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.

This Page Intentionally Left Blank

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--- 100 nt) are used in the binding reaction, the background is often much higher and treatment of the reaction with 1KNase prior to electrophoresis is necessary to resolve a specific complex (22,31). The use of small probes, where possible, is highly recommended once the minimal IKNA binding site has been defined.

C. USE OF COMPETITION ASSAYS TO ESTIMATE RELATIVE AFFINITIES OF DIFFERENT R N A SEQUENCES Competition assays can test the specificity of the complexes and the relative affinity of the R N A binding protein for wild-type and mutant R N A targets. Often multiple complexes will be resolved on the gel, of which only a few will be specific and several may be nonspecific. Often the nonspecific complexes are due to an abundant low-affinity protein, and they are not competed even with a homologous competitor. Competitions are performed by adding excess cold competitor to the

10. Histone mKNA 3' End

169

mobility shift reactions before adding the protein sample. The amount of cold competitor needed to compete the shift must be determined empirically for each system. This is accomplished by adding increasing concentrations of competitor ranging from 1-fold to as high as 10,000-fold over the concentration of the probe (Fig. 2A, lanes 1-6). Since the mobility shift assay requires a relatively high affinity to allow formation of a complex that is stable during electrophoresis, many mutant substrates give no detectable complex formation when tested directly in the mobility shift assay. However, these mutants often have specific affinity for the protein, and the affinities of different mutant RNAs differ. It is possible to use the competition assays to estimate the relative affinity of the mutant RNAs (9). This is done by using a wild-type sequence as a probe and the different mutant sequences as competitors (Fig. 2A, lanes 7-12). By comparing the amount of wild-type competitor R N A necessary to compete binding 50% with the amount of mutant R N A necessary to compete binding to the wild-type probe 50%, it is possible to estimate the relative affinity of different mutant RNAs. This allowed us to distinguish between mutations that reduced the binding affinity 20-fold from mutants (AI'3SL) that reduced the affinity several orders of magnitude (2'OCH3SL), even though neither of the mutant probes formed a detectable complex in a mobility shift assay (9) (Fig. 2B).

D. SUPERSHIFTS If antibodies to the protein are available, it is possible to determine which complexes contain which specific proteins. The R N A binding protein (RBP)R N A complex, when incubated with antibody, forms an A b - R B P - R N A complex that migrates more slowly on the gel. The original complex is no longer detected (Fig. 3, lanes 5 and 11). Antibodies sometimes block the binding of the protein to the RNA, abolishing complex formation. The supershift is performed as described for the mobility shift above, except that after the 10-min incubation at 4~ purified antibodies or crude sera are added to the reaction and the reaction is incubated at room temperature for 5 rain. The complexes are then resolved on a nondenaturing gel. With the use of antibodies raised to a peptide corresponding to the C terminus of SLBP, the complex observed in the mobility shift assay is specifically shifted to a slower migrating form, indicating that the protein detected in the mobility shift assay is recognized by the anti-SLBP antibody (Fig. 3, lanes 5 and 11). To demonstrate the specificity of the supershift, the formation of the antibody complex can be competed by preincubation of the antibody with the antigen. The preincubated antibody should no longer shift the complex. Similarly, antibodies directed against another protein should not affect the complex. Both of these control experiments are essential since often the R N A binding proteins can be

U U

U

U C

UA CG UA CG GC AAAAA G C ACCCA WT SL

Competitor

U

U

C

UA CG UA CG GC GAA G C A5' A3' SL

Competitor

Figure 2 Binding of the SLBP measured by the mobility shift assay. (A) Fifteen femtomoles of radiolabeled stem-loop probe was incubated with 15/.,g of protein from a nuclear extract. The amounts of unlabeled competitor R N A (WT, lanes 1-6 or a mutant with deletions 5' and 3' of the stem-loop, lanes 7-12) are indicated above each lane. The structure of the competitors is shown. No mobility shift was observed if the mutant R N A was labeled. (13) The ability of different competitor RNAs to compete for binding of the SLBP to the wild-type stem-loop was determined by PhosphorImager (Molecular Dynamics, Sunnyvale, CA) analysis of data obtained as in panel A. The structure of the competitors is shown below the graph. The 2'OCH3 competitor had the same sequence, with 2'OCH3 ribonucleotides at each base.

171

10. Histone mtLNA 3' End

B O

100

......

O ......

O

8O B

60 t'n B

40 20 t

2'OCH3

..... 0

o..-.-o

~. . . . . . . . . 1

e I,

10

,

D

~

I

~-.-

I-

10 2

10 3

10 4

Competitor

uu c

u

UA

i

c

AU

uA co

CG GC

uc

9 i

AAAAA G C ACCCA A1, 3 SL Competitor Figure 2

GAAIC

ACCCA

Reverse Stem SL Competitor (C0ntinued)

"sticky" and it is possible to get nonspecific shifts. Thus the complex is not shifted by the preimmune serum (Fig. 3, lanes 4 and 10) or by an antibody to the C terminus of cyclin D3 (Fig. 3, lanes 3 and 9). The supershift can be competed by preincubation of the antibody with 1 /zg of the C terminal peptide prior to the supershift assay (Fig. 3, lanes 6 and 12). Identical results were obtained with SLBP extracted from nuclei (Fig. 3, lanes 1-6) and polyribosomes (Fig. 3, lanes 7-12).

F i g u r e 3 Supershift of the S L B P - R N A complex. A mobility shift assay was performed using a nuclear (lanes 2-6) or polyribosomal extract (lanes 8-12) and the radiolabeled stem-loop probe. In lanes 3 and 9 antisera to mouse cyclin D3 was added to the reaction, and in lanes 4 and 10 preimmune sera for the anti SLBP was added. In lanes 5 and 11, antibody against the C terminus of SLBP was added. In lanes 6 and 12, 1 /~g of peptide was added to the anti-SLBP antibody prior to addition to the reaction. Lanes 1 and 7 are the radiolabeled probe.

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E. DETECTING PROTEINS BY U V CROSS-LINKING The proteins that interact with specific 1KNA sequences can be identified by UV cross-linking. This allows one to identify the apparent molecular weights of the polypeptides that contact the 1KNA. Methods for UV cross-linking 1KNA to proteins have been extensively reviewed elsewhere (32,33). Briefly, the 1KNA probe is labeled with [ol-32p]UTP (labeling with UTP is necessary since uridine is cross-linked more efficiently to protein by UV light), incubated in the extract, and then exposed to UV light. After digestion with RNase, the cross-linked proteins are resolved by sodium dodecyl sulfate (SDS)-gel electrophoresis. By including specific and nonspecific 1KNAs as competitors, one can demonstrate that the cross-linked proteins are bound specifically. We initially identified SLBP as a protein that migrated at 45 kDa on SDS-polyacrylamide gels by UV crosslinking (22). In cases where multiple specific complexes are detected in the mobility shift assay, it is possible to UV cross-link the complexes in the gel to determine which proteins are bound to the 1KNA in each complex. After resolution of the complexes by electrophoresis in a nondenaturing gel, the gel is exposed to film and the position of the complexes is determined. The gel is then placed on a low-wavelength UV light box or in a Stratalinker (Stratagene, La Jolla, CA) and irradiated for 5-10 min. The region of the gel containing each complex is cut out and the 1KNAprotein complex eluted. We use electroelution routinely. The proteins are then treated with 1KNase, resolved on an SDS-polyacrylamide gel, and detected by autoradiography.

F. DETECTING PROTEINS BY NORTHWESTERN BLOTS The proteins that bind specific 1KNA sequences can sometimes be detected by Northwestern blots. In this procedure, the proteins are transferred to a nitrocellulose membrane and then incubated with a radiolabeled probe. This procedure is probably not routinely applicable for identifying an unknown 1KNA binding protein, since the concentrations of the different proteins in the sample are very different and abundant general IKNA binding proteins may overwhelm the signal. The fact that we had previously identified the SLBP by UV cross-linking as a protein that migrated at 45 kDa was very helpful in developing the conditions for the Northwestern assay, and we would be hesitant to use this assay for the initial identification of an unknown 1KNA binding protein. However, we have used it successfully for detecting SLBP (34), particularly in samples that contain large amounts of histone m R N A where SLBP is sequestered by binding to the histone messenger ribonucleoprotein

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(mR.NP) and not detected in a mobility shift assay. We found that renaturing the proteins on the blot greatly improved the detection of SLBP (34). The Northwestern procedure worked only when we used very low concentrations of probe (increasing the probe concentration resulted in a greatly increased number of "nonspecific" bands). Again, using a mutant probe that does not bind the protein of interest (as determined by mobility shift or UV crosslinking assays) helps identify the specific bands. A detailed procedure for the Northwestern procedure that was used successfully in the detection of SLBP is given in reference 34. Based on the intensity of the bands observed with a mobility shift assay and a Northwestern assay on the same sample, we estimate that at most 10% of the SLBP on the filter was available to bind to the probe, further reducing the sensitivity of this assay.

III. I N V I T R O P R O C E S S I N G OF H I S T O N E P R E - m R N A Endonucleolytic cleavage to produce the 3' end of the histone pre-mKNAs is the only processing step leading to translationally active histone mRNAs. Processing requires two sequence elements: the highly conserved stem-loop structure located almost immediately upstream from the cleavage site and a purine-rich region usually located about 10 nucleotides downstream from this site. The cleavage site is generally after a CA, with the site of cleavage determined by U7 small nuclear R N A (snP,.NA) acting as a ruler to specify where cleavage occurs (35). The upstream element is recognized by a factor termed the hairpin binding factor (HBF) (36,37), which is or contains the SLBP (30,38,39). The downstream element is recognized by the U7 small nuclear ribonucleoprotein (snRNP) (40,41). In addition, the cleavage reaction may require a third trans-acting factor, the heatlabile factor, characterized primarily by its sensitivity to a mild heat treatment. The heat-labile factor has not been well characterized and could be one of the components of the active U7 snRNP (42). U7 snP,.NP is indispensable for histone prem R N A processing since its absence or inactivation leads to a complete inhibition of the cleavage reaction (30,40,41). Unlike the U7 snR.NP, HBF is not absolutely required for processing in vitro but only increases the efficiency of the cleavage reaction (30,43,44). The requirement for the stem-loop (and hence probably SLBP) in processing varies for different histone pre-mRNAs, from a high degree of dependence on the SLBP to almost complete independence. This is a function of the cells used and the number of base pairs the U7 snKNP can form with the downstream element (43). One role of HBF may be to help recruit and stabilize U7 snRNP binding. Development of an in vitro system capable of accurate and efficient 3' cleavage of histone pre-m_KNAs has made it possible to study various aspects of this unique

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processing reaction. An in vitro processing system requires two components: nuclear extract and synthetic radiolabeled pre-mlKNA. The procedure used in our laboratory differs slightly from previously published procedures (40,45).

A. PREPARATION OF NUCLEAR EXTRACT FROM MOUSE MYELOMA CELLS Components of the processing machinery reach maximal levels during S phase when the bulk of histone m R N A is produced. It is therefore important that cell cultures are maintained in exponential growth phase and contain a high percentage of dividing cells. Cells with short generation times will have the highest percentage of S-phase cells in an exponentially growing culture (4-6 • 10 s cells/ ml for myeloma cells, which will grow to a density of 1.5 • 106 cells/ml with a generation time of 16 hr). The nuclear extract is prepared according to the procedure of Shapiro et al. (46), modified for preparation of efficient processing extracts (47). All the steps are conducted on ice using cold solutions. 1. Collect cells by centrifugation (800 g). 2. Wash once with phosphate-buffered saline (PBS). 3. Measure the size of the cell pellet (use a conical graduated centrifuge tube) and resuspend the cells in four volumes of hypotonic buffer by gently pipetting. 4. Allow the cells to swell for 5-15 rain and lyse them in a Dounce homogenizer by several strokes with the tight pestle. Homogenization is stopped when more than 90% of the cells are broken, as monitored by phase-contrast microscopy. Both the swelling and lysis of the cells are monitored carefully. The time required for swelling and the number of strokes required for lysis of the cells vary for different cell types. 5. Add 1/10 volume of the restore buffer and spin at 800 g for 2 min. The restore buffer is necessary to keep the nuclei from breaking during the centrifugation. 6. Gently remove the supernatant (cytosolic fraction containing polyribosomes; see Section IV), measure the size of the nuclear pellet, and gently resuspend in 1/2 volume of low-salt extraction buffer. 7. While swirling the tube, add dropwise high-salt extraction buffer until the first evidence of nuclear lysis is observed (final salt concentration is approximately 0.25 M for myeloma cells; for different cell types you may be able to add more or less high-salt buffer).

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William F. Marzluffet al. 8. Extract the nuclei by gently stirring the suspension for 1 hr. The nuclei should not lyse during the extraction. 9. Remove the nuclei by centrifugation at 10,000 g for 30 min. 10. Dialyze the supernatant against 500 volumes of dialysis buffer for 3 - 4 hr, changing the buffer once. 11. Remove the precipitate by centrifugation at 10,000 g for 20 min, aliquot the nuclear extract, and store at - 8 0 ~ or in liquid N2. The extracts are stable for at least 1 year.

From 1 liter of cells (6 X 108) this protocol yields 1-2 ml of nuclear extract with a protein concentration of 5-10 mg/ml. The ability to process in vitro synthesized histone pre-mRNA varies from preparation to preparation and can be as high as 75%. The above protocol is equally suitable for preparation of nuclear extract from other mammalian cell lines. However, conditions need to be optimized for different cells to give maximum processing activity. The parameters to be varied include the final extraction salt concentration, the time of incubation in the hypotonic buffer, and the number of strokes required for sufficient cell lysis. These extracts are also active in splicing and polyadenylation but are not active in transcription by R N A polymerase II. Extracts prepared by extraction at higher salt concentrations that are active in transcription contain an inhibitor of histone pre-mRNA processing, possibly a nonspecific 1KNA binding protein or a nuclease. Addition of excess tlKNA can partially overcome this inhibition (48).

B. In Vitro SYNTHESIS OF 32p-LABELED HISTONE P R E - M R N A The standard in vitro transcription procedure using bacteriophage tLNA polymerase and linear DNA templates allows synthesis of high-quality 32p-labeled premtLNA substrates. In a majority of our studies, a construct was used containing part of the mouse histone H2a-614 gene (49) extended from the PstI site at codon 85 to the HpalI site 35 nucleotides past the U7 binding site. This construct was cloned into pGEM3 under the control of a T7 promoter and linearized at the HindlII site. Transcription of this template by T7 1LNA polymerase results in a 320-nucleotide R N A substrate corresponding to the 3' end of the H2a-614 prem R N A and 19 nucleotides of polylinker preceding the HindlII site. The correctly processed histone m R N A is 266 nt, and the pre-mRNA and processed R N A are readily resolved by gel electrophoresis. Substrates as small as 70 nt have been used, and they are also processed with similar efficiency. Note that the processing substrates have moderate specific activity. The reaction requires a reasonable concentration of the processing substrate, and hence high specific activity substrates are not necessary. These moderate specific activity substrates are stable for at least

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3 weeks after preparation, while high specific activity substrates rapidly undergo radiolysis.

1. P r o t o c o l

1. Prepare the template by cleaving DNA with an appropriate restriction enzyme. N o t e : It is important that KNase not be added during purification of DNA subsequently used for in vitro transcription. The method that relies on ethidium bromide/cesium chloride gradient centrifugation is strongly recommended as a good source of pure plasmid DNA, although crude DNA mini-preparations containing high amounts of contaminating RNA can also be used for in vitro transcription. An alkaline lysis protocol in which the high molecular weight RNA is removed by precipitation with 3 M ammonium acetate and the low molecular weight RNA is removed by chromatography on Sepharose CL4B (Pharmacia, Piscataway, NJ) or Agarose A5M (BIORAD, Richmond, CA) also gives excellent results.

2. Extract the digested DNA twice with phenol equilibrated with TrisHC1 (pH 8) and precipitate with 2.5 volumes of 95% ethanol. 3. Rinse the pellet with 70% ethanol, dry in Speed Vac (Savant Instruments, Farmingdale, NY), and dissolve in sterile water at a final DNA concentration of 1 /xg//xl. 4. The transcription reaction is carried out in a final volume of 25/,1 containing 2.5 /xg of linear DNA template, 40 mM Tris-HC1 pH 7.9, 6 mM MgCI2, 2 mM spermidine, 10 mM DTT, 100/xg/ml bovine serum albumin, 0.5 mM each UTP, ATP, and GTP, 0.05 mM CTP, 20/xCi of [oe-32p]CTP, 25 U ofRNase inhibitor (RNasin), and 25 U of T7 RNA polymerase. Essentially identical results have been obtained with both capped and uncapped transcripts. 5. Incubate at 37~ for 1 hr. 6. Remove unincorporated [32p]CTP by spinning the reaction through a 1-ml G50 Sephadex column. 7. Collect the excluded volume and determine the radioactivity in 1/xl using a scintillation counter. 8. Analyze the transcription product on a sequencing (6-8% acrylamide/ 7 M urea) gel. The resulting labeled transcript is a homogeneous product free of prematurely terminated fragments and does not require further gel purification. The above

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protocol can be used for synthesis of riboprobes by other commonly used and commercially available bacteriophage polymerases (T3, SP6) from constructs containing the appropriate promoters.

C. In Vitro HISTONE P R E - M R N A PROCESSING REACTION Processing in vitro occurs in the absence of divalent ions, and inclusion of high concentrations of EDTA suppresses nuclease activity in the extract. The temperature of incubation and the extract concentration have been optimized for the mouse extracts. An example of a processing reaction is shown in Figure 4. Over 60% of the input pre-mRNA is processed (Fig. 4, lane 2). Processing is dependent on the stem-loop, is greatly reduced by including a 30-mer R N A encoding the stem-loop (Fig. 4, lane 3), and is abolished by an antisense oligonucleotide against U7 snRNA (Fig. 4, lane 4). Heating the extract to 50~ also abolishes processing.

1. Protocol 1. Set up a reaction on ice in a final volume of 20/xl containing 2500 counts per minute (0.33 pmol) of 32p-labeled pre-mlKNA substrate, 20 mM EDTA pH 8, and 17/xl of nuclear extract. Mix by gentle pipetting. 2. Incubate at 32~ for 30 to 45 rain.

Note: The time course experiments revealed that the in vitro cleavage reaction proceeds without a signficant lag time and that virtually all of the final product is generated in the first 30 min of incubation. Extension of this time does not result in further accumulation of the final product but instead could lead to nonspecific degradation of the RNA. 3. Stop the reaction by placing it on ice and adding 400/xl of cold 0.3 M NaOAc. 4. Extract exhaustively once with an equal volume of Tris-equilibrated phenol (pH 8). 5. Precipitate the R N A by adding 2.5 volumes of 95% ethanol and wash the resulting pellet with 70% ethanol. 6. Dry the pellet and dissolve it in 10-15/xl of loading dye (7 M urea, 0.02% bromophenol blue, 0.02% xylene cyanol). 7. Run a 3- to 5-/xl sample on a 6-8% sequencing gel containing 7 M urea (30:1 acrylamide to bisacrylamide).

10. Histone mRNA 3' End

Y U

UGA End

N20.45

179

U N

U-A C-G U-A C-G G-C CCAAAG-CACCCA

PY6 AAAGAGAGCU 3' u u u c u c u c g a U7 snRNP

5'

Figure 4 Biochemical requirements for processing of H2a-614 histone pre-mRNA. 32p-Labeled mouse histone pre-mRNA was synthesized in vitro with the use of T7 R N A polymerase and an H2a614 based template (lane 1). The template was incubated in a nuclear extract from mouse myeloma cells for 30 rain at 32°C, resulting in accurate cleavage of approximately 65% of the input R N A (lane 2). To demonstrate the dependence of processing on the SLBP, a 40-fold molar excess of RNA containing the stem-loop structure was added to the reaction (lane 3). To demonstrate the dependence on U7 snRNP, 1 /xg of an 2'-O-methyl oligoribonucleotide complementary to the 5' end of U7 snRNA was added to the in vitro reaction, resulting in complete inhibition of processing (lane 4).

D. DEPLETION OF PROCESSING FACTORS FROM THE NUCLEAR E X T R A C T Two

o f t h e t h r e e factors r e q u i r e d for p r o c e s s i n g o f h i s t o n e p r e - m R N A s ,

H B F (SLBP) a n d U 7 s n R N P , can b e specifically r e m o v e d f r o m t h e n u c l e a r extract

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by addition of biotinylated oligoribonucleotides followed by incubation of the sample with streptavidin agarose. For depletion of the SLBP, a short, biotinylated oligoribonucleotide containing the wild-type stem-loop structure is used. Depletion of U7 snRNP can be achieved using a biotinylated oligonucleotide complementary to the 5' end of U7 snRNA (50). 2'-O-Methyl oligoribonucleotides are nuclease resistant and form stable double-stranded complexes with complementary sequences. Therefore, for depletion of U7 snRNP, they should be used instead of unmodified oligoribonucleotides. Alternatively, U7 snRNA can be destroyed with the use of a DNA oligonucleotide directed against the 5' end of U7 snRNA. This will direct cleavage of the U7 snRNA by endogenous RNase H in the extract (51). The SLBP can also be depleted by a specific antibody (38), and in principle the U7 snRNP (together with many other snRNPs) can be depleted with the anti-Sm antibody or anti-2,2,7-trimethyl guanosine antibody.

1. P r o t o c o l

1. Rinse the streptavidin agarose beads several times with an excess of ice-cold dialysis buffer (see above), each time spinning the gel suspension in the microcentrifuge for a few seconds. 2. Incubate the agarose beads with several volumes of nuclear extract, continuously mixing at 4~ N o t e : This step prevents possible nonspecific streptavidin-protein

interactions during the subsequent depletion reaction. Presumably any nonspecific binding sites on the beads will be coated with proteins from the nuclear extract. 3. Remove the extract and rinse the pelleted gel with several volumes of cold dialysis buffer. Streptavidin agarose is now ready for the depletion experiment. 4. Use 1/~g of the synthetic 3' biotinylated R N A containing the histone-specific stem-loop or 0.2/a,g of 2'-O-methyl oligonucleotide complementary to the 5' end of U7 snRNP per each 100/~1 of the nuclear extract. Both types of biotinylated R N A can be synthesized on an Applied Biosystems Synthesizer with the use of reagents from Glen Research (Herndon, VA). They are synthesized on a CPG column to which the biotin has been attached. N o t e : Prior to preparation of a depleted extract, a pilot competition

experiment should be performed to determine the smallest amount of each oligonucleotide sufficient for maximal inhibition of an in vitro processing reaction.

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5. Incubate the sample at room temperature for 5-15 min and then at 4~ rocking for 2 hr.

Note: The time and temperature of incubation should be determined empirically, depending on the purpose of the experiment. Prolonged incubation at higher temperatures results in more efficient interaction between the oligonucleotide and a specific factor but at the same time leads to a gradual decrease in processing activity due to nonspecific inactivation of other components of the processing machinery. This could preclude successful complementation assays relying on addition of depleted factor. 6. Add 10/xl of pretreated streptavidin agarose to each 100/xl of extract containing the oligonucleotide. 7. Incubate with continuous rocking at 4~ for 3 hr to absorb the biotinylated complex. 8. Remove the streptavidin agarose and bound biotinylated complex by. centrifugation at 1000 g for 2 min. 9. Check the efficiency of depletion by mobility shift assay and a processing reaction and keep the extract frozen in small aliquots at -80~ Even relatively short exposure of the extract to room temperature or incubation with streptavidin agarose at 4~ results in the loss of approximately 25-30% of processing activity. The extent of the nonspecific effects leading to a decrease in processing efficiency must be determined by performing a control experiment in parallel in which a nonspecific oligonucleotide (with no affinity for known processing factors) is used. The "mock-depleted" extract, done exactly as above but with the use of a nonspecific oligoribonucleotide, is used as a control in all subsequent assays.

IV. I S O L A T I O N

OF POLYRIBOSOMES

After being processed in the nucleus, the mature histone m R N A is transported to the cytoplasm, where it is loaded onto polyribosomes. The transport of histone m R N A to the cytoplasm and loading onto polyribosomes is dependent on the histone 3' stem-loop (13) and presumably on the SLBP that is bound to the stem-loop. The isolation of polyribosomes by subcellular fractionation allows the detection of R N A binding proteins that are a component of the histone m R N P that may play a role in the cytoplasmic metabolism of histone mRNA, including translation and degradation. Several factors that may participate in histone m R N A metabolism have been isolated from polyribosomes, including the SLBP (34) and a 3 ' - 5 ' exonuclease that degrades histone m_RNA (52). The analysis of R N A

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binding proteins from polyribosomes allows us to determine whether the complexes formed on the target 1KNA by polyribosomal proteins are the same as those formed by nuclear proteins (22). Furthermore, using functional assays such as in vitro processing assays (see above), we can ask whether the protein bound to the stemloop is the same protein that functions in histone pre-mlKNA processing in the nucleus (30). In addition, polyribosomal extracts have been used to characterize the degradation machinery of different eukaryotic mRNAs (53,54). The procedure for isolation of polyribosomes is adapted from the procedure of 1Koss (55).

A. PROTOCOL 1. Reserve the cytosol obtained after spinning out the nuclei, described in the preparation of nuclear extract (Step 6 in the nuclear extract procedure, Section III,A). Transfer the cytosol to a CO1KEX tube and spin at 10,000 g in a swinging bucket rotor to remove mitochondria. 2. Transfer the supernatant to a clean tube and add Triton X-100 to a final concentration of 0.1% to release polyribosomes from membranes. The cytosol is then carefully layered over a 6-ml 30% sucrose cushion of Buffer B in a 30-ml centrifuge tube. The polyribosomes are pelleted by centrifugation at 130,000 g for 4 hr in a fixed-angle rotor (e.g., Ti65). 3. The $130 supernatant is removed without disturbing the interface between the supernatant and the sucrose cushion, aliquoted, and stored at -80~ 4. Carefully remove the sucrose cushion and wash the polyribosomes with several milliliters of Buffer A. The polyribosomes are then resuspended in 0.5 ml Buffer A per 5 X 108 cells by pipetting with a pipette tip. The polyribosomes are transferred to a 7-ml homogenizer and homogenized with a loose pestle until the suspension is homogeneous. The polyribosomes are then snap-frozen and stored in liquid nitrogen. 5. The ribosomal salt wash is prepared by adding 4 M KC1 dropwise to the polyribosome suspension, with shaking, to a final concentration of 0.8 M. The polyribosomes are then incubated for 30 rain on a rotator at 4~ This procedure removes loosely bound proteins from the polyribosomes without removing significant amounts of ribosomal proteins. AFter extraction, the polyribosomes are removed by centrifugation at 130,000 g. The supernatant is then collected, dialyzed against Buffer D, and frozen in small aliquots.

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V. I S O L A T I O N A N D C H A R A C T E R I Z A T I O N OF RNA-BINDING PROTEINS USING THE YEAST THREE-HYBRID SYSTEM The yeast three-hybrid system recently devised by Marv Wickens and Stan Fields and co-workers can be used to select 1KNA binding proteins in yeast (20). This new technology is a variation of the yeast two-hybrid system, and we used it successfully in our laboratory to clone a novel R N A binding protein, SLBP, which binds the highly conserved stem-loop at the 3' end of histone mlKNAs. The arrangement of the hybrid components adapted from Wickens and co-workers is shown in Figure 5. The yeast strain (L40-coat) contains a fusion gene that expresses the LexA DNA binding domain and the phage MS-2 coat protein, along with two reporter genes with the LexA DNA binding site in their promoters. A vector expressing a hybrid 1LNA from the ILNase P promoter was introduced into the yeast. This hybrid IKNA contains the MS2 protein binding site and the target 1KNA sequence of interest, in this case the histone stem-loop. The target 1LNA sequence can be cloned in either 5' or 3' of the MS2 protein binding sites. We cloned the histone stem-loop 3' of the MS2 binding sites since the histone stem-loop is naturally at the 3' end of histone mlKNA (38). Finally, a cDNA fused to the Gal4 activation domain is introduced into the yeast, which is a typical two-hybrid library with an appropriate selectable marker, different from that introduced on the plasmid expressing the 1KNA. Expression of a fusion protein from the library that interacts with the target 1KNA sequence will result in the activation of transcription of the reporter genes. SLBP is an excellent candidate for using this system to select a specific 1KNA binding protein, since the histone 3' stem-loop is present only in metazoans and the binding of the stem-loop to SLBP is highly specific. The selection procedure we used is basically identical to the yeast two-hybrid manual from Clontech (Palo Alto, CA). Several tips concerning using this system are presented in the following sections, including maintaining the strain's activity, transformation of the cDNA library to the strain that already contains the bait, and strategies for the identification of specific positive clones. A. POSITIVE CONTROLS The pIII/IRE-MS2 and pAD-IR.P plasmids express the IRE bait lkNA and IRP protein, respectively. The transformation of these two plasmids into the L40coat strain can cause transcriptional activation of His3 and LacZ (20). For assaying the activation of His3 transcription, dropout plates containing 5 mM 3-aminotriazole were used to suppress the basal expression of His3. We used Whatman filter paper to perform colony color assays to detect the Gal4 transcriptional activation (Section V,E).

uc

CU U U

it

C

G A

C

G

G

C

RNas~ P leader

3! nt

U AU

A

U

CCAAAG

MS2

C

C

GC WT

CACCCAUUUUUU

A U

RM

GC CG 5 ....

CCAAAC

GACCCAUUUUUU

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B . CONSTRUCTION OF THE PLASMID BAIT Since the hybrid R N A is expressed from the R N A polymerase III promoter, a stretch of four or more T's can function as an R N A polymerase III terminator. A well-defined bait with k n o w n binding specificity and stoichiometry to the u n k n o w n protein is helpful. Since it is likely that the longer the R N A molecule the more likely it is to interact with nonspecific R N A binding proteins, we took care to use a minimal R N A binding site. SLBP has been successfully cloned with the R N A bait constructed with the target either before (39) or after (38) the MS2 binding sites, so the slightly reduced affinity of SLBP for its target w h e n the s t e m - l o o p is not at the 3' end of the R N A did not prevent efficient selection. W e also designed a mutant bait, with the s t e m - l o o p sequence reversed, which would not bind SLBP. This was used to test each of the positive clones from the initial selection for specific binding. The specific R N A - b i n d i n g protein should not activate/~-galactosidase w h e n introduced into strains containing the mutant bait. Thus each plasmid was transfected into two yeast strains, one containing the bait and the other the mutant bait (and selected only on Leu- media and not on Leu- His- media). W h e n the false positives were eliminated with a mutant bait (which varied from 20% to 60% of the primary colonies in different screens), all the clones that interacted with the bait but did not interact with the mutant bait encoded SLBP.

C. MAINTENANCE OF THE YEAST STRAINS AND PHENOTYPE VERIFICATION 1. The L40-coat strain can survive for up to half a year in a sealed tube at 4~ If the strain is a frozen stock, it can be restored by streaking the strain onto a Y P D agar plate first to revitalize the strain and then restreak it onto the selective dropout media to maintain the selection for plasmids. The yeast colonies should appear slightly pink at the top from the Ade2 mutation. 2. Always verify the nutritional requirement phenotypes of the yeast strains if you use the strain for the first time. It is r e c o m m e n d e d that you restreak the yeast from the original stock w h e n e v e r you see some unexplained results as the strain grows on the SD medium, or if it

Figure 5 Diagram of the yeast three-hybrid system. The strategy for isolation of SLBP using the yeast three-hybrid system is shown. The structure of the wild-type (WT) "bait" RNA and the mutant reverse (RM) RNA used to screen out the potential positives is shown below the diagram.

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William F. Marzluffet al. grows too quickly or too slowly, or if it does not look like the yeast strain you originally streaked. A fresh colony that has the selective nutritional phenotype can be used to make competent cells. D . TRANSFORMATION OF T w o - H Y B R I D VECTORS INTO

COMPETENT YEAST CELLS As in performing yeast two-hybrid screening, you must transform the two fusion genes that express the hybrid 1KNA and library fusion proteins into the L40-coat strain that contains the reporter genes. We used sequential transformation and the L i A c transformation method to obtain a higher transformation frequency (56,57). In our hands, 100 /xg of library cDNA was sufficient to obtain 1 million transformants. A small-scale transformation is recommended for checking the background and transformation efficiency before scaling up for library transformation. The competent cells are good for 2 weeks if they are stored at 4~ In sequential transformations, the cells containing the bait plasmids are cultured in dropout media to prevent loss of the bait plasmid. However, we transfer the cultured cells from the dropout media to YPD liquid media [with dropout media and YPD media (1:1 ratio)] in the last step before making competent cells. The cells are cultured for an extra 3 hr in this media to increase their density and increase their growth rate. Typically, 200/xl of competent cells is transformed with 2.5/xg of DNA and 200/xg of salmon sperm carrier DNA per large plate (150 mm). After 3-5 days on dropout plates at 30~ large colonies can be picked up and assayed for /3-galactosidase activity by Whatman filter paper assay. The cartier DNA is prepared by sonicating and phenol extracting the DNA several times until the 260/280 reading of the cartier DNA is over 1.8 and the average size of the DNA is about 300 bp. Denaturing the cartier DNA repeatedly increases the transformation efficiency. Chemicals, especially the components of the dropout media, should be kept separate from lab chemicals. Only the highest quality chemicals and water should be used in order to increase the efficiency of transformation. Instead of making a large batch of transformations (2 ml of competent cells and 100 /xg of library DNA), we perform a dozen small-scale transformations to avoid the mixing and heat-shock problems caused by a large-scale transformation.

