The Enzymes, Vol XIV: Nucleic Acids, Part A, 3rd Edition

The Enzymes VOLUME M V NUCLEIC ACIDS Part A Third Edition CONTRIBUTORS DAVID G. BEAR THOMAS BONURA BRUCE K. DUNCAN B...

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The Enzymes VOLUME M V

NUCLEIC ACIDS Part A Third Edition

CONTRIBUTORS DAVID G. BEAR THOMAS BONURA BRUCE K. DUNCAN BRIAN ENDLICH ERROL C. FRIEDBERG MALCOLM L. GEFTER MARTIN GELLERT STANLEY HATTMAN RONALD D. KLEIN ARTHUR KORNBERG STEPHEN C. KOWALCZYKOWSKI I. ROBERT LEHMAN STUART LINN ZVI LIVNEH JACK D. LOVE KEVIN McENTEE CHARLES McHENRY STANFORD MOORE

HOWARD A. NASH BARBARA H. PHEIFFER ERIC H. RADANY ROBERT L. RATLIFF CHARLES C. RICHARDSON C. K. SINGLETON JOSEPH SPERLING BETSY M. SUTHERLAND KAREN M. TELANDER MUS KAVITCH INDER M. VERMA PETER H. VON HIPPEL JAMES C . WANG GEORGE M. WEINSTOCK BERNARD WEISS ARTHUR WEISSBACH ROBERT D. WELLS STEVEN B. ZIMMERMAN

ADVISORY BOARD MARTIN GELLERT I. ROBERT LEHMAN CHARLES C. RICHARDSON

THE ENZYMES Edited by PAUL D. BOYER Department of Chemist/ and Moleculur Biology Institute University of California Los Angeles, Culifornia

Volume XIV NUCLEIC ACIDS Part A

THIRD EDITION

198 1

ACADEMIC PRESS A Subsidiuty of Hurcourt Bruce Jovanovich, Publishers

New York London Toronto Sydney San Francisco

COPYRIGHT @ 1981, BY ACADEMIC PRESS, INC. ALL RIGHTS RESERVED. NO PART OF THIS PUBLICATION MAY BE REPRODUCED OR TRANSMITTED IN ANY FORM OR BY ANY MEANS, ELECTRONIC OR MECHANICAL, INCLUDING PHOTOCOPY, RECORDING, OR ANY INFORMATION STORAGE AND RETRIEVAL SYSTEM, WITHOUT PERMISSION IN WRITING FROM THE PUBLISHER.

ACADEMIC PRESS, INC.

111 Fifth Avenue, New York, New

York 10003

United Kingdom Edition published by ACADEMIC PRESS, INC. (LONDON) LTD. 24/28 Oval Road, London NW1

IDX

Library of Coqress Cataloging i n Publication Data Main entry under t i t l e : The Enzymes.

Includes b i b l i q r a p h i c a l references. CONTENTS: v. 1. Structure and control. --v. 2. Kinetics and mechanism.--v. 3. Hydrolysis: 14. Mxleic acid, pt. A. peptide bonds.--[etc.I--v. I. Enzymes. I. Bo er, Paul 0. ed. [DNLM:

PuU5 ~ h e l 574.19'25 ISBN 0-12-122714-6 ( V . 14)

1. Enzymes.

QP601. €523

75-117107 AACfil

PRINTED IN THE UNITED STATES OF AMERICA 81828384

9 8 7 6 5 4 3 2 1

Contents List of Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

xiii xvii

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Section 1 DNA Polymerases and Related Enzymes

1

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DNA Polymerases-A

Perspective

ARTHURKORNBERG

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I Introduction . . . . . . . . . . . . . . . . . I1 . Variable Properties of Polymerases . . . . . . 111. Problems and Prospects in Polymerase Research IV. Conclusion . . . . . . . . . . . . . . . . . .

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4 5 9 12

2 . DNA Polymerase I of Escherichia coli

I . ROBERTLEHMAN I . Purification . . . . . I1 . Physical Properties . I11. Reaction Catalyzed . IV. Biological Role . . . V. Research Applications

3

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16 16

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17 29 35

DNA Polymerase 111 Holoenzyme

CHARLES MCHENRY A N D ARTHURKORNBERG 1. Introduction . . . . . . . . . . I1 . DNA Polymerase I11 . . . . . . I11. DNA Polymerase I11 Holoenzyme 1V.Summary . . . . . . . . . . . . Note Added in Proof . . . . . .

V

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39 40 43 49 49

vi 4

CONTENTS

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1-Phage DNA Polymerase

I . ROBERTLEHMAN I. Introduction . . . . I1. T4 DNA Polymerase I11. T5 DNA Polymerase IV. T7 DNA Polymerase .

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51 52

60 62

Cellular and Viral-Induced Eukaryotic Polymerases

ARTHURWEISSBACH I . Introduction and Perspective . . . . . . . . . . . . . . . . . . . . . . I1. DNAPolymerasea . . . . . . . . . . . . . . . . . . . . . . . . . . I11. DNA Polymerase p . . . . . . . . . . . . . . . . . . . . . . . . . . IV. DNA Polymerase . . . . . . . . . . . . . . . . . . . . . . . . . . V. Herpes Simplex Virus-Induced DNA Polymerase . . . . . . . . . . . . . VI . Vaccinia Virus-Induced DNA Polymerase . . . . . . . . . . . . . . . . VII . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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6

67 69 73 76 80 83 86

Reverse Transcriptow

INDERM . VERMA I . Introduction . . . . . . . . . . . . . . . . . . I1. Purification and Properties . . . . . . . . . . . . 111 Biosynthesis of the Reverse Transcriptase . . . . IV. Mechanism of Reverse Transcription . . . . . . V. Applications to Molecular Biology . . . . . . .

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87 88 94 95 99

Terminal Deoxynucleotidyltransferase

ROBERTL . RATLIFF I . Introduction . . . . . . I1 Purification and Properties I11. The Reactions Catalyzed . IV. Practical Applications . . V Biological Role . . . . .

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105 107 109

114 118

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Section II DNA Nucleases and Related Enzymes

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Deoxyribonucleases: Survey and Perspectives

STUARTLINN I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1. Occurrence. Purification. and Molecular Properties . . . . . . . . . . .

122

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124

vii

CONTENTS 111. I V. V. VI . VII.

9

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Specificity . . . . . . . . . . . . . . . . . . . . . . . . . Assays: Designing Proper Substrates and Detection Procedures Biological Role . . . . . . . . . . . . . . . . . . . . . . . Control of Activities . . . . . . . . . . . . . . . . . . . . . Research Applications . . . . . . . . . . . . . . . . . . . .

. . . . . 124 . . . . . . 129 . . . . . 130 . . . . . 133 . . . . . 134

Type I Restriction Enzymes

BRIANENDLICH A N D STUART LINN I . Introduction . . . . . . . . . . . . . . . . . . . I1. Biological Role . . . . . . . . . . . . . . . . . . I11. Genetics . . . . . . . . . . . . . . . . . . . . . IV. Purification and Properties of Type I Enzymes . . . V. Reactions Catalyzed . . . . . . . . . . . . . . . . VI.Assays . . . . . . . . . . . . . . . . . . . . . . VII . On the Mechanisms of Cleavage-A Model Scheme VIII . Conclusions . . . . . . . . . . . . . . . . . . .

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138 142 143 144 148 150

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156

Type II Restriction Enzymes

ROBERTD . WELLS. RONALDD . KLEIN.

AND

C . K . SINGLETON

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I1 . General Properties . . . . . . . . . . . . . . . . . . . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Ionic Strength and Solvent Effects on Enzyme Specificity . . . . . . . . . V. Synthetic Oligonucleotides as Substrates . . . . . . . . . . . . . . . . . VI . Substituted DNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . VII . Inhibitor Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . VIII . Influence of Drugs and Other Ligands on Cleavage Specificities . . . . . . IX. Cleavage of Single-Stranded DNA Substrates by Certain Restriction Endonucleases . . . . . . . . . . . . . . . . . . . . . . . X . Cleavage of DNA-RNA Hybrids . . . . . . . . . . . . . . . . . . . . XI . Insolubilized Restriction Enzymes . . . . . . . . . . . . . . . . . . . XI1. Crystallization of Restriction Endonucleases . . . . . . . . . . . . . . . XI11 . Genes for Restriction Endonucleases . . . . . . . . . . . . . . . . . . XIV. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

158 159 167

172 176

177 181 183 185

187 188 188 189 191

1 1. Endonucleases Specific for Single-Stranded Polynucleotides

I . ROBERTLEHMAN I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Neurosporu c'rassa Endonuclease . . . . . . . . . . . . . . . . 111. Nuclease S1 . . . . . . . . . . . . . . . . . . . . . . . . . IV. Mung Bean Endonuclease . . . . . . . . . . . . . . . . . . . V. Research Applications of Single-Strand-Specific Endonucleases . .

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193 194 198 199 201

viii 12.

CONTENTS

Exodeoxyribonucleases of Escherichiu coli

BERNARD WEBS

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203 206

I. Introduction . . . . , . . . . , . . . . . . . . . . . . . . . . . . . 11. Biological Role . , , , . . . . , . . . . . . . . . . . . . . . . . . . 111. Purification and Biophysical Properties , , . , . . , . , . . . , . . , . IV. The Reactions Catalyzed . . . . . . , . . . . . . . . . . . . . . . . . V. Models for the Mechanism of Action of the ATP-Dependent Double-Stranded DNA Exonuclease . , , . , . . , . . , . , . . . . . . . . . , . . . . VI. Conclusion . . . . , , . . . . , . . . . . . . . . . . . . . . . . . .

