Moss, B. (1989a). The structure of vaccinia virus early promoters. J. Mol. Biol. 210, 749-769. Davison, A. J., and Moss, B. (1989b). The structure of vaccinia virus late promoters. J. Mol. Biol. 210, 771-784. Deng, L., and Gershon, P. D. (1997). Interplay of two uridylate-specific RNA binding sites in the translocation of poly(A) polymerase from vaccinia virus. E M B O J., 16, 1103-1113. Earl, P. L., and Moss, B. (1991). Preparation of cell cultures and vaccinia virus stocks. In "Current Protocols in Molecular Biology" (F. M. Ausubel, R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl, eds.), Vol. 2, pp. 16.16.1-16.16.7. Wiley (Interscience), New York.

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Edmonds, M. (1990). Polyadenylate polymerases. In "Methods in Enzymology" (J. Dahlberg and J. Abelson, eds.), Vol. 181, pp. 161-170. Academic Press, San Diego, CA. Fuerst, T. R., Niles, E. G., Studier, F. W., and Moss, B. (1986). Eukaryotic transient-expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 83, 8122-8126. Gershon, P. D., and Moss, B. (1992). Transition from rapid processive to slow non-processive polyadenylation by vaccinia virus poly(A) polymerase catalytic subunit is regulated by the net length of the poly(A) tail. Genes Dev. 6, 1575-1586. Gershon, P. D., and Moss, B. (1993a). Stimulation of poly(A) tail elongation by the VP39 subunit of the vaccinia virus-encoded poly(A) polymerase. J. Biol. Chem. 268, 2203-2210. Gershon, P. D., and Moss, B. (1993b). Uridylate-containing RNA sequences determine specificity for binding and polyadenylation by the catalytic subunit of vaccinia virus poly(A) polymerase. E M B O J. 12, 4705-4714. Gershon, P. D., and Moss, B. (1996). Expression, purification, and characterization of vaccinia virusencoded RNA and poly(A) polymerases. In "Methods in Enzymology" (L. C. Kuo, D. B. Olsen, and S. S. Carroll, eds.), Vol. 275, pp. 208-227. Academic Press, San Diego, CA. Gershon, P. D., Ahn, B.-Y., Garfield, M., and Moss, B. (1991). Poly(A) polymerase and a dissociable polyadenylation stimulatory factor encoded by vaccinia virus. Cell (Cambridge, Mass.) 66, 1269-1278. Guan, K. L., and Dixon, J. E. (1991). Eukaryotic proteins expressed in Escherichia coli: An improved thrombin cleavage and purification procedure of fusion proteins with glutathione S-transferase. Anal. Biochem. 192, 262-267. Hercyk, N., Horikami, S. N., and Moyer, S. A. (1988). The vesicular stomatitis virus L protein possesses the m R N A methyltransferase activities. Virology 163, 222-225. Hodel, A., Gershon, P. D., Shi, X., and Quiocho, F. A. (1996). The 1.85A structure of vaccinia protein VP39: A bifunctional enzyme that participates in the modification of both m R N A ends. Cell (Cambridge, Mass.) 85, 247-256. Holm, L., and Sander, C. (1995). DNA polymerase /3 belongs to an ancient nucleotidyltransferase superfamily. Trends Biochem. Sci. 20, 345-347. Kates, J., and Beeson, J. (1970). Rhbonucleic acid synthesis in vaccinia virus. II. Synthesis ofpolyriboadenylic acid. J. Mol. Biol. 50, 19-33. Kates, J., and McAuslan, B. R. (1967). Poxvirus DNA-dependent RNA polymerase. Proc. Natl. Acad. Sci. U.S.A. 58, 134-141. Kealey, J. T., Gu, X., and Santi, D. V. (1994). Enzymatic mechanism oftRNA (m5U54)methyltransferase. Biochimie 76, 1133-1142. Kossykh, V. G., Schlagman, S. L., and Hattman, S. (1993). Conserved sequence motifDPPY in region IV of the phage T4 Dam DNA-[N6-adenine]-methyltransferase is important for S-adenosyl-Lmethionine binding. Nucleic Acids Res. 21, 3563-3566. Kovacs, G. R., and Moss, B. (1996). The vaccinia virus H5R gene encodes late gene transcription factor 4: Purification, cloning and overexpression. J. Virol. 70, 6796-6802. Langberg, S. R., and Moss, B. (1981). Post-transcriptional modifications of mRNA. J. Biol. Chem. 256, 10054-10060. Lingner, J., and Keller, W. (1993). 3'-End labeling of RNA with recombinant yeast poly(A) polymerase. Nucleic Acids Res. 21, 2917-2920. Manley, J. L. (1995). Messenger RNA polyadenylylation: A universal modification. Proc. Natl. Acad. Sci. U.S.A. 92, 1800-1801. Martin, G., and Keller, W. (1996). Mutational analysis of mammalian poly(A) polymerase identifies a region for primer binding and a catalytic domain, homologous to the family X polymerases, and to other nucleotidyltransferases. E M B O J . 15, 2593-2603. Martin, S. A., Paoletti, E., and Moss, B. (1975). Purification o f m R N A guanylyltransferase and rnRNA (guanine-7-)-methyltransferase from vaccinia virions. J. Biol. Chem. 250, 9322-9329.