E. fl-GALACTOSIDASE FILTER ASSAY AND ELIMINATION OF THE FALSE POSITIVES If you obtain several hundred H i s 3 positives on your primary screening, streaking all the colonies on fresh plates and performing the filter-lifting assay

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with nitrocellulose is very time-consuming and inefficient (Clontech Matchmaker Manual). We found that directly transferring a portion of a colony (which is usually 1-3 mm in diameter) to a dry Whatman No. 5, No. 1, or No. 541 paper with a toothpick (you can draw a grid directly on the paper with a pencil to keep track of the colonies) and then performing the color assay is sufficiently sensitive to detect the AS-galactosidase activity when performing the filter assay as long as quantitation of/~-galactosidase activity is not required. Only the colonies that can activate both His3 and LacZ reporters are streaked into single colonies, and total yeast D N A containing the library plasmids is extracted. The D N A is purified with either Qiagen or any standard D N A purification kit and then used to transform Escherichia coli HB101 (which is Leu-). We generally obtain 10-50 transformants on M9 plates with one fifth of the mini-prep DNA preparation. It is important to construct a mutant bait 1KNA to act as a negative control that can eliminate nonspecific interactions when performing the secondary screen. We further characterize only the clones encoding proteins that interact with the wild-type bait but fail to interact with the mutant 1KNA. Ira clone that passes all the genetic tests has been selected, in vitro biochemical assays using the expressed protein (as described in the following section) can provide definitive evidence that you have cloned the intended target protein.

F. In Vitro TRANSLATION AND MOBILITY SHIFT ASSAYS USING THE RETICULOCYTE LYSATE In vitro translation in reticulocyte lysates is a convenient method to determine whether the protein that you have cloned is indeed the protein of interest. The gene of interest is cloned behind the T7, T3, or SP6 promoter and 1KNA transcribed in vitro with the use of standard procedures (the "Message Machine"; Ambion, Austin, TX). The 1KNA is then translated in the reticulocyte lysate, and proteins are detected either by autoradiography or by the mobility shift assay. Sufficient protein for detection in a mobility shift assay is synthesized. In addition, one can synthesize the full-length protein and compare its mobility with that of the authentic cellular protein (detected by UV cross-linking or Northwestern blotting). Since 1KNA binding proteins often have aberrant mobilities on SDS-polyacrylamide gels, this procedure is often very helpful. For example, SLBP, which migrates at 45 kDa on SDS-polyacrylamide gels, is actually only 32 kDa. The in vitro translated protein is likely to be properly processed in the reticulocyte lysate (as long as it isn't processed in a membrane-dependent fashion) and is much more likely to yield a functional protein than expressing the protein in bacteria. These analyses can be done rapidly with only small amounts of in vitro translated protein.

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The human SLBP gene encodes a protein that has a predicted molecular weight of 32,000 in contrast to the 45,000 estimated from the mobility on S D S polyacrylamide gels (22,34). Using reticulocyte lysates, we showed that the protein cloned with the yeast three-hybrid system migrates at an apparent molecular weight of 45,000 on SDS-polyacrylamide gels (Fig. 6A, lane 1) and binds the histone 3' stem-loop in a specific manner, forming a complex with mobility identical to that observed in cell lysates. Truncation of the protein at the terminus by 13, 52, 72, or 90 amino acids yielded smaller protein products, as expected (Fig. 6A, lanes 2-6). The in vitro translated proteins can also be used to measure the ability of the tuncated form of SLBP to bind 1KNA, effectively mapping the 1KNA binding site. Figure 6B shows that the in vitro translated SLBP does indeed bind to the 1KNA and forms a complex similar to that formed by the SLBP from polyribosomes. Deletion of 52 or 72 amino acids from the C terminus of the protein does not affect the ability of SLBP to bind 1KNA (Fig. 6A, lanes 4 and 5), but deletion of 90 amino acids abolishes the 1KNA binding activity (lane 6). A small amount of endogenous SLBP is present in the reticulocyte lysate (Fig. 6B, lane 7). A total of 124 amino acids can be deleted from the N terminus of the protein without affecting binding activity (not shown; 38). In vitro translated SLBP binds the stem-loop, forming a complex that comigrates with the complex formed in polyribosomal extracts (Fig. 6C). This complex can be competed with excess cold stem-loop 1KNA (Fig. 6C, lane 3) but is not competed by a reverse stem 1KNA or a nonspecific 1KNA (Fig. 6C, lanes 4 and 5). Similar results are obtained with the 110-amino acid SLBP containing only the 1KNA binding domain, which lacks 87 amino acids from the amino terminus and 72 amino acids from the carboxyl terminus. Binding to this truncated protein is competed by the wild-type 1KNA (Fig. 6C, lane 7) but not by a reverse stem or nonspecific 1KNA (Fig. 6C, lanes 8 and 9). Thus the 110-amino acid protein binds the stem-loop with a specificity similar to that of the full-length protein (38).

APPENDIX:

CELL CULTURE

In order to study histone mR.NA metabolism, we have used MPC-11 mouse myeloma cells that have been adapted to continuous culture, as previously described (58). The mouse myeloma cells were grown in stationary suspension culture in Dulbecco's Modified Eagle's Medium (DMEM) with 10% horse serum, 100 U / m l penicillin, and 100/xg/ml streptomycin (GIBCO-BRL, Gaithersburg, MD). Mouse myeloma cells are both an economical and a convenient source of material from which both nuclei and polyribosomes can be prepared in quantity. Typically, myeloma cells were grown to a density of 5 • 10s cells/ml and harvested for subcellular fractionation. Up to 38 liters can be grown at once in a large spinner

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flask (Bellco, Vineland, NJ), but for the most active preparations, 6-8 liters of cells is recommended because of the time required for processing large preparations.

A. SOLUTIONS PBS

137 mM NaC1, 2.7 mM KC1 4.3 mM Na2HPO4 97 H20 1.4 mM KH2PO4

Hypotonic buffer

10 mM HEPES-KOH, pH 7.9 1.5 mM MgC12 10 mk//KC1

0.5 mM DTT 0.75 mM Spermidine 0.15 mM Spermine Restore buffer

67.5% Sucrose

Low-salt extraction buffer

20 mM HEPES-KOH, pH 7.9 1.5 mM MgCI2 20 mM KCl 0.5 mM DTT 0.2 mM EDTA, pH 8.0 0.75 mM Spermidine 0.15 mM Spermidine 25% Glycerol

High-salt extraction buffer

20 mM HEPES-KOH, pH 7.9 1.5 mM MgC12 1.2 M KCl 0.5 mM DTT 0.2 mM EDTA, pH 8.0 0.75 mM Spermidine 0.15 mM Spermine 25% Glycerol

Dialysis buffer

20 mM HEPES-KOH, pH 7.9 100 mM KCl 0.5 mM DTT 0.2 mM EDTA, pH 8.0 20% Glycerol

N o t e : DTT, spermidine, and spermine should be added to solutions just prior

to use.

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19 1

Figure 6 In vitro translation of SLBP and determination of the R N A binding domain. (A) Translation of human SLBP and SLBP deletions in the reticulocyte lysate. The cloned DNAs were transcribed by T7 R N A polymerase, and the R N A was translated in the reticulocyte lysate for 90 min in the presence of 35S-methionine. The products were analyzed by electrophoresis on 12% polyacrylamide gels and detected by autoradiography. Lane 1: full-length SLBP; lanes 2-5, SLBP truncated 13, 52, 72, or 90 amino acids from the C terminus; lane 6; SLBP with the first 87 and the last 13 amino acids removed. (B) Binding of the stem-loop by the C-terminal deletions of SLBP. Fifteen femtomoles ofradiolabeled stem-loop (lane 1) was incubated with 5/zg of the polyribosomal extract (lane 2) or with 12.5/a,1 of reticulocyte lysate programmed with R N A encoding the full-length (lane 3) or C-terminal deletion mutants (lanes 4-6). The mutants used are indicated above each lane. Lane 7 is an incubation of 12.5 /a,1 of the unprogrammed reticulocyte lysate with the probe. The arrow indicates the complex with full-length SLBP, and "NS" indicates a nonspecific complex formed in the reticulocyte lysate. (C) A 110-amino acid R N A binding fragment of SLBP specifically binds the stem-loop. In lane 1 the probe was incubated with a nuclear extract. In vitro-translated full-length SLBP (lanes 2-5) and the 110amino acid fragment A87NA72C (lanes 6-9) were incubated with 15 fmol of labeled 30-nt R N A containing the stem-loop. The complexes were resolved by electrophoresis on a 10% polyacrylamide gel. In lanes 3 and 7, a 100-fold excess of the 30-nt R N A was included, in lanes 4 and 8 a 100-fold excess of an 1KNA with the stem sequence reversed was included, and in lanes 5 and 9 a 100-fold excess of a nonspecific IKNA (AAAUACCUACUUCAUACAA) was included. The arrows indicate the specific complexes formed. The unbound probe has been run off the gel to allow good resolution of the complex with the 110-amino acid protein.

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REFERENCES 1. Marzluff, W. F., and Pandey, N. B. (1988). Trends Biochem. Sci. 13, 49. 2. Wang, Z.-F., Tisovec, R., Debry, R. W., Frey, M. R., Matera, A. G., and Marzluff, W. F. (1996). Genome Res. 6, 702. 3. Wang, Z.-F., Krasikov, T., Frey, M. R., Wang, J., Matera, A. G., and Marzluff, W. F. (1996). Genome Res. 6, 688. 4. Doenecke, D., Albig, W., Bouterfa, H., and Drabent, B. (1994). J. Cell. Biochem. 54, 423. 5. Allen, B. S., Stein, J. L., Stein, G. S., and Ostrer, H. (1991). Genomics 10, 486. 6. Harris, M. E., B6hni, R., Schneiderman, M. H., Ramamurthy, L., Schtimperli, D., and Marzluff, W. F. (1991). Mol. Cell. Biol. 11, 2416. 7. Ltischer, B., Stauber, C., Schindler, R., and Schtimperli, D. (1985). Proc. Natl. Acad. Sci. U.S.A. 82, 4389. 8. Gick, O., Kr~irner, A., Keller, W., and Bimstiel, M. L. (1986). E M B O J . 5, 1319. 9. Williams, A. S., and Marzluff, W. F. (1995). Nudeic Acids Res. 23, 654. 10. Jackson, R. J., and Standart, N. (1990). Cell (Cambridge, Mass.) 62, 15. 11. Williams, A. S., Ingeldue, T. C., Kay, B. K., and Marzluff, W. F. (1994). Nucleic Acids Res. 22, 4660. 12. Eckner, R., Ellmeier, W., and Birnstiel, M. L. (1991). E M B O J . 10, 3513. 13. Sun, J.-H., Pilch, D. R., and Marzluff, W. F. (1992). Nucleic Acids Res. 20, 6057. 14. Gallie, D. R., Lewis, N. J., and Marzluff, W. F. (1996). Nucleic Acids Res. 24, 1954. 15. Pandey, N. B., and Marzluff, W. F. (1987). Mol. Cell. Biol. 7, 4557. 16. Stauber, C., and Schtimperli, D. (1988). Nucleic Acids Res. 16, 9399. 17. DeLisle, A. J., Graves, R. A., Marzluff, W. F., and Johnson, L. F. (1983). Mol. Cell. Biol. 3, 1920. 18. Heintz, N., Sive, H. L., and Roeder, R. G. (1983). Mol. Cell. Biol. 3, 539. 19. Sittman, D. B., Graves, R. A., and Marzluff, W. F. (1983). Pro& Natl. Acad. Sci. U.S.A. 80, 1849. 20. SenGupta, D. J., Zhang, B. L., Kraemer, B., Prochart, P., Fields, S., and Wickens, M. (1996). Proc. Natl. Acad. Sci. U.S.A. 93, 8496. 21. Pandey, N. B., Williams, A. S., Sun, J.-H., Brown, V. D., Bond, U., and Marzluff, W. F. (1994). Mol. Cell. Biol. 14, 1709. 22. Pandey, N. B., Sun, J.-H., and Marzluff, W. F. (1991). Nucleic Acids Res. 19, 5653. 23. Roberts, S. B., Emmons, S. W., and Childs, G. (1989).J. Mol. Biol. 206, 567. 24. Draper, D. E. (1995). Annu. Rev. Biochem. 64, 593. 25. Burd, C. G., and Dreyfuss, G. (1994). Science 265, 615. 26. Hall, K. B. (1994). Biochemistry 33, 10076. 27. Oubridge, C., Ito, N., Evans, P. R., Teo, C.-H., and Nagai, K. (1994). Nature (London) 372,432. 28. Allain, F. H. T., Gubser, C. C., Howe, P. W. A., Nagai, K., Neuhaus, D., and Varani, G. (1996). Nature (London) 380, 646. 29. Milligan, J. F., Groebe, D. R., Witherell, G. W., and Uhlenbeck, O. C. (1987). Nucleic Acids Res. 15, 8783. 30. Dominski, Z., Sumerel, J., Hanson, R. J., and Marzluff, W. F. (1995). R N A 1, 915. 31. Ratnasabapathy, R., Hwang, S.-P. L., and Williams, D. L., (1990). J. Biol. Chem. 265, 14050. 32. Greenberg, J. R. (1979). Nucleic Acids Res. 6, 715. 33. Greenberg, J. R. (1980). Nucleic Acids Res. 8, 5685. 34. Hanson, R. J., Sun, J.-H., Willis, D. G., and Marzluff, W. F. (1996). Biochemistry 35, 2146. 36. Scharl, E. C., and Steitz, J. A. (1994). E M B O J . 13, 2432. 36. Vasserot, A. P., Schaufele, F.J., and Birnstiel, M. L. (1989). Proc. Natl. Acad. Sci. U.S.A. 86, 4345. 37. Melin, L., Soldati, D., Mital, R., Streit, A., and Schtimperli, D. (1992). E M B O J . 11, 691. 38. Wang, Z.-F., Whitfield, M. L., Ingledue, T. I., Dominski, Z., and Marzluff, W. F. (1996). Genes Dev. 10, 3028.

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39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58.

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Martin, F., Schaller, A., Eglite, S., Schumperli, D., and Muller, B. (1997). E M B O J . 16, 769. Mowry, K. L., and Steitz, J. A. (1987). Science 238, 1682. Soldati, D., and Schtimperli, D. (1988). Mol. Cell. Biol. 8, 1518. Gick, O., Kr~imer, A., Vasserot, A., and Birnstiel, M. L. (1987). Proc. Natl. Acad. Sci. U.S.A. 84, 8937. Streit, A., Koning, T. W., Soldati, D., Melin, L., and Schtimperli, D. (1993). Nucleic Acids Res. 21, 1569. Mowry, K. L., Oh, R., and Steitz, J. A. (1989). Mol. Celt. Biol. 9, 3105. Stauber, C., Soldati, D., Ltischer, B., and Schtimperli, D. (1990). In "Methods in Enzymology" (J. E. Dahlberg and J. N. Abelson, eds.), Vol. 181, p. 74. Academic Press, San Diego, CA. Shapiro, D., Sharp, P. A., Wahli, W., and Keller, M. (1988). D N A 7, 47. Dominski, Z., and Kole, R. (1993). Proc. Natl. Acad. Sci. U.S.A. 90, 8673. Gu, X. H., and Marzluff, W. F. (1996). Nucleic Acids Res. 24, 3797. Hurt, M. M., Chodchoy, N., and Marzluff, W. F. (1989). Nudeic Acids Res. 17, 8876. Smith, H. O., Tabiti, K., Schaffner, G., Soldati, D., Albrecht, U., and Birnstiel, M. L. (1991). Proc. Natl. Acad. Sci. U.S.A. 88, 9784. Black, D. L., Chabot, B., and Steitz, J. A. (1985). Cell (Cambridge, Mass.) 42, 737. Caruccio, N., and Ross, J., (1994).j. Biol. Chem. 269, 31814. Ross, J., Kobs, G., Brewer, G., and Peltz, S. W. (1987). J. Biol. Chem. 262, 9374. Bernstein, P. L., Herrick, D. J., Prokipcak, R. D., and Ross, J. (1992). Genes Dev. 6, 642. Brewer, G., and Ross, J. (1990). In "Methods in Enzymology" (J. E. Dahlberg andJ. N. Abelson, eds.), Vol. 181, p. 202. Academic Press, San Diego, CA. Ito, H., Fukada, Y., Murata, K., and Kimura, A. (1983). J. Bacteriol. 153, 163. Gietz, D., St. Jean, A., Woods, R. A., and Schiestl, R. H. (1992). Nudeic Acids Res. 20, 1425. Laskov, R., and Scharff, M. D. (1970). J. Exp. Med. 131, 515.

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Analysis of mRNP Complexes Assembled in Vitro Stephen A. Liebhaber Howard Hughes Medical Institute Departments of Genetics and Medicine University of Pennsylvania School of Medicine Philadelphia, Pennsylvania 19104

Martin Holcik* Howard Hughes Medical Institute Departments of Genetics and Medicine University of Pennsylvania School of Medicine Philadelphia, Pennsylvania 19104

Analysis of sequence-specific interactions between R N A and proteins is essential for defining the biochemistry of the posttranscriptional steps in gene expression and in understanding a broad range of corresponding genetic controls. Sequential steps in R N A synthesis and expression in which RNA-protein interactions are central include transcript splicing, 3' processing, nuclear-to-cytoplasmic transport, subcytoplasmic localization, translational modulation, and determination of m R N A stability (Surdej et al., !994; McCarthy and Kollmus, 1995). The gel retardation or electromobility shift assay is the simplest and perhaps the most widely used method for investigating protein-nucleic acid interactions in vitro (Leibold and Munro, 1988; Weeks and Crothers, 1991). This assay is based on the observation that binding of proteins to an R N A fragment usually leads to a reduction in the electrophoretic mobility of the R N A in a nondenaturing polyacrylamide gel. Moreover, protection of a subsegment of the R N A from RNase degradation by the bound protein(s) in the complex provides an approach to identifying proteinbinding sites, and the assembly of RNA-protein complexes in vitro can be exploited for affinity purification of the respective binding proteins. * Present Address: Molecular Genetics Laboratory, University of Ottawa, Children's Hospital of Eastern Ontario, Ottawa, Ontario K1H 8L1 Canada. 195 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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I. C O M P L E X

FORMATION

A. PREPARATION OF CYTOPLASMIC $ 1 0 0 EXTRACTS Total cell extracts can be used directly for RNA-protein complex assembly. However, crude extracts contain many sequence-specific and nonspecific 1KNAbinding proteins. Fractionation of cell extracts may reduce the nonspecific background. For in vitro analysis ofRNA-protein interactions thought to occur in the cytoplasm, postnuclear fractions can be prepared by gentle cell lysis and removal of nuclei by low-speed centrifugation. The cytoplasmic fraction can then be further cleared by high-speed centrifugation to sediment large macromolecular complexes (e.g., polysomes). It is important to perform all procedures at 0~ to minimize protein degradation. Various protease inhibitors [e.g., c~-p-nitrophenylglycerol (PNPG), phenylmethylsulfonyl fluoride (PMSF), leupeptin, aprotinin, pepstatin] are included in the homogenization buffer for the same purpose. Selective use of particular protease inhibitors must be determined empirically. The salt concentration and pH used during the preparation of cell lysates are less critical than those used during the subsequent 1KNA binding steps. Therefore, lysis of cells in relatively neutral buffer conditions (low salt concentration, neutral pH) facilitates the adjustment of the cytoplasmic fraction to the desired RNA-protein binding conditions. The particular salt concentration, presence or absence of divalent cations, and optimal pH are determined empirically for each set of proteins. Following high-speed centrifugation of the extract, glycerol is added to the S 100 supernatant and the protein concentration is measured by standard colorimettic methods. The protein concentration should be adjusted to 8-10 mg/ml. Storage of the extract in glycerol and at high concentrations reduces the loss of binding activity resulting from protein degradation and adherence to the tube. In addition, concentrated protein stocks allow the assembly of small-volume, highconcentration, complex assembly reactions. Proteins can be conveniently concentrated using spin-concentrators (e.g., Amicon, Beverly, MA); the molecular weight cutoff value of the concentrator should be determined empirically to ensure that the protein(s) of interest are not removed at this step. The concentrated $100 fraction is aliquoted (30-50/zl) into small Eppendorf tubes and flash-frozen in dry ice-ethanol or a liquid nitrogen bath. Aliquots should be stored at - 80~ Repeated freeze-thawing of the extract should be avoided since it may affect the binding activity. The following protocol describes preparation of the cytoplasmic $100 fraction from cells grown in suspension (K562, MEL) with the use of conditions that have been optimized for generation of the ol-globin RNA-protein complex (described in Wang et al., 1995).

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1. Reagents

Homogenization buffer: 10 mM Tris-HC1 pH 7.4, 1.5 mM MgC12, 10 mM KC1, 0.5 mM dithiothreitol (DTT), 0.1 mM PMSF, 10/xg/ml leupeptin 1KNA-binding buffer: 10 mM Tris-HC1 pH 7.4, 1.5 mM MgC12, 150 mM KC1, 0.5 mM DTT, 0.1 mM PMSF, 10/xg/ml leupeptin 1 M KC1 100 mM PMSF 10 mg/ml Leupeptin Phosphate-buffered saline (PBS) (137 mM NaC1, 4.3 mM Na2HPO4, 2.7 mM KC1, 1.4 mM KH2PO4) Glycerol

2. Protocol 1. Harvest cells during the exponential growth phase by centrifugation from medium at 0~ 2. Wash cells twice with ice-cold PBS. 3. Resuspend cells at 5 • 10 7 c e l l s / m ] in homogenization buffer. 4. Lyse cells with a Dounce homogenizer with 20 strokes, using a loose-fitting glass piston. The homogenate can be monitored by light microscopy for the state of lysis; cells should be disrupted and the nuclei should remain intact. 5. Centrifuge the cell lysate for 10 min at 2000/g to pellet the nuclei. 6. Carefully remove the supernatant and adjust it with KC1 to reach the R N A binding buffer conditions (also see Section I,C). 7. Centrifuge the cytoplasmic fraction for 60 min at 100,000 g. 8. Carefully remove the supernatant ($100 fraction) and add glycerol to it to achieve a final concentration of 5%. 9. Adjust the protein concentration of the $100 fraction to 8-10 mg/ml. 10. Aliquot the protein extract into small Eppendorf tubes (30-50/xl/ tube) and flash-freeze in a dry ice-ethanol bath. 11. Store at -80~

B. PREPARATION OF THE R N A PROBE Several methods are available to prepare internally labeled or end-labeled R.NA probes suitable for R.NA-protein complex analysis. The most straightforward method is to clone a double-stranded DNA segment representing the R N A region of interest into a plasmid vector that contains a bacteriophage promoter suitable

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for in vitw transcription. Several such vectors are available that offer a choice ofT3, T7, or SP6 promoters [e.g., pGEM or pSP series (Promega, Madison, WI), pC1K series (InvitroGen, Carlsbad, CA)]. Cloning into these vectors may also allow synthesis ofantisense transcripts since these vectors often have two different and oppositely oriented promoter sequences on either side of a polylinker cloning region. Such antisense 1KNAs can be used as specificity controls in the binding studies. To synthesize transcripts of defined length, the plasmid construct should be linearized by restriction digestion at a site distal (3') to the transcription unit. If it is desirable to avoid a cloning step, an alternative method is to generate the DNA template for direct in vitro transcription by the polymerase chain reaction (PC1K). In this case, the 5' PC1K primer must contain a promoter consensus sequence (T7: TAATA CGACT CACTA TAGGG CGA; T3: TTATT AACCC T C A C T AAAGG GAAG; SP6: ATTTA GGTGA CACTA TAGAA TAC). In vitro transcription from either the cloned or PCR-generated double-stranded DNA (dsDNA) is performed with the use of purified 1KNA polymerase, 0.5-1 /xg of DNA template, a mixture of rNTPs (rUTP, rATP, rGTP, rCTP) and [32p]rNTP (N is the most common nucleotide in the transcribed region to ensure high specific activity of the probe). Recently, several commercial in vitro transcription kits have been introduced (by Ambion, Austin, TX, Promega, and others). These kits offer a convenient method to synthesize large amounts oflkNA probes of high quality and high specific activity. Small 1KNA fragments can be directly generated on an oligonucleotide synthesizer. This synthetic probe can then be end-labeled with [~/-32p]ATP and T4 polynucleotide kinase and gel purified prior to incubation with extract. This method is, however, still rather costly and is practical only for small 1KNA probes.

C. R N A BINDING AND INCUBATION PROCEDURES The particular conditions for a given RNA-protein binding reaction must be determined empirically. The protocol outlined below is generally useful and has been employed successfully to characterize ribonuclear protein (1KNP) complexes formed on a variety of 1KNA probes (Kiledjian et al., 1995; Wang et al., 1995; Wang and Liebhaber, 1996). Several investigators prefer to carry out both 1KNA binding reactions and gel electrophoresis at 4~ to increase the stability of 1KNP complexes. It can be argued, however, that such conditions do not reflect in vivo conditions. While electrophoresis at 4~ does offer better band resolution and increased sharpness, the 1KNA binding reactions in our laboratory are routinely performed at room temperature. The critical step in successful 1kNA-protein complex formation is determination of the optimal salt concentration (Record et al., 1981). As mentioned above, different 1KNA binding proteins may require different concentrations of salts and/or the presence of divalent cations. It is generally assumed that increased ionic strength 1KNA binding buffer (0.5-1.5 M) will

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select for the formation of stable complexes. Hence, the initial reactions should be performed in medium-low ionic strength buffer (50-150 mM). Preference for a particular type of salt (NaC1 vs. KC1) should also be tested, as well as the requirement for divalent cations (e.g., Mg 2+) (Diekmann, 1987). Often, heparin, a polysulfated anion, is added during incubation to reduce the nonspecific background (see below). Another critical step is prevention of R N A probe degradation by endogenous RNases. Several RNase inhibitors are available from different vendors. RNase inhibitors have different potency and specificity of action. Their use and concentration should therefore be tested empirically and dictated by the specific goals of the experiment. In our laboratory we have been using PRIME RNase inhibitor (5 Prime --+ 3 Prime, West Chester, PA), which inhibits RNases A, B, and C but not T (1,2), H, CL3, or U (1,2). Heparin is also a very effective RNase inhibitor.

1. Reagents 32p-Labeled R N A probe (---10,000 cpm/0.1-0.5 ng) Unlabeled specific and nonspecific competitor RNAs (100 ng//,l) RNase inhibitor RNase mix: 1 U//,I RNase T1, 1 0 / , g / m l RNase A 50 mg/ml Heparin

2. Protocol 1. Quickly thaw an aliquot of cytoplasmic $100 extract. 2. Add RNase inhibitor (1-2 U / 3 0 / x l of extract) and 2-mercaptoethanol (final concentration of 2%). Incubate for 20-30 rain at room temperature. 3. Mix the RNA-binding reaction in 10-15/,1 total volume as follows: 10-30 ~g of S 100 protein extract 1-5 • 104 cpm of R N A probe Unlabeled nonspecific competitor R N A (if desired) or Unlabeled specific competitor R N A (for competition studiesmsee Section II,C) Concentrated RNA-binding buffer diluted to a final concentration of 1• 4. Incubate at room temperature for 30 min. 5. Add 1 ~1 RNase mix and incubate at room temperature for 10 min. 6. Add heparin (5 mg/ml final) and incubate at room temperature for 10 min.

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Martin Holcik and Stephen A. Liebhaber 7. Resolve the P,.NP complexes on a nondenaturing polyacrylamide gel (see Section II,A).

II. D E T E C T I O N

OF RNP COMPLEXES

A. ELECTROPHORETIC MOBILITY SHIFT ASSAY RNA-protein complexes are detected by a change (shift) in the mobility of the R N A probe on a nondenaturing acrylamide gel after incubation of the probe with protein extract. The detection of the complex depends on the separation of the RNA-protein complex from the free R N A and on the stability of the complex within the gel. Addition of RNases to the binding reaction after the complex has formed will degrade any unbound R N A probe and will enhance the resolution of the complex on the gel. Analysis of DNA-protein complexes indicates that they are more stable during electrophoresis than expected from their kinetic stability in solution (Fried and Crothers, 1981). This finding can be extrapolated to R N A protein complexes. Care must be taken, however, not to run electrophoresis under conditions that destabilize R N P complexes (e.g., high voltage, high gel concentration, small pore size). Typically, 3-6% polyacrylamide gels should resolve most R N P complexes. Low ionic strength buffers [e.g., Tris-borate-EDTA (TBE) or Trisacetate-EDTA (TAE)] are preferred as electrophoresis buffers because they generate less heat and increase the speed ofmigration (Fried and Crothers, 1981; Kleinschmidt et al. , 1991).

1. Reagents 30% Polyacrylamide mix (acrylamide : bisacrylamide = 29.2: 0.8) 5• TBE: 450 mM Tris-borate, 10 mM EDTA (pH 8.0) N,N,N',N'-Tetramethylethylenediamine (TEMED) 10% Ammonium persulfate 2. Protocol 1. To prepare 30 ml of 6% polyacrylamide gel, mix: 6 ml of 30% polyacrylamide 3 m 1 5 • TBE 0.5 ml 10% Ammonium persulfate 20.5 ml H20 15/,1 TEMED 2. Pour the gel and let the acrylamide polymerize at room temperature for 30-60 min.

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3. Prerun gel in 0.5X TBE for 30-60 min at 8-9 V/cm. 4. Load samples and run the gel at 6-7 V/cm (loading dye = 0.25% bromophenol blue, 0.25% xylene cyanol, 30% glycerol in water). 5. After the indicator dye has migrated to 2-5 cm from the bottom of the gel, stop electrophoresis, transfer the gel to Whatman 3MM paper, and dry on a vacuum gel dryer with heat for 45-60 min. 6. Expose overnight at - 8 0 ~ on XAR film with an intensifying screen.

B. ELIMINATION OF NONSPECIFIC BINDING All nucleic acid binding proteins display some degree of nonspecific interaction with nucleic acid probes. Even proteins that exhibit very strict target-sequence requirements will bind to some extent to targets with nonconsensus sequences. The band smearing and multiple bands that result from such nonspecific binding complicate the interpretation of gel retardation analysis. Several procedures can be used to reduce nonspecific binding and to enhance discrimination. Often the addition of salt to the binding buffer will increase selective binding by disrupting nonspecific ionic bonds (Lane et al., 1992). However, addition of salt may also reduce binding of specific R N A binding proteins. Hence, a more general approach is to add an excess (10-500 • of nonspecific, unlabeled competitor RNA. The amount of nonspecific competitor must be determined empirically since too much may affect specific complex formation as well. Transfer R N A (tRNA) is commonly used as a nonspecific competitor. Caution should be exercised, however, since tRNA may contain sequences or secondary structures that may be recognized by R N A binding proteins. Synthetic single-stranded polymers such as poly(A-T) or poly(I-C) may also be used. As with tRNA, these nonspecific competitors may also sometimes compete for complex formation since some R N A binding proteins have a preference for poly(A-T), poly(I-C), or homoribopolymers [e.g., heterogeneous nuclear R N P (hnRNP) K and o~CP [poly(C) binding protein] for poly(C) and PABP [poly(A) binding protein] for poly(A)]. Another commonly used nonspecific competitor is polysulfated carbohydrate heparin (Chodosh, 1988).

C. SPECIFICITY STUDIES As mentioned earlier, various nucleic acid binding proteins preferentially form complexes on specific R N A templates. Competition studies with sequencespecific competitors help to characterize complex formation and to obtain insight into the nature of the protein(s) involved in the R N P complex assembly. Sequence specificity of a given RNA-protein interaction is initially confirmed by sensitivity to self-competition. Generation of unlabeled competitor R N A

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for this purpose is essentially the same as that of the 32p-labeled probe, except that radiolabeled rNTP is excluded from the reaction. Specificity of the complex formation can also be demonstrated by lack of competition by unrelated sequences. The most powerful approach, however, is to generate a series of point mutations within the R N A probe that are subsequently tested for R N P complex formation (Wang et al., 1995). This approach confirms the sequence specificity of the observed complex and provides initial clues to the structure of the binding site. Synthetic homoribopolymers [poly(A), poly(C), poly(G), poly(U)] can also be tested for their ability to disrupt complex formation. Competition experiments with homoribopolymers often identify complex components that bind preferentially to one type ofpolynucleotide. This information can be then exploited for further identification (see Section III) or biochemical purification of 1KNA binding proteins.

III. ANALYSIS

OF RNP

ELECTROPHORETIC

COMPLEXES MOBILITY

IDENTIFIED

BY

SHIFT ANALYSIS

Separation of R N P complexes by gel electrophoresis is an initial step in the identification of R N P complex components (both R N A and proteins). Further characterization can be accomplished by Western or Northwestern analysis of proteins transferred from the gel to a solid support membrane or by excision of a gel fragment containing the R N P complexes of interest, followed by isolation and analysis of the R N A or protein constituents.

A. DIRECT ANALYSIS OF PROTEIN CONSTITUENTS OF R N P COMPLEXES BY TRANSFER TO MEMBRANES During electrophoretic separation on a nondenaturing gel, R N A binding proteins that are incorporated in an R N P complex shift from their native position to a new position dictated by the size and conformation of the R N P complex. This shift in the electrophoretic migration can be used to identify protein constituents of an R N P complex by Western or Northwestern analysis. Both methods involve electrotransfer of proteins and R N P complexes from the gel to a nitrocellulose membrane. The transferred proteins are subsequently identified by binding of a specific antibody (Western) or by a sequence-specific R N A (or ssDNA) probe (Northwestern). 1. Reagents For electrotransfer of proteins: Nitrocellulose membrane

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Transfer buffer: 25 m M Tris, 192 m M glycine, 15% methanol (Although methanol does not need to be used, it facilitates transfer of large proteins to the membrane. Also, some membrane manufacturers require soaking of membranes in methanol prior to transfer. In those cases, methanol should be present in the transfer buffer as well.)