234 235 238 240

I. General Properties . 11. Specific Exonucleases

13.

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recBC-like Enzymes: Exonucleuse V Deoxyribonucleases

KARENM. TELANDER MUSKAVITCH A N D STUART LINN

14.

Enzymes That Incise Damaged DNA

ERROLc. FRIEDBERG, THOMASBONURA,ERICH. JACK D. LOVE

RADANY, A N D

I. Introduction . . , . . . . . . . . . . , . . . . . . . . . . . . . . . 11. Enzymes That Attack Phosphodiester Bonds in DNA Following Hydrolysis of N-Glycosylic Bonds (AP Endonucleases) . . . . . , . , . , . . , . . 111. Enzymes that Attack Phosphodiester Bonds in Damaged DNA with Intact N-Glycosylic Bonds . . . . . , . . . . , . . . . . . . . . . . . Note Added in Proof . . . . . . , , . , , . . . . . . . . . . . . . .

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247 250

251 252 274 279

Pancreatic DNase

STANFORD MOORE I. Introduction . , . . , , . . . . . , . . . 11. Purification. . . , . . . . . . . . . . . . 111. Chemical Structure . . . . . . . . . . . . IV. Catalytic Properties , . . . . . , . . . . . V. Actin as an Inhibitor of DNase I , , . . . . VI. Research Applications . . . . , . , , . . .

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281 282 285 288 293 295

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Section 111. DNA Modification 16.

Bacteriophage T4 Polynucleotide Kinare

CHARLES C. RICHARDSON I. Introduction

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11. Isolation and Physical Properties

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ix

CONTENTS

I11 . Catalytic Properties . . . . . . . . . . . . . . . . . . . . . . . . . . 305 I V. Role of Polynucleotide Kinase in Vivo . . . . . . . . . . . . . . . . . . 312 V. Research Applications . . . . . . . . . . . . . . . . . . . . . . . . . 313

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17

Eukaryotic DNA Kinaser

STEVENB . ZIMMERMAN A N D BARBARA H . PHEIFFER I . Introduction and Perspectives . . . . . . . . . . . . . . . . . . . . . I1. Purification and Properties . . . . . . . . . . . . . . . . . . . . . . . 111. The Catalytic Reaction . . . . . . . . . . . . . . . . . . . . . . . . 1V. Comparison of the DNA Kinases with RNA Kinase and Polynucleotide Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . V. Biological Role . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Research Applications . . . . . . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . . . . . . . . . .

315 316 318 326 327 329 329

18. Type I DNA Topoisomerarer

JAMES C . WANG I. I1 . 111. IV.

Introduction . . . . . . . . . . . . . . . . . Purification and Properties . . . . . . . . . . . The Reactions Catalyzed by the Enzymes . . . Biological Roles . . . . . . . . . . . . . . . . V . Research Application . . . . . . . . . . . . .

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DNA Gyrase and Other Type II Topoisomerases

MARTINGELLERT I . Introduction and Perspectives . . . . . . . . . . . . . . . . . . . . . I1 . Definitions and General Methods . . . . . . . . . . . . . . . . . . . . I11 . DNAGyrase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . IV. Other Type I1 Topoisomerases . . . . . . . . . . . . . . . . . . . . . V. Biological Role . . . . . . . . . . . . . . . . . . . . . . . . . . . . VI . Research Applications . . . . . . . . . . . . . . . . . . . . . . . . .

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345 347 348 359 361 366

DNA Unwinding Enzymes

MALCOLM L . GEFTER I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . I1 . Purification and Properties of the rep Protein . . . . . . . . . . I11. Isolation and Characterization of Helicase I11 . . . . . . . . . IV. Mechanism of Action of rep Protein and Helicase 111 . . . . . . V. The Biological Role of Enzymes That Catalyze Unwinding of DNA

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CONTENTS

X

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21

Single-Stranded DNA Binding Proteins

STEPHENC . KOWALCZYKOWSKI. DAVID G . BEAR.A N D PETERH . VON HIPPEL I . Introduction and Overview . . . . . . . . . . . . . . . . . . . . . . . I1. Theoretical and Experimental Considerations . . . . . . . . . . . . . . 111. Protein Isolation and Purification: Procedures and Strategies . . . . . . . IV. Structure. Properties. and Nucleic Acid Binding Interactions of Several Single-Stranded DNA Binding Proteins . . . . . . . . . . . . . . . . V. DNA Binding Proteins as Research Tools . . . . . . . . . . . . . . . VI . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

22

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374

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. 388 . 441 442

The recA Enzyme of Escherichia coli and Recombination Assays

KEVINMCENTEEA N D GEORGEM. WEINSTOCK I. Introduction . . . . . . I1. Purification . . . . . . . I11. Physical Properties . . . IV. Reactions Catalyzed . . . V. Assays for Recombination VI . Biological Role . . . . . VII . Research Applications . .

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445 447 448 453 464 465 470

23. Site-Specific Recombination Protein of Phage lambda

HOWARDA . NASH I . Introduction and Perspectives I1. Purification and Properties . . 111. Reactions Involving Int . . . IV. Biological Role . . . . . . . V. Research Applications . . . .

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471 473 474 479 479

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482 485 493 510 512

Photonractivating Enzymes

BETSYM . SUTHERLAND I . Introduction . . . . . . I1 . Purification and Properties 111. The Reaction . . . . . . IV. Biological Role . . . . . V . Research Applications . .

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25 . DNA Methylation

STANLEYHATTMAN I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 517 I1 Nature of Methylated Bases and Methods of Analysis . . . . . . . . . . . 518

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xi

CONTENTS

I11 . DNA Methylases . . . . . . . IV. Distribution of Methylated Bases V. Other DNA Modifications . . . VI . Biological Roles . . . . . . . . VII . Concluding Remarks . . . . .

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521 531 536 537 547

26. DNA Base-Insertion Enzymes (Insertases)

ZVI LIVNEH A N D JOSEPH SPERLING I . Introduction . . . . . 1I.Assay . . . . . . . . 111. Purification . . . . . . IV. Properties . . . . . . V. Mechanism of Insertion VI . Biological Role . . . .

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549 551 554 555 559 560

DNA Glycorylases

BRUCE K . DUNCAN 1. I1 . 111. IV.

Introduction . . . . Physical Properties . Enzymatic Properties Physiological Role . . V. Research Applications Note Added in Proof

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565 568 569 578 584 586

Author Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

587

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622

Subjecr Inc1e.r

Contents of Orher Volumes

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

DAVID G. BEAR (373), Institute of Molecular Biology and Department of Chemistry, University of Oregon, Eugene, Oregon 97403 THOMAS BONURA (25 l), Laboratory of Experimental Oncology, Department of Pathology, Stanford University, Stanford, California 94305 BRUCE K. DUNCAN (565), Institute for Cancer Research, Fox Chase Cancer Center, Philadelphia, Pennsylvania 191 1 1 BRIAN ENDLICH (137), Department of Biochemistry, University of California, Berkeley, California 94720 ERROL C. FRIEDBERG (251), Laboratory of Experimental Oncology, Department of Pathology, Stanford University, Stanford, California 94305 MALCOLM L. GEFTER (367), Department of Biology, Massachusetts Institute of Technology, Cambridge, Massachusetts 02 139 MARTIN GELLERT ( 3 4 3 , Laboratory of Molecular Biology, National Institute of Arthritis, Metabolism and Digestive Diseases, National Institutes of Health, Bethesda, Maryland 20205 STANLEY HATTMAN (5 17), Department of Biology, University of Rochester, Rochester, New York 14627 RONALD D. KLEIN (157), Department of Biochemistry, College of Agricultural and Life Sciences, University of Wisconsin, Madison, Wisconsin 53706 ARTHUR KORNBERG (3, 39), Department of Biochemistry, Stanford University Medical School, Stanford, California 94305 STEPHEN C. KOWALCZYKOWSKI' (373), Institute of Molecular Biology and Department of Chemistry, University of Oregon, Eugene, Oregon 97403 Present address: Department of Molecular Biology, Northwestern University Medical and Dental Schools, Chicago, Illinois 60611.

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xiv

LIST OF CONTRIBUTORS

I. ROBERT LEHMAN (15, 51, 193), Department of Biochemistry, Stanford University Medical School, Stanford, California 94305 STUART LINN (121, 137, 233), Department of Biochemistry, University of California, Berkeley, California 94720 ZVI LIVNEH (549),2Department of Organic Chemistry, The Weizmann Institute of Science, Rehovot, Israel JACK D. LOVE ( Z l ) , Laboratory of Experimental Oncology, Department of Pathology, Stanford University, Stanford, California 94305 KEVIN McENTEE (445),3 Department of Biochemistry, Stanford University School of Medicine, Stanford, California 94305 CHARLES McHENRY (39), Department of Biochemistry and Molecular Biology, University of Texas Medical School, Houston, Texas 77025 STANFORD MOORE (281), The Rockefeller University, New York, New York 10021 HOWARD A. NASH (4711, Laboratory of Neurochemistry, National Institute of Mental Health, National Institutes of Health, Bethesda, Maryland 20205 BARBARA H. PHEIFFER (315), Laboratory of Molecular Biology, National Institute of Arthritis, Metabolism and Digestive Diseases, National Institutes of Health, Bethesda, Maryland 20205 ERIC H. RADANY (25 l), Laboratory of Experimental Oncology, Department of Pathology, Stanford University, Stanford, California 94305 ROBERT L. RATLIFF (105), Genetics Group, Life Sciences Division, Los Alamos Scientific Laboratory, University of California, Los Alamos, New Mexico 87545 CHARLES C. RICHARDSON (299), Department of Biological Chemistry, Harvard Medical School, Boston, Massachusetts 021 15 C. K. SINGLETON (1571, Department of Biochemistry, College of Agricultural and Life Sciences, University of Wisconsin, Madison, Wisconsin 53706

* Present address: Department of Biochemistry, Stanford University, Stanford, California 94305. Present address: Department of Biological Chemistry, UCLA Medical School, Los Angeles, California 90024.