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Moss, B. (1990). Regulation of vaccinia virus transcription. Annu. Rev. Biochem. 59, 661-688. Moss, B. (1996). Poxviridae: The viruses and their replication. In "Fields Virology" (B. N. Fields, D. M. Knipe, and P. M. Howley, eds.), pp. 2637-2671. Lippincott-Raven, Philadelphia. Moss, B., and Rosenblum, E. (1974). Vaccinia virus polyriboadenylate polymerase: Covalent linkage of the product with polyribonucleotide and polydeoxyribonucleotide primers. J. Virol. 14, 86-98. Moss, B., Rosenblum, E. N., and Paoletti, E. (1973). Polyadenylate polymerase from vaccinia virions. Nature (London), New Biol. 245, 59-63. Moss, B., Rosenblum, E. N., and Gershowitz, A. (1975). Characterization of a polyriboadenylate polymerase from vaccinia virions. J. Biol. Chem. 250, 4722-4729. Moss, B., Barbosa, E., and Keith, J. M. (1979). Specificity ofmRNA methyltransferases. In "Transmethylation" (E. Usdin, R. T. Borchardt, and C. R. Creveling, eds.), pp. 373-380. Elsevier/NorthHolland, New York. Moss, B., Elroy-Stein, O., Mizukami, T., Alexander, W. A., and Fuerst, T. R. (1990). New mammalian expression vectors (product review). Nature (London) 348, 91-92. Moss, B., Ahn, B.-Y., Amegadzie, B., Gershon, P. D., and Keck, J. G. (1991). Cytoplasmic transcription system encoded by vaccinia virus (minireview). J. Biol. Chem. 266, 1355-1358. Munyon, W. E., Paoletfli, E., and Grace, J. T.J. (1967). RNA polymerase activity in purified infectious vaccinia virus. Pro& Natl. Acad. Sci. U.S.A. 58, 2280-2288. Nevins, J. R. and JoHik, W. K. (1975). Poly(A) sequences ofvaccinia virus messenger RNA: Nature, mode of addition and function during translation in vitro and in vivo. Virology 63, 1-14. Nevins, J. R., and Joklik, W. K. (1977). Isolation and partial characterization of the poly(A) polymerases from HeLa Cells infected with vaccinia virus. J. Biol. Chem. 252, 6939-6947. O'Shannessy, D. J., Brigham-Burke, M., and Peck, K. (1992). Immobilization chemistries suitable for use in the BIAcore surface plasmon resonance detector. Anal. Biochem. 205, 132-136. Rossmann, M. G., Moras, D., and Olsen, K. W. (1974). Chemical and biological evolution of a nucleotide-binding domain. Nature (London) 250, 194-199. Rottman, F. M., Bokar, J. A., Narayan, P., Shambaugh, M. E., and Ludwiczak, R. (1994). N6-Adenosine methylation in mRNA: Substrate specificity and enzyme complexity. Biochimie 76, 1109-1114. Schluckebier, G., O'Gara, M., Saenger, W., and Cheng, X. (1995). Universal catalytic domain structure of AdoMet-dependent methyltransferases. J. Mol. Biol. 247, 16-20. Schnierle, B. S., Gershon, P. D., and Moss, B. (1992). Cap-specific mRNA (nucleoside-O2'-)methyltransferase and poly(A) polymerase stimulatory activities of vaccinia virus are mediated by a single protein. Proc. Natl. Acad. Sci. U.S.A. 89, 2897-2901. Schnierle, B. S., Gershon, P. D., and Moss, B. (1994). Mutational analysis ofa multifunctional protein, with mRNA 5' cap-specific (nucleoside-2'-O-)-methyltransferase and 3'-adenylyltransferase stimulatory activities, encoded by vaccinia virus. J. Biol. Chem. 269, 20700-20706. Segal, D. M., and Eichler, D. C. (1989). The specificity of interaction between S-adenosyl-L-methionine and nucleolar 2'-O-methyltransferase. Arch. Biochem. Biophys. 275, 334-343. Shi, X., Yau, P., Jose, T., and Gershon, P. D. (1996). Methyltransferase-specific domains within VP39, a bifunctional protein which participates in the modification of both mRNA ends. R N A 1, 88-101. Shuman, S. (1995). Capping enzyme in eukaryotic mRNA synthesis. Prog. Nucleic Acid Res. Mol. Biol. 50, 101-129. Shuman, S., and Moss, B. (1988). Vaccinia virus poly(A) polymerase: Specificity for nucleotides and nucleotide analogs. J. Biol. Chem. 263, 8405-8412. Shuman, S., and Moss, B. (1990). Purification and use of vaccinia virus messenger RNA capping enzyme. In "Methods in Enzymology" (]. E. Dahlberg and J. N. Abelson, eds.), Vol. 181, pp. 170-180. Academic Press, San Diego, CA. Thomson, J. G., and Gershon, P. D. (1995). Use ofvaccinia virus poly(A) polymerase for RNA 3'end labeling with either a chain-terminating nucleotide or a short 3' homopolymer tract. BioTechniques 19, 416-425.