2. P r o t o c o l

Semidry electrotransfer of proteins: 1. Measure the gel and cut the nitrocellulose membrane and three pieces of Whatman paper of the same size. It is critical that the membrane and Whatman paper not be bigger than the gel. 2. First wet the nitrocellulose membrane in water, then in transfer buffer. 3. Place one piece of the Whatman paper, wetted, in the transfer buffer on the bottom of the blotter apparatus. 4. Place the membrane on top of the Whatman paper and remove air bubbles by rolling a pipette over the membrane. 5. Place one piece of the Whatman paper on the gel and peel the gel off the glass plate. 6. Place the Whatman paper with gel on the membrane (gel side against the membrane) so that all sides and corners are aligned and remove all air bubbles with a pipette. 7. Wet two pieces of Whatman paper in the transfer buffer, place them on top of the membrane/gel/Whatman paper sandwich, and remove all air bubbles. 8. Wet the top plate of the blotter with plenty of transfer buffer and close the electrotransfer chamber tightly. 9. Transfer the proteins at a rate of 100-150 mA for 1-2 hr. Actual current and time depend on the size and thickness of the gel and on the size of the proteins of interest (small proteins transfer more easily than larger ones). 9 W e s t e r n analysis

1. R e a g e n t s

PBS (+0.04% NP-40 or 0.5% Tween 20) Blocking solution: 5% dried milk in PBS (+0.04% NP-40); the presence of NP-40 in the blocking solution helps to reduce the background substantially Primary and secondary antibodies

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2. P r o t o c o l * 1. Remove the nitrocellulose membrane from the electrotransfer apparatus and briefly rinse it in PBS. 2. Gently shake the membrane in blocking solution for 1-2 hr at room temperature or overnight at 4~ 3. Wash the membrane in PBS (+0.04% NP-40). 4. Add the primary antibody in suitable volume of blocking solution and incubate for 1-2 hr at room temperature. Generally, primary antibodies can be used in dilution from 1:1000 to 1:10,000; appropriate dilutions are determined empirically for each particular antibody. 5. Wash the membrane two or three times in PBS (+0.04% NP-40). 6. Wash the membrane for 5-10 min in blocking solution. 7. Add the secondary antibody diluted (1:5000 to 1 : 7000) with blocking solution. Incubate for 1-2 hr at room temperature with shaking. (If the enhanced chemiluminescence (ECL) detection method is used, avoid the use of sodium azide in the antibody solution, as it can interfere with the ECL reaction.) 8. Wash the membrane three times for 20 rain or longer in PBS (+0.04% NP-40). 9. The membrane is ready for detection. The detection method will depend on the type of enzyme to which the secondary antibody is conjugated. 9 Northwestern analysis 1. Reagents

32p-Labeled R N A probe Renaturing solution: 10 m M Tris-HC1 (pH 7.4), 50 m M NaC1, 1 m M EDTA (pH 8.0), 1• Denhardt solution, 1 m M D T T Hybridization solution: 10 m M Tris-HC1 (pH 7.4), 50 mM NaC1, 1 m M EDTA (pH 8.0), 1• Denhardt solution, t R N A (20/,g/ml), heparin

(5 ~g/ml) Washing buffer: 10 m M Tris.C1 (pH 7.4), 50 m M NaC1, 1 m M EDTA (pH 8.0), 1• Denhardt solution. 2. P r o t o c o l 1. Remove the nitrocellulose membrane from the electrotransfer apparatus and briefly rinse it in PBS. * For detailed protocols on Western analysis of proteins, see Harlow and Lane (1988).

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2. Renature proteins in situ by incubating the membrane in renaturing solution for 2 hr at room temperature with gentle shaking. 3. Decant the renaturing solution. Add just enough hybridization solution to cover the membrane. 4. Add the 32p-labeled R N A probe (---20,000 cpm/ml of hybridization solution). 5. Incubate at room temperature for 2 hr with gentle shaking. 6. Wash the membrane in a series of washes (5-10 min each) in the washing buffer at room temperature. Monitor the membrane between washes for the amount of remaining radioactivity. When the background signal is low, wrap the membrane in plastic wrap and expose it to X-ray film with an intensifying screen. The signal of Northwestern hybridization is usually weak, and extended exposure time may be needed.

B. ANALYSIS OF RESOLVED COMPLEXES EXCISED FROM THE GEL Separation of RNA-protein complexes by gel retardation is a powerful initial step in identification of individual components of the R N P complexes. It can be used to map the region of the RNA probe that is protected by R N P proteins from nucleolytic cleavage. This is especially advantageous if the initial R N A probe was large. Mapping of the termini of the R N A probe can be accomplished by RNase H mapping (Keller and Crouch, 1972). RNase H specifically degrades R N A - D N A hybrids. Hence, hybridization of specific oligonucleotides to eluted RNA, followed by RNase H cleavage and gel electrophoresis, will identify the boundaries of the R N A fragment that is covered by R N A binding protein(s) and therefore protected against nuclease cleavage. The termini can be further defined by primer extension analysis, using end-labeled primers internal to the protected region. The protein constituents of the R N P complexes can be extracted from gel-resolved complexes and further characterized by sodium dodecyl sulfatepolyacrylamide gel electrophoresis (SDS-PAGE), Western, or Northwestern analysis.

C. EXTRACTION OF THE R N A

FROM ISOLATED GEL SLICES

1. Reagents Elution buffer: 0.5 M ammonium acetate, 0.1% SDS, 1 mM EDTA (pH8.0), 10% methanol (Maxam and Gilbert, 1977)

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Martin Holcik and Stephen A. Liebhaber Phenol Chloroform : isoamylalcohol (24 : 1) Absolute ethanol 70% Ethanol 2. Protocol 1. Following gel electrophoresis, wrap gel in plastic wrap (do not dry) and expose to X-ray film. 2. With the autoradiogram as a template, cut out gel slices that correspond to the positions of the R N P complexes. 3. Transfer the weighed gel slices into an empty Poly-Prep Chromatography column [Bio-Rad (Hercules, CA) or similar] and grind the slices with a siliconized glass rod. 4. Add three or four volumes of elution buffer and soak overnight at room temperature. 5. Spin columns to collect the elution buffer. 6. Wash columns with 100/xl of elution buffer and repeat the centrifugation. 7. Extract the eluent twice with an equal volume of phenol and once with an equal volume of chloroform : isoamylalcohol. 8. Precipitate the eluted R N A with three volumes of ice-cold absolute ethanol. Incubate at - 2 0 ~ for 15-30 rain. 9. Pellet the precipitated 1KNA by centrifugation at 4~ for 30 min. 10. Wash the pellet in 70% ethanol, air dry, and resuspend the R N A in a suitable volume of RNase-free H20. 11. Analyze the recovered R N A on a urea/polyacrylamide gel.

D.

MAPPING OF THE TERMINI OF THE PROTECTED REGION BY R N A S E H CLEAVAGE

The following protocol is a modified version of the standard RNase H protection assay of Keller and Crouch (1972). 1. Reagents 40% Urea-polyacrylamide mix (acrylamide : bisacrylamide = 19: 1; 42% urea) 5• TBE: 450 mM Tris-borate, 10 mM EDTA (pH 8.0) 10 X Hybridization mix: 0.25 M Tris-HC1 (pH 7.5), 10 mM EDTA (pH 8.0), 0.5 M NaC1

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2• RNase H buffer: 40 m M Tris-HC1 (pH 7.5), 20 m M MgC12, 100 m M NaC1, 2 m M DTT, 60/xg/ml BSA, RNase H (0.2-0.5 U//xl final) Formamide loading dye: 5 m M EDTA, 0.25% bromophenol blue, 0.25% xylene cyanol in 95% formamide 2. Protocol

To prepare 30 ml of 12% denaturing gel mix: 9 ml of 40% urea-polyacrylamide mix 3 m 1 5 • TBE 0.5 ml 10% Ammonium persulfate 17.5 ml H20 15/xl TEMED After the gel has polymerized, prerun it in 0.5 X TBE for 45-60 rain at 20-25 V/cm before loading samples. 1. Mix eluted R N A and approximately 300 ng of specific oligonucleotide in a total volume of 9/xl; add 1 ~1 of 10 X hybridization mix. 2. Heat at 95~ for 10 rain and cool slowly to 30~ 3. Add 2/xl of 2X RNase H buffer and mix well. 4. Incubate at 30~ for 60 min. 5. Add 2/xl of loading dye, heat for 3 min at 100~ and load on denaturing polyacrylamide gel. 6. Run the gel in 0.5X TBE at 20-25 V/cm for the desired time. 7. Transfer the gel to a sheet of Whatman 3MM paper, dry it in a vacuum gel dryer (urea-containing gels must be dried or wrapped in plastic wrap in order to prevent them from sticking to film), and expose it to X-ray film overnight at - 8 0 ~ with an intensifying screen.

E. ISOLATION OF PROTEINS FROM THE EXCISED RNP

COMPLEXES

The following protocol was originally described by Ossipow et al. (1993) for the rapid extraction and renaturation of DNA binding proteins from SDSPAGE gels. Unlike more laborious methods described previously (e.g., Hager and Burgess, 1980), one-step extraction-renaturation allows for fast recovery of an amount of active protein(s) of interest sufficient for Western or Northwestern analysis. We have successfully applied this method to the study of R N A binding protein in our laboratory.

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1. Reagents Protein elution buffer: 1% Triton X-100, 20 mM N-2hydroxyethylpiperazine-N'-2-ethanesulfonic acid (HEPES) (pH 7.6), 1 mM EDTA, 100 mM NaC1, 5/xg/ml bovine serum albumin (BSA), 2 mM DTT, 0.1 mM PMSF, 0.1% aprotinin

2. Protocol 1. Following gel electrophoresis, wrap the gel in plastic wrap (do not dry) and expose it to X-ray film. 2. With the autoradiograrn as a template, excise gel slices that correspond to the positions of the tLNP complexes. 3. Transfer weighed gel slices into empty microfuge tubes and crush the slices with a siliconiszed glass rod. 4. Add five volumes of protein elution buffer and incubate at 37~ for 4-20 hr. 5. Spin the tubes in a microcentrifuge for 10 min to pellet the polyacrylamide residues. 6. Transfer the supernatant into a new microfuge tube and repeat the centrifugation. 7. Transfer the supernatant into a new microfuge tube. 8. To precipitate the proteins, add one or two volumes of ice-cold 100% acetone, chill at -20~ and repeat the centrifugation. 9. Pellet the precipitated proteins by centrifugation for 15 rain. Wash the pellet once with ice-cold 80% acetone. 10. tLesuspend the proteins in buffer that is compatible with the next analysis step.

ACKNOWLEDGMENTS This work was supported in part by a grant from the National Institutes of Health (HL38632). Dr. Liebhaber is an Investigator in the Howard Hughes Medical Institute. The authors thank Julia Morales and Faith Cash for critical reading of the manuscript.

REFERENCES Chodosh, L. A. (1988). Mobility shift DNA binding assay using gel electrophoresis. In "Current Protocols in Molecular Biology" (F. M. Ausubel, IL. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl, eds.), 12.2.1-12.2.11. Wiley (Interscience), New York. Diekmann, S. (1987). Temperature and salt dependence of the gel migration anomaly of curved DNA fragments. Nucleic Acids Res. 15, 247-265.

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Fried, M.G., and Crothers, D.M. (1981). Equilibria and kinetics of lac repressor-operator interactions by polyacrylamide gel electrophoresis. Nucleic Acids Res. 9, 6505-6524. Hager, D. A., and Burgess, R. R. (1980). Elution of proteins from sodium dodecyl sulfate-polyacrylamide gels, removal of sodium dodecyl sulfate, and renaturation of enzymatic activity: Results with sigma subunit of Escherichia coli RNA polymerase, wheat germ DNA topoisomerase, and other enzymes. Anal. Biochem. 109, 76-86. Harlow, E., and Lane, D. (1988). "Antibodies. A Laboratory Manual." Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. Keller, W., and Crouch, R. (1972). Degradation ofDNA RNA hybrids by ribonuclease H and DNA polymerases of cellular and viral origin. Pro& Natl. Acad. Sci. U.S.A. 69, 3360-3364. Kiledjian, M., Wang, X., and Liebhaber, S. A. (1995). Identification of two KH domain proteins in the c~-globin mRNP stability complex. E M B O J . 14, 4357-4364. Kleinschmidt, C., Tovar, K., and Hillen, W. (1991). Computer simulations and experimental studies of gel mobility patterns for week and strong noncooperative protein binding to two targets on the same DNA: Application to binding of Tet repressor variants to multiple and single tet operator sites. Nudeic Acids _Res. 19, 1021-1028. Lane, D., Prentki, P., and Chandler, M. (1992). Use of gel retardation to analyze protein-nucleic acid interactions. Microbiol. Ret,. 56, 509-528. Leibold, E. A., and Munro, H. N. (1988). Cytoplasmic protein binds in vitro to a highly conserved sequence in the 5' untranslated region of ferritin heavy- and light-subunit mRNAs. Proc. Natl. Acad. Sci. U.S.A. 85, 2171-2175. Maxam, A. M., and Gilbert, W. (1977). A new method for sequencing DNA. Proc. Natl. Acad. Sci. U.S.A. 74, 560-564. McCarthy, J. E., and KoUmus, H. (1995). Cytoplasmic mRNA-protein interactions in eukaryotic gene expression. Trends Biochem. Sci. 20, 191-197. Ossipow, V., Laemmli, U. K., and Schibler, U. (1993). A simple method to renature DNA-binding proteins separated by SDS-polyacrylamide gel electrophoresis. Nucleic Acids Res. 21, 6040-6041. Record, M. T., Mazur, S. J., Melacon, P., Roe, J. H., Shaner, S. L., and Unger, L. (1981). Double helical DNA: Conformations, physical properties, and interactions with ligands. Annu. _Reu. Biochem. 50, 997-1024. Surdej, P., RAedl, A., and Jacobs-Lorena, M. (1994). Regulation of mRNA stability in development. Annu. Rev. Genet. 28, 263-282. Wang, X., and Liebhaber, S. A. (1996). Complementary change in cis determinants and trans factors in the evolution of an mRNP stability complex. EMBO.]. 15, 5040-5051. Wang, X., Kiledjian, M., Weiss, I. M., and Liebhaber, S. A. (1995). Detection and characterization of a 3' untranslated region ribonucleoprotein complex associated with human oe-globin mRNA stability. Mol. Cell. Biol. 15, 1769-1777. Weeks, K. M., and Crothers, D. M. (1991). RNA recognition by Tat-derived peptides: Interaction in the major groove? Cell (Cambridge, Mass.) 66, 577-588.

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Analysis of RNA Binding Proteins Using in Vitro Genetics Ite A. Laird-Offringa Departments of Surgery and of Biochemistry and Molecular Biology University of Southern California School of Medicine Los Angeles, California 90033

R N A binding proteins play an important role in the R N A metabolism of all living organisms. They are involved in such diverse processes as R N A synthesis, packaging, processing, and translation. A characteristic of R N A binding proteins that is critical for their function is their ability to recognize and bind to their proper R N A targets. Our knowledge of the mechanisms governing the strength and specificity of RNA-protein interactions is limited. A genetic system for studying these interactions would be an invaluable tool for rapidly increasing our understanding of RNA-protein recognition. This chapter describes an in vitro genetic system that allows the selection of R N A binding proteins that bind tightly to a given R N A target sequence. The system utilizes phage display to physically link R N A binding proteins to their genes. The R N A binding proteins are displayed as fusions on the outside of filamentous bacteriophage particles and can be trapped on R N A attached to beads. Thus, protein variants with certain RNA-binding characteristics can be selected from a library of mutants through iterative rounds of binding and amplification. Even very rare clones can be retrieved in this way. Because the selective step occurs in vitro, binding conditions can be completely controlled. Accessory proteins involved in binding can be added to the binding reaction, so that complex RNA-protein interactions involving numerous components can be studied. The incorporation of a mutagenic step in the cycles of selection should allow the evolution of R N A binding proteins with new binding specificities. 211 m R N A F o r m a t i o n and Function Copyright 9 1997 by Academic Press. All fights of reproduction in any form reserved.

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I. I N T R O D U C T I O N Posttranscriptional gene regulation is essential for the correct expression of genes. Processes such as tLNA splicing, mR.NA export from the nucleus to the cytoplasm, translational efficiency, m R N A half-life, and mtLNA localization can have a profound effect on the amount, type, and location of the final protein product. To increase our understanding of the mechanisms of posttranscriptional gene regulation, we must know more about the proteins involved in these processes. R N A binding proteins are critical components of RNA-based gene regulation. They are involved in functions as diverse as transcriptional elongation and termination, (viral) R N A packaging, (differential) splicing, 5' and 3' end formation, and translation (Burd and Dreyfuss, 1994). Their ability to bind to the correct tLNA sequence is an essential characteristic of these proteins. To understand the function of R N A binding proteins and the processes in which they are involved, we must gain insight into how they specifically recognize their proper R N A targets. Until recently, there was no widely applicable genetic system to study R N A binding proteins. However, the desperate need for such a system was obvious from the recent flurry of activity that has resulted in the establishment of several very different genetic methodologies aimed at the study of R N A binding proteins (Harada et al., 1996; Jain and Belasco, 1996; Laird-Offringa and Belasco, 1996; SenGupta et al., 1996; Stripecke et al., 1994). Each of these methods has advantages and disadvantages, and each may be optimally suited for the study of a particular group of R N A binding proteins. The method I will discuss in this chapter is based on in vitro genetics. In other words, the selective step (the binding of a protein to RNA) occurs in the test tube. The principle of in vitro genetic analysis of R N A binding proteins is depicted schematically in Figure 1. An in vitro genetic system for analyzing the RNA-binding function of proteins requires a method of selecting, from a population of mutant R N A binding proteins (Fig. 1A), those proteins with the capacity to bind to a particular target RNA. This can be accomplished by using an R N A target that is attached to a solid support, such as a bead (Fig. 1B). In addition, one needs a method to amplify the population of bound proteins, allowing the selection process to be repeated (Fig. 1E). Lastly, a means to determine the identity of the selected "binders" is required (Fig. 1F). Since the binders consist of protein, this would require cumbersome technologies such as protein sequencing, unless the bound proteins are physically coupled to the genes that encode them. Such a physical link can be accomplished by using a method called phage display. Phage display involves the insertion of a foreign gene fragment into a filamentous bacteriophage genome in such a way that the foreign gene is fused to one of the phage coat protein genes (Smith, 1985). The foreign protein is then "displayed" as a fusion on the outside of the viral particle, while the corresponding gene is carried along inside the particle. This

12. In Vitro Genetics of RNA Binding Proteins

213

Figure 1 Schematicdepiction of in vitro genetic analysis of RNA binding proteins (RNA-BPs). (A) A library of mutated RNA-BPs is generated. (B) Proteins that can bind to a particular RNA target are bound selectively to RNA on beads. (C) Proteins that cannot bind are washed away. (D) Bound proteins are eluted. (E) The bound proteins are amplified to provide a new population for a subsequent round of selection. (F) Following several rounds of selection, the best binders are identified.

coupling of gene to protein is what makes in vitro genetic selection of proteins feasible. Figure 2 shows the result of the incorporation of phage display technology into the scheme depicted in Figure 1. The details of the system will be explained in the following paragraphs. The in vitro genetic system described in this chapter provides a powerful m e t h o d for the analysis of R N A - p r o t e i n interactions. In combination with existing in vitro genetic approaches aimed at studying nucleic acids (SELEX) (Gold et al., 1995), this in vitro genetic approach should rapidly increase our understanding of the mechanisms by which proteins specifically recognize certain R N A structures or sequences. The in vitro genetic system has several important advantages over other genetic methods. First, the target R N A by itself can be studied; it does not need to be attached to a long reporter R N A that could interact with the target sequence and s o m e h o w influence its structure. Second, the selective conditions

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Figure 2 In vitro selection of RNA binding protein variants that bind tightly to a target RNA. (A) A combinatorial library of mutagenized cDNA fragments encoding the RNA-binding domain of the protein of interest is inserted into a phagemid vector such that the resulting proteins become fused to the gene III coat protein of filamentous bacteriophage. The library is transformed into an E. coli host. (B) A pool of phage displaying a library of RNA-binding protein variants is generated by infecting the host with helper phage. (C) This phage pool is allowed to bind to the RNA target annealed to a biotinylated oligodeoxynucleotide. RNA-bound phage are captured on streptavidin-coated beads. (D) After washing, the phage bound to the beads are used to infect E. coli, allowing the cycle of amplification and selection to be repeated. (Adapted from Laird-Offringa and Belasco, 1996, Fig. 1, p. 150.)

can be completely controlled because they are occurring outside o f a living organism. Thus, buffer conditions, salt concentrations, detergents, etc., can be modified at will. M o r e important, any accessory proteins that may influence the binding reaction can be added. This should allow the study o f 1 L N A - p r o t e i n interactions in c o m p l e x systems involving n u m e r o u s components, such as the spliceosome. Third, due to the iterative nature o f the selection process, even rare clones can be amplified and isolated after a n u m b e r o f rounds o f selection. Successful display o f 1LNA binding proteins on phage is the most critical step o f the in vitro genetic system. Therefore, the emphasis o f this chapter will be on phage display: the theory b e h i n d it, the techniques used to p r o d u c e phage displaying 1LNA binding proteins, ways to measure w h e t h e r the displayed protein can actively bind to R N A , and ways to i m p r o v e a p o o r display. T h r o u g h o u t I will use the spliceosomal protein U 1 A as an example. This protein is a c o m p o n e n t o f the U1 small nuclear r i b o n u c l e o p r o t e i n ( R N P ) , which forms part o f the spliceo-

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215

some (Sillekens et al., 1987). U1A is a representative of the largest family of R N A binding proteins, those carrying so-called R N A recognition motif (RRM) domains, (also referred to as R N P domains) (Burd and Dreyfuss, 1994; Nagai et al., 1995). These ---100 amino acid domains are the RNA-binding motifs most commonly found in nature. They fold into a globular structure consisting of a four-stranded /3 sheet (the RNA-binding surface) supported by two o~ helices. They can be identified by two highly conserved regions (RNP-1 and RNP-2) present in the middle two strands of the ~3 sheet. Proteins carrying one or more R R M domains are found in all organisms, from viruses to higher eukaryotes, are involved in a wide variety of regulatory processes, and can bind to many different R N A structures and sequences. The U1A protein has two R R M domains, of which the N-terminal one allows it to bind tightly and specifically to a stem-loop structure in the U1 R N A called U1 hairpin II (UlhplI) (Lutz-Freyermuth et al., 1989; Scherly et al., 1989). The in vitro genetic method for the analysis of RNA-BPs was developed using the U1A N-terminal R R M as a model system (Laird-Offringa and Belasco, 1996). In the following sections, information relevant to the different steps of the in vitro genetic system will be discussed. The final section consists of detailed protocols describing the experimental procedures.

II. S U C C E S S F U L L Y M A K I N G D I S P L A Y P H A G E A. RELEVANT FACTS ABOUT PHAGE MORPHOGENESIS Phage display is a critical element of the in vitro genetic approach. Therefore, it is essential that the protein one wishes to study (from here on called X) is efficiently displayed. In order to be displayed, protein X must be fused to a bacteriophage protein that is on the surface of the viral particle. Filamentous phage (such as M13, fd, and fl) are long, thin packages of viral proteins surrounding a single-stranded DNA genome (Denhardt, 1975). Filamentous phage retard host growth but do not kill their host. This offers an advantage over display on lyric phage, since filamentous phage preparations are not heavily contaminated with debris from dead host cells. Filamentous phage particles are formed when the capsid proteins are assembled around a single-stranded viral DNA genome as it is extruded through the inner bacterial membrane. The two capsid proteins commonly used for display are the gene III and gene VIII proteins (Smith, 1993; Smith and Scott, 1993). Protein VIII is the major component of the viral particle and is present in about 2800 copies per virion. The actual number depends on the length of the viral particle and thus on the length of the encapsidated viral DNA. The gene III protein is present in five copies at the pointed end of the particle, which is assembled last. The protein is also called A, for attachment, and is responsible for the ability of the phage to bind to the Escherichia coli host pili. The N termini

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Ite A. Laird-Offringa

of both protein VIII and protein III are on the outside of the particle and can be used for the insertion of foreign polypeptides. During phage reproduction, both proteins are inserted in the bacterial membrane, where they await assembly of the virion, with their N termini protruding into the periplasm and their C termini remaining in the cytoplasm. Thus, in order to be successfully displayed at the N terminus of either protein, the foreign protein X must be able to pass through the bacterial membrane. While the fusion protein awaits assembly, it is exposed to interactions with periplasmic proteins, including proteases. Indeed, proteolytic cleavage of displayed proteins is often observed (McCafferty et al., 1991; Parmley and Smith, 1988). The extent of proteolysis seems to depend on many factors, including the host strain and the amino acid sequence of the displayed protein. Limited proteolysis is not usually a problem, since very large numbers of phage are easily generated (1012 particles per milliliter of bacterial supernatant).

B. VECTORS USED FOR PHAGE DISPLAY The first phage displaying a protein fragment was made by inserting a foreign sequence into the otherwise wild-type genome of a filamentous phage (Smith, 1985). However, working with phage has a number of disadvantages, such as low yields of DNA and the need to work with plaques instead of colonies. A very popular alternative is to use a phagemid instead. This is a plasmid carrying a filamentous phage origin of replication that allows it to replicate and be packaged in the presence of helper phage. Generally, a helper phage is tagged with antibiotic resistance and is partially replication deficient, allowing preferential replication and packaging of the phagemid [e.g., M13K07, carrying a kanamycin resistance marker (Vieira and Messing, 1987)]. pUC119 is a common phagemid on which a large number of current display vectors, including our own, are based (Vieira and Messing, 1987). The vector we use for display, pDISPLAY-B (Fig. 3), was derived from vector pHEN1 (Hoogenboom et al., 1991). It carries a gene III fusion cassette, into which gene X can be inserted. As mentioned in Section II,A, proteins can be displayed as either protein III or protein VIII fusions. Phagemids carrying gene VIII fusions allow the display of many copies of the fusion protein along the full length of the viral particle, whereas phagemids carrying gene III fusions will lead to phage with very few copies (mostly one or fewer; see below) at the tip of the virion (Bass et al., 1990). When trying to select for 1KNA binding proteins that can bind tightly and selectively to a specific 1KNA target, it is best to avoid multivalent phage. Therefore, a gene Ill-type vector is used. Vectors of the gene VIII type will not be further discussed. The fusion gene on pDISPLAY-B is controlled by the lac promoter and carries a signal sequence at its 5' end. This is followed by a polylinker region for insertion of gene X, a hexahistidine tag for protein purification, a MYC epitope tag, and the fd phage gene III. Bacteria carrying this vector produce virions only

12. In Vitro Genetics of R N A Binding Proteins

B

217

Sfi I Ncol Pst I Sal I 1 JI+ 1 G.GCC.CAG.CCG.GCC.ATG.GCC.CAG.GTG.CAG.CTG.CAG.GTC.GAC. Xba I Barn HI Not l (His)6-tag TCT.AGA.GGA.TCC.CCG.GTT.GCG.GCC.GCA.CAT.CAT.CAC.CAT.CAT.CAC. MYC-tag GTG.GCC.GCA.GAA.CAA.AAA.CTC.ATC.TCA.GAA.GAG.GAT.CTG.AAT.GGG. I

gene !11

GCC.GCA.AGA.TCT.GTT.GAA

Bgln Figure 3 The phage display vector pDISPLAY-B. (A) Schematic diagram of the pDISPLAY-B phagemid. This vector was derived from phagemid pHEN1 (Hoogenboom et al., 1991). Aside from a plasmid origin of replication, an ampicillin resistance gene, and the M13 origin of replication, the vector carries a gene III fusion cassette into which foreign genes can be inserted. The fusion cassette consists of the tac promoter, a signal sequence, a polylinker region, a hexahistidine tag (used for protein purification after mutants of interest have been transplanted to expression vectors), a MYC epitope tag (so that a single antibody can be used to recognize different fusion protein products), and gene III from bacteriophage fd. Restriction sites of interest are marked, with unique sites shown in boldface. (B) The polylinker and tag region of pDISPLAY-B. Unique restriction sites are shown in boldface. The signal sequence cleavage site is marked by a downward arrow. The hexahistidine tag, MYC epitope tag, and beginning of gene III are marked. The correct reading frame is indicated by dots between the codons.

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after they have been infected with helper phage. The produced particles contain either a helper phage genome or a phagemid in their interior and carry a mix of wild-type protein III (from the helper phage) and fusion protein III (from the phagemid) on their exterior. Of these, only particles that can bind to R N A and yield ampicillin-resistant colonies are studied. These particles, which carry the R N A binding protein on their exterior and the phagemid in their interior, will be referred to as display phage. All other particles are counterselected, either during the binding assay or during growth on selective plates. Aside from reducing the chance of getting "multivalent" phage, the mix of wild-type and fusion protein Ill ensures the full infectivity of the particles, since fusions to the gene III protein have been known to affect attachment. So far as I know, there is no limit to the size of the fragment that can be cloned into pDISPLAY-B. Possibly very large inserts might lead to unstable virions. The largest fragment we have inserted to date is a 1.4-kb fragment from the yeast poly(A)-binding protein gene (Sachs et al., 1986), encoding four R R M domains and part of the C-terminal end of the protein (also see Section V,B,4).

C. EFFICIENT DISPLAY OF R N A BINDING PROTEINS The most critical step in obtaining an efficient display is establishing conditions that allow the fusion protein to pass through the bacterial inner membrane. This is not a problem when displaying proteins such as antibodies, growth factors, or other naturally secreted proteins. But it may be difficult when trying to display intracellular proteins that have not been evolutionarily selected to be able to pass easily through the inner bacterial membrane. One can target proteins to the secretory pathway in E. coli by adding signal peptides to their N termini (Gennity et al., 1990). The signal peptide is removed by cleavage during secretion. However, addition of a signal peptide does not guarantee successful export. A protein may fold too quickly or may carry too many charged residues to pass through the bacterial membrane (Boyd and Beckwith, 1990; Model and Russel, 1990). Charged residues (especially positive ones) can interfere with export, in particular if they are near the signal sequence cleavage site. To reduce the possible effect of premature folding of protein X, one can lower the temperature of the bacterial culture. The U1A N-terminal R R M will not be displayed unless the bacteria are grown at 30~ In difficult cases, one could lower the growth temperature even further. Problems with charge may be avoided by choosing a region of gene X that does not encode many charged amino acids or by placing any charged regions as far away from the signal peptide cleavage site as possible. In cases where many charged residues are present throughout the region of interest, one can consider putting a spacer between the signal sequence and gene X. Since it may often be difficult to predict whether a protein will be exported, a derivative ofpDISPLAY-B was developed that can be used to measure the export

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of protein X before display is attempted (pDISPLAYblue-B; Fig. 4). A gene fragment inserted into the polylinker region of this vector will be fused at its 5' end to the signal sequence and at its 3' end to the bacterial alkaline phosphatase (phoA) gene. This will result in the formation of a fusion protein, directed to the secretory pathway. Alkaline phosphatase is a periplasmic enzyme that becomes activated only upon entry into the periplasm, where disulfide bonds essential for its function are formed. It can be used as a convenient marker for protein location (San Millan et al., 1989), since its activity can be visualized by growing bacteria on plates containing the chromogenic substrate 5-bromo-4-chloro-3-indolyl phosphate (BClP), which makes the bacteria turn blue. If the N-terminally fused protein X prevents alkaline phosphatase from being secreted, resulting in white colonies, one can experiment with reduced growth temperature or modification of the sequences surrounding the signal sequence cleavage site. Once conditions have been found that allow export of protein X, the phoA cassette can be removed by a simple NotI deletion and religation. This automatically puts gene X in flame with gene III. When using the phoA assay to determine secretion levels, it is useful to plate bacteria containing pDISPLAYblue-B (without insert X) and pDISPLAY-B (lacking phoA) for comparison. The former should give rise to bright blue colonies, whereas the latter should produce colonies that are white to very pale blue (the endogenous level ofph0A derived from the single copy on the bacterial chromosome is too low to significantly affect colony color), pDISPLAYblue containing an insert will usually give rise to colonies of an intermediate blue color, depending on the effect of the protein X fragment on secretion. For example, a construct containing a fragment encoding the N-terminal P,.RM of U1A leads to the formation of medium-blue colonies, whereas a construct containing most of the yeast poly(A) binding protein, including four R R M domains, yields colonies that are pale blue. If a more accurate quantitation of the phosphatase activity is desired, a liquid assay using cell extracts can be performed (Manoil, 1991).

D. HOST STRAIN REQUIREMENTS The E. coli host strain used for the production of display phage must be able to produce pill in order to be infected by helper phage. Consequently, it must carry an F episome, on which the genes for pili formation are located. (In order to form pili, the host must be grown at 37~ prior to infection.) In addition, the bacteria must contain the components to shut off expression of the gene III fusion. This is necessary for two reasons. First, expression of protein III immunizes the host against superinfection, ensuring that a single host can produce only a single kind of phage (Boeke et al., 1982). This means that expression of gene III from the phagemid would effectively immunize the host against helper phage infection. Second, we have found that the secretion of normally intracellular proteins can

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B

$fi I

Nco !