LIST OF CONTRIBUTORS

xv

JOSEPH SPERLING (549),4 Department of Organic Chemistry, The Weizmann Institute of Science, Rehovot, Israel BETSY M. SUTHERLAND (481), Biology Department, Brookhaven National Laboratory, Upton, New York 11973 KAREN M. TELANDER MUSKAVITCH (233), Department of Biochemistry, University of California, Berkeley, California 94720 INDER M. VERMA (87), The Salk Institute, San Diego, California 92138 PETER H. VON HIPPEL (373), Institute of Molecular Biology, Department of Chemistry, University of Oregon, Eugene, Oregon 97403 JAMES C. WANG (331), Department of Biochemistry and Molecular Biology, Harvard University, Boston, Massachusetts 021 15 GEORGE M. WEINSTOCK (445),6 Department of Biochemistry, Stanford University School of Medicine, Stanford, California 94305 BERNARD WEISS (203), Department of Microbiology, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21205 ARTHUR WEISSBACH (67), Department of Cell Biology, Roche Institute of Molecular Biology, Nutley, New Jersey 071 10 ROBERT D. WELLS (157), Department of Biochemistry, College of Agricultural and Life Sciences, University of Wisconsin, Madison, Wisconsin 53706 STEVEN B. ZIMMERMAN (3 15), Laboratory of Molecular Biology, National Institute of Arthritis, Metabolism and Digestive Diseases, National Institutes of Health, Bethesda, Maryland 20205

Present address: Department of Biochemistry, Stanford University, Stanford, California 94305. Present address: The Biological Laboratories, Harvard University, Cambridge, Massachusetts 02138. ' Present address: Frederick Cancer Research Center, Frederick, Maryland 21701.


Preface This volume marks a change in organization, but not in the basic objectives, of “The Enzymes.” Until this volume, the Third Edition of “The Enzymes” has been organized as an updated companion to the First and Second Editions ; namely, a multivolume treatise that covers the present knowledge of groups of enzymes that catalyze similar reactions. But as the Third Edition progressed, as the field grew ever more extensive, and as molecular explanation of biological function continued to emerge, a different format seemed more appropriate. We felt that users of this treatise would be better served if enzymes were grouped by their biological function instead of by the nature of the reaction catalyzed. The objective for each enzyme type presented remains the same: Outstanding authorities in the field present important information about the molecular nature of enzymes and of the reactions they catalyze, together with a perspective of their biological role. The information contained thus far in this edition serves as a base for later volumes that will follow in an open-ended series. The first topic to be covered under the new format is the enzymology of nucleic acids. Volume XIV is one of two on this topic. The volumes have been planned with the help of a fine Advisory Board composed of I. Robert Lehman, Martin Gellert, and Charles C. Richardson. Volumes XIV and X V will give authoritative coverage of the enzymes that make, modify, cleave, recombine, repair, and degrade the nucleic acids. This enzymology makes possible the remarkable isolation and restructuring of DNA and RNA and allows penetrating experimentation about their biological function. Such achievements are at the core of a biological revolution. Volume XIV covers all enzymes with DNA as the primary substrate or product except DNA ligases, which will be treated with the RNA ligases xvii

xviii

PREFACE

in Volume XV. As with previous volumes, the quality of the present volume is gratifying and was achieved because nearly all of the first-choice contributors agreed to participate. Also, in keeping with a tradition already established for “The Enzymes,” the volume is timely. All manuscripts were received within a period of several months. Volume XIV was ready for distribution in less than a year from the time the first manuscript was received. The splendid cooperation of the authors and the publisher made this possible. Preparation of the volume was greatly facilitated by the capable participation of Lyda Boyer as Assistant Editor. Also, I take pleasure in acknowledging the warm and professional interest and cooperation of the staff of Academic Press. Paul D. Boyer

Section I

DNA Pulymerases and Related Enzymes


DNA Polymerases . A Perspective ARTHUR KORNBERG

I . Introduction . . . . . . . . . . . . . . . . . . . A . Invariant Rules of Polymerase Action . . . . . . I1. Variable Properties of Polymerases . . . . . . . . . A . Organization and Size . . . . . . . . . . . . . B . Template Preference . . . . . . . . . . . . . . C . Primer Preference . . . . . . . . . . . . . . . D . Tolerance for Substitution of a Nucleotide Analog . E . Fidelity . . . . . . . . . . . . . . . . . . . . F. Processivity . . . . . . . . . . . . . . . . . . G . Catalytic Efficiency . . . . . . . . . . . . . . . H . Nick Translation and Strand Displacement . . . . 1. Multiplicity and Abundance . . . . . . . . . . . J . Physiological Functions . . . . . . . . . . . . . K . Optima! Conditions . . . . . . . . . . . . . . . 111. Problems and Prospects in Polymerase Research . . . A . Isolation of a Scarce Enzyme . . . . . . . . . . B . Physical and Functional Properties . . . . . . . . C . The Template-Primer for Assay . . . . . . . . . D . Linkage to Related Replication Proteins . . . . . E . Biosynthesis and Regulation . . . . . . . . . . . IV. Conclusion . . . . . . . . . . . . . . . . . . . .

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Copyright 0 1981 by Academic Press InC . All rights of reproduction in any form reserved ISBN 0-12-122714-6

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1. Introduction

DNA polymerases are found in nature wherever DNA is made (1-3). Assembly of a DNA chain by these deoxyribonucleotidyltransferases invariably follows certain rules, whether synthesis is extensive as in replication of a chromosome or brief as in the repair of a gap in the chain. A. INVARIANT RULESO F POLYMERASE ACTION(4) 1. The Primer

A nucleotide is added to the 3’-hydroxyl moiety of the terminal nucleotide of a preexisting chain, the primer. DNA polymerases, unlike RNA polymerases ( 4 7 , 5 ) cannot start a chain.

2. The Template The nucleotide added is selected in response existing DNA chain, the template, to which the base-paired double helix, the template-primer . tidyltransferase (6 ), which assembles a random direction, is not a polymerase by definition.

to directions by a preprimer is annealed in a Terminal deoxynucleochain without template

3 . The Nucleotide Substrate

The nucleotide added to the template-primer is a 2’-deoxyribonucleoside-5’-triphosphate chelated with a divalent metal, generally Mg’+. 4. Base Pairing

Selection of the nucleotide, dictated by the template chain, follows the Watson and Crick rules of pairing adenine with thymine, guanine with cytosine. Generally there is no recognition of a specific sequence of bases in the template. An animal cell polymerase copies bacterial DNA and the reverse is equally true. 1. T. Kornberg and A. Kornberg, “The Enzymes,” Vol. X, p. 119, 1974. 2. L. A. Loeb, “The Enzymes,” Vol. X, p. 174, 1974. 3. A. Kornberg, “DNA Replication.” Freeman, San Francisco, 1980. 4. Polymerase in this chapter will refer only to DNA polymerase although the term applies as well to RNA polymerases, the ribonucleotidyltransferases that assemble RNA chains by DNA template direction. 4a. P. Chambon, ”The Enzymes,” Vol. X, p. 261, 1974. 5 . M. J. Chamberlin, “The Enzymes,” Vol. X, p. 333, 1974. 6. F. J. Bollum, “The Enzymes,” Vol. X, p. 145, 1974.

I.

DNA POLYMERASESA PERSPECTIVE

5

5. Mechanism Nucleophilic attack by the 3'-hydroxyl group of the primer terminus (7), on the innermost (or a ) phosphorus of the nucleotide selected for addition, produces the 3'-5' phosphodiester bond of the DNA backbone and eliminates inorganic pyrophosphate.

6 . Polarity of Chain Growth Because the mechanism entails a nucleotidyl transfer to a 3'-hydroxyl primer terminus, DNA chain growth is in the 5' + 3' direction. The polarity of the newly synthesized chain is opposite to that of the template; the growing chain and template are antiparallel. 7. Reversal of the Reaction Pyrophosphorolysis of a DNA chain, and exchange between inorganic pyrophosphate and the p ,y group of a deoxynucleoside triphosphate during a synthetic step, are reversals of polymerization; as such, these reactions require a primer terminus base-paired with a template. An exchange rate more rapid than pyrophosphorolysis may be due to attack b y inorganic pyrophosphate at an enzyme site more favorable than that occupied by the primer terminus in pyrophosphorolysis. II. Variable Properties of Polymerases

DNA polymerases, isolated from a single cell or from different cells and organisms, vary widely in certain properties. A.

ORGANIZATION AND SIZE

They vary from a single polypeptide of 40,000 daltons (mammalian /3 polymerase) (8) to a seven-subunit complex of about 500,000 daltons (E. coli DNA polymerase I11 holoenzyme) (9). Inasmuch as multisubunit forms dissociate easily, a polymerase isolated as a single polypeptide may be part of a larger assembly in the cell.