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Verdine, G. (1994). The flip side o f D N A methylation. Cell (Cambridge, Mass.) 76, 197-200. Wei, C. M., and Moss, B. (1975). Methylated nucleotides block 5'-terminus ofvaccinia virus mRNA. Proc. Natl. Acad. Sci. U.S.A. 72, 318-322. Whoriskey, S. K., Usman, N., and Szostack, J. W. (1995). Total chemical synthesis of a ribozyme derived from a group I intron. Proc. Natl. Acad. Sci. U.S.A. 92, 2465-2469. Yue, D., Maizels, N., and Weiner, A. (1996). CCA-adding enzymes and poly(A) polymerases are all members of the same nucleotidyltransferase superfamily: Characterization of the CCA-adding enzyme from the archeal hyperthermophile Sulfolobus shibatae. R N A 2, 895-908.

A Genetic Approach to Mapping Coding Region Determinants of mRNA Instability in Yeast Aidan N. Hennigan

Allan Jacobson Department of Molecular Genetics and Microbiology University of Massachusetts Medical School Worcester, Massachusetts, 01655

Department of Molecular Genetics and Microbiology University of Massachusetts Medical School Worcester, Massachusetts, 01655

This chapter describes a method for mapping coding region sequences in yeast genes that promote rapid messenger R N A (mRNA) decay. The approach exploits the link between the function of such sequence elements and their translation, as well as the dependence of nonsense-mediated m R N A decay on the activity of the UPF1 gene product. The feasibility of the method has been tested with the MA Tot 1 m R N A since an instability determinant was previously localized to a 65nt segment of its coding region. Systematic insertion of in-frame translational termination codons into the MA Tot 1 coding sequence demonstrated that termination of translation 5' to this element stabilized the MATOtl m R N A two- to threefold relative to wild-type transcripts. Translation through the element was sufficient to restore rapid m R N A decay, while internal termination resulted in different extents of m R N A stabilization, depending on the precise location of ribosome stalling. An extension of the mapping protocol to include a detailed mutagenesis of the MA Tot1 instability element (IE) has provided insight into the mechanism by which these sequences act in cis to destabilize mRNA.

I. I N T R O D U C T I O N m R N A turnover is a regulated process dependent on specific trans-acting factors and cis-acting sequences (1-4). The latter include sequences that promote either m R N A stability or instability, and their localization generally involves the construction of chimeric genes, composed of segments encoding both stable and 149 m R N A Formation and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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unstable mRNAs, and the analysis of in vivo decay rates of the resulting chimeric mRNAs. In Saccharomyces cerevisiae this procedure, combined with deletion and mutational analyses, has successfully localized IEs to the coding regions of the M A T c z l (5,6), HIS3 (7), STE3 (8), S P 0 1 3 (9), and RPL2 (10) mRNAs, the 3' untranslated regions of the STE3 (8) and M F A 2 (11) mRNAs, and the 5' untranslated regions of the PPR1 (12) and SDH2 (13) mRNAs. While this approach is useful, its limitations include its labor intensiveness and the possible generation of significant perturbations to overall m R N A structure (see Table I). Development of a new IE mapping protocol, which we term nonsense codon mapping, was made possible by the identification of gene products required for nonsense-mediated m R N A decay (14-20) and prior demonstration of an intimate link between m R N A decay and translation. The latter is exemplified by the location of some IEs to m R N A coding regions (5,7,8,21), the accelerated degradation of mRNAs promoted by premature translational termination (19,22,23), the association of trans-acting degradation factors with ribosomes (2427), and the stabilization of mRNAs that occurs when translation is inhibited (1,28-33). A particularly relevant link that formed the basis for the development of nonsense codon mapping was a set of experiments showing that normally unstable chimeric PGK1-MATczl and A C T 1 - M A T o l l mRNAs were stabilized when an in-frame translational termination codon was inserted at the junction of the sequences from the respective stable and unstable m R N A components of the chimeras (6). Since MA Tcz I IE activity required its translation, we hypothesized that translation-dependent IEs could, in general, be mapped by insertion of in-frame nonsense codons (as depicted for a genetic m R N A in Fig. 1). Those inserted 5' to the element should stabilize the transcript, while downstream codons should be inconsequential to the m R N A half-life. To circumvent activation of the nonsensemediated m R N A decay pathway, half-lives of allelic transcripts could be measured in a strain deficient for Upflp, a trans-acting factor essential for activity of this decay pathway (17-19). We have tested the feasibility of this new mapping method with the use of the MA Tczl gene since the precise coding sequence location of its 65-nt IE has been defined (5,6). We find that, in addition to mapping the element, nonsense codon mapping provides insight into the mechanism oflE functioning and facilitates further understanding of the interrelationships of m R N A decay and translation.