1 ~+ 1

Pst I

Sal I

G.GCC.CAG.CCG.GCC.ATG.GCC.CAG.GTG.CAG.CTG.----C-AG.GTC.---GAC.

Xba I

Bam HI

Not l

pho A

TCT.AGA.GGA.TCC.CCG.GTT.GCG.GCC.GCA.ACT.

C

(His)6-tag MYC-tag Not l GCG. G CC. G CA. CAT. CAT. CAC. CAT. CAT. CAC. GTG. G CC. G CA.G AA. CAA.AAA MYC-tag gene III .._ I -'CTC.ATC.TCA.GAA.GAG.GAT.CTG.AAT.GGG.GCC.GCA.AGA.TCT.GTT.GAA Bgl II

Figure 4 The phage display vector pDISPLAYblue-B. (A) This vector is identical to pDISPLAY-B, except for the insertion of an E. coliphoA gene segment [a PstI-NotI fragment of pPHO7, encoding the mature alkaline phosphatase gene (Gutierrez and Deved`jian, 1989)]. This insertion allows the efficiency of export of the fusion protein to be measured before display is attempted. A NotI site was incorporated.just downstream of the pkoA gene so that the pkoA cassette could be removed by a single digestion. (]3) The polylinker region upstream of the pkoA gene. Unique restriction sites are shown in boldface. The signal peptide cleavage site is marked by a downward arrow. The correct reading flame is marked by dots. (C) Sequence of the region downstream of the phoA insert. The unique BglII site allows inserts to be removed with the tags attached, so that they can be inserted into expression vectors. The correct reading flame is indicated by dots. (Adapted from Laird-Offringa and Belasco, 1996, Fig. 2, p. 154.)

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221

make E. coli sick, presumably because the inefficient export of proteins perturbs the bacterium's secretion machinery. This could lead to selection against fusion constructs. For these reasons, the fusion gene, under control of the lac promoter, is kept off at all times unless expression is required for phage production. This is achieved by the presence of high amounts of lac repressor (lacI q) inside the host (Bass et al., 1990). In addition, the bacteria are grown in 2% (w/v) glucose, which further represses the lac promoter (Hoogenboom et al., 1991). During phage production, glucose is left out of the medium, providing enough of the fusion transcript to allow the fusion protein to be displayed on phage. Any further induction [e.g., by the addition of small quantities of isopropyl-/3-D-thiogalactopyranoside (IPTG)] is deleterious and will not lead to an increase in displayed protein (Bass et al., 1990; Hoogenboom et al., 1991; our unpublished observations). The need to repress the fusion gene is another reason why using a phagemid vector is preferable to using a phage vector. In phage, the fusion gene is under the control ofthe authentic phage promoter, which does not allow it to be turned off. The E. coli strain TG1 carries both an F factor and the lacI q and is frequently used for display (Hoogenboom et al., 1991). A recombination-deficient strain (IAL-3) was derived from TG1 and was used for the U1A experiments. A disadvantage of this strain is that it can suppress amber codons. This is undesirable because it can lead to the accumulation of spontaneous amber codons in the fusion construct which will be preferred by the host because these codons lead to a slight reduction in fusion gene expression (Laird-Offringa and Belasco, 1995). Therefore, we have recently switched to strain DH12S (GIBCO-BRL, Gaithersburg, MD) which does not suppress nonsense codons. In addition, this strain can be made highly electrocompetent, a useful trait when large libraries are studied. DH12S produces titers of phage comparable to those provided by IAL-3; however, the percentage ofphage displaying R N A binding proteins is consistently twofold lower than that displayed when IAL3 is used (our unpublished observations, 1996). We suspect that DH12S may have more active periplasmic proteases. A characteristic that may make a strain better suited for the display of positively charged proteins is the introduction of the prlA 4 mutation (Derman et al., 1993). This is a mutation in one of the membrane components involved in secretion and may be added by P1 transduction (Sternberg and Maurer, 1991).

III. B I N D I N G

OF DISPLAYED RNA BINDING PROTEINS TO RNA

A. PREPARATION OF THE R N A TARGET The R N A targets that are used to selectively retain proteins are generated by in vitro transcription from linearized plasmid templates. They are attached to

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streptavidin-coated paramagnetic beads (Dynal, Lake Success, NY), which can be pulled to the side of a tube using a powerful magnet. The attachment of the R N A to the beads is indirect; the R N A target itself is not biotinylated, but is annealed to a biotinylated oligonucleotide complementary to the nucleotides at the 3' end of the R N A , derived from transcribed polylinker sequences. Figure 5 shows the R N A sequence and complementary oligonucleotide used as a target for U1A. The advantages of annealing the R N A to a biotinylated oligo instead of incorporating biotin into the R N A are that (1) the biotin is not located in the target area and will not interfere with binding, (2) the yield of the in vitro-transcribed R N A is high, and (3) the same biotinylated oligonucleotide can be used to link many different target R N A s to the beads. Binding can proceed in two ways. The R N A / D N A duplex can be bound to the beads first, followed by incubation of the beads with the display phage. This method is applied when an excess of R N A is used for binding, for example, when the percentage of phage that display the R N A binding protein is measured. Alternatively, the R N A / D N A duplex can be bound to the displayed protein first, followed by subsequent trapping of the bound R N A on beads. The latter procedure allows phage and R N A to bind in solution and provides better control over the actual R N A concentration. This method is used when more stringent binding conditions are desired, for example, when one tries to select the strongest binders from phage preparations containing many binders of similar strength (Laird-Offringa and Belasco, 1995). A disadvantage of this approach is that some R N A - b o u n d phage may be lost if the oligonucleotide is not 100% biotinylated. (Unpurified oligonucleotides may be contaminated with up to 50% nonbiotinylated oligos.)

UA CG GC

AU

~

5'- GGGAGACCCAGCAGGUCGACUCUAGAGGAUCCCCGG- 3' 3'- CTGAGATCT CCTAGGGGCCA-@

Figure 5 The UlhpII target RNA. The region corresponding to the sequence of wild-type UlhpII RNA is shaded. The sequence of the complementary oligodeoxynucleotidebiotinylated at the 5' end (circled B) is shown. An arrow marks the 3' end of a shorter transcript used for binding assays with purified UIA variants. (From Laird-Offringa and Belasco, 1996, Fig. 4, p. 159.)

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B. BINDING AND WASHING CONDITIONS The conditions used for binding displayed protein to 1KNA depend on the protein studied. In the case of U1A, salt and buffer conditions were based on published work (Hall and Stump, 1992; Scherly et al., 1990). Magnesium was omitted from the binding buffer to minimize possible 1KNase activity, and a little detergent (0.5% Triton X-100) was added to reduce background binding. Gel shift analyses showed that these modifications did not affect the ability of U1A to bind tightly to the UlhpII target IKNA. Ideally, the capacity of a protein to bind to its 1KNA target in the display binding buffer should always be tested beforehand. Once RNA-bound phage is trapped on the beads, the free phage must be washed away. We use an excess of binding buffer for this purpose. Since the bound phage can be released during washing, it is important to wash as quickly as possible. This is facilitated by the paramagnetic beads, which can be pulled to the side of a microfuge tube within 30-60 sec. Extended washing steps can be used to select for protein variants that release their 1KNA target very slowly.

C. DETERMINING THE PERCENTAGE OF PHAGE DISPLAYING AN ACTIVE R N A - B I N D I N G DOMAIN When starting to work with a new displayed protein, it is necessary to determine whether the displayed 1KNA-binding domain is active and what percentage of the phage carry an active domain. It is also useful to determine what the background level of binding is. Both of these questions can be answered by a single binding experiment. Display phage are incubated with an excess of 1KNA target attached to beads, allowing binding to occur. The beads are then washed, and the number of phage before and after binding selection is determined. It is convenient to include an internal control of "bare" phage to determine background binding levels. The control bare phage are made from a pDISPLAY-B derivative tagged with a lacZ gene (Laird-Offringa and Belasco, 1996). This phage will lead to the formation of blue colonies on indicator plates containing X-gal. For a typical experiment, an excess of lacZ-tagged phage is mixed with display phage. The mixture is titered to determine the input number of blue and white colonies. Following a single round of binding selection, the mixture is titered again. The ratio between the number of blue colonies after and before binding, multiplied by 100, will give the percentage background binding. This is usually between 0.0005% and 0.005%. In contrast, when the U IA 1KRM is displayed, 1-5% of the phage are retained. The number of white colonies relative to blue ones should increase after binding selection, indicating the fold purification per round of selection. Based on colony counts, a single round of selection leads to a severalhundred-fold purification of the U IA-carrying phage. This purification factor is

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an underestimate, since only 5% or less of the colonies counted before binding actually display an active RNA-binding domain, while all of the colonies counted after binding selection display one. Thus it is estimated that "binders" can be purified by over four orders of magnitude during a single round of binding selection. The relatively low percentage display seen with U1A is comparable to the percentage display observed by others using different proteins and is a result of partial proteolysis of the displayed protein and the presence of wild-type protein III derived from the helper phage. The presence of fewer than one RNA-binding domain per phage particle ensures that all binding phage are monovalent. Background binding levels can also be determined by incubating display phage with beads carrying no R N A or an irrelevant RNA. Background binding levels determined this way are comparable to those determined using the bare lacZ-tagged phage. The number of both white and blue colonies should decrease dramatically, and no change in the ratio of blue to white colonies should be observed.

IV. A P P L I C A T I O N O F In Vitro G E N E T I C S T O T H E STUDY OF RNA BINDING PROTEINS A. ANALYSIS OF THE INTERACTION BETWEEN KNOWN R N A BINDING PROTEINS AND THEIR R N A TARGETS Once protein X has been shown to be present in its active RNA-binding form on the outside of the phage, the stage is set for in vitro genetic experiments. The in vitro system is ideally suited for the detailed analysis of small regions of known 1KNA binding proteins. The procedure is very similar to the binding experiment described in Section III, except that instead of a single clone, a library of phagemid clones is used to generate a library of displayed proteins. The size of the libraries that can be studied by this method is limited by the amount of DNA bacteria can take up. Very highly electrocompetent cells can yield 101~ colonies per microgram of supercoiled DNA. Thus, the maximal library size that can be analyzed is about 108-109 per microgram ofligated vector. A completely randomized region of seven amino acids represents a library of-> 1 X 109 possible clones, which is approximately equal to randomization of a stretch of 30 amino acids to two possible amino acid identities. The best way to generate a phagemid library that is mutated in a defined region is to replace parts of gene X by randomized oligonucleotides. One approach is to use an oligonucleotide that is randomized in the middle and flanked at either side by constant regions containing unique restriction sites. The oligonucleotide is first made double-stranded by hybridizing a small oligo to its constant 5' end and extending this primer using DNA polymerase. A prerequisite for this approach

12. In Vitro Genetics of RNA Binding Proteins

225

is the presence of restriction sites at convenient locations. These can be generated beforehand by site-directed mutagenesis. However, the codon usage in the area may not be suitable for the insertion of silent restriction sites. An alternative approach is to use a partially randomized oligonucleotide as a polymerase chain reaction (PCR) primer. The constant 3' region would serve to prime the "forward" reaction, and a small, constant oligonucleotide hybridizing downstream would prime the "backward" reaction. In this case, only one restriction site close to the mutagenized region is needed; the other site may lie far away. Care must be taken when choosing a template for such a P C R reaction because differential priming may lead to the production of a biased library. The P C R approach was used to generate a library aimed at determining the importance of each residue in a nineamino acid loop in the N-terminal R R M domain of U i A for binding to UlhplI (Laird-Offringa and Belasco, 1995). In that particular experiment, the nine amino acids were each randomized to two possible identities using a randomized oligonucleotide. After every round of binding selection, the selection procedure can be repeated by taking the bound phage and infecting the host to produce a new display phage library enriched for mutants that bind to the target RNA. At any time, the progress of the selections can be determined by analyzing the phage population. One convenient way of looking for changes is sequence analysis of the whole phage population. In the experiment with U1A mentioned earlier, the importance of the identity of each amino acid was assessed by observing the rate and position of sequence changes in the pool of phage, which reflected preferences for certain amino acids. Residues important for binding quickly dominated at certain positions, whereas irrelevant positions showed very little preference for a particular amino acid. The number of selective rounds that are carried out depends on the size of the starting library and deserves careful consideration. Large libraries will require more rounds of selection than smaller ones. The starting pool of the U1A experiment had only 512 members, and after merely two rounds of selection, marked changes were seen at several positions. Once a group of clones with very similar binding characteristics is established, certain clones may take over the population. A single clone cannot provide any information about the relative importance of each amino acid residue in a mutagenized area, since it is unclear which of the residues exerted the strongest selective effect and which residues simply colocalize with the important ones. Due to the in vivo propagation of the display phage library, it must be taken into account that minor biological advantages may start to play a role at this point. Continuing the selection for too many rounds will result in a loss of information. This happened in the U I A experiment when a single clone took over the population after four rounds of selection. The clone bound slightly less well to the target R N A than did clones conforming to the consensus determined from the second round of selection. It was determined that

226

Ite A. Laird-Offringa

the final clone had a slight biological advantage due to a fortuitous amber mutation outside of the mutagenized region (Laird-Offringa and Belasco, 1995). This example illustrates the importance of confirming biochemically the results of the genetic selection. Due to the limits on display library size, randomization of large protein fragments is not an option. One possible method for the study of large protein domains is the selection of compensatory mutations in the P, NA binding protein using an R N A target carrying a defined mutation. Chemical mutagenesis (Meyers, 1987) or error-prone PCP, (Cadwell and Joyce, 1994) can be used to introduce mutations in to the R N A binding protein. Mutagenesis of the signal sequence must be avoided to prevent the introduction of mutations that may provide biological advantages to certain clones.

B. PROSPECTS AND FUTURE APPLICATIONS A possible application of the in vitro genetic system is the identification of genes encoding R N A binding proteins that bind to known R N A sequences. The fact that the selection process can be repeated many times, allowing rare clones to be amplified, is a big advantage of using in vitro genetics as opposed to traditional methods. A disadvantage is that complementary DNA (cDNA) libraries inserted in pDISPLAY-B must be in frame at both ends and may contain stop codons that would prevent the protein fusion from being formed. Thus, only one out of nine codin2 region inserts would be in the right frame. In addition, there is no guarantee that the sought protein can be adequately secreted. The use of a display vector in which cDNAs can be fused C-terminally, such as pJuFo, may offer a solution (Crameri et al., 1994). The displayed protein is fused C-terminally to a FOS leucine zipper, which attaches to the phage via a J U N leucine zipper fused N-terminally to the gene III protein. In addition to reducing the number of possible reading frames and the interference of stop codons, this vector may allow better secretion, since the cDNA is separated from the signal sequence by the leucine zipper encoding fragment. One of the most exciting applications of the in vitro genetic system is the evolution of R N A binding proteins with new binding affinities. By incorporating a mutagenic step into the selection cycles, allowing new changes to be introduced during each round, it may be possible to evolve proteins that can bind to new target RNAs. Such tailor-made proteins could be very useful tools for the manipulation of gene expression and may one day be useful as therapeutic agents that target undesirable RNAs for destruction. Experiments to determine the feasibility of R N A binding protein evolution by phage display are underway.

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V. PROTOCOLS

A. BACTERIAL STRAINS AND REAGENTS 1. E. coli Strains

IAL-3 is a recA::cat derivative of TG1 (K12, A(lac-pro), supE, thi, hsdD5/F' traD36, proA+B+'lacP, lacZAM15) (Hoogenboom et al., 1991). DH12S is d?80dlacZAM15, mrcA, A(mrr-hsdRMS-mcrBC), araD139, A(ara, leu)7697, AlacX74, galU, galK, rpsL, deoR, nupG, recAl/F'proAB +, lacI q ZAM15 and was obtained from GIBCO-BRL.

2. Reagents

Ampiciltin: 1000X Stock solution of 100 mg/ml in water, filter sterilized, stored at - 2 0 ~

BCIP: 5-Bromo-4-chloro-3-indoyl phosphate (Sigma, St. Louis, MO), stored at - 2 0 ~ as a 40 mg/ml (1000• concentrated) stock solution in N,N-dimethylformamide Glucose stock solution (20% w/v): Dissolve glucose in distilled water, filter sterilize LB: 10 g Bacto-tryptone, 5 g Bacto-yeast extract, and 10 g NaC1 per liter of distilled water, sterilized. For plates, add 15 g Bacto-agar per liter medium Kanamycin: 1000• Stock solution of 70 mg/ml water, filter sterilized, stored at - 2 0 ~ Polyethylene glycol (PEG)/ammonium acetate: 20% Polyethylene glycol 8000, 3.5 M ammonium acetate Protease inhibitors: Phenylmethylsulfonyl fluoride (PMSF), 100 mM stock in ethanol, stored at -20~ used at 500 tzM; pepstatin, 1 mg/ml stock in methanol, stored at 4~ used at 0.7/zg/rnl; leupeptin, 1 mg/ml stock in water, stored at -20~ used at 0.5/zg/ml TE: 10 mM Tris-HC1 pH 8.0, 1 mM ethylenediaminetetraacetic acid (EDTA), autoclaved (Sigma) 1• TENT buffer: 10 mM Tris-HC1 pH 8.0, 1 mM EDTA, 250 mM NaC1, 0.5% (v/v) Triton X-100 Top agar: LB containing 7 g Bacto-agar per liter of medium, autoclaved tRNA: From E. coti, 20 mg/ml stock solution in water (Sigma) 2 • YT: 16 g Bacto-tryptone, 10 g Bacto-yeast extract, 5 g NaC1 per liter of distilled water

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Ite A. Laird-Offringa

X-gal: 5-Bromo-4-chloro-3-indolyl-/3-D-galactopyranoside (Research Products International Corp., Mount Prospect, IL), 40 mg/ml stock solution in N,N-dimethyl formamide, stored at - 2 0 ~

B. METHODS 1. Preparation of Constructs in pDISPLAYblue and Analysis on Indicator Plates When a protein is used for display for the first time, it is wise to insert the encoding gene fragment into the polylinker of pDISPLAYblue-B (Fig. 4) so that the ability of the protein to be secreted can be assessed. Unique sites in the polylinker are PstI, SalI, and XbaI. SfiI is unique but lies in the signal sequence. Later constructs can be inserted into the vector lacking the phoA cassette (pDISPLAY-B, Fig. 3), which has two additional unique sites in the polylinker: NotI and NcoI. However, NcoI lies just upstream of the signal sequence cleavage site. It can be used for cloning when the resulting amino acid at the - 1 position is small and uncharged. Naturally, care must be taken to insert gene X fragments in frame with the upstream signal sequence and the downstream gene III. For the reasons outlined in Section II,D, bacteria containing the constructs should be grown in the presence of 2% glucose at all times except during phage production. Once the desired pDISPLAYblue-B constructs are obtained, bacteria containing the plasmid are streaked on indicator plates containing 100/~g/ml ampicillin, 2% glucose, and 40 /zg/ml BCIP and are incubated at 30~ for at least 18 hr. To best compare colony color, bacteria containing pDISPLAYblue-B and pDISPLAY-B should also be streaked on the same plates. No increase in blue color compared to the negative control could mean that a cloning error was made (construct not in frame), that a stop codon is present, or that protein X is interfering with secretion. If the colonies are blue, the phoA cassette is deleted out of the construct by digestion with NotI and religation. Infection of the bacteria with helper phage will now allow display phage to be produced. When a phagemid library containing a mutagenized fragment is prepared, enough colonies to represent the whole phagemid library multiple times are plated on large plates. These colonies are scraped and pooled in a minimal amount of LB containing 20% glycerol. The stock is stored at -80~ A small aliquot is used to inoculate a culture for display phage preparation. If the library is too large to be plated, the cells may be grown in liquid culture. To avoid changes in the composition of the library, growth times should be minimized.

2. Preparation of Helper Phage Stock Aerosol-blocking pipette tips should be used when working with phage or phagemid stocks. This will help prevent cross-contamination.

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229

The E. coli host strain is inoculated from a glycerol stock or a fresh plate into LB and shaken overnight at 37~ 250 rpm. The next day, the culture is diluted 1:50 into 10 ml of 2• YT and incubated with shaking (250 rpm) for 30 min at 37~ (A 1:500 dilution of host in LB is also made and is grown for ---6 hr at 37~ with shaking. These cells will be used to determine the helper phage titer.) Helper phage is added to a final concentration of 1.108 plaque-forming units (pfu)/ml [we use M13K07 (Vieira and Messing, 1987); Stratagene (La Jolla, CA) sells VCSM13]. After incubation with shaking for 30 min at 37~ kanamycin is added to 7 0 / , g / m l to select for cells infected by the helper phage. The culture is shaken at 37~ 250 rpm, for 4 hr. Several 1.3-ml aliquots are pipetted into 1.5-ml microfuge tubes and centrifuged for 5 min at full speed. Then 1-ml aliquots of supernatant are taken to a new tube (leaving the bacterial pellet untouched) and centrifuged again. A 900-p.1 supernatant is taken from each tube, deposited into a clean tube, and heated at 65~ for 10 min to kill any remaining cells. The helper phage stock is stored at 4~ The titer is determined by making serial 1 : 100 dilutions (5/,1 plus 495/~1) in LB, adding 100/.d of the 10 -6 and 10 -8 dilutions to 200/.d of host cells each, pipetting into 3 ml of warm top agar (50~ mixing, and pouring onto an LB plate. After the top agar has solidified, plates are incubated overnight at 37~ Plaques will be visible as small, circular areas of retarded growth on a lawn of cells. Host cells alone and phage alone are plated as controls. Typical helper phage titers are 1-2.1011 pfu/ml.

3. Preparation of Display Phage The fusion phage construct(s) are transformed into the host strain. The next day, a fresh colony is inoculated into 2 ml 2• YT, 100 b~g/ml ampicillin, and 2% glucose and shaken overnight at 37~ 250 rpm. The overnight culture is diluted 1:200 into 20 ml of 2• YT, 100/~g/ml ampicillin, and 2% glucose and shaken at 37~ 300 rpm, until an OD600 of---0.5 is reached. When a library is used, growth times are shortened to minimize changes in the population. In that case, an aliquot (e.g., 20 N1) of the frozen glycerol stock of the bacteria containing the phagemid library is inoculated into 20 ml of 2• YT, 1 0 0 / , g / m l ampicillin, and 2% glucose and grown for approximately 2 hr until an OD600 of---0.5 is reached. Then 10 ml of the culture is pipetted into a 30-ml centrifuge tube, and helper phage is added to 1.101~ pfu/ml. After standing at 37~ for 30 min, the mixture is centrifuged for 10 min at room temperature, 3500 rpm, in a Sorvall SS34 rotor. As much supernatant as possible is removed, and the bacterial pellet is resuspended in 20 ml 2• YT, 100 ~g/ml ampicillin, and 70/.~g/ml kanamycin and shaken overnight at 30~ 300 rpm. (A host cell culture is also inoculated for growth overnight, to be used for titer determination. This stock is diluted 1:500 in LB the next day and grown for 6 hr. Freshly grown stock produces the best results.) The overnight culture is centrifuged for 20 min at 5000 rpm, 4~ in a

230

Ite A. Laird-Offringa

Sorvall SS34 rotor. Then 16 ml of supernatant is transferred to a new tube, to which 4 ml of PEG/ammonium acetate is added. The solution is mixed by inversion and set on ice for 1 hr, then centrifuged for 15 min at 12,000 rpm, 4~ in a Sorvall SS34 rotor. The supernatant is discarded, and the pellet is resuspended in 1 ml TE containing protease inhibitors and transferred to a sterile 1.5-ml microtube. After centrifugation for 2 min at maximal speed in a microfuge to remove remaining bacteria, the supernatant is transferred to a clean tube, 250/,1 PEG/ammonium actetate is added, and the mixture is incubated on ice for 1 hr. It is then centrifuged for 5 min at maximum speed. The supernatant is discarded, and the pellet is resuspended in 100/,1 TE containing protease inhibitors. (When larger amounts of display phage are desired, the preparation can simply be scaled up.) The titer is determined by making serial dilutions in LB (5 ~1 plus 495/.,1 LB for each 100fold dilution), taking 100/,1 of the 10 -8, 10 -1~ and 10 -12 dilutions, adding 100 ~1 host to each, standing at 37~ for 30 min, and plating on LB plates containing 100 ~g/ml ampicillin and 2% glucose. To control for contamination of phage preparations with bacteria or for infection of the host with phage, host alone and phage alone (10 -2 dilution) are always plated as well. These preparations of fusion phage consist of a mix of phage displaying no protein or protein X on their exterior and carrying the helper phage genome or the phagemid in their interior. The helper phage does not interfere with the experiments because it does not give rise to ampicillin-resistant colonies. Whether protein X is displayed can be assessed by Western blot analysis (see the following section). Whether the displayed protein is functional must be determined by performing an R.NA-binding experiment with an excess of tLNA (see Section V,B,6).

4. Western Blot Analysis of Display Phage It is useful to prepare a stock ofpDISPLAY-B (no insert) phage for comparison by Western blot analysis. The "empty" vector produces a band migrating at about 65 kDa (the actual size is about 45 kDa). A single I0.5%) and concentrated Tris buffers (i.e., >30 mM) in these reactions because they scavenge hydroxyl radicals and severely inhibit cleavage efficiency (Celander and Cech, 1990; Tullius et al., 1987). Sodium phosphate buffer (pH 7.0), on the other hand, significantly increases the amount of cleavage (Tullius et al., 1987). Tullius et al. (1987) have provided a detailed, quantitative analysis of the effect of various buffer components and reaction conditions on radical-induced cleavage of DNA. In addition, Celander and Cech (1990) have examined Fe(II)-EDTA-catalyzed cleavage of an P,.NA substrate under varying cation and Tris buffer concentrations. These studies provide a useful resource for adapting binding conditions to suit a particular RNA-protein interaction while maintaining efficient levels of radical-induced cleavage. Binding and Fe(II)-EDTA cleavage reactions were performed as described by Ansel-McKinney et al. (1996). It is extremely important to minimize P,.NA storage time after radiolabeling. For the best results, we use RNAs labeledonly a few days prior to the cleavage reactions because a notable level of R N A hydrolysis is evident 1-2 weeks after labeling. The resulting bands complicate the analysis of hydroxyl radical-induced cleavage sites and, by increasing the background (Tullius et al., 1987), can significantly reduce the level of detectable protection. Prior to incubation with protein, 5' end-labeled P, NA is renatured in a buffer containing 10 mMsodium phosphate (pH 6.5), 50 mMNaC1, 3 mMMgC12, and 0.1 m M EDTA by heating at 68~ for 5 rain, followed by slow cooling over ---15-30 min to less than 35~ For binding reactions, about 1 • 10 6 cpm of

15. IkNA-Protein Interactions by Footprinting

293

R N A is incubated in the absence or presence of protein (i.e., 1 /xM) or peptide (i.e., 10-20/xM) in an Fe(II)-EDTA binding buffer (10 m M sodium phosphate (pH 6.5), 50 mMNaC1, 1 m M E D T A , 0.15/xg//xl yeast tRNA) at room temperature for 30 min (Ansel-McKinney et al., 1996). Fe(II)-EDTA-catalyzed strand scission is performed as reported by Latham and Cech (1989), except that cleavage is for 1 hr at room temperature in 2 m M Fe(II), 4 m M EDTA, and 10 m M DTT A 20 m M ammonium iron(II) sulfate hexahydrate (Aldrich, Milwaukee, W I ) / 40 m M EDTA solution and a 100 m M DTT solution are freshly prepared, and 1/xl of each solution is added to the 8/xl binding reactions by careful pipetting. The reaction is quenched by adding thiourea, an effective hydroxyl radical scavenger (Tullius et al., 1987), to a final 10 m M concentration (1/xl of a 100 m M thiourea solution), followed by a 5 min incubation. Gel loading buffer (11/xl of 10 M urea in 0.25 • TBE buffer, plus dyes) is added, and the solution is heated at 68~ for 5 min before 3.3 • 10 s cpm per reaction is loaded onto a 20% polyacrylamide-urea gel to resolve the chemically cleaved products. Due to the high percentage of polyacrylamide and accompanying complications of gel drying, autoradiography is routinely performed on wet gels, with an exposure time of---6-12 hr. Drying is, however, recommended for gels containing acrylamide concentrations -70%) required between the defective IRES and the complementing molecule for efficient complementation to occur (Stone et al., 1993; Roberts and Belsham, 1997). It is important to emphasize that the complementation is quite efficient; it is observed without any selection process and occurs without detectable recombination between the two molecules. In a typical experiment, it is possible to enhance the activity of a mutant EMCV IRES from less than 1% of wt activity to about 25% of wt IRES activity without any sign of a recombinant molecule being generated, as .judged by reverse transcription-polymerase chain reaction (RT-PCR) analysis (Roberts and Belsham, 1997). These results are in accord with the estimate (Evans et al., 1988) of about one recombination event occuring every 45 kbp between plasmid DNAs transfected into vaccinia-infected cells during a 24-hr period in the absence of any selection. It is only through the use of the vaccinia/T7 R N A polymerase system that this property of IRES-directed translation has been observed. It may be argued that it is an artifact of the system, but this seems unlikely for several reasons. First, it is worth noting that picornaviruses like EMCV can translate and replicate very efficiently in vaccinia-infected cells (Whitaker-Dowling and Younger, 1986), so there is no adverse effect of the vaccinia virus infection on the ability of this IRES to function. Second, as already mentioned, where similar analyses of either mutant forms of the IRES or studies to determine the boundaries of the IRES elements have been performed, there has been essentially complete agreement between the results obtained using the vaccinia/T7 system and other assay systems, such as

17. Picornavirus IRES Function

337

RRL. It is likely that the complementation effect is a result of the high levels of R N A transcripts achieved within the cell. However, it is worth mentioning that in a picornavirus infection, up to 5% of the total cellular R N A can be represented by the viral RNA, i.e., as much as all the cellular mRNA. This is not the only example in which the activity of a complex R N A structure can be complemented in trans by another R N A molecule. Several studies have shown that the activity of the Tetrahymena thermophilia Group I self-spicing intron ribozyme can be regenerated by assembling the separated domains (van der Horst et al., 1991; Doudna and Cech, 1995), and non-Watson-Crick interactions between separate R N A fragments have also reconstituted Escherichia coli M1 R N A catalytic activity (Guerrier-Takada and Altman, 1992). Although we do not think that picornavirus IRES elements usually function in trans, we believe these observations are significant for understanding the mechanism by which the IRES directs the ribosome to the initiation site of protein synthesis. Further studies, facilitated by the use of the transient expression system described here, should help in this process.

ACKNOWLEDGMENTS I thank Rachael Seamons for excellent technical assistance, and also Lisa Roberts, Morwenna Robertson, and Rachael Seamons for helpful comments on the manuscript.

REFERENCES Agol, V. I., Drozdov, S. G., Ivannikova, T. A., Kolesnikova, M. S., Korolev, M. B., and Tolskaya, E. A. (1989). Restricted growth of attenuated poliovirus strains in cultured-cells of a human neuroblastoma. J. Virol. 63, 4034-4038. Belsham, G.J. (1992). Dual initiation sites of protein synthesis on foot-and-mouth disease virus RNA are selected following internal entry and scanning of ribosomes in vivo. EMBO_J. 11, 1105-1110. Belsham, G.J., and Brangwyn, J. K. (1990). A region of the 5' noncoding region of foot-and-mouth disease virus R N A directs efficient internal initiation of protein synthesis within cells: Involvement with the role of L protease in translational control. J. Virol. 64, 5389-5395. Belsham, G.J., and Sonenberg, N. (1996). RNA-protein interactions in regulation of picornavirus R N A translation. Microbiol. Reg. 60, 499-511. Borman, A., and Jackson, R.J. (1992). Initiation of translation of human rhinovirus RNA: Mapping the internal ribosome entry site. Virology 188, 685-696. Brown, E. A., Day, S. P., Jansen, R. W., and Lemon, S. M. (1991). The 5' nontranslated region of hepatitis A virus RNAmsecondary structure and elements required for translation in vitro. J. Virol. 65, 5828-5838. Chen, C.-Y., and Sarnow, P. (1995). Initiation of protein synthesis by the eukaryotic translational apparatus on circular RNAs. Science 268, 415-417. Das, S., Kenan, D. J., Bocskai, D., Keene, J. D., and Dasgupta, A. (1996). Sequences within a small yeast RNA required for inhibition of internal initiation of translation: Interaction with La and other cellular proteins influences its inhibitory activity..J. Virol. 70, 1624-1632.