PREFERENCE B. TEMPLATE The enormous variations in size, structure and composition of the template provide the basis for the clearest display of the variable proper7. P. M. J. Burgers and F. Eckstein, JBC 254, 6889 (1979). 8. D. Korn, P. A. Fisher, J. Battey, and T. S.-F. Wang, CSHSQB 43, 613 (1978). 9. C. McHenry and A. Kornberg, JBC 252, 6478 (1977).

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ARTHUR KORNBERG

ties of polymerases. Extending a chain end at a nick, at a short gap or a long gap, displacing a strand, and even switching templates from one strand to the other, are all template-directed capacities that distinguish one polymerase from another.

C. PRIMERPREFERENCE

RNA priming is the general mechanism for starting DNA chains but probably not the universal one (10); a specific deoxynucleotidyl protein appears to provide the Y-start and the primer terminus of each chain of the adenoviral DNA duplex (11). Polymerases encoded by phages T7 and T4 extend tetra- and pentanucleotide RNA primers of nearly fixed composition; mammalian RNA primers for polyoma, SV40, and chromosomal replication are decanucleotides of relatively random composition (12). Escherichia coli primase, responsible for priming phage 6x174 and chromosomal DNA synthesis, incorporates deoxy- as well as ribonucleotides into chains only afew residues long that can be extended by DNA polymerase I11 holoenzyme (13). It seems likely that polymerases differ in their capacity and efficiency to use primers of varying size and composition, and that they may also interact directly with primases responsible for primer synthesis.

D. TOLERANCE FOR SUBSTITUTION OF A NUCLEOTIDE ANALOG An alteration in the base, sugar, or phosphate may be accepted in varying degrees. The same is true for comparable alterations in the primer and template and for substitution of Mn2+for Mg”. E. FIDELITY Insertion of the correct nucleotide into the chain is determined not only by the strictness of selection in the addition step, but also by availability of a subsequent proofreading step to remove a mismatched nucleotide from the primer terminus. The latter step can be accomplished by a 3’ + 5‘ exonuclease domain of the polymerase polypeptide, or perhaps by an 10. A. Kornberg, I n “RNA Polymerases” (R. Losick and M. Chamberlin, eds.), p. 331, Cold Spring Harbor Laboratory, Cold Spring Harbor, 1976. 11. M. D. Challberg, S. V. Desiderio, and T. J. Kelly, PNAS 77, 5105 (1980). 12. R. Eliasson and P.Reichard, JBC 253, 7469 (1978). 13. L. Rowen and A. Kornberg, JBC 253, 770 (1978).

1.

DNA POLY MERASES-A

PERSPECTIVE

7

associated exonuclease. Thus, correct base pairing may be required at the primer terminus as well as in its covalent extension when the incoming nucleotide is matched to the template.

F. PROCESSIVITY When the polymerase remains associated with the template-primer after the covalent addition of a nucleotide it is regarded as processive. (The opposite term is distributive .) The working polymerase molecule, if distributive, may equilibrate with free polymerase molecules after every addition event or, if processive, may not so do until chains, many thousands long, have been completed. The degree of processivity is determined by the state and nature of the polymerase, the template-primer, the nucleotide, the metal, and the reaction conditions (14).

G. CATALYTIC EFFICIENCY The rate of chain growth depends on the rate of the many events that comprise a nucleotide addition step: complex formation with primer terminus and template, binding of the correct deoxynucleoside triphosphate and rejection of incorrect ones, formation of the diester bond, elimination of inorganic pyrophosphate, and movement along the template-primer to the new primer terminus or dissociation from it. Turnover numbers, measured in vitro, vary from several hundred to many thousand residues polymerized per enzyme molecule per minute.

H. NICKTRANSLATION AND STRAND DISPLACEMENT A polymerase, such as E. coli polymerase I, has the capacity to start synthesis on duplex DNA at a nick in one of the strands and coordinately degrade the 5' end of the chain with its 5' -+ 3' exonuclease activity ( 3 ) . This concurrent polymerization and hydrolysis moves (linearly translates) the nick along the duplex without change in mass of the DNA. The nicktranslation capacity of E. coli polymerase I may be a manifestation of its repair function in removing an RNA primer from the 5' end of a chain, or in excising other uncommon nucleotides, such as a mismatched sequence in that location. Should the 5' chain end at the nick escape cleavage, E. coli polymerase I can displace that chain for even great distances simply by progressive polymerization. Most polymerases need the assistance of helicases and the expenditure of ATP energy to destabilize duplex DNA 14.

p. J. Fay, K.

0. Johanson, C. McHenry, and R. Bambara, J5C 256, 976 (1981).

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ARTHUR KORNBERG

and achieve strand displacement; the presence of several distinctive helicases inE. coli (15) may be responsible for strand displacement associated with specialized functions, and may also provide for specific interactions with polymerases. I.

MULTIPLICITYA N D ABUNDANCE

Eukaryotic cells have one or a variety of polymerases in the nucleus; polymerases are also in cytoplasmic locations, as in mitochondria, chloroplasts, and certain viral factories. Even a prokaryotic cell, such as E. coli, has three distinctive polymerases. The molecular abundance of polymerases may be low, as for those associated with replicative forks, or many times higher for those designed for gap-filling and repair functions. Polymerases encoded by viruses are relatively numerous because they are produced to sustain rapid viral multiplication unregulated by strict copynumber controls, as is the host chromosome.

J. PHYSIOLOGICAL FUNCTIONS Certain polymerases seem designed for the extensive, highly processive, rapid chain growth of chromosome replication. In this role, the polymerase associated with other proteins that contribute to the progress of the replication fork may perhaps be part of a larger entity. Should the existence of such an organized complex become established, it might be called a replisome . It seems likely that the properties of a polymerase that is an integral part of such a larger complex will be markedly different from those observed in the purified enzyme. Other polymerases are designed for gap filling and serve in the excision and repair of mismatched regions, completion of chains undergoing recombination, and the removal and replacement of sections of RNA that served as primers to initiate DNA synthesis. While specialized in function, polymerases are also adaptable and fill other roles demanded by exigencies of mutation, cellular stresses, and invasion by viruses and plasmids. Thus a polymerase may provide an auxiliary or alternative means of synthesis to assist or partially replace a polymerase deficient in numbers or in function.

K. OPTIMALCONDITIONS In view of the multiplicity of polymerase forms, locations, and functions, there is (as might be expected) great variation in the physical and chemical IS. B. Kuhn, M. Abdel-Monern, and H. Hoffrnann-Berling, CSHSQB 43, 63 (1978).

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DNA POLYMERASESA PERSPECTIVE

9

components that define an optimal condition for DNA synthesis, and the range about this optimum that can be tolerated. Sharp distinctions are based on sensitivity to ionic strength, sulfhydryl-blocking agents, temperature, and pH. Especially noteworthy are the influences of DNA-binding agents such as polyamines, histones, specialized DNA-binding proteins, and DNA helicases.

111.

Problems and Prospects in Polymerase Research

A. ISOLATION OF A SCARCE ENZYME Lack of an adequate quantity of a homogeneous enzyme has been, and remains, the greatest obstacle to progress in understanding the nature and action of a polymerase. Chemical and physical studies with impure preparations have been of minimal value, and biochemical studies of such preparations have generally been misleading. The two major difficulties in obtaining a homogeneous preparation of a polymerase are its naturally low abundance and, especially in the case of multisubunit enzymes, their instability. DNA polymerase 111 holoenzyme ofE. coli, for example, is present in only ten to twenty copies per cell and activity is lost upon gentle manipulations (9). Yields of about 3 mg of a preparation, only about 60% pure, are obtained from 3 kg of cell paste after a laborious procedure consuming several weeks. The difficulties in isolating mammalian polymerases from cell cultures and tissues are even more formidable. Whereas skillful and patient application can circumvent enzyme instability, the pursuit of a trace-quantity enzyme can be a forbidding enterprise. No wonder that so little enzymology is done by so few people on so important a subject. To overcome the difficulties of isolating polymerases, two approaches can be taken. One is to locate a relatively abundant source; the other is to amplify the polymerase gene in order to overproduce the enzyme. The natural abundance of a virus-encoded polymerase is high because many copies of the viral chromosome are produced to facilitate rapid and extensive viral multiplication. Thus the phage T4- and T7-encoded polymerases have been isolated in pure form in relatively large amounts and studied extensively (16, 17). An attractive eukaryotic system is the Drosophifu embryo which doubles its DNA content every 10 minutes; 16. C. C. Liu, R. L. Burke, U. Hibner, J. Barry,and B. Alberts, CSHSQB 43,469 (1978). 17. C. C. Richardson, L. J. Romano, R . Kolodner, J. E. LeClerc, F. Tamanoi, M. J. Engler, F. B . Dean, and D. S . Richardson, CSHSQB 43, 427 (1978).