II. E X P E R I M E N T A L

METHODOLOGY

A standard nonsense codon mapping protocol, as applied to the M A T a l gene, is as follows:

Table I Comparison o f Methods for Identification o f c/s-Acting Determinants o f m R N A Instability Method

Advantages

Disadvantages

Chimeric gene approach

Demonstrated ability for mapping IEs

May perturb overall m R N A structure Requires construction of chimeric genes Requires further deletion and/or point mutational analysis to define IE

Deletion/mutational analysis

Demonstrated ability for mapping IEs Simple to execute

Requires prior knowledge of IE's general location within gene May be dependent on chimeric gene approach

Nonsense codon mapping

Minimal perturbations to overall m R N A structure Does not require construction of chimeric genes Demonstrated ability for mapping IEs Simple to execute

May be unable to distinguish different coding sequence IEs present in a single gene Identifies only those IEs whose activity require translation Defining 5' and 3' boundaries of IE will require further deletion analysis

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mRNA

stability:

S

S

S

?

U

U

Figure I The nonsense mapping approach to localizing coding region instability elements. In separate constructs, in-frame nonsense codons are systematically inserted throughout the coding region of a gene encoding an unstable mR.NA. In a upflA strain, nonsense codons that are 5' to a coding region instability element are expected to stabilize the mRNA, whereas those that are 3' to the element are expected to have no effect on the inherently rapid decay of the mR.NA. The effects of nonsense codons within the element are discussed in the text. S, mP,.NA stabilized; U, mt(NA remains unstable.

1. A series of MA T a 1 alleles are generated in which single in-flame translational termination codons are located either 5', internal to, or 3' to the coding region IE. While any one of several site-directed mutagenesis protocols may be used, we have had success with the m e t h o d of Kunkel et al. (34). 2. Mutations are verified through D N A sequencing (35), and wild-type and nonsense-containing alleles are then subcloned into an appropriate yeast expression vector. Plasmids used in this study were all derived from pGALMA Tc~ 1 (provided by G. Caponigro and R. Parker; 36), in which expression of the M A T o l l gene from a G A L 1 U A S is regulatable in a substrate-specific manner. 3. Standard yeast transformation protocols (37) are used to introduce the individual pGALMA To~ 1 recombinants into a upflA strain. O u r experiments have successfully used strain yAH01 (MA Ta rpb 1-1 ura3-52 leu2-3,112 upfl : : LEU2; 36) in which transcription of the genomic MA To~l locus is repressed. 4. Half-lives of the allelic MAToll m R N A s are determined according to the methodology of Herrick et al. (30) and Decker and Parker (38). Essentially, after a short burst of m R N A synthesis, transcription is inhibited by simultaneous repression of the G A L l U A S (controlling M A T o l l ) and thermal inactivation of a temperature-sensitive R N A polymerase II. Cells growing in m e d i u m containing 2% raffinose are supplemented with the more readily utilizable substrate galactose, which induces expression of GAL-driven genes, including MA Tee 1. The resulting pulse of transcription is then inhibited by the addition of glucose and the rapid elevation of the culture temperature to 36°C. The latter step inhibits transcription because the cells harbor the rpb 1-1 allele (30). Cells are harvested both prior to the transcriptional pulse and at 0, 3, 6, 9, 12, 18, 25, and 35 min after the inhibition of transcription. Cell pellets are frozen on dry ice and stored at - 7 0 ° C to await P,.NA extractions.

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5. Cellular 1LNA is isolated, for example, by the hot phenol method, and mlLNA decay rates are measured as described elsewhere (39). Electrophoresed tLNA is transferred to Zeta Probe membranes (Bio tLad, Hercules, CA), and blots are probed using a MA To~1 D N A fragment labeled by the use of a random priming protocol (40). To verify efficient inhibition of transcription at the nonpermissive temperature, we simultaneously probe blots for a short-rived transcript, such as the STE2 mtLNA. 6. Northern blots are quantitated with a blot analyzer such as a Betascope (Betagen, Waltham, MA) or a PhosphorImager (Bio tLad, Hercules, CA; Molecular Dynamics, Sunnyvale, CA). Half-life values are obtained by expressing data as the log10 of the percentage of each m R N A remaining versus time at 36~ To ensure a meaningful interpretation of the data, all values presented should represent the average ___1 standard deviation of at least two half-life determinations for each of at least two independently isolated transformants.

III. E X P E R I M E N T A L R E S U L T S A. THE CODING SEQUENCE LOCATION OF THE m a T a l IE is IDENTIFIABLE WITH THE USE OF IN-FRAME TRANSLATIONAL TERMINATION CODONS The MATo~I gene encodes an inherently unstable m R N A having a halflife of approximately 5 min (4,6,30). Chimeric gene and deletion analyses previously defined a 65-nt IE, spanning nt 201 to 266, that is capable of destabilizing both the m R N A and a heterologous gene in cis (5). The IE is bipartite, containing a 33-nt segment highly enriched for rare codons preceding a 32-nt, predominantly AU-rich stretch, and both segments are necessary for element activity (5). To establish the feasibility of our new approach to mapping coding region IEs, we first determined the effect that inserting in-flame nonsense codons at positions 18, 52, and 139 had on the MATo~I mlKNA half-life (Fig. 2A). The first two nonsense codons prevent translation of the IE, whereas that at position 139, which follows the IE, allows translation of approximately 79% of the MA Toz 1 coding region. In the upflA strain, in which nonsense-mediated mlKNA decay is blocked, nonsense codons at positions 18 and 52 stabilized the MATo~I transcript almost threefold to half-lives o f ~ 1 3 . 8 and ~ 12.5 min, respectively. However, ribosome translation to codon 139 resulted in an unstable mlKNA having a wild-type half-life of ---5.2 min (Fig. 2A). In all strains, the STE2 mlKNA had a half-life of "--5 to 6 min (data not shown), ruling out the possibility that extended half-lives for the MATUAG18 and MATUAG52 alleles were a consequence of incomplete inhibition of transcription. These results therefore demonstrate the ability of nonsense codon insertion to localize an IE within the coding sequence. Similar mapping experiments with the yeast HIS3 gene also provide evidence for at least one