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Devaney, M. A., Vakharia, V. N., Lloyd, R. E., Ehrenfeld, E., and Grubman, M. J. (1988). Leader protein of foot-and-mouth disease virus is required for cleavage of the p220 component of the cap binding protein complex. J. Virol. 62, 4407-4409. Dorner, A. J., Semler, B. L., Jackson, R. J., Hanecak, R., Duprey, E., and Wimmer, E. (1984). In vitro translation of poliovirus RNA: Utilization of internal initiation sites in reticulocyte lysate. J. Virol. 50, 507-514. Doudna, J. A., and Cech, T. R. (1995). Self-assembly of a group I intron active site from its component tertiary structural domains. R N A 1, 36-45. Drew, J., and Belsham, G.J. (1994). trans-Complementation by RNA of defective foot-and-mouth disease virus internal ribosome entry site elements. J. Virol. 68, 697-703. Elroy-Stein, O., Fuerst, T. R., and Moss, B. (1989). Cap-independent translation o f m R N A conferred by the encephalomyocarditis virus 5' sequence improves the performance of the vaccinia/bacteriophage T7 hybrid expression system. Proc. Natl. Acad. Sci. U.S.A. 86, 6126-6130. Etchison, D., Milburn, S., Edery, I., Sonenberg, N., and Hershey, J. W. B. (1982). Inhibition of HeLa cell protein synthesis following poliovirus infection correlates with the proteolysis of a 220,000 dalton polypeptide associated with eukaryotic initiation factor 3 and a cap-binding complex. J. Biol. Chem. 257, 14806-14810. Evans, D. H., Stuart, D., and McFadden, G. (1988). High levels of genetic recombination among cotransfected plasmid DNAs in pox-virus infected mammalian cells. J. Virol. 62, 367-375. Fuerst, T. R., Niles, E. G., Studier, F. W., and Moss, B. M. (1986). Eukaryotic transient expression system based on recombinant vaccinia virus that synthesizes bacteriophage T7 R N A polymerase. Proc. Natl. Acad. Sci. U.S.A. 83, 8122-8126. Garcia-Sastre, A., Muster, T., Barclay, W. S., Percy, N., and Palese, P. (1994). Use of a mammalian internal ribosomal entry site element for expression of a foreign protein by a transfectant influenza virus. J. Virol. 68, 6254-6261. Gingras, A.-C., Svitkin, Y., Belsham, G. J., Pause, A., and Sonenberg, N. (1996). Activation of the translational suppressor 4E-BP1 following infection with encephalomyocarditis virus and poliovirus. Proc. Natl. Acad. Sci. U.S.A. 93, 5578-5583. Glass, M.J., and Summers, D. F. (1993). Identification ofa trans-acting activity from liver that stimulates hepatitis A virus translation in vitro. Virology 193, 1047-1050. Glass, M. J., Jia, x. Y., and Summers, D. F. (1993). Identification of the hepatitis-A virus internal ribosome entry site--in vivo and in vitro analysis of bicistronic RNA's containing the HAV 5' noncoding region. Virology 193, 842-852. Guerrier-Takada, C., and Altman, S. (1992). Reconstitution of enzymatic activity from fragments of M1 RNA. Proc. Natl. Acad. Sci. U.S.A. 89, 1266-1270. Haghighat, A., Mader, S., Pause, A., and Sonenberg, N. (1995). Repression of cap-dependent translation by 4E-binding protein-l--competition with p220 for binding to eukaryotic initiation factor 4E. E M B O J . 14, 5701-5709. Hellen, C. U. T., Pestova, T. V., and Wimmer, E. (1994). Effect of mutations downstream of the internal ribosome entry site on initiation of poliovirus protein-synthesis. J. Virol. 68, 6312-6322. Hinnebusch, A. (1996). Translational control of GCN4: Gene specific regulation by phosphorylation of elF2. In "Translational Control" (J. w . B. Hershey, M. B. Mathews, and N. Sonenberg, eds.), pp. 199-244. Cold Spring Harbor Lab. Press, Cold Spring Harbor, New York. Jackson, R.J., and Kaminski, A. (1995). Internal initiation of translation in eukaryotes: The picornavirus paradigm and beyond. R N A 1, 985-1000. Jang, S. K., Krausslich, H.-G. Nicklin, M. J. H., Duke, G. M., Palmenberg, A. C., and Wimmer, E. (1988). A segment of the 5' non-translated region of encephalomyocarditis virus R N A directs internal entry of ribosomes during in vitro translation. J. Virol. 62, 2636-2643. Kaminski, A., Howell, M. T., and Jackson, R. J. (1990). Initiation of encephalomyocarditis virus RNA translation: The authentic initiation site is not selected by a scanning mechanism. E M B O J . 9, 3753-3759.

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Kaminski, A., Hunt, S. L., Patton, J. G., and Jackson, IL. J. (1995). Direct evidence that polypyrimidine tract binding protein (PTB) is essential for internal initiation of translation of encephalomyocarditis virus 1LNA. R N A 1, 924-938. Kozak, M. (1989). The scanning model for translation: An update. J. Cell Biol. 108, 229-241. Kratisslich, H.-G., Nicklin, M.J., Toyoda, H., Etchison, D., and Wimmer, E. (1987). Poliovirus protease 2A induces cleavage of eucaryotic initiation factor 4F polypeptide p220. J. Virol. 61, 2711-2718. Kuge, S., Kawamura, N., and Nomoto, A. (1989). Genetic variation occurring on the genome of an in vitro insertion mutant poliovirus type 1. J. Virol. 63, 1069-1075. LaMonica, N., and Racaniello, V. IL. (1989). Differences in replication of attenuated and neurovirulent polioviruses in human neuroblastoma cell line SH-SY5Y. J. Virol. 63, 2357-2360. Lamphear, B.J., Kirchweger, IL., Skern, T., and Rhoads, R. E. (1995). Mapping of functional domains in eukaryotic protein synthesis initiation factor 4G (elF4G) with picornaviral proteases. J. Biol. Chem. 270, 21975-21983. Macejak, D. G., and Sarnow, P. (1991). Internal initiation of translation mediated by the 5' leader of a cellular mRNA. Nature (London) 353, 90-94. Mader, S., Lee, H., Pause, A., and Sonenberg, N. (1995). The translation initiation factor elF-4E binds to a common motif shared by the translation factor eIF-4T and the translational repressors 4E-binding proteins. Mol. Cell. Biol. 15, 4990-4997. Martinez-Salas, E., Saiz, J.-C. Davila, M., Belsham, G.J., and Domingo, E. (1993). A single nucleotide substitution in the internal ribosome entry site of foot-and-mouth disease virus leads to enhanced cap-independent translation in vivo. J. Virol. 67, 3748-3755. Martinez-Salas, E., Regalado, M. P., and Domingo, E. (1996). Identification of an essential region for internal initiation of translation in the aphthovirus internal ribosome entry site and implications for viral evolution. J. Virol. 70, 992-998. Merrick, W. C. (1992). Mechanism and regulation of eukaryotic protein synthesis. Microbiol. Rev. 56, 291-315. Merrick, W. C., and Hershey, J. W. B. (1996). The pathway and mechanism of eukaryotic protein synthesis. In "Translational Control". (J. W. B. Hershey, M. B. Mathews, and N. Sonenberg, eds.), pp. 31-69. Cold Spring Harbor Lab. Press, Cold Spring Harbor, N.Y. Molla, A., Paul, A. V., and Wimmer, E. (1991). Cell-free, de novo synthesis of poliovirus. Science 254, 1647-1651. Oh, S. K., Scott, M. P., and Sarnow, P. (1992). Homeotic gene antennapedia messenger-lLNA contains 5'-noncoding sequences that confer translational initiation by internal ribosome binding. Genes Dev. 6, 1643-1653. Pause, A., Belsham, G.J., Gingras, A.-C., Donze, O., Lin, T.-A., Lawrence, J. C., Jr., and Sonenberg, N. (1994a). Insulin dependent stimulation of protein synthesis by phosphorylation of a novel regulator of 5'-cap function. Nature (London) 371, 762-767. Pause, A., Methot, N., Svitkin, Y., Merrick, W. C., and Sonenberg, N. (1994b). Dominant negative mutants of mammalian translation initiation factor elF-4A define a critical role for elF-4F in capdependent and cap-independent initiation of translation. E M B O J . 13, 1205-1215. Pelletier, J., and Sonenberg, N. (1988). Internal initiation of translation of eukaryotic mRNA directed by a sequence derived from poliovirus RNA. Nature (London) 334, 320-325. Pelletier, J., Flynn, M. E., Kaplan, G., Racaniello, V. R., and Sonenberg, N. (1988). Mutational analysis of upstream AUG codons of poliovirus RNA. J. Virol. 62, 4486-4492. Percy, N., Belsham, G. J., Brangwyn, J. K., Sullivan, M., Stone, D. M., and Almond, J. W. (1992). Intracellular modifications induced by poliovirus reduce the requirement for structural motifs in the 5' non-coding region of the genone involved in internal initiation of protein synthesis.J. Virol. 66, 1695-1701. Pilipenko, E. V., Blinov, V. M., Romanova, L. I., Sinyakov, A. N., Maslova, S. V., and Agol, V. I. (1989a). Conserved structural domains in the 5'-untranslated region of picornaviral genomes: An analysis of the segment controlling translation and neurovirulence. Virology 168, 201-209.

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Pilipenko, E. V., Blinov, V. M., Chemov, B. K., Dmitrieva, T. M., and Agol, V. I. (1989b). Conservation of the secondary structure elements of the 5'-untranslated region of cardio- and aphthovirus RNAs. Nucleic Acids Res. 17, 5701-5711. Reynolds, J. E., Kaminski, A., Kettinen, H. J., Grace, K., Clarke, B. E., Carroll, A. R., Rowlands, D. J., and Jackson, R.J. (1995). Unique features of internal initiation of hepatitis-C virus-RNA translation. EMBOJ. 14, 6010-6020. Roberts, L. O., and Belsham, G.J. (1997). Complementation of defective picomavirus internal ribosome entry site (IRES) elements by the co-expression of fragments of the IRES. Viwlogy 227, 53-62. Skinner, M. A., RacanieUo, V. R., Dunn, G., Cooper, J., Minor, P. D., and Almond, J. W. (1989). New model for the secondary structure of the 5' non-coding RNA of poliovirus is supported by biochemical and genetic data that also show that RNA secondary structure is important in neurovirulence. J. Mol. Biol. 207, 379-392. Stone, D. M., Almond, J. W., Brangwyn, J. K., and Belsham, G.J. (1993). trans-Complementation of cap-independent translation directed by the poliovirus 5' noncoding region deletion mutants: Evidence for R N A / R N A interactions. J. Virol. 67, 6215-6223. Svitkin, Y. V., Meerovitch, K., Lee, H. S., Dholakia, J. N., Kenan, D. J., Agol, V. I., and Sonenberg, N. (1994). Internal translation on poliovirus RNA: Further characterization of La function in poliovirus translation in vitro. J. Viwl. 68, 1544-1550. Tsukiyama-Kohara K., Ilzuka, N., Kohara, M., and Nomoto, A. (1992). Internal ribosome entry site within hepatitis C virus RNA.J. Virol. 66, 1476-1483. van der Horst, G., Christian, A., and Inoue, T. (1991). Reconstitution of a group I intron self splicing reaction with an activator RNA. Proc. Natl. Acad. Sci. U.S.A. 88, 184-188. van der Velden, A., Kaminski, A., Jackson, R. J., and Belsham, G.J. (1995). Defective point mutants of the encephalomyocarditis virus internal ribosome entry site can be complemented in trans. Virology 214, 82-90. Whitaker-Dowling, P., and Younger, J. S. (1986). Vaccinia-mediated rescue of encephalomyocardits virus from the inhibitory effects of interferon. Virology 152, 50-57.

1KNA Traffic and Localization Reported by Fluorescent Molecular Cytochemistry in Living Cells T h o r u Pederson Cell Biology Group Worcester Foundation for Biomedical Research University of Massachusetts Medical School Shrewsbury, Massachusetts 01545

Marty R. Jacobson Cell Biology Group Worcester Foundation for Biomedical Research University of Massachusetts Medical School Shrewsbury, Massachusetts 01545

I. I N T R O D U C T I O N Once released from the transcriptional machinery, the various cellular RNAs are accurately segregated to the appropriate subcellular domains where they function. Understanding this remarkable phenomenon is important for an intellectually complete and coherent picture of gene expression. Unfortunately, there are few methods that allow the direct visualization of tLNA dynamics inside living cells. The intracellular distribution of discrete 1LNA populations between the nucleus and cytoplasm was classically defined by cell fractionation experiments (e.g., Allfrey and Mirsky, 1959; Zimmerman et al., 1963; Scherrer et al., 1963; Penman et al., 1963; Penman, 1966). Combining cell fractionation with pulsechase and hybridization selection or immunoselection methods permits the analysis of specific 1LNA redistribution over time between subcellular compartments (e.g., Madore et al., 1984). However, this combination of methods potentially suffers from possible redistribution of K N A or cross-contamination of components during cell fractionation (even though these problems can be shown to be negligible in some cases; e.g., Holland et al., 1980). Currently, in situ nucleic acid hybridization is most frequently used to directly visualize the subcellular localization of specific RNAs inside cells (Pardue and Gall, 1969; Angerer and Angerer, 1981; Lawrence and Singer, 1986). Although very sensitive, in situ hybridization typically utilizes fixed cells and therefore does not allow the analysis of dynamic KNA movements in uivo.

341 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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Marty R. Jacobson and Thoru Pederson

We have developed a method for studying the dynamic movement and localization of various RNAs inside living cells, which we term fluorescent RNA cytochemistry (Wang et at., 1991; Jacobson et al., 1995, 1997a,b). In this chapter we describe this method in detail, emphasizing its unique advantages. We also provide examples of this method as an in vivo reporter of R.NA dynamics and describe how fluorescent P, NA cytochemistry can be coupled with other techniques to define the R.NA translocation/localization step in eukaryotic gene readout.

II. F L U O R E S C E N T L A B E L I N G OF R N A A. R N A SYNTHESIS Several methods have been described for coupling affinity or fluorescent groups to chemically synthesized nucleic acids (e.g., Agrawal et al., 1986; Temsamani et al., 1991; Agrawal, 1994) or in vitro transcribed P, NAs (Wells and Cantor, 1977, 1980; Langer et al., 1981; Wang et al., 1991; Ainger et al., 1993; Lucas et al., 1995). We have found particularly expedient the method described by Langer et al. (1981), in which 5-(3-aminoallyl)UTP (obtained from Sigma, St. Louis, MO) is incorporated into P, NA during in vitro transcription. A standard transcription reaction, which typically yields 10-20/xg P,.NA//xg template DNA, is carried out as follows: 1. All reagents and tubes should be RNase-free prior to beginning. Bring all reagents (except enzymes) to room temperature prior to transcription. In a microcentrifuge tube, mix the reagents in the following orderl: H20 5 X Transcription b u f f e r 2 0.1 M Dithiothreitol (DTT) R.Nasin (20-40 U / t ,I ; Promega, Madison, WI) 10 X rNTP mix (10 mM each ATP, CTP, GTP, UTP) 10 mM 7meGpppG (New England Biolabs, Beverly, MA) 1 mM 5-(3-aminoallyl)UTP

105/,1

100/,1 100/~1 20/,1 50/,1 50 ~1 50/,1

~The DNA template can be precipitated by higher concentrations of spermidine (present in the 5 • transcription buffer), resulting in a low R N A yield. Mixing the reagents in this order should alleviate this problem. 2 5• Transcription buffer: 200 mM Tris-HC1 (pH 7.5), 30 mM MgC12, 10 mM spermidine, 50 mM NaC1.

18. FluorescentRNA Cytochemistry

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Linearized plasmid DNA (1 mg/ml) R N A polymerase (T3, T7, or SP6; "-'20 U//zl)

20 p,1 5/.d Total:

500/xl

2. Incubate the reaction at 30~176 for 1-2 hr. 3 3. Add 5/.d of RQ1 DNase (1 U//.d,; Promega) and incubate at 30~176 for an additional 10 min. 4. Purify the product R N A by extracting with an equal volume of phenol-chloroform-isoamyl alcohol (25:25:1; v/v/v). 5. Ethanol precipitate the R N A by adding 0.1 volume of 3 M sodium acetate (pH 5.2) and 2-3 volumes of 95-100% ethanol. 6. Resuspend the aminoallyl-labeled R N A pellet in 155/.d of water (RNase-free). Although 5-(3-aminoallyl)UTP (AA-UTP) is efficiently incorporated into R N A by the bacteriophage T3, TT, or SP6 R N A polymerases (Langer et al., 1981; J. Wang and T. Pederson, unpublished results, 1990), we add AA-UTP to our standard in vitro transcription reaction at 1/10th the concentration of UTP, as shown above. We find that this ratio of AA-UTP/UTP in the transcription reaction results in derivatized RNAs that, when coupled with either rhodamine or fluorescein (see Section II,B), are efficiently spliced in the case of fluorescently labeled pre-mRNA (Wang et al., 1991) or accurately assembled into ribonucleoprotein particles, as observed for both fluorescently labeled RNase MRP R N A (Jacobson et al., 1995) and RNase P R N A (Jacobson et al., 1997a).

B. FLUORESCENT LABELING OF AMINOALLYI~U SUBSTITUTED R N A The aminoallyl-U substituted R N A is next conjugated with either tetramethylrhodamine-5-isothiocyanate (TRITC) or fluorescein-5-isothiocyanate (FITC) (both from Molecular Probes, Eugene, OR) essentially as described previously (Langer et al., 1981; Agrawal et al., 1986; Wang et al., 1991), with only minor modification. 1. Mix the following in a 1.5-ml microcentrifuge tube: Water (RNase-free) R N A (aminoallyl-labeled from the above transcription)

70/xl 150/xl

3If incomplete transcripts accumulate in the reaction, lowering the reaction temperature to 30~ will often increase the proportion of full-length transcript produced (Krieg and Melton, 1987).

344

Marty R. Jacobson and Thoru Pederson 1 M Sodium carbonate-bicarbonate buffer (pH 9.0) TILITC or FITC (10 mg/ml in N, N- dime thylfo rmami de)

30/xl 50/xl Total:

300/xl

2. Incubate at room temperature for 4-16 hr with continuous shaking. 3. The fluorochrome-coupled ILNA is purified by gel filtration through a 5- to 10-ml Bio-Gel P-60 (Bio-iLad, Hercules, CA) or a Sephadex G50 (Sigma) column (with comparable results). If necessary, the ILNA can be electrophoretically purified by denaturing polyacrylamide gel electrophoresis (8-10% polyacrylamide/8.3 M urea). 4. Pool the peak ILNA fractions, ethanol precipitate twice, and dissolve the ILNA pellet in sterile 5 mM Tris-acetate buffer (pH 7.0) at ---50/xg o f R N A per milliliter. 5. Centrifuge the resuspended, fluorescently labeled ILNA at 25,000 rpm in a Beckman 42.2 Ti rotor (100,000 g) for 30 min to remove any aggregated material. The supernatant is then carefully removed and can either be used immediately for microinjection or stored at - 2 0 ~ until needed. As shown in Figure 1 (panel A), fluorescently labeled tLNA synthesized as described above can be visualized directly by fluorescence following denaturing polyacrylamide gel electrophoresis. Fluorescently coupled ribonucleoside triphosphates are now commercially available (Boehringer Mannheim, Indianapolis, IN; DuPont/NEN, Boston, MA; Molecular Probes, Eugene, OIL) and can be employed for direct incorporation into ILNA during in vitro transcription, obviating a subsequent fluorochrome coupling reaction. However, these fluorescent ribonucleoside triphosphates are incorporated into tLNA by bacteriophage ILNA polymerases rather inefficiently, and we therefore continue to use the aminoallyl-UTP approach.

III. CELL C U L T U R E , M I C R O I N J E C T I O N , A N D DIGITAL IMAGE PROCESSING The considerations and requirements for culturing cells on the microscope stage, as well as the design of cell culture chambers and microscope incubators, have been discussed and described in considerable detail (McKenna and Wang, 1989). Here we will briefly cover these points as they pertain to the description of the particular microscopy workstation we use for our microinjection studies.

A. CULTURING CELLS FOR MICROINJECTION Although the requirements for maintaining different cell types in culture can vary, mammalian cell cultures are typically grown in an appropriate medium

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Figure 1 Purification of rhodamine-labeled tLNA. Human U3 small nucleolar P,.NA (lanes 1-3) and human U8 small nucleolar tLNA (lanes 4-6) were transcribed in vitro in the presence of 5-(3aminoallyl)UTP (lanes 1 and 4), labeled with rhodamine (lanes 2 and 5), and purified by fractionation over a Bio-Gel P60 column (lanes 3 and 6), as previously described (Wang et al., 1991; Jacobson et al., 1995, 1997a) and detailed in this chapter. Panel A: Unstained P,NAs, illuminated by 254-r/rn light to excite rhodamine. Panel B: Same gel as in panel A visualized by illumination with 254-r/m light after ethidium bromide staining.

(pH " 7 . 4 ) in a humidified incubation chamber at ---37~ with 5 - 1 0 % CO2. In our studies (Wang et al., 1991; Jacobson et al., 1995, 1997a; M. 1K. Jacobson and T. Pederson, unpublished, 1995), cells are cultured in F-12K m e d i u m (J1KH Bioscience, Lenexa, KS) containing 10% fetal bovine serum (J1KH Bioscience), 1 m M L-glutamine, 50 /zg/ml streptomycin, and 5 0 / z g / m l penicillin. W e find this cell culture m e d i u m particularly useful for fluorescent 1KNA cytochemistry studies because of its intrinsically low autofluorescence. Approximately 3 6 - 4 8 hr prior to microinjection, the cells are plated in a cell culture chamber consisting of a 45 X 5 0 - m m glass coverslip assembled onto a 70 X 47 X 6 - m m Plexiglas or Teflon plate with a 3 5 - m m annulus drilled through the center, as previously described (see Fig. 4 in M c K e n n a and Wang, 1989).

B. MICROSCOPE INCUBATION CHAMBER As described above, the microscope incubation chamber must be capable of maintaining a constant temperature, humidity, and CO2 level (pH) for prolonged incubation of cells on the microscope stage. Furthermore, the chamber must allow the investigator easy access for microinjection or micromanipulation while

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maintaining these constant conditions. Unfortunately, few microscope chambers capable of meeting these requirements are commercially available. The incubator we use was built in the Worcester Foundation machine shop to encase the stage and objectives of a Leica DMItLB inverted microscope (Leica, Deerfield, IL), as shown in Figure 2. Access to the inside of the chamber is provided by doors on the left and fight sides. R o u n d Plexiglas ports are used as inlets and outlet (Fig. 2; in and out, respectively) for w a r m air flow. These ports are connected to a heating chamber (Fig. 2, hc) containing an adjustable hair dryer set at 600 W (Fig. 2, hd), by ---2-inch flexible aluminum pipes (Fig. 2, hp). The air temperature is controlled by regulating the current to the hair dryer using a YSI Model 72 proportional temperature controller (Fig. 2, ysi; Yellow Springs Instruments, Yellow Springs, OH), which receives signals from a YSI Model 421 surface probe (Fig. 2, sp; Yellow Springs Instruments) attached to the underside of the microscope stage with silicone glue. Humidification is accomplished by having the air coming into the incubation chamber first pass over a water chamber built into one of the air inlet ports (Fig. 2, wc). CO2 is supplied to the chamber through tubing (Fig. 2, ci) that is held near the cell culture chamber (Fig. 2, cc). The flow is controlled by a gas flow regulator (Fig. 2, fr) that is adjusted to maintain ---5% CO2 inside the microscope chamber to sustain the p H of the cell culture medium. Using this setup, we have successfully cultured cells on the microscope stage for as long as 2 - 3 days (though such prolonged culture is not necessary for most types of experiments).

C. MICROINJECTION OF CELLS WITH FLUORESCENTLY LABELED R N A s All experiments are done with actively growing cells in subconfluent cultures. Approximately 30 rain prior to microinjection of cells, the m e d i u m is replaced

Figure 2 Photograph of microscopy workstation and on-stage cell incubation chamber, cc: cell culture chamber. Panels A and B represent two different photographic perspectives, cccd: Photometrics Series 200 cooled charge-coupled device camera; ci: CO2 inlet tubing; dfc: dual filter wheel manual controller; dfw: Metaltek dual filter wheel equipped with a Uniblitz shutter (Vincent Assocs., Rochester, NY); dr: incubation chamber door; fr: CO2 flow regulator; hc: hair dryer heat chamber; hd: hair dryer; hic: Hamamatsu C2400 controller; hlh: quartz-halogen lamp housing for epi-illumination; hp: aluminum heat pipes; iccd: Hamamatsu Model C2400 intensified charge-coupled device camera; in: incubation chamber heat/air inlet; lps: Lambda Model LLS-6018 power supply; mm: Leica mechanical micromanipulator; nh: microinjection needle holder; out: incubation chamber heat/air outlet; pit: pneumatic microinjector tubing; pk: 2PK+ generator/pneumatic regulator; sc: microscope stage x-y manual control; sfE z-axis stepping motor manual fine-focus controller; sp: YSI Model 421 surface probe; wc: water chamber/air inlet; xlh: xenon arc lamp housing for epi-illumination; ysi: YSI Model 72 proportional temperature controller; zsm: z-axis manual/computer-controlled stepping motor.

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with fresh, prewarmed medium. Microinjection needles are prepared by pulling 100 m m borosilicate capillary tubing (1.2 m m OD • 0.9 mm ID; FHC, Brunswick, ME) on a Kopf Model 720 vertical pipette puller (David Kopf Instruments, Tijunga, CA). Just prior to microinjection, the fluorescently labeled R N A (50/xg/ml) is heated at 65~176 for ---3 min and immediately drawn into a pulled Pasteur pipette by capillary action. This pulled Pasteur pipette containing the fuorescent IKNA sample is then used to back-fill the microinjection needle (see Fig. 3). The microinjection needle is assembled into the needle holder, mounted on the micromanipulator (Fig. 2, mm; Leica mechanical micromanipulator) and connected to a pneumatic regulator (Fig. 2, pk; 2PK+ generator, ALA Scientific Instruments,

Figure 3 Loading a microinjection needle (mn) using a flame-drawn/pulled Pasteur pipette (pp) containing the fluorescent RNA sample, by backfilling. Panel B is a closer view of panel A.

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349

N e w York, NY) that controls the continuous flow rate of the fluorescent R N A solution to be microinjected. With the use of a 40• N.A., Ph 2 objective, the tip of the microinjection needle is next carefully brought into the field of view and positioned so that it is immediately above or barely touching the surface of the cell to be microinjected. The cell is microinjected by gently tapping the zaxis fine control knob of the micromanipulator. Once the cell is microinjected, the needle is quickly removed from the area. If necessary, the medium in the culture chamber can be replaced after the microinjection is completed to remove any fluorescent 1KNA outside the cell. However because of the dilution effect in the surrounding medium, we rarely find this to be a problem. Considerations in microinjecting cells as described above are (1) the viscosity of the R N A solution to be microinjected, (2) the shape and size of the microinjection needle, and (3) the flow rate produced by the pneumatic controller. These factors are interrelated, and it is not uncommon that fine adjustments of one or more of these parameters must be made before microinjection is successful. Another important consideration is the cellular site of microinjection. In general, the needle used for cytoplasmic microinjections has a larger tip opening than one used for the more difficult nuclear microinjections. For our nuclear microinjections, we typically use a needle with a tip --~1.5/xm in diameter. Using this size needle and fluorescent RNAs at a concentration of--~50 /xg/ml typically requires a setting of ~ 2 0 mm Hg (---0.4 psi) on our 2PK+ pneumatic generator, resulting in ~0.01 pl of solution (typically ---0.5 fg RNA) injected per nucleus.

D. MICROSCOPY AND DIGITAL IMAGE PROCESSION Our microscopy workstation consists ofa Leica DMIR]3 inverted fluorescent microscope (see Fig. 2) equipped with a phase contrast long working distance condenser, a PL Fluotar 10• NA Ph 2 objective, an N Plan L40• / 0.55 NA Ph 2 objective, a PL Fluotar 25• NA Ph 2 oil objective, a PL Fluotar 40 • NA Ph 3 oil objective, and a Plan-Apo 100 • / 1.40 NA Ph 3 oil objective. Either a 100-W quartz-halogen lamp or a 75-W xenon arc lamp is used as the light source for epi-illumination. For live cell studies requiting low light levels the 100-W quartz-halogen lamp is controlled by a Lambda power supply, Model LLS-6018 (Lambda Electronics, New York, NY; Fig. 2, lps). Leitz filter systems 13 (FITC) and M2 (TRITC) and a Chroma 61000 Series triple band filter set ( D A P I / F I T C / T R I T C or Cy3 TM;Chroma Technology, Brattleboro, VT) with excitation filters and neutral density filters mounted in a Metaltek dual filter wheel equipped with a Uniblitz shutter (Fig. 2, dfw) are used for fluorescence observation. Fluorescence images are acquired with either a Photometrics Series 200 cooled charge-coupled device (CCCD) camera (Fig. 2, cccd; Photometrics, Tucson, AZ) or a Hamamatsu Model C2400 Intensified-CCD camera (Fig. 2, iccd;

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Marry R. Jacobson and Thoru Pederson

Hamamatsu Photonics, Hamamatsu-City, Japan), linked to a Universal Imaging workstation equipped with a MetaMorph 2.5 image processing system (Universal Imaging, West Chester, PA) for data acquisition, processing, storage, and display. Digital optical sections are obtained with the use of a computer-controlled stepping motor (Fig. 2, zsm) adjustable to a 0.1 /xm step size. Removal of out-of-focus light in optical slices is achieved with the use of a nearest-neighbor algorithm (Castleman, 1979; Agard, 1984; Shaw and Rawlings, 1991) and point spread functions obtained with our optical system. Visualization of images in threedimensional space (either x-y stereo pairs or video loops) is accomplished by using the MetaMorph image processing and analysis software mentioned above.

IV. E X A M P L E S A. PRE-M~SSENGER R N A s LOCALIZE AT SPLICING FACTOR-RICH SITES IN THE NUCLEUS In our initial application of this method, we found that when fluorescent pre-messenger RNAs (mRNAs) were microinjected into the nucleus of cultured rat embryonic kidney (NRK) cells, they became progressively concentrated in discrete loci scattered throughout the nucleoplasm, which we demonstrated to also contain pre-mRNA splicing factors (Fig. 4; Wang et al., 1991). Note in panel b of Figure 4 the application of immunocytochemistry, after initial observation of the microinjected fluorescent pre-mRNA, to demarcate intranuclear sites at which pre-messenger splicing factors are concentrated.

B. RI~ASE M R P R N A , REQUIRED FOR PRE-RIBOSOMAL R N A PROCESSING, LOCALIZES IN THE NUCLEOLUS As shown in Figure 5b, when fluorescent RNase M R P R N A is microinjected into the nucleus of N R K cells, it rapidly becomes localized in nucleoli (Jacobson et al., 1995). This is consistent with the initial characterization of this R N A (also known as 7-2 RNA) as a nucleolar species (Reddy et al., 1981). As shown in Figure 5d and f, at later times signal also appeared in cytoplasmic structures. These numerous cytoplasmic foci observed 3 hr after nucleus microinjection (Fig. 5d, f) were highly motile, as visualized by time-lapse fluorescence microscopy (M. R. Jacobson and T. Pederson, unpublished, 1993). These observations demonstrate the potential of fluorescent R N A cytochemistry in examining dynamic R N A movements inside living cells. We have also combined the fluorescent R N A cytochemistry approach with immunocytochemistry to spatially relate the initial nucleolar localization of RNase

18. Fluorescent KNA Cytochemistry

351

Figure 4 Co-localizationofnucleus-microinjected, rhodamine-labeledpre-rntkNA and endogenous pre-mtkNA splicing factors. (a) Localization of a human/3-globin pre-mRNA 30 min after nucleus microinjection. (b) Immunocytochemicallocalization of splicing factors in the same cell using the Sm rnonoclonal antibody Y12, which is specific for proteins common to several splicing-related small nuclear ribonucleoprotein particles (Lerner et al., 1981). Bar = 10/zm. (Reprinted from Figure 4 (ab) in Wang et al., 1991, by permission of the Proceedings of the National Academy of Sciences.)

M R P R N A specifically to the dense fibrinar component of the nucleolus (Fig. 5g, h; Jacobson et al., 1995). This is the region of the nucleolus known to be the site of pre-ribosomal R N A processing (for a review see Spector, 1993), a function in which RNase M R P has been implicated (Schmitt and Clayton, 1993; Chu et al., 1994; Lygerou et al., 1994). Moreover, we found that a mutant RNase M R P R N A that lacked the binding site of a known nucleolar protein, the " T o " antigen (Yuan et al., 1991), did not localize in the nucleolus following nuclear microinjection (Jacobson et al., 1995). Thus these combined results provide a consistent picture and reinforce confidence in the reliability of the fluorescent R N A cytochemistry method.

V. W H A T IS T H E B I O L O G I C A L A U T H E N T I C I T Y FLUORESCENT RNA?

OF

The presence of a rhodamine or other fluorochrome groups on an ILNA molecule at 1 out of every 2 5 - 3 3 nucleotides along the polynucleotide chain (our routine level of substitution) could theoretically influence an R.NA's overall intramolecular structure and activity. W e found that pre-mR.NA labeled with rhodamine at this level is functional in a p r e - m R N A splicing assay (Wang et al., 1991). Other fluorescent tkNAs we have prepared are functional by the criterion of

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Figure 5 Dynamic localization of rhodamine-labeled human RNase MRP RNA after nucleus microinjection. MRP RNA was transcribed in vitro, labeled with rhodamine, purified, and microinjected into the nucleus of NRK cells. Panels a-f: Localization of MRP RNA visualized in the same living cell at 4 min (b), 25 min (c), and 3 hr (d, f). Phase contrast images of the cell at 4 min and 3 hr after nuclear microinjection are shown in (a) and (e), respectively. Note that the cell is binucleate (panels a and e) and that the nuclei have reoriented during the course of observation. Panels g and h: An NRK cell, nucleus microinjected with rhodamine-labeled RNase MRP RNA, fixed ---1 min after microinjection, permeabilized with 0.8% Triton X-100, and the dense fibrillar component of the nucleolus stained with anti-fibrillarin monoclonal antibody 72B9 (Reimer et al., 1987). The antibody-antigen complexes were detected with fluoresceinconjugated anti-mouse IgG. (g) Rhodamine-labeled RNase MRP RNA; (h) endogenous fibriUarin. Bar (b, d, f, and g) = 5 /.~m. (Reprinted from Fig. 3 and panels a and b of Fig. 7 in Jacobson et al., 1995, by permission of The Rockefeller University Press.)

t h e i r efficient assembly i n t o i m m u n o l o g i c a l l y distinctive r i b o n u c l e o p r o t e i n particles r e c o g n i z e d b y specific a n t i b o d i e s (Jacobson et al., 1995, 1997a; M . R . J a c o b s o n a n d T. P e d e r s o n , u n p u b l i s h e d results, 1996). A l t h o u g h it is o b v i o u s l y essential to

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investigate this point of each new R N A being studied, our experience with several different classes of R N A s so far (pre-mRNAs, small nuclear RNAs, nucleolar 1LNAs, small cytoplasmic RNAs, etc.), indicates that the presence of a rhodamine group at every 25-33 nucleotides does not impair functional R N A processing or ribonucleoprotein assembly.