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ARTHUR KORNBERG

from 1 kg of embryos, 1 mg of a nearly homogeneous a polymerase can be obtained (18). As a general method for obtaining an abundant source, the approaches offered by genetic engineering techniques are compelling. By identifying a chromosomal segment containing the polymerase gene or the messenger RNA transcribed from it, recombinant DNAs with the gene in viral or plasmid vectors can create major cellular factories for producing the enzyme. In this way, the abundance of polymerase I of E. coli (19) and the y subunit of polymerase I11 holoenzyme (20) have been raised a hundredfold and the yields of pure protein nearly a thousandfold. Similar avenues are now open for overproduction of eukaryotic as well as prokaryotic polymerases. PRGPERTIES B. PHYSICAL AND FUNCTIONAL

For even the most intensively investigated polymerases virtually nothing is known about amino acid sequence or three-dimensional structure. Based on a facile proteolytic cleavage of E. coli polymerase I into two functionally distinct fragments, the presence of an exposed hingelike region in this multifunctional enzyme can be inferred (I). Also a limited amount of information about the active sites has been obtained with chemical probes. However, the assembly and arrangement of subunits in the complex polymerases are unexplored, and the major harvest of knowledge from X-ray ditFraction analysis of crystal structure has yet to be made. With regard to functional properties, enough is known to formulate the invariant rules of polynucleotide assembly that have been cited. Yet the several component stages that comprise the template-directed, covalent addition of a nucleotide have not been dissected and analyzed. The number and nature of substrate binding sites are known in only a few instances. Allosteric effects have not been discovered or established. Nor has the role of zinc, possibly a universal component of polymerases, been explained. How each of the polypeptides of the multisubunit polymerases contributes to the catalytic efficiency, specificity, fidelity, and processivity is still a mystery. With the imminent availability of adequate quantities of homogeneous polymerases and sensitive methods for tracing them, there can be no excuse for neglecting the intensive and widespread investigation of their physical and functional properties. 18. G. Villani, B. Sauer, and I. R. Lehman, JBC 255, 9479 (1980). 19. W. S. Kelley, K. Chalmers, and N . E. Murray, PNAS 74, 5632 (1977). 20. U. Hiibscher and A. Kornberg, JBC 255, 11698 (1980).

I.

DNA POLYMERASES-A

c.

PERSPECTIVE

11

THETEMPLATE-PRIMER FOR ASSAY

That homogeneous polymerases are essential for serious studies has already been emphasized. “Not wasting clean thinking on dirty substances” is, after all, an elementary principle of chemistry. Yet the application of this dictum to the template-primer in the assay of a polymerase has not been sufficiently appreciated. The standard DNA preparations derived from natural sources and most synthetic homopolymers in general use have been a frequent source of confusion. Obtaining a clean, intact E. coli chromosome, namely the circular, duplex DNA molecule of four million base pairs, is still impossible. What can be obtained is a collection of random fragments with nicks, gaps, and variable ends. A comparably battered substrate is calf thymus DNA, often “activated” for assay use by further degradation with pancreatic DNase. Such templates provide targets for many adventitious actions by a polymerase and serve unpredictably as substrates for a large variety of nucleases, recombinases, and other DNA-directed enzymes. Inhomogeneities in size and structure of synthetic DNA polymers bedevil their use, as with most natural DNAs. Sources of intact natural DNA can be found in the form of viral chromosomes and plasmids and have proved to be attractive substrates for the template-primer role for polymerases. Although such molecules are generally inert for polymerases directly, they can serve in the discovery of the natural priming systems that make them susceptible to polymerase action. Improved techniques for isolating the DNAs of even large viruses and plasmids will soon make their use feasible too. Advances in the organic synthesis of relatively long DNA sequences are likely to provide the variety of templates and primers of defined primary and secondary structure needed for refined studies. D.

LINKAGE TO RELATED REPLICATION PROTEINS

It seems plausible that a DNA polymerase would interact in a precise way and for a sustained period with proteins that precede, facilitate, and follow its actions. Several examples can be suggested: (i) Polymerases can use very short RNA primers because they are held in place by the primases that make them (13, 21). (ii) Single-strand binding proteins that destabilize helical structures in some instances stimulate replication only by particular polymerases (16). (iii) Progress at the replicating fork by continuous synthesis by phage T4 polymerase is enhanced by the gene 41 and 61 proteins responsible for the priming actions in the discontinuous 21. G. Hillenbrand, G. Morelli, E. Lanka, and E. Scherzinger, CSHSQB 43,449 (1978).

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phase of synthesis (16); similarly in renlication of the duplex form of phage 4x174, a coupling may exist between DNA polymerase III holoenzyme and the complex of priming proteins, called the primosome (22, 23). (iv) Associated nucleases may contribute proofreading or lesion-excision functions (3, 24). (v) Machinery for biosynthesis of the deoxynucleoside triphosphate substrates may be part of the fabric that contains the polymerase (25). While still imaginary, a large replisome with functional responsibilities that go beyond replication to chromosome orientation and segregation, is an attractive investigative goal.

E. BIOSYNTHESISAND REGULATION A major current question in biology and medicine is what determines the initiation of a cycle of chromosome replication and thus the factors that govern the patterns of DNA synthesis in resting or dividing cells. Despite all that is known about the cell cycle from studies of its biology, it seems unlikely that a genuine understanding of the control of replication will emerge without a detailed knowledge of its biochemistry. More information about the identity of the polypeptides responsible for DNA synthesis and the genetic loci that encode them will make it possible to determine their levels as a function of the cell cycle, their assembly into larger complexes, and the mechanisms for regulation of their biosynthesis. In such studies, plasmid-bearing strains that overproduce the polypeptides, and mutants that produce defective polypeptides, will be of special value.

IV.

Conclusion

Fashion dictates the ebb and flow of scientific activity as much as it does other human social activities. Current attention to manipulating DNA, examining the immune system, and explaining physiologic and pathologic events has taken its toll in solving other problems, including some basic ones. Enzymology is one of the victims, particularly the study of complex and scarce enzymes such as the DNA polymerases. Fortunately there are now opportunities and forces that may direct more effort toward these neglected goals. An adequate start has been made in iden22. 23. 24. 25.

N . Arai and A. Kornberg, JBC 256, 5294 (1981). R. L. Low, K. Arai, and A. Kornberg, PNAS 78, 1436 (1981). D. W. Mosbaugh and R. R. Meyer, JBC 255, 10239 (1980). J . B. Flanegan and G. R. Greenbcrg, JBC 252, 3019 (1977).

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13

tifying the subunit structure of the polymerases and the genes that encode them. Remarkable techniques are provided by genetic engineering for amplifying the levels of these scarce enzymes: Novel methods are available for isolating enzymes, such as the use of monoclonal antibodies resulting from cell fusions; isotopic and staining methods, a hundredfold more sensitive, can be used for tracing the enzymes; and improved techniques are in hand for obtaining well-defined DNAs as template-primers for assay. Intensive studies of polymerase enzymology will be repaid by insights into the protein-nucleic acid interactions fundamental to gene expression and by a better understanding of the factors that control replication, recombination, and repair of DNA.


DNA Polymerase I of Escherichia coli I. ROBERT LEHMAN

. . . . .

I. Purification . . . . . . . . . . . . . . . . . . . . .

. . . . . . . . . , , . . . . . A. Multiple Functions of DNA Polymerase I . . . . . B. General Features of the Polymerization Reaction . C. Processivity of Polymerization . . . . , . . . . . D. 3’ + 5‘ Exonuclease , . . . . . . . , . . . . . E. 5’ + 3’ Exonuclease . . . . . . . , , . . . , . IV. Biological Role . . . . . , . . . . , , . . , . . . A. DNARepair . . . . , . . . . . , . . . . . . . B. DNA Replication . . . . . . . . . . . . . . . . V. Research Applications . . . . . . . . . . . . . . . A. Preparation of Highly Radioactive DNA Probes . . B. Molecular Cloning . . . . . . . . . . . . . . . C. DNA Sequencing . , . . . , . . , , . . . , . .

11. Physical Properties 111. Reaction Catalyzed

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DNA polymerase I (pol I) of E. coli catalyzes the polymerization of nucleotides at the direction of a nucleic acid template. It was the first such erizyme to be discovered ( I ) . Since Pol I was the subject of a detailed review in 1974, in Volume X of this series (2), the major focus of this chapter is on knowledge of the enzyme that has developed during the ensuing 6 1. A. Kornberg, I. R. Lehman, M. Bessman, and E. S. Simrns, BBA 21, 197 (1956). 2. T.Kornberg, and A. Kornberg, “The Enzymes,” 3rd ed., Vol 10,p. 119, 1974. 15 THE ENZYMES, Vol. XIV Copyright 0 1981 by Academic Press. Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

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years; in particular, information regarding the processivity of polymerization, and the function of pol I in vivo as deduced from the analysis of mutant enzymes. 1.

Purification

Purification of pol I, previously a difficult task because of the relatively low concentration of the enzyme in wild-type E. coli (400 molecules per bacterium), has been greatly simplified by the molecular cloning of the polA gene in a lambda transducing bacteriophage (3). Extracts prepared from cells infected with this phage are up to one hundredfold enriched in pol I activity, and homogeneous enzyme can be obtained after approximately one hundredfold purification (4). This is to be compared with the five thousandfold purification previously required to achieve homogeneous enzyme (S). Pol I purified by this procedure is indistinguishable from that isolated from wild-type E. coli in its molecular weight and amino acid composition. II. Physical Properties

The amino acid sequence and three-dimensional structure of pol I have not yet been determined. However, considerable information is available regarding its structure and physicochemical properties (6 1, This information may be briefly summarized as follows. DNA polymerase I consists of a single polypeptide with a molecular weight 109,000 and an sz0,,+, value of 5.5-5.6 (5). It appears to be folded into two domains joined by a protease-sensitive peptide linker (7). The protein is approximately spherical, with a diameter of 65 A as determined from its hydrodynamic properties (8) and from electron microscopy (9). As judged by ORD measurements it contains a significant amount of alpha-helical structure (1%45%) (5). Pol I contains a single free sulfhydryl W. S. Kelley, K. Chalmers, and N. E. Murray, PNAS 74, 5632 (1977). W. S. Kelley and K. H. Stump, JBC 254, 3206 (1979). T. M. Jovin, P.T. Englund, and L. Bertsch, JBC 244, 6996 (1969). A. Kornberg, “DNA Replication,” Freeman, San Francisco, 1980. D. Brutlag, M. R. Atkinson, R. Setlow, and A. Kornberg, BBRC 37, 982 (1%9); H. Klenow and I. Henningsen, PNAS 65, 168 (1970). 8. P. T. Englund, M. P. Deutscher, T. Jovin, R. B. Kelly, N. R. Cozzarelli, and A. Kornberg, CSHSQB 33, 1 (1968). 9. J. Gnffith, J. A. Huberman, and A. Kornberg, J M B 55, 209 (1971). 3. 4. 5. 6. 7.