154

F i g u r e 2 Effects of inhibiting translation 5', internal to, and 3' to the MAToll IE. (A) Schematics, m R N A half-fives, and Northern blots for the wild-type MAToll gene and six nonsense alleles, as measured in the upflA strain. The locations of single in-frame translational termination codons relative to the bipartite IE (shaded regions) are shown. MATc~ 1 alleles are named according to the nucleotide triplet changed to a nonsense codon. (B) Summary of the consequences of premature translational termination on the decay of the M A T s 1 m R N A .

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coding sequence IE, located in the gene's 3' proximal end (A. H. and A. J., unpublished results). We further developed nonsense codon insertion as a general approach to mapping coding region IEs by determining the consequences on m R N A stability of inhibiting translation within an IE. In-frame translational terminators were inserted at codons 67, 79, and 85 of the MA Toel gene (Fig. 2A). MA 7UAG67 terminates translation immediately 5' to the IE; MA TUGA79 changes the first codon in the 3' 32-nt AU-rich sequence to a terminator, such that ribosomes translate only the rare codon segment of the IE; and MA TUAG85 terminates translation after ribosomes progress through 60% of the AU-rich sequence, equivalent to 80% of the entire element. Ribosome stalling at codons 67 and 79 yielded transcripts with half-lives of 9.8 and 9.4 min, respectively, in the upflA strain, indicating an approximately twofold stabilization relative to wild-type m R N A decay (Fig. 2A). However, translation to codon 85 yielded an m R N A half-life of---4 min, indicating that ribosome progression had been sufficient to promote normal decay of the MA Toe1 transcript (Fig. 2A). Hence internal termination resulted in different extents o f m R N A stabilization, depending on the precise location of ribosome stalling, and indicated that while insertion of nonsense codons identifies a general localization of an IE, delimiting the actual 5' and 3' boundaries may require deletion analyses in the vicinity of the IE. Figure 2B summarizes the IE mapping experiments, plotting the half-lives of the six MA Toe 1 transcripts against their coding sequence locations. The abrupt transition to more rapid m R N A decay defines the IE location by virtue of the dependence of its phenotypic expression on translation.

B. NONSENSE CODON MAPPING FACILITATES ANALYSES OF THE MECHANISM OF I E ACTIVITY The mapping procedure described here also provides insight into the interrelationships o f m R N A decay and translation and the mechanism by which the MA Toe 1 IE functions as an m R N A destabilizer. The abrupt twofold decrease in m R N A stability that occurred as ribosomes traversed the IE (see Fig. 2B) was investigated further in an attempt to map this stability transition more precisely. To this end, constructs with single in-frame UAG translational termination codons at positions 76, 77, 78, 80, 81, 82, 83, and 84 were constructed by oligonucleotide site-directed mutagenesis (Fig. 3A). This set of constructs, together with MA TUGA79 and MA 7UAG85 (Fig. 2A), allows ribosomes translating the respective mRNAs to progress through the IE in increments of one codon. The effects of such ribosome progression on m R N A decay rates were determined in the upflA strain. Half-lives of-'--9 to 10 min were obtained for the allelic MATUAG76, MATUAG77, and MAgUAG78 mRNAs (Fig. 3A), similar to the previously determined value for the MA TUGA79 transcript (Fig. 2A), and an intermediate decay of---7.5 min was obtained for the MAgUGA80 transcript (Fig. 3A). In

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contrast, the MATUAG81, MATUAG82, MATUAG83, and MATUAG84 mR.NAs were unstable, with half-lives of 4 to 5 min (Fig. 3A), similar to that determined for the MA 7UAG85 allele (Fig. 2A). These results are summarized in a bar plot of the half-lives obtained (Fig. 3B), which clearly indicates a transition in m R N A stability as translation proceeds over a two-codon interval of the IE, codons 80 and 81. The transition region is bordered by an upstream segment wherein translation termination stabilizes the mR.NA and by a downstream segment in which termination has no effect on mR.NA decay. The results demonstrate at the molecular level a clear relationship between mR.NA stability and m R N A translation, supporting previous studies that have linked the two processes (2,3,5,7,8,18,19,22,30,37). We speculated that this two-codon transition in MA Tol 1 mR.NA half-life reflected a requirement for the ribosome (or a ribosome-borne factor) to interact with a downstream sequence, or bound factor, and that this interaction provided the signal for rapid decay of the transcript. Hence, in experiments designed to test this hypothesis and further extend the mapping protocol, we deleted or duplicated codon 82 of the nonsense alleles that span the stability transition region, i.e., MATUAG78, MATUGA79, MATUAG80, and MA TUAG81. Half-life determinations for the respective transcripts in the upflA strain indicated the stability transition to have shifted in the 5' direction in the deletion mutants, while a 3' directional shift was evident with the insertion mutants (36). These results support the proposed model of an IE-mediated interaction between a translating ribosome and a downstream element and demonstrate the utility of this mapping protocol for high-resolution analysis of the mechanism oflE functioning.