VI. T E C H N I C A L P R O B L E M S A N D THEIR RESOLUTION The following difficulties are sometimes encountered. Inefficient in vitro transcription (of course, not unique to fluorescent R N A cytochemistry) is usually attributable to a defective D N A template, a subactive R N A polymerase, nuclease contamination of a reagent or solution, or loss of the transcribed R N A during purification. Stickiness of the fluorescent R N A to the glass microinjection needles usually is due to excessive concentration of the fluorescent R N A . Pitfalls in microinjection include broken needles (when the microinjection is too vigorous or accompanied by needle movement oblique to the angle of approach) or introduction of an excess volume (we typically introduce ---0.01 pl of R N A solution into a nucleus). It is also important that cells exist in ideal culture conditions both prior to and during the microinjection and subsequent period of observation. Physiologically nonoptimal cells are less likely to withstand microinjection in our experience. Finally, there can be autofluorescence (e.g. from mitochondria) and other wellknown complications inherent in fluorescence microscopy, which have been discussed elsewhere (Wang, 1989). An important consideration in the use of fluorescent R N A cytochemistry is the ratio between the amount ofmicroinjected R N A and the existing endogenous concentration of that R N A species. In our work so far, we have introduced amounts of fluorescent R N A that correspond to a few hundred to 1000 molecules per cell, so that we are likely to be in a "tracer" situation for abundant and moderately abundant R N A s (i.e. 104-106 molecules per cell). For lower abundance R N A s (1-103 molecules per cell), the situation is more challenging since the demonstrated reliability of low-level fluorescent labeling cannot readily be reconciled with the need for a critical threshold of fluorescent groups in relation to the photon detection capability of the microscopy system. We have not systematically investigated higher levels of R N A fluorescent labeling, but we believe that at some point the sensitivity of detection would be compromised by alterations of R N A structure. The alternative is to microinject larger amounts of low-level substituted fluorescent R N A (i.e., our standard two to three fluorochromes per 100 nucleotides), but this will, at some point, begin to appreciably increase the total intracellular

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concentration of a given R N A and thus could, as a mass action effect, alter the intracellular pathways of movement and localization observed. At our present level of experience with fluorescent R N A cytochemistry, we consider it prudent to label R N A at a level of two to three fluorochrome groups per 100 nucleotides and to dedicatedly operate in the "tracer" range, i.e., the microinjected amount of R N A should approximate 1-10% of the endogenous concentration.

VII. I N T E G R A T I N G F L U O R E S C E N T R N A CYTOCHEMISTRY WITH OTHER METHODS A. MUTAGENESIS OF R N A One of the most significant attributes of fluorescent R N A cytochemistry is its ability to introduce mutant RNAs into cells and compare their dynamic behavior and localization with those of the normal R N A species. For example, we employed various mutant RNAs to establish that the "speckled" nucleoplasmic localization of p r e - m R N A was related to the presence of a functional intron in the introduced R N A (Wang et al., 1991). Mutants ofRNase M R P R N A (Jacobson et al., 1995) and RNase P R N A (Jacobson et al., 1997a) were useful in defining the sequence elements required for nucleolar localization of these RNAs. This capacity to employ mutant RNAs is clearly a key advantageous feature of the fluorescent R N A cytochemistry method.

B. IMMUNOCYTOCHEMISTRY Once the microinjected fluorescent R N A has been localized and recorded in the living cell, immunocytochemistry can be carried out with the use of standard procedures. Examples of the combined application of fluorescent R N A cytochemistry and immunocytochemistry are presented in our published studies (Wang et al., 1991;Jacobson et al., 1995, 1997a) and shown in Figures 4 and 5 of this chapter.

C. VITAL DYES SELECTIVE FOR INTRACELLULAR ORGANELLES The method of fluorescent R N A cytochemistry is ideally suited for the simultaneous labeling of specific organelles in living cells by vital dyes if the latter's fluorescence can be differentiated from that of the microinjected fluorescent RNA. Our experience so far with RNase M R P R N A labeled with rhodamine and the mitochondrion-selective dye MitoTracker-Green TM (Molecular Probes) indicates that this is an attractive general approach (M. R. Jacobson, Q. Huang,

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and T. Pederson, unpublished results, 1996). Similar applications are underway in our laboratory with respect to the cytoplasmic traffic of fluorescent 7SL RNA, a component of the signal recognition particle that directs protein secretion, in relation to the endoplasmic reticulum of living cells, as revealed by membraneselective fluorescent vital dyes (Terasaki and Reese, 1992).

D. In Situ HYBRIDIZATION Once a given fluorescent R N A has been microinjected and its localization observed in the living cell, standard in situ hybridization can be carried out (Johnson et al., 1991) to detect the RNA's endogenous counterparts. For example, the fact that in situ hybridization reveals a nucleolar localization of endogenous RNase MRP R N A (Li et al., 1994; Jacobson et al., 1997a) bolsters confidence in the similar nucleolar localization of nucleus-microinjected fluorescent RNase MRP R N A (Fig. 5).

VIII. W H A T D O E S F L U O R E S C E N T CYTOCHEMISTRY

ACTUALLY

RNA

MEASURE?

An important aspect of this method is that it is capable of reporting both dynamic and static aspects of R N A in living cells, as demonstrated in Figure 5. In much of our work to date, we have introduced a given R N A at an intracellular site that is spatially distinct from the ultimate locus of its dynamic accumulation (Wang et al., 1991; Jacobson et al., 1995, 1997a). This allows one to determine whether a given R N A is or is not capable of moving from point A to point B. For example, we found that although some of the RNase M R P R N A injected into one nucleus of a binucleate cell enters the cytoplasm, it does not move into the other nucleus (]acobson et al., 1995; see Fig. 5e, f). We have also found that RNase MRP R N A is capable of rapidly moving, within a single nucleus, from the nucleoplasm to the nucleolus. This is new and important information, reported in living cells. It is also likely that fluorescent R N A cytochemistry will be effective in revealing R N A movements in differentiated cell types, such as oligodendrocytes (Ainger et al., 1993) or neurons, or in other systems such as oocytes, early embryos, or plants (e.g., see Lucas et al., 1995). At the same time, fluorescent R N A cytochemistry reveals high-affinity intracellular binding sites at which a microinjected R N A ultimately becomes concentrated (Wang et al., 1991; Jacobson et al., 1995, 1997a). This can be thought of as a "staining" feature of the method, analogous to classical cytochemistry (which is based on the affinity of various dyes for specific chemical groups available for noncovalent conjugation in fixed cells), and is part of the reason we use the term fluorescent R N A cytochemistry. The latter feature of the

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method can be corroborated by independent methods such as in situ hybridization, adding to confidence in the fluorescence results. At the same time, the spatial pathway by which a given R N A is observed to arrive at its final intracellular destination is properly accorded considerable confidence because it is observed in living cells.

XI. NEW FRONTIER

METHODS

The emerging confidence in the reliability of microinjecting fluorescent R N A into living cells opens the door to new approaches to the molecular dynamics and mobility of R~NA in vivo. We are presently exploring the potential of fluorescence recovery after photobleaching (Cardullo et al., 1991) and other biophysical methods to measure R N A mobility inside living cells, as well as the application of nonradiative fluorescence resonance energy transfer (Cardullo et al., 1988) to assess intermolecular nucleic acid associations in vivo, as potentially important advances from the underlying platform of the fluorescent R N A cytochemistry method we have introduced. We also believe it may be possible to integrate the use of fluorescent R N A cytochemistry into mechanistic investigations of R N A movement, both in studies conducted with cellular structural frameworks in vitro and in studies in living cells.

X.

CLOSING

REMARKS

Fluorescent R N A cytochemistry has considerable potential for advancing the cell biology of gene expression. As we have described, the preparation of fluorescently labeled R N A is now quite straightforward. However, there is no escaping the fact that the method's requirement for expertise in and facilities for cell microinjection (under strict physiological conditions of temperature, CO2, and pH), video microscopy, and digital image processing cannot be promptly assembled in laboratories not already engaged in such work. We are therefore always very pleased to respond to technical inquiries or welcome visitors to our laboratory to explore the potential of fluorescent R N A cytochemistry to their particular cell types and research interests.

ACKNOWLEDGMENTS The initial fluorescent RNA cytochemistry studies in our laboratory were carried out by Jin Wang, a graduate student in our laboratory, in collaboration with Long-guang Cao, a graduate student in the laboratory of Yu-li Wang at the Worcester Foundation. We gratefullyacknowledgethe important

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collaborative contributions of Yu-li Wang in the evolution of these methods. The work presented in this chapter was supported by NIH grant GM-21595-21 to T.P. and funds awarded to the Worcester Foundation by the G. Harold and Leila Y. Mathers Foundation.

REFERENCES Agard, D. A. (1984). Optical sectioning microscopy: Cellular architecture in three dimensions. Annu. Rev. Biophys. Bioeng. 13, 191-219. Agrawal, S. (1994). Functionalization of oligonucleotides with amino groups and attachment of amino specific reporter groups. In "Methods in Molecular Biology" (S. Agrawal, ed.), Vol. 26, pp. 93-120. Humana Press, Totawa, NJ. Agrawal, S., Christodoulou, C., and Gait, M.J. (1986). Efficient methods for attaching non-radioactive labels to the 5' ends of synthetic oligodeoxyribonucleotides. Nucleic Acids Res. 14, 6227-6245. Ainger, K., Avossa, D., Morgan, F., Hill, S. J., Barry, C., Barbarese, E., and Carson, J. H. (1993). Transport and localization of exogenous myelin basic protein mRNA microinjected into oligodendrocytes. J. Cell Biol. 123, 431-441. Allfrey, V. G., and Mirsky, A. E. (1959). Contrasts in the metabolic stability of different nucleotides in the ribonucleic acids of-isolated nuclei. Proc. Natl. Acad. Sci. U.S.A. 45, 1325-1344. Angerer, L. M., and Angerer, R. C. (1981). Detection ofpolyA + RNA in sea urchin eggs and embryos by quantitative in situ hybridization. Nucleic Acids Res. 9, 2819-2840. Cardullo, R. A., Agrawal, S., Flores, C., Zamecnik, P. C., and Wolf, D. E. (1988). Detection of nucleic acid hybridization by nonradioactive fluorescence resonance energy transfer. Proc. Natl. Acad. Sci. U.S.A. 85, 8790-8794. Cardullo, R. A., Mungovan, R. M., and Wolf, D. E. (1991). Imaging membrane organization and dynamics. In "Biophysical and Biochemical Aspects of Fluorescence Spectroscopy" (T. G. Dewey, ed.), pp. 231-260. Plenum, New York. Castleman, K. R. (1979). "Digital Image Processing." Prentice-Hall, Englewood Cliffs, NJ. Chu, S., Archer, R. H., Zegel, J. M., and Lindahl, L. (1994). The RNA ofRNase MRP is required for normal processing of ribosomal RNA. Proc. Natl. Acad. Sci. U.S.A. 91, 659-663. Holland, C. A., Mayrand, S., and Pederson, T. (1980). Sequence complexity of nuclear and messenger RNA in HeLa cells. J. Mol. Biol. 138, 755-778. Jacobson, M. R., Cao, L.-G., Wang, Y.-L., and Pederson, T. (1995). Dynamic localization of RNase MRP RNA in the nucleolus observed by fluorescent RNA cytochemistry in living cells. J. Cell Biol. 131, 1649-1658. Jacobson, M. R., Cao, L.-G., Taneja, K., Singer, R. H., Wang, Y.-L., and Pederson, T. (1997a). Nuclear domains of the RNA subunit of RNase P. J. Cell Sci. 110, 829-837. Jacobson, M. R., Pederson, T., and Wang, Y.-L. (1997b). Conjugation of fluorescent probes to proteins and nucleic acids. In "Cell Biology: A Laboratory Handbook" (J. E. Celis, ed.). Academic Press, San Diego, CA (in press). Johnson, C. V., Singer, R. H., and Lawrence, J. B. (1991). Fluorescent detection of nuclear RNA and DNA: Implications for genome organization. Methods Cell Biol. 35, 73-99. Krieg, P. A., and Melton, D. A. (1987). In vitro RNA synthesis with SP6 RNA polymerase. In "Methods in Enzymology" (R. Wu, ed.), Vol. 155, pp. 397-415. Academic Press, Orlando, FL. Langer, P. R., Waldrop, A. A., and Ward, D. C. (1981). Enzymatic synthesis of biotin-labeled polynucleotides: Novel nucleic acid affinity probes. Proc. Natl. Acad. Sci. U.S.A. 78, 6633-6637. Lawrence, J. B., and Singer, R. H. (1986). Intracellular localization of messenger RNAs for cytoskeletal proteins. Cell (Cambridge, Mass.) 45, 407-415.

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Lerner, E. A., Lemer, M. R., Janeway, C. A., and Steitz, J. A. (1981). Monoclonal antibodies to nucleic acid-containing cellular constituents: Probes for molecular biology and autoimmune disease. Proc. Natl. Acad. Sci. U.S.A. 78, 2737-2748. Li, K., Smagula, C. S., Parsons, W. J., Richardson, J. A., Gonzalez, M., Hagler, H. K., and Williams, R. S. (1994). Subcellular partitioning of MRP R N A assessed by ultrastructural and biochemical analysis. J. Cell Biol. 124, 871-882. Lucas, W. J., Bouch6-Pillon, S., Jackson, D. P., Nguyen, L., Gaker, L., Ding, B., and Hake, S. (1995). Selecting trafficking of KNOTTED 1 homeodomain protein and its m R N A through plasmodesmata. Science 270, 1980-1983. Lygerou, Z., Mitchell, P., Petfalski, E., S6raphin, B., and Tollervey, D. (1994). The POP1 gene encodes a protein component common to the RNase MRP and RNase P ribonucleoproteins. Genes Dev. 8, 1423-1433. Madore, S.J., Wieben, E. D., Kunkel, G. R., and Pederson, T. (1984). Precursors of U4 small nuclear RNA. J. Cell Biol. 99, 1140-1144. McKenna, N. M., and Wang, Y.-L. (1989). Culturing cells on the microscope stage. Methods Cell Biol. 29, 195-205. Pardue, M. L., and Gall, J. G. (1969). Formation and detection of R N A - D N A hybrid molecules in cytological preparations. Pro& Natl. Acad. Sci. U.S.A. 63, 378-383. Penman, S. (1966). R N A metabolism in the HeLa cell nucleus. J. Mol. Biol. 17, 117-130. Penman, S., Scherrer, K., Becker, Y., and Darnell, J. E. (1963). Polyribosomes in normal and poliovirusinfected HeLa cells and their relationship to messenger-RNA. Pro& Natl. Acad. Sci. U.S.A. 49, 654-662. Reddy, R., Li, W.-Y., Henning, D., Choi, Y. C., Nohaga, K., and Busch, H. (1981). Characterization and subcellular localization of 7-8S RNAs of Novikoff hepatoma. J. Biol. Chem. 256, 8452-8457. Reimer, G., Pollard, K. M., Penning, C. A., Ochs, R. L., Lischwe, M. A., Busch, H., and Tan, E. M. (1987). Monoclonal autoantibody from a (New Zealand black X New Zealand white) F1 mouse and some human scleroderma sera target an Mr 34,000 nucleolar protein and the U3 R N P particle. Arthritis Rheum. 30, 793-800. Scherrer, K., Latham, H., and Darnel1, J. E. (1963). Demonstration of an unstable R N A and a precursor to ribosomal R N A in HeLa cells. Pro& Natl. Acad. Sd. U.S.A. 49, 240-248. Schmitt, M. E., and Clayton, D. A. (1993). Nuclear RNase MRP is required for correct processing of pre-5.8S rRNA in Saccharomyces cerevisiae. Mot. Celt. Biol. 13, 7935-7941. Shaw, P. J., and Rawlings, D. J. (1991). Three dimensional fluorescence microscopy. Prog. Biophys. Mol. Biol. 56, 187-213. Spector, D. L. (1993). Macromolecular domains within the cell nucleus. Annu. Rev. Cell Biol. 9, 265-315. J Temsamani, J., Agrawal, S., and Pederson, T. (1991). Biotinylated antisense methylphosphonate oligodeoxynucleotides. Inhibition of spliceosome assembly and affinity selection of U1 and U2 small nuclear RNPs. Or. Biol. Chem. 266, 468-472. Terasaki, M., and Reese, T. S. (1992). Characterization of endoplasmic reticulum by co-localization of BiP and dicarbocyanine dyes./. Cell Sci. 101, 315-322. Wang, J., Cao, L.-G., Wang, Y.-L., and Pederson, T. (1991). Localization ofpre-messenger R N A at discrete nuclear sites. Pro& Natl. Acad. Sci. U.S.A. 88, 7391-7395. Wang, Y.-L. (1989). Fluorescent analog cytochemistry: Tracing functional protein components in living cells. Methods Cell Biol. 29, 1-12. Wells, B. D., and Cantor, C. R. (1977). A strong ethidium binding site in the acceptor stem of most or all transfer RNAs. Nucleic Adds Res. 4, 1667-1680. Wells, B. D., and Cantor, C. R. (1980). Ribosome binding by tRNAs with fluorescent labeled 3' termini. Nucleic Acids Res. 8, 3229-3246.

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Yuan, Y., Tan, E., and Reddy, R. (1991). The 40-kilodalton To autoantigen associates with nucleotides 21 to 64 of human mitochondrial R N A processing/7-2 RNA in vitro. Mol. Cell. Biol. 11, 52665274. Zimmerman, E. F., Heeter, M., and Darnell, J. E. (1963). R N A synthesis in poliovirus-infected cells. Virology 19, 400-418.

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Detection of mRNA in Situ" Techniques for Studying Gene Expression in Dwsophila melanogaster Tissues A r m e n S. M a n o u k i a n Division of Cell and Molecular Biology Ontario Cancer Institute Department of Medical Biophysics University of Toronto Toronto, Ontario M5G 2M9, Canada

I. I N T R O D U C T I O N Embryogenesis in Drosophila melanogaster has been shown to be a valuable genetic system for the study of development. The end result of 23 hr ofembryogenesis is the first instar larva, an organism with visible and characteristic segments. This allows one to use mutations to alter the pattern of segmentation. Structures in the first instar larvae can be mapped back to the embryo, which has led to the documentation of an extensive fate map. Ntisslein-Volhard and Wieschaus (1980) using genetic analysis, first identified, a series of genes involved in pattern formation. They used the segmentation of the first instar larva (during embryogenesis) as a model system. Mutations that disrupted this pattern were isolated, and the genes they identified were cloned. It was found that the expression of many of these genes was restricted to specific spatial distribution patterns in the embryo. When the expression patterns of these genes in different mutant embryos were compared, it was discovered that they were affected in some mutants. What has become apparent is that these genes are organized in a temporal and spatial genetic hierarchy (reviewed in Ingham, 1988). Central to these analyses was the technique of in situ hybridization to messenger R N A (mRNA) as a means of studying gene expression. Therefore, in situ hybridization to m R N A has been a powerful tool in the examination of the spatial role of genes during Drosophila development and in the study of pattern formation. Many developmental paradigms were established with the use of this technology. For example, the homeotic gene Antennapedia is required for the proper development of the first two thoracic segments of the first instar 361 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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larva, and its transcripts are expressed in these cells during embryogenesis. The original technique relied on 35S-labeled probes and the preparation of tissue sections (Levine et al., 1983). Two major developments have led to the development of the current procedure outlined in this chapter. Use of digoxigenin (DIG)- and biotin-labeled probes for in situ hybridization has made 35S-labeled probes obsolete (Tautz and Pfeiffle, 1989). Also, the use of whole tissue for processing (wholemount procedure) has led to the preservation of cellular morphology (Tautz and Pfeiffte, 1989; Baumgartner and Noll, 1991). Both of these changes have resulted in a dramatic increase in the resolution of the in situ hybridization technology in Drosophila. This has allowed the Drosophila research community to analyze gene expression in situ at a much more refined level and has led to a great deal of interesting work. The use of DIG- and biotin-labeled probes has allowed the use of color chemical reactions to detect the mlKNA signal. This is certainly preferable to the radioactive signal, which is often too diffuse to determine cellular resolution. Antibodies to DIG and biotin allow detection of in situ signals with the use of immunohistochemical methods. The main advantage of a whole-mount in situ hybridization procedure on Drosophila embryos is the preservation of morphology. To this end, the whole-mount procedure is superior to procedures based on the sectioning of tissue. Together these two advances have allowed the unambiguous detection and analysis of mR.NA abundance at the single-cell level. By extension, they have allowed the examination of development and cell fate determination with greater resolution, allowing the accumulation of a more comprehensive fate map of developing tissues in Drosophila. In this chapter, I outline novel techniques that allow greater resolution and more sophisticated uses of the DIG-based in situ hybridization protocol first developed by Tautz and Pfeiffle (1989).

II. W H O L E - M O U N T

IN SITU

HYBRIDIZATION

TO

mRNA WITH DIG-LABELED PROBES The most important aspect of probe preparation for use in situ is the size of the final labeled DNA/1KNA. Generally, probes 200-300 bases in length are most successful in reaching their mR.NA targets. Shorter probes can contribute to background. Therefore, in all the methods of probe preparation, compromises are necessary for success. For example, some protocols utilize 1KNA probes that then have to be boiled to reduce their size. Unidirectional polymerase chain reaction (PClK)-generated probes are also treated this way. I prefer a random primer labeling method that is very easy to use and produces labeled DNA fragments 200-300 bases long that can encompass the length of the entire complementary DNA (cDNA) of concern. This procedure generates probes that produce excellent signals.

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Other labeling procedures have also been published, including an excellent one for 1KNA probe production (O'Neill and Bier, 1994).

A. PROTOCOL 1. Random-Primer DIG-Labeling Procedure 1. From 50 to 100 ng linear DNA is required (the size is not critical, as one can label fragments of up to 10 kb, although I generally choose fragments of 2-4 kb). 2. Primers used for the random primer labeling reaction are hexameric [pd(N)6], and I recommend a stock solution of 10 mg/ml. When the levels of random primer are boosted, the labeled fragments become smaller and are therefore ideal for in situ hybridization. 3. On ice, set up the following reaction: 1.0/xl DNA (50-100 ng gel-purified restriction fragment) 5.0/~1 pd(N)6 random primers at 10 mg/ml 6.0/xl H20 4. Boil the reaction mixture (12/xl) for 5 min and chill it on dry ice/ ethanol. 5. Add the following on ice: 2.0/xl of 10• Vogel buffer (475/xl 1 M 1,4piperazinebis(ethanesulfonic acid) (PIPES) (pH6.6), 25.0/xl 1 M MgC12, 3.6/xl 2-mercaptoethanol) 2.0/xl 10• dATP, dCTP, and dGTP (each at 1 mM) 1.3/xl 10• dTTP (1 mM) 0.7/xl DIG-dUTP (1 mM) (Boehringer Mannheim, Indianapolis,

IN) 1.5/xl Klenow (2 U//xl) 6. 7. 8. 9. 10.

The total reaction volume is 20.0/xl. Incubate the reaction overnight at 14~ Add 1/xl Klenow (2 U//xl) and incubate for 4 hr at room temperature. Stop the reaction by adding ethylenediaminetetraacetic acid (EDTA) (to a final concentration of 50 mM) and heating at 65~ for 5 min. Add 80/xl H20, 40 b~g glycogen, 40 b~g transfer 1KNA (tRNA), and LiC1 to a final concentration of 300 m M and 300/xl of ethanol. Precipitate DNA for 3 hr in a dry ice/ethanol bath at - 7 0 ~ or overnight at - 20~

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Armen S. Manoukian 11. Resuspend the pellet in 100/~1 hybridization buffer (50% deionized formamide, 5 • SSC, 100/zg/ml sonicated and boiled salmon sperm DNA, 100/zg/ml tRNA, 50/zg/ml heparin, and 0.1% Tween 20). 12. The probe is generally diluted 10• in hybridization buffer. Before use, boil the probe mixture for 20 min and chill it on dry ice/ ethanol.

2. Hybridization to Embryos 1. Collect the embryos in a nylon mesh and rinse with 0.5% NaC1 and 0.01% Triton • solution. 2. Dechorionate the embryos by incubating them for 2-3 min in 50% commercial bleach solution (sodium hypochlorite) and rinse copiously with water. 3. In a glass scintillation vial, prepare a biphasic solution of 5 ml fixative [5% formaldehyde in 1• phosphate buffered saline (PBS)] and 7 ml heptane. Using tweezers, transfer the nylon mesh containing the embryos to the vial. The embryos will all be transferred to the interphase. Fix the embryos by twirling the vial for 20 min. 4. To finish fixing the embryos, transfer them as a plug using a Pasteur pipette to a 1.5-ml Eppendorf tube containing 600/zl heptane and 600/zl methanol. Shake and vortex vigorously for 30 sec. At this point, all the embryos will fall to the bottom of the tube. Remove the aqueous and interphase levels and wash (while shaking) with methanol. After being washed in methanol, the embryos can be stored indefinitely at -20~ 5. To start the hybridization, rinse the embryos in 50% methanol/50% PBT (1• PBS plus 0.1% Tween 20) and then several times with PBT. 6. Fix the embryos for 20 min with 5% formaldehyde in PBT. 7. Wash the embryos 3• (5 min) with PBT. 8. Incubate the embryos with proteinase K (50/.tg/ml in PBT) for 12 min. Every batch of proteinase K will have different activity, so every batch must be tested for the best results (attaining a balance of good morphology and a clean, strong signal). 9. The proteinase reaction is stopped by incubating in 2 mg/ml glycine (in PBT) for 5 min. 10. Wash 2• with PBT (2 min). 11. Fix again for 20 min (5% formaldehyde in PBT). 12. Wash 5• (2 min) in PBT. 13. Wash in 50% PBT/50% hybridization solution and then in 100% hybridization solution. At this point, transfer embryos to a 0.5-ml Eppendorf tube (siliconized tubes are best).

19.

In Situ

365

Hybridization of n~NA

14. Prehybridize for 1-12 hr at 48~ (in 100/zl hybridization buffer) in a water bath. 15. Hybridize at 48~ overnight in a 100-/,1 volume (10/zl heatdenatured probe with 90/zl hybridization buffer). After hybridization, you can reuse solution on other embryos at least five times. Boil and chill the probe before each use. Reused probe is much better! 16. Following hybridization, wash at 48~ for 20 rain each: 400/,1 hybridization solution 400/a,1 50% hybridization solution/50% PBT 5 • with PBT (1-ml aliquots in 1.5-ml Eppendorf tubes) 17. Incubate while shaking for 2 hr at room temperature (or at 4~ overnight) in anti-DIG alkaline phosphatase (AP) antibody (Boehringer Mannheim). The antibody is used at a dilution of 1:2000. 18. Following antibody incubation, wash 5 • in PBT (for 20 min each). 19. Wash 2• in AP reaction buffer (100 m M NaC1, 50 m M MgC12, 100 m M Tris pH 9.5, 1 m M Levamisol, 0.1% Tween 20). 20. The AP reaction is developed in 1 ml AP buffer plus 4.5/zl nitro blue tetrazolium (NBT) reagent (Boehringer Mannheim) and 3.5/zl 5-bromo-4-chloro-3-indolyl phosphate (BCIP) reagent (Boehringer Mannheim). The color (deep purple) will develop within 20 min to several hours and must be monitored. The time required for development of the signal depends on the abundance of mRNA. Stop the reaction with two or three washes of PBT. 21. My favorite mountant is 70% glycerol/30% 0.1 M Tris (pH 8.5). Glycerol greatly enhances morphology, and the embryos may grow as glycerol is infused into the tissue. The reaction product is a deep purple color. Through a dehydration/rehydration series using ethanol, the color changes to dark blue (because the purplish component of the reaction product is ethanol soluble and the rest of the product is not). After the ethanol washes, ending with several washes of PBT, add 150/,1 of the mountant. The embryos immediately rise to the top. Allow the embryos to layer and settle down to the bottom of the tube for a few hours before mounting them on a slide for observation.

III. U S E S O F T H E W H O L E - M O U N T

PROCEDURE

B. EXAMINATION OF WHOLE-MOUNT EMBRYOS Using the outlined procedure, it is possible to produce tissue samples that can be examined for expression of specific genes at the cellular level. For example,

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one can easily see that the gooseberry (gsb) gene is expressed in two-cell-wide stripes in the ventral epidermis of Drosophila embryos (Fig. 1A, B). Use of the glycerolbased mountant provides great tissue morphology, allowing easy interpretation of in situ data.

B.

Q U A N T I T A T I O N OF M R N A

ABUNDANCE IN SlTU

Tissue hybridized with DIG-labeled probes can also be used for quantitative analysis of gene expression. The D I G signal can then be quantitated with the use of anti-DIG-AP antibody and the detection of AP activity colorometrically. This method utilizes a substrate that produces an ethanol-soluble reaction product, which can then be extracted and quantified. Hybridization of probe to embryos is as described in Section II. The number of embryos can be measured by allowing the embryos to settle as a specific volume in Eppendorf tubes (Manoukian and Krause, 1992). For example, 100 /zl of embryos of the same genotype gives equivalent values with this method.

C.

PROTOCOL

1. After in situ hybridization and antibody incubation (anti-DIG-AP), embryos are washed several times in PBT and then rinsed in diethanolamine buffer (10 m M diethanolamine pH 9.5, 0.5 m M MgC12). 2. Incubate the embryos (specific volume in Eppendoff tubes) with 400 p,1 of 0.5 m g / m l p-nitrophenyl phosphate (PNP) in diethanolamine buffer for exactly 1 hr at 25~ 3. Stop the reaction with E D T A to a final concentration of 100 raM.

Figure 1 (A) Stage 11 embryo hybridized with DIG-labeled gsb probe. The blue signal depicts the expression ofgsb in 14 stripes. (]3) Close-up view ofgsb stripes showing that they are two cells wide. (C) Embryos double-stained to detect engrailed protein (En) in brown and wingless transcripts (wg) in blue. En and wg are expressed in adjacent cells. (D) Close-up view of En and wg stripes showing the sharp wg stripes straddling the En-expressing cells. (E) Close-up view of EN and wg stripes focusing on the apex of cells. The wg stripes are localized to the apex of these cells. (F) Embryos double-stained to detect/3-galactosidase protein in brown and wg mlKNA in blue. ~-Galactosidase marks the wgtranscribing cells due to the wg promoter-lacZ fusion transgene present in the genome of the embryo. Therefore, the brown signal marks the wg-expressingcells, clearlydemonstrating the apical localization of wg transcripts (blue signal).

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367

4. The activity of the reaction can then be determined by measuring the A405 of the supernatants. Within the 1-hr incubation time, the activity values are linear with respect to time. The AP enzyme is extremely stable. To make sure that the experiment was a success, the embryos can be stained with NBT/BCIP to produce the insoluble reaction product and to allow visual inspection.

D. OTHER DETECTION METHODS To date, fluorescent detection methods have not been successful in detecting an in situ signal in Drosophila embryos. Such a method would be very desirable, as one could use confocal microscopy to analyze embryos. However, fluorescent signals (as generated by using anti-DIG fluorescein or rhodamine antibodies; Boehringer Mannheim) are far too weak and diffuse to be useful in this analysis. A reproducible method of amplifying these signals needs to be developed. It may also be possible to do electron microscopic in situ hybridization using anti-DIG gold-labeled antibodies; this will, of course, extend in situ methodology to the subcellular level. These improvements would be important advances in the technique.

IV. DETECTION

OF MULTIPLE

GENE

PRODUCTS

Sometimes it is necessary to monitor the expression of more than one gene product in the same tissue. One simple method involves the use of biotin-labeled probes in addition to DIG-labeled ones (O'Neill and Bier, 1994). I have developed two separate protocols for the detection of mR.NA and protein simultaneously.

A. SIMULTANEOUS D E T E C T I O N OF M R N A AND NUCLEAR PROTEINS Preparing samples for in situ hybridization requires a proteinase K incubation step. This step tends to destroy a majority of membrane-associated and cytoplasmic proteins. Nuclear proteins appear to be quite insensitive to this treatment. Therefore, it is possible to detect nuclear proteins in samples that have already been hybridized with DIG-labeled probes. 1. After developing the in situ AP signal and the dehydration series with ethanol (see the protocol in Section III), take off the 100% ethanol solution and wash several times with methanol.

368

Armen S. Manoukian 2. Quench the endogenous peroxidase activity [as the antibody signal will be detected through horseradish peroxidase (HRP) activity] by washing in 3% hydrogen peroxide in methanol for 5 min. 3. Stop the reaction with three or four washes of methanol. 4. Rehydrate the embryos using five washes of PBT, and then block the embryos in PBTM (0.5% nonfat dry milk in PBT) for 45 min. 5. Incubate with primary antibody to any nuclear protein (diluted in PBTM) for 2 hr at room temperature or overnight at 4~ 6. Wash 5 • over 1 hr with PBTM. 7. Incubate with HRP-conjugated secondary antibody (diluted in PBTM) for 1.5 hr (at this point, you can use any HRP-based system to detect your antibody signal). 8. Wash 5 X over 1 hr with PBT. 9. Detect the signal in the substrate solution [1/,g/ml diaminobenzidene (DAB), 0.3% hydrogen peroxide in PBT]. The reaction signal is brown, which complements well the blue signal from the in situ hybridization step. 10. Layer and mount in the glycerol mountant, as in the in situ hybridization protocol.