17

2. DNA POLYMERASE I OF Escherichia coli

group that can form a dissociable complex with HgZ+ (10) and can be carboxymethylated without loss of activity (5). As determined by atomic absorption spectroscopy, pol I contains one Zn2+atom per molecule (/I). As yet there is no direct evidence for the participation of this metal in the catalytic cycle of the enzyme. Indeed, the finding that the Zn2+chelator, o-phenanthroline is a potent inhibitor of pol I can no longer be taken as evidence for the involvement of Zn2+in pol I action. It has been found that the inhibition by phenanthroline is due to an inhibitory phenanthrolineCu(1) chelate, which forms in the presence of the trace contaminants of Cu2+and thiols present in pol I assay mixtures (12). An analysis of pol I by a variety of techniques including equilibrium dialysis and spectroscopic examination (NMR and ESR), as well as kinetic measurements, has yielded a picture of the active center of the enzyme in which there are three closely juxtaposed sites: (1) A DNA template-binding site, (2) a nucleoside-monophosphate-binding site, which presumably represents the site at which the primer and growing chain are bound, and (3) deoxynucleoside triphosphate site, which accommodates all four triphosphates (6 ). 111.

Reaction Catalyzed

A. MULTIPLEFUNCTIONS OF DNA POLYMERASE I (6) DNA polymerase I catalyzes the addition of mononucleotide units from deoxynucleoside 5'-triphosphates to the 3'-hydroxyl terminus of a primer chain. For this reaction, a template is absolutely required and directs the enzyme in its selection of the specific triphosphate according to the Watson-Crick base-pairing rules. This reaction may be written: (dNMP).

+ dNTP

(dNMPL,,

+ PP,

The turnover number for the enzyme in this reaction is 667 nucleotides polymerized per molecule of enzyme per minute at 37". In addition to the polymerization of nucleotides, pol I catalyzes the pyrophosphorolysis of DNA, a reaction which in fact represents the reversal of polymerization. (dNMP).,,

+ PP,

(dNMPX, + dNTP

10. T. M. Jovin, P. T. Englund, and A. Kornberg, JBC 244, 3009 (1969). 11. J. P. Slater, A. S. Mildvan, and L. A. Loeb, BBRC 44, 37 (1971). 12. D. S. Sigman, D. R. Graham, V. D'Aurora, and A. Stem, JBC 254, 12269 (1979).

18

I. ROBERT LEHMAN

It also catalyzes pyrophosphate exchange according to the following reaction: (dNMP),

+ dNTP + PP,* %(dNMP). + dNMP-PP* + PP,

This reaction represents a repetitive sequence of nucleotide addition and pyrophosphorolysis of the newly synthesized phosphodiester bond. In addition to nucleotide polymerization and phosphorolysis, pol I catalyzes the hydrolysis of phosphodiester bonds. There are two such hydrolytic activities; one is a 3‘ + 5‘ exonuclease and the other a 5‘ + 3‘ exonuclease. The two exonuclease activities are associated with the two different domains of the pol I molecule. Thus, exposure to subtilisin cleaves the enzyme into two active fragments, a “large” fragment (MW 76,0001, which contains polymerase and 3‘ + 5’ exonuclease activities, and a “small” fragment (MW 36,000), which contains the 5‘ -+3‘ exonuclease. The small fragment corresponds to the amino terminal portion of the molecule as judged by the identity of the N-terminal amino acid sequences of the intact enzyme and small fragment (13), and the finding that an amber peptide isolated from the polA1 mutant retains the 5‘ 3 3’, but not the polymerase activity of the enzyme (14).

B. GENERAL FEATURES OF THE POLYMERIZATION REACTION(6) DNA polymerase I can catalyze three different modes of polymerization: Gap filling, nick translation, and strand displacement. It is unique among DNA polymerases in its capacity to promote the two latter reactions (Fig. 1). 1. In gap filling, pol I acts to fill in the single-stranded regions of a gapped DNA duplex by nucleotide addition to the 3’-hydroxyl group at the gap. 2. Nick translation consists of the coordinated polymerization of nucleotides at the 3’-hydroxyl terminus and the exonucleolytic removal of nucleotides at the 5’ terminus of a nick in duplex DNA. The result of the concerted polymerization and 5‘ + 3‘ exonuclease action is the propagation or translation of the nick along the DNA duplex. 3. Strand displacement consists of polymerization by pol I uncoupled from 5’ -+ 3’ exonuclease action, so that nucleotide addition at the 3‘hydroxyl terminus at a nick displaces rather than hydrolyzes the 5’terminated strand ahead of it. 13. H. Jacobsen, H. Klenow, and K. Overgaard-Hansen, EJB 45, 623 (1974). 14. I. R. Lehman and J. R. Chien, JBC 248, 7717 (1973).

2. DNA POLYMERASE I OF Escherichia coli NICKEDDNA

U

w

3’

NICK TRANSLATION

19

1I I II I I I I I I I Ill

,;. Yr.LLull \ \ /-

STRAND DISPLACEMENT

GAPPED DNA

GAP FILLING

m

3’

f

FIG.1. DNA polymerase I action at a nick or gap in duplex DNA. Nicked DNA is subject to nick translation or strand displacement; gapped DNA can be filled in to generate a nick, which can then support nick translation or strand displacement. In each of these three modes of pol I action, the basic reaction is the same: Addition of a deoxynucleotidyl unit onto a 3’-hydroxyl terminus with the resultant pairing of the added nucleotides to the complementary nucleotide on the template strand. The extraordinary fidelity of DNA replication by pol I is a consequence of the high degree of selectivity during polymerization and the associated 3‘ -+ 5’ exonuclease (see Sections II1,C and D below). A kinetic analysis of polymerization with a defined series of primer-templates indicates that the presence of the template may facilitate rejection of the noncomplementary nucleotide (15). The template may also increase the a n ity of pol I for the incoming complementary nucleotide. Both of these features of polymerase action could contribute to the fidelity of polymerization. Further to this point, an NMR analysis of the conformation of deoxynucleoside triphosphate-Mn(I1) complexes bound to pol I in the absence of template has shown that there is puckering of the complex such that the base-deoxyribose conformation is close to that found in double helical DNA (16). Formation of such a structure should permit insertion of the nucleotide into the helix in proper alignment for WatsonCrick base pairing and would contribute to the fidelity of replication. The role of the 3’ + 5’ exonuclease component of pol I in ensuring the fidelity of replication is discussed in Section II1,D below.

c.

PROCESSIVITY OF POLYMERIZATION

An important question regarding the mechanism of pol I is whether the polymerization of nucleotides proceeds in a processive or nonprocessive 15. E. C. Travaglini, A. S. Mildvan, and L. A. Loeb, JBC 250, 8647 (1975). 16. D. L. Sloan, L. A. Loeb, A. S. Mildvan, and R. J . Feldman,JBC 250, 8913 (1975).

20

I. ROBERT LEHMAN

(distributive) manner. A processive reaction is one in which many nucleotides are incorporated into the growing chain before the enzyme dissociates. A nonprocessive mechanism is one in which dissociation of the polymerase from the primer-template occurs following addition of each nucleotide. To state this point somewhat differently, a nonprocessive mechanism is one in which dissociation of the enzyme from the primertemplate after each nucleotide addition is an obligatory feature of the catalytic cycle. Three approaches have been taken to determine the processivity of pol I-catalyzed deoxynucleotide polymerization: (1) Template challenge experiments, (2) determination of nucleotide incorporation ratios, and (3) measurement of the decrease in polymerization rate upon removal of one or more of the four deoxynucleoside triphosphates. 1.

Template Challenge

Template challenge experiments measure the rate at which the enzyme can redistribute itself between competing templates. In one such study, an excess of poly(dC) oligo(dG) was added to a reaction mixture in which pol I was replicating poly(dA) oligo(dT) (17). Incorporation of dTTP was promptly inhibited and, correspondingly, incorporation of dGTP began. The conclusion from these studies was that nucleotide polymerization by pol I is distributive. However, the time scale of the experiment was such that 100 nucleotides or more may have been polymerized before template switching occurred. In another experiment in which calf thymus DNA was added as the competing template and the temperature was lowered to 4" to slow polymerization, it was found that within the time required for turnover of one nucleotide the calf thymus DNA could compete effectively with the d(A-T) copolymer that served as the initial template (18). Again, the conclusion reached was that pol I is distributive. In this instance an important limitation to the interpretation is the lack of information regarding the affinity of the enzyme for the different polynucleotides at 4". 2 . Nucleotide Incorporation Ratios This method for the assessment of processivity takes advantage of the nucleotide sequence of the cohesive ends of bacteriophage lambda DNA (Fig. 2). With this DNA a known sequence of nucleotides is incorporated into the cohesive ends, and furthermore the composition of the nu17. L. M. s. Chang,JMB 93, 219 (1975). 18. W. R. McClure and T. M. Jovin, JBC 250, 4073 (1975).