IV. D I S C U S S I O N A N D C O N C L U S I O N S A new method that allows localization of cis-acting coding region IEs in yeast genes encoding inherently unstable m R N A s is described in this chapter. The procedure provides insight into the mechanism by which the IE of the MA Tc~l gene acts as a destabilizer and how decay of the encoded mlLNA is related to its translation. While this approach was developed using a upflA strain, we anticipate that strains disrupted for any nonessential gene coding a trans-acting factor specific to nonsense-mediated m R N A decay will provide an alternative for this protocol (1,4). Localization experiments, as summarized in Figure 2B, demonstrated that insertion of nonsense codons 5' to the MA Toz 1 IE stabilized the transcript twoto threefold, while location of a translation termination codon downstream of the IE, by allowing translation to proceed through the element, had no effect on transcript half-life. Hence, the successful localization of this IE by nonsense codon insertion provides a valuable tool in the search for coding-region IEs in other yeast genes specifying inherently unstable mlKNAs. The m_RNA half-lives obtained in our studies are in agreement with those of previously reported MA Tc~ 1 deletion experiments (5). For example, in experiments by Caponigro et al. (5), in which

158

F i g u r e 3 Mutagenesis of the rare codon/AU-rich boundary of the M A T a 1 IE. (A) Schematic of the eight allelic forms of the IE (each of which contains a single in-frame translational termination codon), their half-lives, and Northern blots as obtained from the upflA strain. (B) Locations of in-frame translational termination codons in the MATc~ 1 IE versus m R N A half-life measured in the upflA strain. The translation-dependent stability transition is bordered by codons 79 and 81.

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the 65-nt element was removed from the gene, the m R N A was stabilized twofold. A threefold stabilization occurred if the AU-rich sequences immediately 3' to the IE were also deleted (5). Furthermore, the extents of message stabilization were similar irrespective of whether the 3' 32-nt AU-rich sequence of the element was physically removed or nonsense codons were used to block its translation (5) (Figs. 2 and 3). The n e w mapping procedure offers certain advantages over the chimeric gene approach in that it causes minimal perturbations to overall m R N A structure, since constructs differ from the wild-type transcript by only 1 to 3 nt. The procedure does not, however, delimit the IE and requires further deletion analyses to map the 5' and 3' boundaries of a localized element. A potential difficulty with nonsense codon mapping may be its inability to distinguish different IEs in a gene containing multiple destabilizing sequences. Furthermore, because of the translational dependency of the mapping protocol, only those elements whose destabilizing activities require ongoing translation may be characterized. Table I provides a summary of the relative advantages and disadvantages of the various IE mapping protocols. Based on our observations, we conclude that the new m e t h o d o l o g y should be most informative w h e n used as a first step in mapping coding region IE(s) within a previously unstudied gene. Subsequent deletion a n d / o r mutational analysis could then facilitate IE characterization. T h r o u g h the identification o f IEs, the n e w mapping protocol should provide insight into the c i s - a c t i n g determinants of m R N A stability.

ACKNOWLEDGMENTS Work in the authors' laboratory is supported by grants to A.J. from the National Institutes of Health (GM27757) and the American Cancer Society (NP-843) and by a senior postdoctoral fellowship to A.H. from the Massachusetts Division of the American Cancer Society.

REFERENCES 1. Jacobson, A., and Peltz, S. W. (1996). Annu. Rev. Biochem. 65, 693-740. 2. Peltz, S. W., Brewer, G., Bernstein, P., and Ross, J. (1991). C R C Crit. Rev. Eukaryotic Gene Expression 1, 99-126. 3. Ross, J. (1995). Microbiot. Rev. 59, 423-450. 4. Peltz, S. W., and Jacobson, A. (1993). In "Control of mRNA Stability" (G. Brawerman and J. Belasco, eds.), pp. 291-328. Academic Press, San Diego, CA. 5. Caponigro, G., Muhlrad, D., and Parker, R. (1993). Mol. Cell. Biol. 13, 5141-5148. 6. Parker, R., and Jacobson, A. (1990). Proc. Natl. Acad. Sci. U.S.A. 87, 2780-2784. 7. Herrick, D., and Jacobson, A. (1992). Gene 114, 35-41. 8. Heaton, B., Decker, C., Muhlrad, D., Donahue, J., Jacobson, A., and Parker, R. (1992). Nucleic Acids Res. 20, 5365-5373.

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28. 29. 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40.