B. SIMULTANEOUS DETECTION OF M R N A AND NONNUCLEAR PROTEINS For simultaneous detection of mRNA and cytoplasmic or membrane-bound proteins, it is desirable to detect the protein before performing in situ hybridization with the proteinase K step. To avoid this problem, I perform the antibody detection of protein and develop the HRP signal prior to in situ hybridization and the development of the AP signal. The DAB signal is quite stable throughout all the manipulations of the in situ hybridization protocol even though the proteins being detected are not. 1. Fixed embryos stored in methanol are incubated in a solution of 3% hydrogen peroxide (in methanol) for 5 min and then washed 3 • with methanol. 2. Wash in 50% methanol/50% PBTH [filter-sterilized PBT (0.2/,m high-protein-binding filter) plus 100/,g/ml heparin and 100/,g/ml (nuclease-free)], tRNA, and 0.05 U/ml RNasin or RNAguard RNase inhibitors). Rehydrate completely with PBTH washes (5 X for 10 rain). 3. Block the embryos for 30 min with PBTHB [0.1% nuclease-ffee bovine serum albumin (BSA) in PBTH].

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369

4. Incubate with primary antibody (to specific protein) at room temperature for 2 hr (do not perform this step overnight). 5. Wash 5 • over 1 hr with PBTHB. 6. Incubate with secondary antibody conjugated to H1LP for 2 hr. 7. Wash 5 • over 1 hr with PBTH. 8. Develop the HILP signal with 1 /zg/ml (DAB) and 0.3% hydrogen peroxide in PBTH. This step may take a long time; allow the signal to develop where some background is observed. Most of the signal will be preserved through the in situ protocol, but some will be lost; therefore, it is prudent to develop a robust H R P (brown color) signal. 9. Proceed in the in situ hybridization protocol, start at step 6 (fixation of embryos with 5% formaldehyde in PBT), and follow the rest of the protocol.

V. S U B C E L L U L A R L O C A L I Z A T I O N

OF

wingless m R N A

To outline the uses of the protocols in this chapter, it is worthwhile to illustrate their use in studying an interesting developmental observation. The wingless (wg) gene belongs to the segment polarity class of genes in Drosophila and is required for the maintenance of all segments during embryogenesis (Baker, 1987). wg is transcribed in single-cell-wide stripes in the ventral portion of the embryo (one stripe per segment) and encodes a secreted protein that is believed to be secreted into neighboring cells to maintain the expression of the engrailed (en) gene (DiNardo et al., 1988; Martinez-Arias et al., 1988; van den Heuvel et al., 1989). en itself is transcribed in two-cell-wide stripes just adjacent to the wg-transcribing cells (Fig. 1C, D). Interestingly, wg m R N A is apically localized within these cells from the onset of its transcription (Fig. 1E, F), perhaps consistent with the notion that the protein is secreted into the extracellular matrix, which is also apically localized. During embryogenesis, wg transcripts are further localized toward the en-expressing cells (Fig. 1D), presumably to aid in the delivery of wg protein to these cells. We are thus able to use the double-staining procedure outlined in this chapter to analyze subcellular localization of gene products in Drosophila embryos. As the in situ hybridization protocol is further inproved, it should be possible to analyze the distribution of wg mlLNA at the electron microscopic level. In this case, it should be possible to determine the subcellular compartment where wg m R N A is localized. We are currently working to develop this technique. This should allow us to study wg distribution and function with more precision.

REFERENCES Baker, N. E. (1987).Molecularcloning of sequences from wingless,a segmentpolaritygene in Drosophila: The spatial distribution of a transcript in embryos. EMBOJ. 6, 1765-1773.

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Baumgartner, S., and Noll, M. (1991). Network of interactions among pair-rule genes regulating paired expression during primordial segmentation of Drosophila. Mech. Dev. 33, 1-18. DiNardo, S., Sher, E., Heemskerk-Jongens, J., Kassis, J. A., and O'Farrell, P. H. (1988). Two-tiered regulation of spatially patterned engrailed gene expression during Drosophila embryogenesis. Nature 332, 604-609. Ingham, P. (1988). The molecular genetics of embryonic pattern formation in Drosophila. Nature (London) 335, 25-34. Levine, M., Hafen, E., Garber, tk. L., and Gehring, W.J. (1983). Spatial distribution of Antennapedia transcripts during Drosophila development. EMBOJ. 2, 2037-2046. Manoukian, A. S., and Krause, H. M. (1992). Concentration-dependent activities of the even-skipped protein in Drosophila embryos. Genes Dev. 6, 1740-1751. Martinez-Arias, A., Baker, N., and Ingham, P. W. (1988). Role of segment polarity genes in the definition and maintenance of cell states in the Drosophila embryo. Development (Cambridge, UK) 103, 157-170. Ntisslein-Volhard, C., and Wieschaus, E. (1980). Mutations affecting segment number and polarity in Drosophila. Nature (London) 287, 795-801. O'Neill, J. W., and Bier, E. (1994). Double-label in situ hybridization using biotin and digoxigenintagged RNA probes. BioTechniques 17, 870-875. Tautz, D., and Pfeiffle, C. (1989). A non-radioactive in situ hybridization method for the localization of specific RNAs in Drosophila reveals translational control of the segmentation gene hunchback. Chromosoma 98, 81-85. van den Heuvel, M., Nusse, tk., Johnston, P., and Lawrence, P. (1989). Distribution of the wingless gene product in Drosophila embryos: A protein involved in cell-cell communication. Cell (Cambridge, Mass.) 59, 739-749.

Differential Display Protocol That Preferentially Identifies mRNAs of Moderate to Low Abundance in a Microscopic System Ognian C. Ikonomov*

Michele H. Jacobt

Neurobiology Group Worcester Foundation for Biomedical Research Shrewsbury, Massachusetts 01545

Neurobiology Group Worcester Foundation for Biomedical Research Shrewsbury, Massachusetts 01545

I. I N T R O D U C T I O N The reverse transcription-polymerase chain reaction (RT-PCR)-based differential display is an attractive method for identifying differences in gene expression in two distinct cell populations, such as cells exposed to a test condition compared to controls. The underlying principle of differential display is the amplification of multiple messenger RNAs (mRNAs) in each R T - P C R reaction run on R N A isolated from test and control cells and the subsequent comparison of the radiolabeled amplification products (the R N A fingerprints) by polyacrylamide gel electrophoresis to distinguish differentially appearing bands adjacent test and control lanes. This pioneering approach was introduced as an m R N A differential display by Liang and Pardee (1992) and as R N A arbitrarily primed P C R (RAP-PCR) by Welsh et al. (1992). To diminish the selectivity of priming in the P C R amplification step and simultaneously amplify multiple mRNAs, a low-stringency P C R annealing temperature (40~176 is used throughout or in the initial cycle. The high sensitivity of polyacrylamide gel electrophoresis enables separation of the multiple products on the basis of size. Differentially appearing bands are detected, isolated, cloned, sequenced to establish their identity, and further analyzed to confirm their differential expression using Northern blot analysis. * Present address: af Present address:

Department of Psychiatry,Wayne State University,Detroit, Michigan48201. Department of Neuroscience, Tufts University School of Medicine, Boston,

Massachusetts 02111. 371 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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The two original differential display methods differ mainly in priming strategy. As an antisense primer to target the 3' end of polyadenylated mRNAs, the m R N A differential display uses oligo(dT)12MN (M represents a degenerate base of A, C, or G; N represents A, C, G, or T). An arbitrary 10-mer (50% GC-rich random sequence) is used as a sense primer. In contrast, in RAP-PCR, a single arbitrary 20-mer (50% GC content) is used as both sense and antisense primer. Both approaches have been used successfully to identify differentially expressed mRNAs (see the references in Liang and Pardee, 1995). Differential display has gained widespread interest since its introduction in 1992 because of the relative speed, sensitivity, small amount of total R N A necessary, and feasibility of the approach for a single person or small laboratory (for recent reviews, see Eberwine et al., 1995; Liang and Pardee, 1995; Livesey and Hunt, 1996; McClelland et al., 1995; Wang et al., 1996). In comparison to the alternative approaches of differential hybridization and subtractive complementary DNA (cDNA) library screening (Hofbauer and Denhardt, 1993), it is less technically demanding, expensive, and time-consuming. However, the practical implementation of both mRNA differential display and RAP-PCR is accompanied by multiple problems, including (1) low reproducibility of the band pattern, (2) frequent isolation of'false-positive cDNAs, and (3) a bias toward mRNAs of high abundance (Welsh et al., 1992; Bertioli et al., 1995; Liang and Pardee, 1995; McClelland et al., 1995; Ikonomov and Jacob, 1996). Numberous modifications have been introduced to address these problems. The primers used in mRNA differential display have been extended by 10 base pairs, including a restriction site, in order to increase reproducibility, reduce false positives, and aid cloning (Liang et al., 1994; Zhao et al., 1995). The consensus in the field is that primer elongation and a balanced GC content are important improvements (Diachenco et al., 1996; Liang et al., 1994; Zhao et at., 1995). Our lab has introduced further modifications. We developed a differential display procedure with selected primers (SPR) that preferentially identifies moderate to low-abundance transcripts, provides good reproducibility of the multiple band pattern with independent R N A extractions, and is applicable to microscopic biological systems (Ikonomov and Jacob, 1996). Increasing the bias in favor of the identification of moderate- to low-abundance transcripts is important in studies of differential gene expression due to the high proportion (>90%) of such transcripts in the total cellular m R N A pool (Bertioli et at., 1995). SPR combines the primer design (50% GC content) and reaction conditions of specific R T - P C R with the sensitivity of amplification product separation on a sequencing gel developed for arbitrarily primed R T - P C R differential display. Specifically, SPR differs from m R N A differential display and R A P - P C R by the use of (1) experimentally selected primer pairs that avoid the amplification of highly abundant transcripts, (2) a higher PCR annealing temperature (50~ rather than 40~ in all cycles, (3) band selection from the sequencing gel on the basis of dependence on the two primers for amplification (in addition to a differential

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appearance in test and control tissue lanes and dependence on the presence of reverse transcriptase), (4) PCR-based tests for reproducibility, SPR amplification of independent sets of R N A extractions, and Southern blot analysis of the products with an isolated radiolabeled clone, and (5) quantitative R T - P C R , instead of Northern blot analysis, to confirm the differential expression of individual cDNAs. Although the total number of amplified products is diminished by the higher stringency conditions, SPR dramatically increases the efficiency of identifying differentially expressed transcripts of moderate to low abundance. In this chapter, we describe the methodology of SPR, including relevant technical tips from the recent literature (Fig. 1).

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

PROCEDURES

A. R N A EXTRACTION SPR works well with small amounts of total RNA, and was originally developed to identify changes in gene expression that underlie synapse formation in the embryonic chick parasympathetic ciliary ganglion, a microscopic biological system (Ikonomov and Jacob, 1996). Ciliary ganglia that have developed in the absence as compared to the presence of synaptic interactions are rapidly dissected from operated and control embryos, respectively, and immediately frozen on dry ice. Typically, four or five ganglia are pooled per tube. On embryonic day (E) 8, the resulting amount of total R N A is approximately 0.5/zg. Total R N A is extracted by either of two different guanidinium isothiocyanate protocols as modified by scaling down volumes and the addition of glycogen (1-5/xl) as cartier and used without DNase treatment (Chomczynski, 1993; Feramisco et al., 1982; Levey et al., 1995).

B. PRIMER

SEQUENCES

Primer selection is the key variable that increases the bias toward amplifying moderate- to low-abundance transcripts in SP1L. Our studies demonstrate that primers that avoid the amplification of highly abundant transcripts must be selected experimentally. This is necessary since only a six-base pair perfect match between the 3' end of the primer and the sequence is sufficient for priming, even with the greater selectivity achieved by using a high P C R annealing temperature (50~ in SPR. The large 16,775-base pair intronless sequence of the chicken mitochondrial genome greatly increases the likelihood of such matches (Desjardins and Morais, 1990). Primers with a high (50-65%) GC content at the 3' end should be avoided since they tend to amplify highly abundant ribosomal and mitochondrial transcripts.

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Extraction of total RNA from test and control tissue

RT-PCR with a selected primer pair

Polyacrylamide gel electrophoresis and band selection A

2

B

C

D

m

Cloning and sequence analysis

Reproducible detection and differential expression: Southern blotting of PCR products from novel RNA extractions

Quantitative RT-PCR for individual cDNAs

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The crucial importance of the 3' end of the primer sequence in influencing the selective amplification of m R N A s in SPR is demonstrated further by the observation that adding six to nine base pairs at the 3' end of the primer results in a different R N A fingerprint. Three distinct types of primer pairs (15-21-mers, 50% GC-rich sequences) worked successfully in SPR, including arbitrarily designed primer pairs, specific primers that target a known m R N A , and arbitrary combinations of the specific primers. Surprisingly, specific primers are effective in amplifying unexpected transcripts. Although the targeted sequence is the predominant, and often the only, amplification product visible on an agarose gel, several additional weaker bands can be detected on a polyacrylamide gel after overnight or longer exposure. Thus, specific primers already known to avoid the amplification of highly abundant sequences can be readily used in target-predicting or arbitrary combinations to identify differentially expressed m R N A s by SPR.

C. RT-PCR

AMPLIFICATION REACTION

R T - P C R methods used in previous differential display protocols were modified to increase selectivity and reproducibility while still enabling the simultaneous amplification of multiple mRNAs. The major change is the use of a higher P C R annealing temperature. Single-stranded c D N A is synthesized from total R N A equivalent to the amount present in a single E8 ciliary ganglion, 0.1 /xg in 2 4 /xl diethylpyrocarbonate (DEPC) water, by using Moloney murine leukemia virus reverse transcriptase (MMLV-RT) and an antisense primer. The 20-/xl reaction contains R N A , 1/~M 3' primer, 20 m M of each deoxyribonucleoside triphosphate (dNTPs), 1 U RNasin (RNAse inhibitor), 10 m M dithiothreitol, 200 U M M L V - R T , and 1 • R T buffer (50 m M Tris-HC1, p H 8.3; 75 m M KC1; and 3 m M MgC12). The R N A and 3' primer are heat denatured at 65~ for 5 min to eliminate secondary structure. The reaction mix is incubated at 42~ for 10 min prior to the addition of reverse transcriptase, after which incubation is continued

Figure 1 Flowchart of the RT-PCR-based differential displayprotocol with selectedprimers (SPR). An autoradiogram is depicted schematically to demonstrate the band selection approach. Lane A, control tissue; lane B, test tissue; lane C, the same as A, except that the 5' primer is omitted in the PCR step; and lane D, the same as A except that reverse transcriptase is omitted in the RT step. Band 1 is dependent on reverse transcription but appears in lane C, suggesting that it contains sequences that are amplified by the 3' primer alone. Band 2 has a differential appearance in test and control tissue lanes and is dependent on the two primers for amplification. Such bands are selected for reamplification, cloning, heterogeneity analysis, and further testing to confirm differential expression of the individual cDNAs (adapted from Fig. 1 of lkonomov and Jacob, 1996).

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at 42~ for 50 min, followed by incubation at 94~ for 5 min. At the end of the cycle, the tubes are spun briefly to collect the condensation. Single-stranded cDNAs are amplified by P C R in a programmable thermocycler. Two-microliter aliquots of the 20-/,1RT mix are used for each P C R reaction, making it possible to use up to 10 different 5' primers to amplify cDNAs from each R T reaction. The 20-/,1 P C R reaction also contains enzyme buffer, 1.875 re_A4 MgC12, 1 /xM of 5' and 3' primer, 5 0 / , M dNTPs, 1.25/xCi [oe-32p]dCTP, and 1 U Taq DNA polymerase. P C R samples are overlayed with mineral oil to prevent evaporation, heat denatured at 94~ for 5 min, and amplified for 30 cycles (denaturation at 94~ for 30 sec, primer annealing at 50~ for 30 sec, and DNA extension at 72~ for 1.5 min), followed by incubation for a final 5 rain at 72~ to complete extension. We use fewer cycles (30 instead of the 40 typically used in differential display protocols) to avoid saturation and remain in the linear phase of amplification for most transcripts, thereby increasing the detection of differentially expressed mRNAs (see also Guimaraes et al., 1995). Controls include the omission of reverse transcriptase in the R T step or cDNA template (RT mixture) in the P C R step. Recent innovations include the use of two thermostable DNA polymerases to obtain larger products, up to 2 kb (Diachenko et al., 1996; also see Haag and Raman, 1994; Hadman et al., 1995, for discussions on DNA polymerases in differential display). Additionally, other radioactive and nonradioactive labels can be used to detect P C R amplification products (Bauer et al., 1993; Ito et al., 1994; Liang and Pardee, 1995; Tokuyama and Takeda, 1995).

D . BAND SELECTION AND REAMPLIFICATION R N A fingerprints from test and control tissues are compared by separating radiolabeled R T - P C R products on a 6% sequencing gel (Liang and Pardee, 1992). The dried gel is exposed to X-ray film, and multiple bands appear after overnight exposure at room temperature. The total number of visible bands ranges from 10 to 80, with a mean of 34 + 7 (SEM) for both specific and arbitrary primer pairs (Fig. 2; Ikonomov and Jacob, 1996). The band patterns are primer sequencedependent, with two general patterns being distinguished on the autoradiograms. Specific primer pairs that target identified genes produce a predominant band of the expected size accompanied by several weaker bands. In contrast, arbitrarily designed primers and specific primers in arbitrary combinations result in multiple bands of similar strength. For each primer pair, the band pattern is reproducible with independent sets of R N A extractions, the same R N A sample in separate experiments, and ciliary ganglion R N A that is extracted by different guanidinium isothiocyanate protocols. Bands are selected on the basis of three criteria: (1) differential appearance in test and control tissue lanes, (2) dependence on both sense and antisense primers

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Figure 2 Autoradiogram demonstrating the differential appearance of bands in adjacent test and control tissue lanes after RT-PCR and polyacrylamide gel electrophoresis. RNA fingerprints of ciliary ganglia at distinct stages of normal development (A and B) and in ganglia that have developed in the absence as compared to the presence of synaptic interactions (C). Arrows indicate some of the differentially appearing bands. A and B, lane 1, E6; lane 2, E9; lane 3, El5 ciliary ganglion. C, lanes 1 and 2, normal developing ciliary ganglia; lane 3, input-deprived ganglion; lanes 4 and 5, target tissuedeprived ganglia; lane 6, both input- and target-deprived ganglion, all at Eg.

for amplification, and (3) d e p e n d e n c e on the addition o f reverse transcriptase to eliminate products amplified from g e n o m i c D N A c o n t a m i n a t i o n o f extracted R N A (Fig. 1). Selected bands are excised after careful alignment o f the autoradiogram to the gel, and D N A is recovered and reamplified with the same primers and P C R p r o t o c o l (Liang et al., 1993; also see K i m et al., 1995, for alignment tips). Bands bigger than 250 base pairs w i t h o u t m a r k e d differences in their appearance in test

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and control lanes are also sampled occasionally because of the frequent presence of multiple co-migrating species in the larger bands, which might mask differences in the levels of a subset of these products (Ikonomov and Jacob, 1996). Moreover, our studies indicate that only 5% of the products that are dependent on the two primers for amplification are highly abundant sequences. In contrast, the majority (60%) of the cDNAs that are amplified by the 3' primer alone are highly abundant mitochondrial, ribosomal, and glyceraldehyde-3-phosphate dehydrogenase transcripts (Ikonomov and Jacob, 1996). Thus, selection of products that are dependent on the two primers greatly increases the bias toward isolating transcripts of moderate to low abundance. This selection can be accomplished by including a P C R reaction in which the 5' primer is omitted. Amplification products that are identical in size and appearance to those generated in the absence as compared to the presence of 5' primer are avoided (Fig. 1). Alternatively, this band selection can be carried out by labeling the 5' primer with the use of the procedure of Hadman et al. (1995) and Tokuyama and Takeda (1995).

E. CLONING, SEQUENCING, AND TESTING FOR BAND HETEROGENEITY Reamplified P C R products are cloned with the use of the TA cloning kit, and plasmid DNA is isolated according to standard methods. Sequence analysis is carried out with a modification of the dideoxy chain-termination method of Sanger et al. (1977) (Sequenase version 2.0) with T7 promoter and M13 reverse primers. Nucleotide and predicted amino acid sequences are tested for homology to known sequences in the DNA and protein databases. Since single bands larger than 250 base pairs often contain multiple comigrating sequences, five or six plasmid DNA minipreps are routinely prepared from the cloning of each such band. After the DNA is sequenced from one miniprep, inserts of the same size from the other minipreps are tested for heterogeneity by restriction endonuclease mapping or Southern blot analysis (see Section II,F). Inserts are excised from the TA vector by E c o R 1 digestion and purified from a low-melt agarose gel. Inserts that are not cleaved by a selected enzyme, those producing unexpected fragment sizes, and those that fail to hybridize to the already identified cDNA clone after Southern blotting are subsequently sequenced. In total, 56 cDNAs have been isolated and sequenced with the use of SPR. None show significant homology to highly abundant transcripts. Twelve of the cDNAs correspond to known sequences of moderate to low abundance, whereas the remaining 44 have little homology to known sequences. Sixty-five percent of these unknown cDNAs have a single open reading frame, suggesting that the amplified fragments are frequently located in the coding region of the respective transcript.

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F. REPRODUCIBLE AMPLIFICATION AND ELIMINATION OF FALSE POSITIVES Selected sequences are analzyed for reproducible amplification and differential expression with the use of Southern blotting. Radiolabeled cDNAs are hybridized to Southern filters of R T - P C R products that are amplified from novel, independent R N A extractions with the same primer pair used to generate the cDNA clone. Basically, the P C R products are separated by agarose gel electrophoresis and transferred to a nylon membrane by alkaline blotting. The Southern filter is hybridized and washed under high stringency conditions with the use of standard procedures. For a probe, the cloned cDNAs are labeled with [c~-32p]dCTP to high specific activity by random priming and purified with the use of the Chromaspin-100 column protocol. As an internal control, R T - P C R with specific primers for cfl4-tubulin is included to establish the quality and presence of equivalent quantities of total R N A from test and control ganglia. Transcript levels of cfl4-tubulin, a neuron-specific form of fl-tubulin (Sullivan et al., 1986), are not altered by synapse deprivation (Levey et al., 1995). Sequences that have different levels in test and control ganglion lanes relative to cfl4-tubulin are subjected to further analysis. Alternative Southern blot approaches for eliminating false positives have been reported (Callard et al., 1994; Li et al., 1994; Mou et al., 1994; Mathieu-Daude et al., 1996; VoegeliLahge et al., 1996).

III. C O N F I R M A T I O N OF DIFFERENTIAL EXPRESSION Differential expression of selected cDNAs is confirmed by Northern blot analysis or quantitative R T - P C R . Northern blot analysis is typically used in most differential display protocols. Total R N A is separated on a 1% agaroseformaldehyde-containing gel (Sambrook et al., 1989) and transferred to a nylon membrane by alkaline blotting. Probes are radiolabeled as described for Southern blotting. However, Northern blot analysis does not provide the sensitivity required to detect the moderate- to low-abundance transcripts identified by SPR, with a few exceptions. The sensitivity of our Northern blots is limited by the low abundance of the transcripts, particularly at E8; by the small size of the probes (usually 200- to 350-base pair P C R products); and by the small amounts of total R N A due to the need for embryonic surgeries ( 2 - 3 / , g R N A extracted from 20-30 ganglia). Specifically, no signal is detectable with most of the isolated cDNAs even after exposure of the blots to X-ray film for 7 days with an intensifying screen at -70~ whereas the same blots hybridized to a 340-base pair 18S ribosomal probe produce

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a strong signal on autoradiograms after a 2 - h r exposure at r o o m temperature. M o r e o v e r , nicotinic acetylcholine receptor cz5 subunit m R N A , included as a control since it is k n o w n to be present at a level o f 200 transcripts copies per E8 ciliary ganglion n e u r o n (Levey et al., 1995), is not detected on these N o r t h e r n filters. These results suggest that the majority o f SPR-identified sequences are expressed at moderate to l o w abundance. T o achieve greater sensitivity, relative R T - P C R is used to confirm differential expression (Fig. 3). Primers and R T - P C R reaction conditions are optimized to amplify only the targeted sequence, as d e t e r m i n e d by agarose gel electrophoresis, restriction endonuclease mapping, and sequence analysis. T h e specificity o f R T P C R is i m p r o v e d by (1) extending the 5' primer inward or replacing it with an internal sequence for the isolated c D N A , (2) increasing the P C R annealing temperature to 63~ from 50~ and (3) using Superscript T M R N a s e H - reverse

Figure 3 Confirmation of the differential expression of a specific transcript by relative RT-PCR. Primers and RT-PCR reaction conditions are optimized to amplify a targeted sequence (A). Band intensity is reduced in gangliathat have developed in the absence of input and/or target tissue interactions relative to normal developing control ganglia. The change in transcript levels per ganglion is normalized to an internal standard, a transcript whose levels are not affected by the absence of synaptic interactions (B). The number of cycles is adjusted to ensure that both transcripts are amplified in the linear range.

20. mlLNA Detection Differential Display

381

transcriptase (BRL), enabling reverse transcription to be carried out at 50~ rather than 42 ~ Aliquots of the same ganglionic R N A sample are simultaneously amplified with the use of the optimized primers for the isolated cDNA and cfl4-tubulinspecific primers (Levey et al., 1995). Care is taken to remain in the exponential phase of amplification for each sequence. Typically, 20 cycles are used for cfl4tubulin amplification, whereas 30 cycles (sometimes with a larger amount of R T mixture) are necessary for the cDNAs of interest. For quantitation, PCtL is carried out in the presence of [o~-32p]dCTP, and the radiolabeled products are separated on a 2% agarose gel. Visible bands are dissected and transferred to a liquid scintillator, and their radioactivity is counted. Corresponding areas of gel from control lanes (samples minus reverse transcriptase) are also counted and used to subtract out background. The ratio between the amount of cDNA in the test versus the control sample is normalized to the ratio for cfl4-tubulin mILNA. In the case of transcript levels that could be measured by both relative tLT-PCR and Northern blot analysis, we found a good correlation between the results, establishing the validity of both methods for quantifying the relative change in mILNA levels.

IV.

CONCLUDING

REMARKS

SPR provides the sensitivity and selectivity required to reproducibly detect differentially expressed transcripts of moderate to low abundance in small amounts of total R N A extracted from a microscopic biological system. SPR differs from other differential display protocols in the use of (1) a higher PCtL annealing temperature, (2) 30 rather than 40 P C R cycles to remain in the linear phase of amplification for most transcripts, (3) both primer and band selection to avoid the amplification and isolation of highly abundant sequences, respectively, and (4) quantitative tLT-PCIL to confirm differential expression of specific transcripts. When all of these modifications are applied, none of the differentially expressed transcripts identified by SPR show significant homology to highly abundant sequences. In contrast, a large proportion of sequenced clones isolated by RAPP C R and mILNA differential display are characterized as highly abundant (Welsh et al., 1992; Bertioli et al., 1995; McClelland et al., 1995). Thus, SPR is a highly reproducible and simple differential expression procedure that has two key distinguishing features: a dramatic increase in the efficiency of isolating moderateto low-abundance transcripts and applicability to microscopic biological systems. Increasing the bias toward isolating moderate- to low-abundance transcripts is essential for the analysis of differential gene expression due to the high proportion (>90%) of these transcripts in the cellular mILNA pool.

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ACKNOWLEDGMENTS This research was supported by NIH Grant NS 21725 (to M.H.J).

REFERENCES Bauer, D., Muller, H., Reich, J., Rdedel, H., Ahrenkiel, V., Warthoe, P., and Strauss, M. (1993). Identification of differentially expressed mRNA species by an improved display technique (DDRTPCR). Nucleic Acids Res. 21, 4272-4280. Bertioli, D.J., Schlichter, U. H. A., Adams, M. J., Burrows, P. R., Steinbiss, H. H., and Antoniw, J. (1995). An analysis of differential display shows a strong bias towards high copy number mRNAs. Nucleic Acids Res. 23, 4520-4523. Callard, D., Lescure, B., and Mazzolini, L. (1994). A method for the elimination of false positives generated by the mRNA differential display technique. BioTechniques 16, 1096-1097. Chomczynski, P. (1993). A reagent for the single-step simultaneous isolation of RNA, DNA and proteins from cell and tissue samples. BioTechniques 15, 532-535. Desjardins, P., and Morais, R. (1990). Sequence and gene organization of the chicken mitochondrial genome: A novel gene order in higher vertebrates. J. Mol. Biol. 212, 599-634. Diachenco, L. B., Ledesma, J., Chenchik, A. A., and Siebert, P. D. (1996). Combining the technique of RNA fingerprinting and differential display to obtain differentially expressed mRNA. Biochem. Biophys. Res. Commun. 219, 824-828. Eberwine, J., Crino, P., and Dichter, M. (1995). Single-cell mRNA amplification: Implications for basic and clinical neuroscience. Neuroscientist 1, 200-211. Feramisco, J. R., Smart, J. E., Burridge, K., Helfman, D. M., and Thomas, J. P. (1982). Co-existence of vinculin-like protein of higher molecular weight in smooth muscle. J. Biol. Chem. 257, 1102411031. Guimaraes, M. J., Lee, F., Zlobnik, A., and McClanahan, T. (1995). Differential display by PCR: Novel findings and applications. Nucleic Acids Res. 23, 1832-1833. Haag, E., and Raman, V. (1994). Effects of primer choice and source of Taq polymerase on the banding patterns of differential display RT-PCR. BioTechniques 17, 226-228. Hadman, M., Adam, B. L., Wright, G. L., and Bos, T. J. (1995). Modifications to the differential display technique reduce background and increase sensitivity. Anal. Biochem. 226, 383-386. Hofbauer, R., and Denhardt, D. T. (1993). Isolation of cDNA clones of low-abundance mRNAs: Alternative strategies in differential and subtractive hybridization. Methods Mol. Genet. 2, 3-25. Ikonomov, O. C., and Jacob, M. H. (1996). Differential display protocol with selected primers that preferentially isolates rnKNAs of moderate- to low-abundance in a microscopic system. Bio Techniques 20, 1030-1042. Ito, T., K. Kito, K., Adati, N., Mitsui, Y., Hagiwara, H., and Sakai, Y. (1994). Fluorescent differential display: Arbitrarily primed R T - P C R fingerprinting on an automated sequencer. FEBS Lett. 351, 321-326. Kim, A., Roffler-Tarlov, S., and Lin, C. S. (1995). New technique for precise alignment of an RNA differential display gel with its film image. BioTechniques 19, 346. Levey, M. S., BrumweU, C. L., Dryer, S. E., and Jacob, M. H. (1995). Innervation and target tissue interactions differentially regulate acetylcholine receptor subunit mRNA levels in developing neurons in situ. Neuron 14, 153-162. Li, F., Bamathan, E. S., and Kariko, K. (1994). Rapid method for screening and cloning cDNAs generated in differential mRNA display: Application of Northern blot for affinity capturing of cDNAs. Nucleic Acids Res. 22, 1764-1765.

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Liang, P., and Pardee, A. B. (1992). Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science 257, 967-971. Liang, P., and Pardee, A. B. (1995). Recent advances in differential display. Curr. Opin. Immunol. 7, 274-280. Liang, P., Averboukh, L., and Pardee, A. B. (1993). Distribution and cloning of eukaryotic mRNAs by means of differential display:refinements and optimizations. Nucleic Acids Res. 21, 3269-3275. Liang, P., Zhu, W., Zhang, X., Guo, Z., O'Connel, R. P., Averboukh, L., Wang, F., and Pardee, A. B. (1994). Differential display using one-base anchored oligo-dT primers. Nucleic Acids Res. 22, 5763-5764. Livesey, F.J., and Hunt, S. P. (1996). Identifying changes in gene expression in the nervous system: mRNA differential display. Trends Neurosci. 19, 84-88. Mathieu-Daude, F., Cheng, R., Welsh, J., and McClelland, M. (1996). Screening of differentially amplified cDNA products from arbitrarily primed PCR fingerprints using single strand conformation polymorphism (SSCP) gels. Nucleic Acids Res. 24, 1504-1507. McClelland, M., Mathieu-Daude, F., and Welsh, J. (1995). RNA fingerprinting and differential display using arbitrarily primed PCR. Trends Genet. 11, 242-246. Mou, L., Miller, H., Li, J., Wang, E., and Chalifour, L. (1994). Improvements to the differential display method for gene analysis. Biochem. Biophys. Res. Commun. 199, 564-569. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). "Molecular Cloning: A Laboratory Manual." Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY. Sanger, F., Nicklen, S., and Coulson, A. R. (1977). DNA sequencing with chain-terminating inhibitors. Proc. Natl. Acad. Sci. U.S.A. 74, 5463-5467. Sullivan, K. F., Havercroft, J. C., Machlin, P. S., and Cleveland, D. W. (1986). Sequence and expression of the chicken/35 and/34-tubulin genes define a pair of divergent/3-tubulins with complimentary patterns of expression. Mol. Cell. Biol. 6, 4409-4418. Tokuyama Y., and Takeda, J. (1995). Use of 33p-labeled primer increases the sensitivity and the specificity ofmRNA differential display. BioTechniques 18, 424-425. Voegeli-Lange, R., Brueckert, N., Boller, T., and Wiemken, A. (1996). Rapid selection and classification of positive clones generated by mRNA differential display. Nucleic Acids Res. 24, 1385-1386. Wang, X., Ruffolo, R. R., and Feuerstein, G. Z. (1996). mRNA differential display: Application in the discovery of novel pharmacological targets. Trends Pharmacot. Sci. 17, 276-279. Welsh, J., Chada, K., Dalal, S. S., Cheng, R., Ralph, D., and McClelland, M. (1992). Arbitrarily primed PCR fingerprinting of RNA. Nucleic Acids Res. 20, 4965-4970. Zhao, S., Ooi, S. L., and Pardee, A. B. (1995). New primer strategy improves precision of differential display technique. BioTechniques 18, 842-850.