21

2. DNA POLYMERASE I OF Escherichiu coli left-hand end 3 5

right-hand end

C C C G C C G C T G G A (G)

GGGCGGCGACCT

3

FIG.2. Terminal sequences of bacteriophage lambda DNA. The letters represent the nucleotides incorporated when the cohesive ends are completely filled in. The bracket indicates the nucleotides incorporated if only dGTP and dCTP are used for synthesis. The letter in parentheses represents the 3'-terminal dG of the left-hand end of native lambda DNA. This residue is exchangeable with dGTP during synthesis in the absence of dATP. The cohesive ends are represented by solid lines. The long double-stranded internal region of the molecule is not shown. cleotides incorporated changes in a predictable way as polymerization proceeds ( 1 9 ) . When dGTP and dCTP are added only the nucleotides enclosed by the bracket are incorporated into the right-hand end, and there is no synthesis at the left-hand cohesive end. If nucleotide polymerization occurs by a processive mechanism, at a low molar concentration of enzyme relative to DNA, the ratio of dG to dC incorporated remains constant and does not change with the extent of reaction. On the other hand, if the mechanism is nonprocessive, early in the reaction a large proportion of the lambda DNA molecules incorporate only 1 or 2 dG residues into their right-hand cohesive end, and the ratio of dG to dC is high. As the reaction proceeds to completion the dG : dC ratio drops to a value of 3, the ratio of the number of dG and dC residues incorporated into the right-hand cohesive end. When this type of analysis was applied to pol I at 6",the ratio of dG to dC incorporated remained close to three throughout a 10-hour period of incubation (Fig. 3). Thus, under these conditions pol I is processive for at least the 12 nucleotide length of the right-hand cohesive end of lambda DNA. In a control experiment in which there was a 60-fold excess of enzyme over 3'-hydroxyl termini, the dG : dC ratio decreased from 17 at 30 seconds to a value of 3 at 5 minutes. An interesting feature of this experiment was the finding that even in the presence of a large molar excess of enzyme over DNA termini, the time required to complete synthesis at the direction of any one cohesive end was considerably shorter than that required to complete polymerization at all ends (20). Thus, regardless of whether pol I was present in excess or was limiting, about 30 to 40% of the enzyme mole19. D. Uyemura, R. Bambara, and I. R. Lehman JBC 250, 8577 (1976). 20. R. A. Bambara, D. Uyemura, and I. R. Lehman,JBC 250, 4090 (1976).

22

I. ROBERT LEHMAN

0

0

1

2

4

10

TIME (HOURS)

FIG.3. Nucleotide incorporation into the cohesive ends of lambda DNA by DNA polymerase I at 6" in the presence of dGTP, dCTP, and limiting enzyme (19). cules began polymerization rapidly, but synthesis on the remaining molecules was significantly delayed (Fig. 4). This delay was not a consequence of a low rate of association between enzyme and primer-termini at the cohesive ends, but rather of the slow onset of polymerization at all ends after association had occurred. A plausible explanation for this effect is that two forms of the enzyme or enzyme-DNA complex exist, only one of which is active. Inasmuch as 30 to 40% of the enzyme molecules begin polymerization immediately, the normal equilibrium between the two forms would give a ratio of inactive to active molecules of two. By this argument the rate of conversion from inactive to active enzyme (or enzyme-DNA complex) is the rate-determining step for the initiation of polymerization. Indeed one consequence of the defect in pol I from strains bearing thepolAI2 mutation (see Section IV,B,l) is apparently to alter the equilibrium such that only a relatively small fraction of active molecules are present at any time. However, the rate of conversion from the inactive to the active form is not significantly altered by the mutation.

23

2. DNA POLYMERASE I OF Eschcrichici coli I

E

0

-B 3000

.

EXCESS ENZYME

0 I-

2

g a

2000

E

a

zn

+a-

LIMITING ENZYME

B 1000

-I

M

-0 n

20

60

120

180

TIME (MINUTES)

FIG. 4. Comparison of rates of nucleotide incorporation into the cohesive ends of lambda DNA at excess and limiting concentrations of DNA polymerase I(20). 3 . Polymerization in the Absence of One or More Deoxynucleoside Triphosphates

Determination of the rate of polymerization in the presence of one, two, or three deoxynucleoside triphosphates compared to the rate in the presence of all four provides a quantitative measure of processivity over a large range of values (one to several hundred). Under these conditions the rate of polymerization is determined in part by processivity; the rate with a full (as compared to a partial) complement of deoxynucleoside triphosphates can be used to distinguish processivity from other factors that affect the rate of polymerization (2 1 ). With a large excess of primer-termini over enzyme molecules, nucleotide polymerization may be considered to be a two-phase cyclic process. In the first phase free enzyme diffuses to and binds the primertemplate. In the second phase the enzyme catalyzes an ordered succession of dNTP binding, nucleotide condensation, and translocation until the enzyme dissociates from the primer-template. The entire cycle is then repeated. The average increment of time during one complete cycle, from the binding and reaction of the DNA polymerase with a 3’-primer terminus, through the dissociation and diffusion steps, to the rebinding of the enzyme to another reactive 3’ terminus, is defined as the “cycling time.” 21. R. A. Bambara, D. Uyemura, and T. Choi,JEC 253, 413 (1978).

24

I. ROBERT LEHMAN

The polymerization rate whenx dNTPs are present may be expressed as follows:

P,

=

ENJT,

(1)

where P, is the rate of polymerization, E the number of active enzyme molecules, N , the average number of nucleotides incorporated per polymerization cycle, and T, the cycling time. This is a general expression that applies both to synthesis with a limited complement of dNTPs (x = 1, 2, or 3) and to synthesis with a complete complement of dNTPs (x = 4). For DNA containing all four bases, N4 is the average number of nucleotides polymerized with a complete complement of dNTPs, and is therefore the processivity of polymerization. The aim of the calculations that follow is to express N4 in terms of parameters that can be measured experimentally. The ratio of polymerization rates with a limited versus a complete complement of dNTPs (PXz4) may be expressed as

or P,:4 = N,T4/N4T,

(x = 1, 2, or 3)

(3)

Values of N , may ultimately be expressed in terms of N4 and two statistical factors. The TJT, ratio must be determined for individual experiments with the use of an inhibitor that binds the DNA polymerase for a short period of time during the first phase of the polymerization cycle. The presence of the inhibitor increases the cycling time, T,, by an increment, Ti such that

Tx,i = Tf + Tx

(4)

where T,,{ is the cycling time in the presence of the inhibitor. The inhibitor should be chosen so that the value of Tf is not affected by the value of x. This requirement can be verified experimentally because processivity dewould not be equivalent unless Ti were indetermined from P I , and pendent of x . Using this definition of Ti, T,/T4 can be expressed in terms of rates of reaction. First, it follows from Eq. (1) that

T, = EN,/P,

(5)

With this form of the equation the cycling times for inhibited and uninhibited reactions can be related directly to measurable rates:

25

2. DNA POLYMERASE I OF Eschcrichia Cali

where Px,i is the rate of reaction when the inhibitor is present. Substituting for Tx,i from Eq. (4)

Ti can be evaluated as follows:

From Eq. (7), when four dNTPs are used in the reaction -T4 +

Ti - &

T4

P4.f

Substituting for Ti in Eq. (9) by using the value of Ti in Eq. ( 8 )

and solving for Tx/T4

Returning to Eq. (3), the measured reaction rates Px, P4, Px,i, and P4,t can be related to the processivity of polymerization:

This expression is rearranged as follows:

and

Equation (13) may also be rewritten

26

I. ROBERT LEHMAN

where P&, = P,:,T,/T,. Since only a single triphosphate is labeled in all experiments, a correction is made to allow use of the ratio of rates of radioactive nucleotide incorporation (R::,). The factor L relates RI ,: to P&, as follows:

R1.4

=

LP;:4

(16)

This expression is incorporated into Eq. (15) to yield N4 = LN,/R::,

(17)

The theoretical relationship between N4 and R::, for calf thymus DNA is shown in Fig. 5. The value of R::, is given by R::, = Rx:4(Tx/T4)

(18)

where R,:, is the ratio of label incorporated in limited (x = I , 2 , or 3) as compared to complete ( x = 4) reactions and T,/T, is calculated from Eq. (11).