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Surosky, R. T., and Esposito, R. E. (1992). Mol. Cell. Biol. 12, 3948-3958. Presutti, C., Villa, T., Hall, D., Pertica, C., and Bozzoni, I. (1995). E M B O J . 14, 4022-4030. Muhlrad, D., and Parker, R. (1992). Genes Dev. 6, 2100-2111. Pierrat, B., Lacroute, F., and Losson, R. (1993). Gene 131, 43-51. Cereghino, G. P., Atencio, D. P., Saghbini, M., Beiner, J., and Scheffler, I. E. (1995). Mol. Biol. Cell 6, 1125-1143. Cui, Y., Hagan, K. W., Zhang, S., and Peltz, S. W. (1995). Genes Dev. 9, 423-436. He, F., and Jacobson, A. (1995). Genes Dev. 9, 437-454. Lee, B.-S., and Culbertson, M. R. (1995). Proc. Natl. Acad. Sci. U.S.A. 92, 10354-10358. Leeds, P., Peltz, S. W., Jacobson, A., and Culbertson, M. R. (1991). Genes Dev. 5, 2303-2314. Leeds, P., Wood, J. M., Lee, B.-S., and Culbertson, M. R. (1992). Mol. Cell. Biol. 12, 2165-2177. Peltz, S. W., Brown, A. H., and Jacobson, A. (1993). Genes Dev. 7, 1737-1754. Peltz, S. W., He, F., Welch, E., and Jacobson, A. (1994). Prog. Nucleic Acid Res. Mol. Biol. 47, 271-298. Shyu, A.-B., Belasco, J. G., and Greenberg, M. E. (1991). Genes Dev. 5, 221-231. Hagan, K. W., Ruiz-Echevarria, M. J., Quan, Y., and Peltz, S. W. (1995). Mol. Cell. Biol. 15, 809-823. Losson, R., and Lacroute, F. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 5134-5137. Atkin, A. L., Altamura, N., Leeds, P., and Culbertson, M. R. (1995). Mol. Cell. Biol. 6, 611-625. Bandyopadhyay, R., Coutts, M., Krowczynska, A., and Brawerman, G. (1990). Mol. Cell. Biol. 10, 2060-2069. Caruccio, N., and Ross, J. (1994).j. Biol. Chem. 269, 31814-31821. Peltz, S. W., Trotta, C., Feng, H., Brown, A., Donahue, J., Welch, E., and Jacobson, A. (1993). In "Protein Synthesis and Targeting in Yeast" (A. J. P. Brown, M. F. Tuite, andJ. E. G. McCarthy, eds.), pp. 1-10. Springer-Verlag, Berlin. Baim, S. B., Pietras, D. F., Eustice, D. C., and Sherman, F. (1985). Mol. Cell. Biol. 5, 1839-1846. Beelman, C. A., and Parker, R. (1994).J. Biol. Chem. 269, 9687-9692. Herrick, D., Parker, R., and Jacobson, A. (1990). Mol. Cell. Biol. 10, 2269-2284. Peltz, S. W., Donahue, J. L., and Jacobson, A. (1992). Mol. Cell. Biol. 12, 5778-5784. Ross, J. (1995). Microbiol. Rev. 59, 423-450. Wisdom, R., and Lee, W. (1990). J. Biol. Chem. 265, 19015-19021. Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987). In "Methods in Enzymology" (R. Wu and L. Grossman, eds.), Vol. 154, pp. 367-382. Academic Press, Orlando, FL. Sanger, F., Nicklen, S., and Coulson, A. R. (1977). Proc. Natl. Acad. Sci. U.S.A. 74, 5463-5467. Hennigan, A. N., and Jacobson, A. (1996). Mol. Cell. Biol. 16, 3833-3843. Schiestl, R. H., and Gietz, R. D. (1989). Curr. Genet. 16, 339-346. Decker, C.J., and Parker, R. (1993). Genes Dev. 7, 1632-1643. He, F., Peltz, S. W., Donahue, J. L., Rosbash, M., andJacobson, A. (1993). Proc. Natl. Acad. Sci. U.S.A. 90, 7034-7038. Feinberg, A. P., and Vogelstein, B. (1983). Anal. Biochem. 132, 6-13.

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Identification of the Protein That Interacts with the 3' End of Histone m R N A William F. Marzluff

Michael L. Whitfield

Program in Molecular Biology and Biotechnology University of North Carolina Chapel Hill, North Carolina 27599

Program in Molecular Biology and Biotechnology University of North Carolina Chapel Hill, North Carolina 27599

Zbigniew Dominski

Zeng-Feng Wang

Program in Molecular Biology and Biotechnology University of North Carolina Chapel Hill, North Carolina 27599

Program in Molecular Biology and Biotechnology University of North Carolina Chapel Hill, North Carolina 27599