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INDEX

A

B

Aconitase, as mRNA binding protein, 238 A CT1-MATo~I mRNA, stabilization studies on, 150 A C T 1 mRNA, decay of, 112 S-Adenosylmethionine, protein interaction with, 131 S-Adenosylmethionine decarboxylase, mRNA encoding, 307 Adenoviruses, poly(A) sites in, 80 Adenovirus major late promoter (AdML), use in HIV-1 transcription reactions, 7, 9, 15 Adenovirus model substrate, splicing of, 36 AdoMet cofactor, 131 protein interactions with, 139-145 Affinity reagent, TAP,. RNA as, 13 Agarose/formaldehyde gel electrophoresis, of RNA, 312 Alfalfa mosaic virus coat protein hydroxyl radical footprinting studies on, 287-288, 293-299 use in translational repression studies, 282 Amine Coupling Kit, 141 Aminoallyl-U substituted RNA, fluorescent labeling of, 343-344 antennapedia gene, of Drosophila, 326, 361 Antibodies RNA targets of, 238, 244-245 use in R B P - R N A complex studies, 169 use in snRNP splicing studies, 40-42 Antisense oligonucleotides, depletion of snRNPs with, 39 Aphthoviruses, IRES elements in, 325-327, 328 Aprotinin, as protease inhibitor, 195 Arginine-rich motif (ARM), in RNA binding proteins, 263, 281,282 Arginine-serine-rich (RS) domains, in splicing factors and regulators, 32 ASF/SF2 protein gene encoding, 42 purification of, 47

Bacteria, plating of, for RNA binding protein studies, 105-106 Bacteriophage. See also Phage Bacteriophage promoter, in plasmid vector, 197-198 Baculovirus, VP39 overexpression by, 135 BamH1 restriction site, 118 B cells, development of, 79 Benesi-Hildebrand plots, 66 Betascope, 153 BIAcore surface plasmon resonance instrument, 140, 141 Biotin-labeled probes, 362-365 Bradford Dye Assay, 30 Branchpoint region, of pre-mRNA, 33 Brome mosaic virus RNA, methylation of, 139

C Caenorhabditis elegans, 238 histone mRNA of, 165 Cap-dependent translation, internal ribosome entry site compared to, 334-355 Cap-protein interaction, surface-plasmon resonance-based assay, 140 Carbodiimide metho-p-toluene sulfonate (CMCT), use in RNA footprinting method, 255 Carbonate alkali ladder, 253 Cardioviruses, IRES elements in, 325-327 CBP 80 (human cap-binding protein), immunodepletion protocol for, 41 CCA-adding enzymes, 130 CCMB procedure, plasmid transformation using, 271-272 CDC2 factor, sequences in, 248 cDNA clones insertion into pDISPLAY-B, 226 of RNA binding proteins, site-specific, 72-75, 99, 100, 105-108, 117-121 use in RNA binding studies, 237 385

386

Index

cDNA expression library screens, use in identification of RNA binding proteins, 72-75, 100 Cell cycle control factors, sequences in, 248 Cell-free system for splicing studies, 57 for Tat-TAR RNP studies, 4 Cellular metabolism, regulation by RNA binding proteins, 238 c-fos gene, Hel-N1 binding to, 239, 248 Chimeric gene approach method, for identification of cis-acfing determinants, 151 Chloramphenicol acetyltransferase (CAT), reporter plasmids encoding for, 328, 331 Circular dichroism, RNA-protein interaction studies using, 66 c/s-acting elements direction of 3' processing by, 79, 80, 93 effects on mRNA instability, 151, 157 Cleavage and polyadenylation specificity factor (CPSF) interaction with cleavage stimulation factor, 80, 85, 90 interaction with pre-l"nRNA, 85, 90, 96 mRNA binding of, 80, 82 Cleavage factors Im and IIm, for pre-mRNA, 80 Cleavage stimulation factor (CstF), interaction with cleavage and polyadenylation specificity factor, 80, 85, 90 Cloning, of RNA binding proteins, 246-247 c-myc gene, Hel-N1 binding to, 239, 248, 249 Common cold virus, as picornavirus, 324 Competition assays, for affinities of RNA sequences, 168-173 Complementary DNA. See cDNA Congenital health disorders, RNA binding protein involvement in, 263 Cordycepin phosphate, use in RNA 3'-end labeling, 135-137 CPSF. See Cleavage and polyadenylation specificity factor (CPSF) Cro repressor protein, protection of operator DNA regions by, 286 CstF. See Cleavage stimulation factor (CstF) 3',5'-Cytidine bisphosphate (pCp), preparation of, in Hel-N1 protein studies, 251-252 Cytidine 3',4'-[5'-3Sp]bisphosphate, preparation of, 113

Cytokine mRNA, regulated stability of, 238 Cytoplasmic $100 extracts, preparation of, 196-197

D Deletion and substitution mutagenesis, in identification of sequence requirements of RNA protein binding, 103, 104 Deletion/mutational analysis, for identification of cis-acting determinants, 151 DH12S strain, electrocompetency of, 221 Differential display protocol, for mRNA identification, 371-383 Dignam protocol, 81 Digoxigenin (DIG)-labeled probes, 362-365 Dimethyl sulfide (DMS), use in RNA footprinting method, 255 Display phage method display phage preparation for, 229-230 RNA binding protein domain determination by, 232-233 RNA target preparation for, 231-232 Western blot analysis in, 230-231,232 Dissociation constants, of RNA binding protein variants, 279-280 Double selection procedure, for RNA binding site characteritzation, 143-144 doublesex gene, 43 Drosophila antennapedia gene expression of, 361 IRES elements in, 326 ELAV, as mRNA binding protein, 239 hunchback gene of, mRNA binding protein role in translation of, 238 in situ detection of mRNA in, 361-370 splicing factors of, 42, 43 wingless mRNA of, 369 DT40, use in splicing factor studies, 42 Dyes, for fluorescent molecular cytochemistry, 354

E E complex, formation of, 69, 71 ELAV family Hel-N1 protein in, 239, 247-250 as mRNA binding proteins, 239 Electrophoretic mobility bandshift assay (EMSA), hydroxyl radical footprinting combined with, 297, 299

Index

Electrophoretic time-course assay, for poly(A) polymerase, 136 elF-2-~ factor, sequences in, 248 elF4 factors, role in mRNA translation, 324, 334 Elongation complex formation, in HIV-1 RNA transcripts, 18-21 Encephalomyocarditis virus (EMCV), IRES elements in, 325-327, 328, 334 Escherichia coli

mRNA translation in, 264 reporter plasmid for lacZ expression in, 270 RNA binding protein expression in, 269, 272-273 use in display phage studies, 219, 227, 229, 234 use in genetic method for RNA binding proteins, 263-284 strains used in, 267-268 VP39 overexpression by, 134-135 Exons, splicing together of, 31, 55

F Ferritin mRNA, translation regulation of, 238 F factor, in E. coli strain TG1,221 Filamentous bacteriophage genome, use in phage display method, 212, 216 5' end, labeling of, 251,291 5' processing, of mRNA, 79-98 Fluorescein, anti-DIG, 367 Fluorescence enhancement, calculation of, 63 Fluorescent molecular cytochemistry actual measurements in, 355-356 digital image processing in, 349-350 dyes used in, 355 integration with other methods, 354-355 microscope incubation chamber for, 345-346 potential uses of, 356-357 of RNA traffic and localization, 341-359 technical problems in, 353-354 Foot-and-mouth disease virus (FMDV) IRES elements in, 326, 334 L protein of, 327 as picornavirus, 324 Footprinting, of RNA-protein complexes with hydroxyl radicals, 285-303

G Gal4 activation domain, introduction into yeast, 183

387

fl-Galactosidase assay, 186-187, 266, 267 in translation repression studies, 273-274 Gel electrophoresis, of spliceosomes, 68, 69 Gel filtration chromatography, of spliceosomes, 68, 69 Gel mobility shift analysis problems with, 168 of protein-nucleic acid interactions, 85, 87-88, 102-103, 195, 200-202 of stem-loop binding protein binding to RNA, 167-168, 169 of 3' processing of mRNA, 80, 165 using reticulocyte lysate, 187-188 Gel purification, of end-labeled transcripts, 252, 254 GenBank database, 248 Gene III, expression of, 219 Gene(s) expression, RNA binding protein role in, 238 posttranslational regulation of, 212 Genetic methodologies application to HIV-1,281-282 for RNA binding protein studies, 212, 263-284 troubleshooting in, 280 Gene X, phage display studies of, 218, 224 9G8 (human SR protein), immunodepletion protocol for, 41 Glutathione S-transferase (Gst) Tat fusion protein of, 10, 11 use for Tat purification, 5 VP-55 fusions of. See GST-VP39 Granulocyte-macrophage colony-stimulating factor (GM-CSF), Hel-N1 binding to, 239, 248 Growth factors, sequences in, 248 GST-VP39, E. coli overexpression of, 134-135

H Hairpin binding factor removal from nuclear extract, 179-181 stem-loop binding proteins and, 174, 175 H2a-614 pre-mRNA, preparation of, 176 H complex, formation of, 69 Heat inactivation, snRNP depletion by, 39-40 HeLa cells growth of, 25 VP39 and VP55 overexpression in, 133-134

388

Index

HeLa nuclear extracts, 25-30 3' processing complex studies using, 85-90 U2AF purification from, 33 use in mRNA 3' processing studies, 80-81, 82-84 Hel-N1 protein, 239 minimal binding site of, 250-257 sequences bound by, 247-250 Helper phage, for display phage method, 228-229 Hepatitis virus(es) IRES elements in, 326, 328, 333 poly(A) sites in cores of, 80 Heterogeneous nuclear RNP. See hnRNP HIS3 mRNA, turnover studies on, 149, 153 Histone mRNA biosynthesis of, 163 regulated stability of, 238 three-dimensional structure of, 163, 164 3' end of, 163-193 Histone pre-mRNA 32p-labeled, 176-177 in vitro processing of, 174-181 Histone proteins, mRNAs encoding, 79 HIV-1 cross-linking to cleavage and polyadenylation specificity factor, 91 gene expression for, 1 long terminal repeat of, 16 poly(A) sites in, 80 pre-mRNA, 3' processing complex formation on, 85, 88, 94 promoter of, transcription activity of, 6, 7, 9, 10 Rev protein of, RNA recognition by, 264, 281-282 hnRNP alternative splice sites in, 47 pre-mRNA association with, 69 RNA binding protein preference for, 201 Human ELAV-like neuronal proteins, as m R N A binding proteins, 239 Human immunodeficiency virus type 1. See HIV-1 Humans, stem-loop binding proteins of, 188 hunchback gene, of Drosophila, m R N A binding protein role in translation of, 238 Hydroxyl radical footprinting advantages of, 287, 299 binding and cleavage reactions in, 292-293

combination with other methods, 297-299 data from, 293-299 flowchart for, 289 of RNA-protein complexes with, 285-303 RNA template preparation for, 288-292 specificity assessment in, 295-297

I IgM heavy chain, regulation of m R N A synthesis of, 79 Immunocytochemistry, use with fluorescent molecular cytochemistry, 354-355 Immunodepletion, use in snRNP splicing studies, 40-42 Immunoglobulin heavy chain binding protein, IRES elements in, 326 Immunoinhibition, use in snRNP splicing studies, 40-42 Interleukins, sequences in, 248 Internal ribosome entry site (IRES) cap-dependent translation compared to, 334-355 defective, 335-337 in picornavirus RNA, 323, 325-327 Introns excision of, 55 in pre-mRNAs, 31 IRES. See Internal ribosome entry site (IRES) Iron(II), use in production of hydroxyl radicals, 287, 299, 300 Iron-response element, as mRNA binding protein, 238 Isopropyl-/3-thiogalactopyranoside (IPTG), 272, 273 use in phage display method, 221

J

JUN leucine zipper, displayed protein attachment by, 226

K Kaleidagraph software, 66 Kanamycin resistance marker, in M13k07, 216 Kethoxal, use in RNA footprinting method, 255, 256 K-homology (KH) domain, in RNA binding proteins, 263 Kinases, sequences in, 248

Index

L lacZ gene, pDISPLAY-B derivative tagged

with, 223, 224, 233 lacZ reporter mRNA, in genetic method for

RNA binding proteins, 264, 267, 276 Lambda cDNA libraries, filter lifts from, 105-106 Lambda repressor protein, protection of operator DNA regions by, 286 Lariat configuration, of introns, 31 L40-coat yeast strain maintenance of, 185-186 use in yeast three-hybrid system, 183-188 Leucine zipper, displayed protein attachment by, 226 Leupeptin as protease inhibitor, 195, 196 use in Tat purification, 5 LexA DNA binding site, in hybrid RNA, 183 LiAc transformation method, application to two-hybrid vectors, 186 Ligase reaction mix, 114 Little Blue Tank, electroelution using, 167 Luciferase (LUC), open reading frame of, 328, 331,333 Luminescence spectrometer, 62

U Manganese ion, use in polyadenylation studies, 82, 135 M A Toe l

instability element (IE) of, 149, 153-157 mechanism studies, 157-160 mRNA of, turnover studies on, 149, 150, 152 McGhee-von Hippel noncooperative binding equation, 63 Megashortscript kit, 231 "Message Machine," 187 Metal ions, use in polyadenylation studies, 82 Metazoa, pre-mRNA splicing in, 55 2'-O-Methylation, cap-specific, of RNA, 138-139 Methyl group, of AdoMet cofactor, 132 Methyltransferase AdoMet interaction with, 142-143 VP39 regions required for activity of, 131 M F A 2 mKNA, turnover studies using, 149

389

Microinjection, cell preparation for, 344-345, 346-349 Microscope incubation chamber, for fluorescent molecular cytochemistry, 345-346 Microscopy, use in fluorescent molecular cytochemistry, 349-350 Mitotracker-green, use in fluorescent molecular cytochemistry, 355 M13k07, as phagemid, 216, 229 Mono S column chromatography, of poly(A) polymerase, 133 Mountant, for Drosophila embryos, 365-367 Mouse myeloma cells, nuclear extract preparation from, 175-176, 188-191 MPC-11 mouse myeloma cell cultures, maintenance of, 188-191 mRNA biosynthesis of, 79 differential display protocol for, 371-383 5' processing of, 79-98 functional localization in cytosol, 306 poly(A) tail of, 79 in situ detection in Drosophila, 361-370 stability of, 195 synthesis of, 31 3' processing of, 79-98 translation of onto polysomes, 305-306 RNA-protein complex role in, 286 turnover of, 149 mRNP complexes, analysis of, 195-209 MS-2 coat protein, fusion gene for, 183 Mutagenesis, oligonucleotide-directed, 275, 276 Mutations affecting pre-mRNA splicing, 57 affecting 3' processing, 112

Y Ni-NTA resin, use for protein purification, 60 0-Nitrophenylgalactosie (ONPG) (reagent), 274 oe-p-Nitrophenylglycerol (PNPG), as protease inhibitor, 195 Noncoding region (NCR), in mRNAs, 324-325 Nonsense codon mapping protocol, application to M A T o t l gene, 150-160 Northern blot hybridization of RNA, 312-315, 379 3' processing studies using, 112

390

Index

Northwestern analysis identification of cDNA clones of RNA binding proteins by, 99, 105-108 of R B P - R N A complexes, 173-174, 204, 205, 207 Nuclear magnetic resonance (NMR) spectroscopy, RNA-protein interaction studies using, 66, 286 Nuclear-to-cytoplasmic transport, in RNA-protein interactions, 195 Nucleoli pre-ribosomal RNA processing in, 350 RNA binding proteins in, 238 Nucleolin, use in translational repression studies, 282

O Oligo(A) tails, addition to RNA, 137 micro-assay, 138 Oligo(C) tails, addition to RNA, 137-138 Oligodeoxynucleotides, use in spliced-product analysis, 43-47 Oligo(dT) beads, RNA retention on, 96 Oligo(dT)-cellulose chromatography, U2AF depletion by, 33-35 Oligo(G) tails, addition to RNA, 137 Oligo(U) tails, addition to RNA, 137, 138 Oncoproteins, RNA binding proteins and, 239 Open reading frame (ORF), translation by picornavirus internal ribosome entry site, 323

P PAB1, mapping of transcripts of, 119, 120, 121 pap 1-1 gene, 119, 120 mutation of, 112 PCK1-MATa I mRNA, stabilization studies on, 150 pCp labeling method, poly(A) tail labeling by, 112 pDISPLAY-B depiction of, 217 fragment cloning into, 218 fusion gene on, 216 as vector for phage display, 216, 238 pDISPLAYblue-B, as phage display vector, 220 Pepstatin as protease inhibitor, 195 use in Tat purification, 5 pETPSF protein, purification of, 58

pGEX-2TK vector, use for Tat purification, 5 Phage, use in Northwestern screening of R N A binding proteins, 105-108 Phage display method active RNA-binding domain in, 223-224 formation of display phage in, 215-221 for RNA binding protein studies, 212 vectors used in, 216-218 Phagemid, use in phage display method, 216, 221 Phenylalanine, in RNA binding domains, as cross-linking targets, 102 Phenylmethylsulfonylfluoride (PMSF) as protease inhibitor, 195, 227 use in GST-VP39 purification, 134 use in Tat purification, 5 phoA fusion protein, formation of, 219 Phosphorlmager, 57, 58, 69, 153 Picornavirus IRES elements in, 325-327, 328, 335-338 in vivo function, 328-334 R N A translation in, 323-340 pJuFo, displayed protein attachment by, 226 Plasmid bait, construction of, 185 Plasmids, for expression of LacZ reporter, 269 Poliomyelitis virus, as picornavirus, 324 Poly(A)-binding protein gene, from yeast, insertion in pDISPLAY-B, 218 Poly(A) binding protein II, role in poly(A) tail length control, 80 Polyadenylation assay for, 123-125 electrophoretic time-course assay for, 136 phenotypes, in Saccharomyces cerevisiae, 111-125 substrates for, 95-97 Poly(A) polymerase, electrophoretic time-course assay for, 136 Poly(A) polymerase/cap-specific 2 ' - 0 methyltransferase expression and purification of, 132-135 poly(A) tail addition by, 135-136 uses of, 135-139 from vaccinia virus, 127-148 Poly(A) polymerase (PAP) role in 3' processing of mRNA, 80, 93 of vaccinia virus, 129-130 Poly(A) tails (rnRNA) labeling of, 113-117 metabolic importance of, 164

Index

o f m R N A , 79, 111 in vaccinia virus, 128, 129-131 in yeast, 118 Polyethenoadenosine as fluorescence substrate, 62 PTB binding to, 67 Polymerase chain reaction (PCR) polyadenylation site mapping by, 117-121 reverse transcription type (RT-PCR), 81, 371,375-376 use for HIV-1 long terminal repeat (LTR) preparation, 16 use in mRNA synthesis, 81 use in phage display method, 225, 226 use in polyadenylated RNA studies, 97 use in studies of ELAV proteins, 240, 241-243 Polymerase chain reaction (PCR) mutagenesis, of reporter-plasmid regions, 275, 276 Poly(N) tracts, addition to RNA 3' ends, 137-138 Polypyrimidine tract, effects on pre-mRNA splicing, 56-57 Polypyrimidine tract binding protein (PTB) binding experiments with, 63 binding sequences for, 67 combination with U2AF, 60, 62 mutant form of, 334 polyethenoadenosine binding to, 67 preparation of, 58-59 Polysomes extraction and display of, 307-312 fractionation of, 308-310 isolation of, 181-182 mRNA translation on, 305 profiles and translational control of, 305-307 Polysulfated carbohydrate heparin, as nonspecific competitor in RNA binding, 201 pp22 gene, suppression of splicing defect by, 40 PPR1 mRNA, turnover studies using, 149 Preinitiation complex formation, of HIV-1 R N A synthesis, 18 pre-mRNAs, 31 cleavage of, 82-85 fluorescent molecular cytochemistry of, 350 polyadenylation of, 82-85 poly(A) site cleavage of, 82-85 splicing complexes of, 68-72 splicing of, 31-32 alternative, 46

391

factors for, 55-78 in metazoa, 55 regulatory mechanisms, 43-49 Primer extension, use in RNA footprinting method, 256 PRIME RNase inhibitor, 199 Promega protocol, 254 Promega RNase ONE enzyme, 318 Protease inhibitors, 196, 227 Proteinase K, use in in situ hybridization, 367, 368 Proteinase K stop solution, 114 Protein III expression of, 219 in phage capsid, 215, 216 phage display studies of, 218-219, 226 Protein kinase C, sequences in, 248 Protein-ligand interactions, assays for, 139-145 Protein VIII, in phage capsid, 215, 216 Protein X, fusion to bacteriophage protein, 215, 216, 224, 230 PRP20 gene, mutations of, 112 prp-16 (yeast DEAH protein), immunodepletion protocol for, 41 PSF assay for activity of, 38 purification of, 60 PTB. See Polypyrimidine tract binding protein (PTB) pUC 119, as phagemid, 216

R Rabbit reticulocyte lysate (RRJ~) translation system, use in ribosome entry studies, 323 Regulators, functional analysis of, 31-45 Reticulocyte lysate, mobility shift assays using, 187-188 Retroviruses, biology of, 1 Reverse transcriptase (RT), 119 Reverse transcription, of RNA, 245, 256 Reverse transcription-polymerase chain reaction (RT-PCR), 371 amplification reaction of, 81,375-376 differential display based on, 371 Rev protein, in HIV-1 cells, 281 Rhinovirus, likES elements in, 325-327, 328 Rhodamine, use in fluorescent molecular cytochemistry, 352, 353, 367

392 Ribonuclease dynamic localization of, 352, 353 protection analysis of, 315-320 Ribonuclease !II enzyme, RNA substrate interaction, hydroxyl radical footprinting of, 286 Ribonucleoproteins, RNA binding proteins in, 238 Ribonucleoprotein complexes, detection of, 200-202 Ribosomes loading onto mRNA, 305-321 RNA binding proteins in, 238 RNA binding assays for, 140 extraction of, 310-312, 373 from isolated gel slices, 205-206 fluorescent labeling of, 342-344 authenticity of, 351-355 in living cells, fluorescent molecular cytochemistry of, 341-359 mutant forms of, fluorescent molecular cytochemistry of, 354 of picornavirus, 323 preparation of, 101,221-222 protein interactions with sequences in, 237-261 synthesis of, 342-343 sequential steps, 195 3'-end labeling of, 136-137 transcription from PCR product, 243-244 in vitro synthesis of, 166-167 R N A 14 and R N A 15 genes, mutations of, 112 RNA binding proteins binding of displayed, 221-224 classes of, 238-239 defective variants of, 276-278 display on phage, 214 function of, 211,238, 263 genetic studies of, 211-236 identification and cloning of, 99-109 identification by cDNA expression, 72-75 interaction with pre-mRNA, 85, 211 with new binding affinities, evolution of, 226 phage display of, 218-226 rapid genetic method for, 263 RNA complexes of, supershift studies, 169-173 RNA recognition motif in, 238-239

Index

sequences bound by, 247-250 sequence-specific, 99-109, 263 suppressor variants of, 278-279 dissociation constants, 279-280 translation repression by, 265-267 in vitro genetics applied to, 224-234 RNA footprinting, chemical modification of, 253-257 RNA ladder-binding protocol, use in Hel-N1 protein studies, 250-253 RNA methyltransferase, three-dimensional structure of, 131 RNA polymerase II transcription complex, 4 RNA polymerases SP6 and T7, pre-mRNA synthesis by, 81 RNA probe, preparation of, 197-198 RNA processing, RNA-protein complex role in, 286 RNA-protein complexes analysis of, 202-208 detection of, 200-202 excision of, 207-208 footprinting with hydroxyl radicals, 285-303 formation of, 196-200 sequence specificity, 201-202 sequential steps, 195 function of, 286 identification of, 102-103 RNA sequence requirements for, 103-108 stability of, gel mobility shift assay of, 88 RNA-protein interaction analysis of, 55-78 fluorescence spectroscopy of, 62-68 UV cross-linking studies of, 58-62 RNA recognition motif (RRM) in RNA binding proteins, 238-239, 263 in U1A spliceosomal protein, 215 RNase A, use in micro-assay of poly(A) tail length, 138 RNase H cleavage, termini mapping by, 206 RNase H mapping, in identification of sequence requirements of RNA protein binding, 103, 104 RNA template, for hydroxyl radical footprinting, 288-292 RNP-consensus motif (RNP-CS), in splicing factors and regulators, 32 R P L 2 mRNA, turnover studies using, 149 R R E targets, of Rev protein, 281 RRAVI. See RNA recognition motif (RRAVI)

Index

RS domain, in splicing factors and regulators, 32, 36 Runoff transcription, RNA preparation by, 101

S Saccharomyces cerevisiae

mRNA turnover studies on, 149-161 polyadenylation phenotypes in, 111-125 poly(A) polymerase of, 136 Schistosoma japonicum, glutathione S-transferase from, 5 SC35 (human SR protein), immunodepletion protocol for, 41 scr-kinase, sequences in, 248 SDH2 mRNA, turnover studies using, 149 SELEX method, for nucleic acid study, 213, 265 Sequencing, of RNA binding proteins, 246-247 Shine-Dalgarno (S/D) region, in mRNA, 264 SLBPs. See Stem-loop binding proteins (SLBPs) Small nuclear ribonucleoproteins. See snRNPs snRBPs, interactions with splice sites, 49 snRNPs cleavage of histone pre-mRNA directed by, 174 depletion of with antisense oligonucleotides, 39 by heat inactivation, 39-40 in vivo procedures, 42 in spliceosomes, 32 structure of, 32 Solid-phase assay, for protein-AdoMet interaction, 142-145 Sonication Buffer, 60 Southern blot method, use in differential display protocol, 378 Spin-concentrators, for proteins, 198 Splice junction oligos, use in spliced-product analysis, 43-47 Spliceosomes activity of, 55 gel-filtration analysis of, 68, 69 RNA binding proteins in, 238 structure of, 32 Splicing complementation assays, of U2AF, 35-36 Splicing factors depletion systems for, 32-42 functional analysis of, 31-53

393

level changes of, 4-48 regulatory mechanisms for. See Splicing regulators Splicing regulators assay of, 45-47 tissue- or stage-specific, 43-47 SPO I3 mRNA, turnover studies using, 149 SP6 promoter, 198 SR proteins, depletion of, 38-39 Stem-loop binding proteins (SLBPs) acting on 3' end of histone mRNA, 164, 165, 166, 169, 171 bait-encoded, 185 binding domain of, 191 cloning of, 183 gel mobility of, 187 of humans, 188 Northern blot detection of, 174 removal from nuclear extract, 174-181 in vitro translation of, 191 Stem-loop structure, of histone mRNA ends, 165, 166 Stem-loop template, use in studies of ELAV proteins, 240 STE3 mRNA, turnover studies using, 149 Stratalinker, 173 Streptavidin-coated magnetic beads, 14, 141, 180, 222 TAR binding to, 15 Streptavidin-paramagnetic particles, use in poly(A) tail micro-assay, 138 Subplasmic localization, in RNA-protein interactions, 195 Sucrose gradient centrifugation, of ribosomes, 308-310 Supershifts, of R B P - R N A complexes, 169, 172 Surface-plasmon resonance-based assay, for cap-protein interaction, 140 SV40 late virus, poly(A) sites in, 80 Sxl, as possible antagonist of U2AF, 43 T TAP. cofactor for, 4 mutated element of, 10 RNA of as affinity reagent, 13-16 use in characterization of Tat-TAR RNP, 13-16

394

Index

T A R (continued) RNA stem-loop structure of, 88 tat binding to, 1 transcriptional function of, 4 TAR loop, cellular factor binding to, 4 Tat binding to TAP., 1 cofactor for, 4 as HIV-1 viral protein, 1 human coactivator of, 13 in purification of HIV-1 cofactors, 10-13 purified samples of, 4-5 role in HIV-1 transcription, 1 model for, 3 trans-activation of, 5-10, 11, 12 solid-phase based system studies on, 16-21 Tat-TAK ribonucleoprotein (RNP) in regulation of HIV-1 gene expression, 1-23 role in HIV-1 transcription, 4 TCM1, mapping of transcripts of, 119, 120, 121 T4 DNA polymerase, inhibition of expression of, 264 TENT buffer, 227, 233 Tetrahymena thermophilia, Group I self-splicing intron ribozyme of, 337 TG1 strain, use in phage display method, 221 3' ends of histone n ~ N A metabolic importance of, 164-165 protein interaction with, 163-193 labeling of, 251-252 of RNA, poly(N) tract addition to, 137-138 3' processing complex formation in, 85-87 complex stability in, 88-90 of mRNA, 79-98 mutations affecting, 112, 121 in RNA-protein interactions, 195 3' processing factor p r e - n ~ N A interactions with, ultraviolet (UV) cross-linking studies, 90-97 sequences directing, 93-97 3' untranslated regions, in mRNA, 111 3' U T R sequences ELAV protein binding to, 239, 248 libraries of, RNA selection from, 257-259 Tobacco mosaic virus, mRNA binding protein role in translation of, 238

Tobacco streak virus coat protein, hydroxyl radical footprinting studies on, 287-288, 293-299 T3 promoter, 198 T7 promoter, 198, 240, 249, 277, 288, 323, 329 Tram-acting factors, 150 effect on histone mRNA, 150 identification of, 60 Tram-action, of RNAs with proteins, 237 Tram-activation response element. See TAlK Transcription of RNA binding proteins, 246 RNA-protein complex role in, 286 Transcript splicing, in RNA-protein interactions, 195 Transferrin mRNA, regulated stability of, 238 Translational modulation, in RNA-protein interactions, 195 Translation factors, sequences in, 248 Translation repression by heterologous R N A binding proteins, 265-267 testing for, 273 TRA protein, role in splicing, 43 TRA-2 protein, role in splicing, 43 T1 ribonuclease ladder marker, 253 T7 RNA polymerase, preparation of, 14 Tryptophan fluorescence, in studies of protein-nucleic acid interactions, 67 T7 transcription reactions, 140-141 r mRNA studies on, 379 U U2AF splicing factor, 32 alternative splicing sites for, 48 binding sequences for, 67 depletion of, by oligo(dT)-cellulose chromatography, 33-35 immunodepletion protocol for, 41 preparation of, 58-59 RS domain in, 36 splicing complementation assays of, 35-36 Sxl as possible antagonist of, 43 U1A proteins, use in translational repression studies, 282 U1-A snRNP proteins, as R N A binding proteins, 239

Index

U1A spliceosomal protein, binding studies of, 214-215 U1A strain, use in phage display method, 222, 223, 225, 233 UlhplI strain, use in phage display method, 222, 225 U1-70K snRNP proteins, as RNA binding proteins, 239 Ultraviolet (UV) cross-linking in identification of sequence-specific RNA binding proteins, 99, 102-104, 173 pre-mRNA splicing complex studies using, 68-72 RNA binding protein studies using, 57, 58-62 in studies of 3' processing of mRNA, 80 3' processing factor-pre-mRNA interactions studies by, 90-97 Untranslated regions (UTRs), in mRNA, 111 UPF1 gene product, effect on mRNA decay, 149 Up/lp, as trans-acting factor, 150 Uridylates, role in VP35-RNA interaction, 143-144 U1 RNA, stem-loop II of, 241 U2 RNA, stem-loop IV of, 241 U7 snRNPs cleavage of histone pre-mRNA directed by, 174 removal from nuclear extract, 179-181 V Vaccinia/T7 RNA polymerase expression system, characteristics and uses of, 327-328, 336 Vaccinia virus gene expression by, 127-128 mRNA cap formation in cells infected by, 128-129 poly(A) polymerase/cap-specific 2 ' - 0 methyltransferase from, 127-148 poly(A) polymerase from, 130, 136, 137

395

Virus assembly, RNA-protein complex role in, 286 VP55 enzyme, of vaccinia virus, overexpression in HeLa cells, 133-134 VP39 enzyme (vaccinia virus), 130-131, 321 assay, 139 binding assay, 140 cap-protein interaction of, 140 overexpression in HeLa cells, 133-134 structure-function relationships of, 131-132 VP55 enzyme (vaccinia virus), interaction with RNA, 140 V protein, RNA operator bound by, 265

W Watson-Crick base-pairing positions, chemical modification of, 254, 255 Western blot analysis of display phage, 230-231,232 of protein expression, 327, 331 of R B P - R N A complexes, 173-174, 203, 204, 205, 207 wingless mRNA, of Drosophila, 369 WM1 transformants, use in translation repression studies, 272, 274, 277 X X-gal (reagent), 228 X-ray crystallography, of RNA-protein interactions, 286 Y Yeast. See also Saccharomycescerevisiaen-~NA instability in, 149-161 Yeast three-hybrid system, isolation of RNA binding proteins using, 183-188

Z Zeta Probe membrane, 153 Zymolase, 121

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