I. I N T R O D U C T I O N The replication-dependent histone mtLNAs are unique among metazoan mtLNAs in that they are the only mtLNAs that do not have poly(A) tails. Histone genes are also the only genes that as a class lack introns (1). The replicationdependent histone genes for all five histone proteins are tightly linked and in mammals are present in two distinct clusters (2-5). There are a total of 50-60 genes for the replication-dependent histone mtLNAs, and these mtkNAs are coordinately regulated during the cell cycle. The concentration ofhistone mtLNAs increases 30to 50-fold as cells enter S phase and decreases again in G2 prior to mitosis (6). Much of this regulation is posttranscriptional and is mediated by the 3' end of the histone mtLNA (7). The biosynthesis of histone mtLNA is much simpler than the synthesis of other mtLNAs, since these genes lack introns. The only processing step necessary is the cleavage of the pre-mtLNA to release the mtLNA from the chromatin and form the mature histone 3' end (8). The 3' end of the histone mtLNA has been conserved in all metazoans. The conserved region is the last 26 nt of the mtLNA, and includes a six-base stem and four-base loop (Fig. 1). The sequence of the stem has been particularly conserved, and the stem sequence is essential for the function ofhistone mtLNA (9). Thus it is likely that this conserved sequence forms a distinct three-dimensional structure that is recognized by the trans-acting factors. 163 m R N A F o r m a t i o n and Function Copyright 9 1997 by Academic Press. All rights of reproduction in any form reserved.

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y2(~

(~

N4

Iu AI YR YR YR 5' C C A A A G

CACCCA

3'

Figure 1 The consensus sequence for the histone stem-loop present in all metazoan replicationdependent histone genes is shown, with the invariant residues boxed.

A. T H E 3 ' E N D OF H I S T O N E M R N A

Is INVOLVED IN ALL

ASPECTS OF HISTONE MRNA METABOLISM Since the poly(A) tail plays a critical role in several metabolic steps in general m R N A metabolism (10), it is logical that the 3' end ofhistone m R N A has assumed these roles for the histone mRNAs. Following processing of the pre-mtLNA, the histone m R N A must be transported to the cytoplasm, and the 3' end is required for efficient transport (11). Indeed, transport of histone m R N A may be directly coupled to the processing reaction (12). Like the poly(A) tail, the 3' end ofhistone m R N A is necessary for efficient localization of the m R N A to polyribosomes (13,14). Finally, an important regulatory step in histone mlLNA metabolism is regulation of the half-life of the histone mRNA. For proper regulation of histone m R N A half-life, the 3' end of the m R N A is necessary and sufficient (15). During the cell cycle there is regulation of both histone m R N A 3' end formation (6,16) and histone m R N A half-life (6). As cells progress from G1 to S phase, there is an increase in the efficiency of histone m R N A 3' end formation (6,16). Thus, in G1 the histone genes are transcribed at about 30% of the transcription rate in S phase (17,18) but only a small percentage of the transcripts are processed, while the processing efficiency increases dramatically as cells enter S phase (6). At the end of S phase the half-life of histone m R N A is reduced from 45-60 rain to about 10 min. (6) This change in half-life can be induced by treating S-phase cells with inhibitors of DNA replication (19). Since the 3' end of the histone m R N A is involved in so many aspects of histone mRA'qA metabolism, we have focused our efforts on identifying and understanding the function of the factor(s) that interact with the 3' end of replicationdependent histone mRNAs. In this chapter, we discuss the methods used to isolate and characterize the factor(s), i.e., the stem-loop binding protein (SLBP), that interact with the 3' end

10. Histone mRNA 3' End

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of histone mRNA. W e start with the initial detection of a factor using mobility shift assays and ultraviolet (UV) cross-linking to identify the SLBP polypeptide and end with the cloning of the SLBP using the yeast three-hybrid system (20).

II. D E T E C T I N G S P E C I F I C PROTEIN-RNA INTERACTIONS The replication-dependent histone mR.NAs end in a highly conserved stemloop structure. Using mutagenesis and mobility shift assays, we determined the sequence requirements for interaction of the SLBP with the stem-loop (9). The stem-loop contains a potential 6-bp stem and a 4-bp loop. This structure is highly conserved among metazoans, with the base pairs at the bottom and top of the stem being invariant (Fig. 1, boxed nucleotides). The invariant conserved nucleotides are the two GC base pairs at the base of the stem and the UA base pair at the top of the stem. The stem is a critical structure necessary for recognition of the R N A by SLBP. Changing the two G C / G C base pairs at the base of the stem to C G / C G reduces the binding affinity more than 30-fold (9) and abolishes the expression of the histone m R N A in vivo (21). Reversing the stem while leaving the loop unchanged, creating a reverse stem mutant used in many of our studies, also abolishes binding activity (9). There is also a well-conserved consensus sequence for the loop, although single insertions and mutations in the loop are tolerated, reducing the affinity for SLBP only 5- to 10-fold (9,22). The second and fourth bases of the loop are variable between different histone genes. The first position of the loop is always uridine, except in Caenorhabditis elegans, whose histone genes have a cytosine in the first position of the loop (23). The second position in the loop is usually a uridine, although there is sometimes a cytosine or an adenosine. The third position in the loop is invariant (a U) in all histone genes examined. The fourth position in the loop can be either uridine, cytosine, or adenosine. Interestingly, there is no guanosine in the loop of any of the histone genes (and guanosine is never found in the five nucleotides on either side of the s