An important limitation to the analysis is the possibility that the average length of the template is shorter than the extent of processivity. If so, the observed rate of polymerization with a full complement of dNTPs will be lowered and the ratio of polymerization rates will be increased, leading to an artifactually low processivity. The average length of template available to a DNA polymerase at each 3' terminus can be determined when there is an excess of enzyme molecules over primer-termini, and extents rather than rates of polymerization are measured. An equation similar to that used in the calculation of processivity can be used. In this case A,:, = LA,/A,

( x = I , 2 , or 3)

(19)

where Ax:,is the ratio of extents of nucleotide incorporated with a limited and a complete complement of dNTPs, and A, and A, are the average maximum number of nucleotides that can be added to a 3' terminus with a limited and complete complement of dNTPs, respectively. The relationship between A,:,, A,, and A, is the same as that between R,:,, N,, and N4 (see above). IfA, = N4 it may be assumed that template length limits the measured processivity of DNA polymerization. If A, > N4 the processivity may be assumed to be a consequence of the intrinsic tendency of the DNA polymerase to dissociate under the conditions of measurement. An analysis of the processivity of pol I by this method has shown that the enzyme can be highly processive, with values as high as 200, depending upon the primer-template and the conditions of polymerization. However, the processivity decreases at high ionic strength and at low temperatures. With nicked DNA templates at 37" and ionic strength 0.085,

2. DNA POLYMERASE I OF Escherichia coli

27

200 100

> 20

f

10

E

5

0

2 1

0.01

0.1

1.o

RATIO OF INCORPORATION RATES (R',,q)

FIG.5 . Theoretical relationship between RiZ4and processivity ( N ) , with activated calf thymus DNA as a primer-template. Curves are shown for five combinations of deoxynucleoside triphosphates in the limited reaction. The abscissa is the ratio of label incorporated @?::4) and the ordinate is the processivity ( N ) . The curves represent limited reactions containing (1) PHIdTTP: (2) PHIdTTP + dCTP or [3H]dTTP+ dGTP; (3) PHIdlTP + dCTP; (4) rH]dTTP + dCTP + dGTP; and ( 5 ) l3H]d=P + dATP + dCTP or PHJdlTP + dATP + dGTP ( 2 1 ) . processivity is 15 t o 20. With gapped DNA under the same conditions, the processivity increases to 40 to 50. With poly d(A-T), the processivity is 188. However, by increasing the ionic strength and decreasing the temperature, this value is reduced to 3. Of interest in this regard is the finding that the enzyme isolated from the polA5 mutant gives processivity values that are only one-fifth of wild-type pol I. To summarize then, pol I binds rapidly t o its primer-template. This is followed by a significant delay in the onset of polymerization because of a slow shift in the equilibrium between the inactive and active forms of the enzyme. Polymerization is processive, the extent of processivity being determined by a variety of factors (temperature, ionic strength) that appear t o influence the structure of the primer-template and its interaction with the enzyme.

D.

3'

+

5' EXONUCLEASE

The 3' + 5' exonuclease associated with pol I acts specifically on single-stranded DNA and hydrolyzes it to nucleoside 5'-monophosphates

28

1. ROBERT LEHMAN

(22). Of particular importance is its ability to remove a non-base-paired terminus. Thus, given a primer-template with one or more mismatched nucleotides at the 3' terminus, pol I removes the mismatched nucleotides by virtue of its 3' +. 5' exonuclease component before initiating polymerization. The removal of mismatched nucleotides by the 3' + 5' exonuclease thus represents a proofreading mechanism that contributes to the high fidelity of DNA replication (23). Following infection with phage T4, a DNA polymerase is induced with a similar 3' + 5' exonuclease, and defects in the balance of polymerase and exonuclease in various mutant phages have been correlated with mutator and antimutator effects observed in vivo (24,251. Thus far, no such effects have been observed for pol I isolated from E. coli polA mutant strains (see below). As described in the preceding section, the selective orientation of the incoming nucleotides by the template and the induced puckering of the deoxyribose-base bond may also be important in maintaining the fidelity of replication at the level of proper base selection. In fact, a "mutator" pol I isolated from Salmonella typhimurium appears to be defective in base selection rather than its 3' + 5' exonuclease (26).

E. 5'

+

3' EXONUCLEASE

The 5' + 3' exonuclease function of pol I degrades duplex DNA in the same direction as polymerization, starting at the 5' terminus of a nick in the duplex (27). In contrast to the 3' 5' exonuclease of pol I, it is inactive on single-stranded DNA; furthermore, it produces 5' phosphoryl-terminated oligonucleotides in addition to mononucleotides (28).The 5' -+3' exonuclease can excise damaged nucleotides from DNA following incision of the DNA duplex at or near the site of damage (29), and it can degrade the RNA component of a DNA RNA hybrid (30). As

+.

22. I. R. Lehman and C. C. Richardson, JBC 239, 233 (1964). 23. D. Brutlag and A. Kornberg, JBC 247, 241 (1972). 24. N . Muzyczka, R. L. Poland, and M . J. Bessman, JBC 247, 7116 (1972). 25. F. D. Gillin and N . G . Nossal, JBC 251, 5219, 5225 (1976). 26. M. Engler, and M. J. Bessman, C S H S Q B 43, 929 (1978). 27. R. P. Klett, A. Cerarni, and E. Reich, PNAS 60, 943 (1968). 28. N . R. Cozzarelli, R. B. Kelly, and A. Kornberg, JMB 45, 513 (1969). 29. R. B. Kelly, M. R. Atkinson, J. A . Huberman, and A. Komberg, Nature (London) 224, 495 (1969). 30. D. Baltimore, and D. F. Smoler,JBC 36, 185 (1972); J. P. Lies, I. Berkower, and J. Hunvitz, PNAS 70,466 (1973);0. Westergaard, D. Brutlag, and A. Kornberg,JBC 248, 1361 (1973).

2. DNA POLYMERASE I OF Escherichiu coli

29

indicated previously, 5‘ -+ 3’ exonuclease action is coupled with nucleotide polymerization at a nick in DNA, resulting in propagation or translation of the nick along the duplex (31). The process of nick translation ensures that long single-stranded stretches do not accumulate within a DNA duplex that is undergoing repair or replication. Although it was believed that the 5’ += 3’ exonuclease requires a 5’ terminus to initiate hydrolysis, it is now clear that under certain circumstances pol I can catalyze an endonucleolytic scission even in the absence of a terminus. This has been observed with negatively supertwisted circular DNA into which a single-stranded DNA or RNA segment has been assimilated to form a D loop. When such a structure is used by pol I as a primer-template the primer is elongated, resulting in an enlargement of the loop. In addition, however, endonucleolytic cleavage of the circular DNA occurs (32). This may be a consequence of 5’ 4 3’ exonuclease action at the strand not paired to the primer. The replication fork generated by polymerase action leads to a distortion in the duplex, which resembles that introduced by, for example, a thymine dimer and, hence, may become a site for the endonucleolytic action of the 5‘ + 3’ exonuclease.

IV.

Biological Role

Strains of E. coli with mutations in the polA gene are abnormally sensitive to ultraviolet irradiation and to radiomimetic agents (.33),and they are defective in chromosomal DNA replication (34). DNA polymerase I is therefore required for both the repair and the replication of DNA in vivo . Pol I is also essential for the replication of certain plasmids, for example the colicinogenic factor, ColE1 (35). A.

DNA REPAIR

The capacity of pol I to promote nick translation is clearly a key feature in its repair function. Thus, following incision at or near the site of damage (for example, a thymine dimer introduced following UV irradiation), pol I 31. R. B. Kelly, N . R. Cozzarelli, M. P. Deutscher, I. R. Lehman, and A. Kornberg, JEC 245, 39 (1970). 32. L. F. Liu and J. C. Wang, In “DNA Synthesis and Its Regulation” (M. Goulian and P. Hanawalt, eds.), p. 38. Benjamin, Menlo Park, 1975. 33. P. L. De Lucia, and J. Cairns, Nature (London) 224, 1164 (1%9). 34. P. L. Kuempel, and G . W. Veomett, BBRC 41, 973 (1970); R. Okazaki, M. Arisawa, and A. Sugino, PNAS 68, 2954 (1971). 35. D. T. Kingsbury, and D. R. Helinski. BBRC 41, 1538 (1970).

30

I. ROBERT LEHMAN

can catalyze the concerted 5' + 3' exonucleolytic removal of a DNA segment that includes the dimer, and the 5' + 3' polymerization required to restore the DNA duplex (31). Inasmuch as polA mutants are only moderately sensitive to UV irradiation, it is clear that alternative excision-repair mechanisms must exist that can substitute for pol I when the latter is defective (36). Indeed, pol I1 and/or pol I11 (which has an intrinsic 5' + 3' exonuclease) have been implicated in what has been termed "long-patch" repair, as opposed to the "short-patch" repair involving little degradation and the limited resynthesis characterization of wild-type strains with a fully functional pol I (36). B. DNA REPLICATION The joining of nascent DNA fragments is retarded in allpolA mutants that have been examined, indicating that pol I is required for the discontinuous replication of the E. coli chromosome (37). The severity of the defect differs considerably from one mutant strain to another; however, it is most pronounced at restrictive temperatures in conditionally lethal, temperature-sensitive mutants (37, 38). As in excision repair, the unique capacity of pol I to catalyze nick translation is the crucial feature of its function in discontinuous DNA replication. Thus, pol I can promote the coordinated 5' + 3' exonucleolytic removal of DNA primers at the 5' termini of Okazaki fragments, and the filling in of the gaps thus created (31). This coupled reaction permits the ligation of discontinuously synthesized DNA fragments to the growing DNA chain. polA mutants have been identified whose pol I is defective in the polymerase, the 5' + 3' exonuclease, and the capacity to coordinate the two. In each instance, there is an abnormal persistence of nascent DNA (Okazaki) fragments. The latter two mutant enzymes have been purified to homogeneity and are considered below. 1. Nonlethal PolA I2 Mutant

Strains bearing thepolA12 mutation show a temperature sensitive repair defect; that is, they are defective in the repair of DNA damage at 43", but not at 30" (39). They are, however, viable at both temperatures. The polAI2 polymerase is extremely thermosensitive. It is also rapidly dena36. P. Hanawalt, A. Burrell, P. Cooper, and W. Masker, In "DNA Synthesis and Its Regulation" (M. Goulian and P. Hanawalt, eds.), p. 774. Benjamin, Menlo Park, 1975. 37. I. R. Lehman, and D. Uyemura, Science 193, %3 (1976). 38. D. Uyemura, D. C. Eichler, and I. R. Lehman, JBC 251, 4078 (1976). 39. M. Monk, and J. Kinross, J . Bacteriol. 109, 971 (1972).

2. DNA POLYMERASE I OF Escherichirr coli

31

tured by even brief exposure to ionic strengths less than 0.1. Thus, purification of the mutant enzyme requires procedures that avoid or minimize exposure to solutions of low ionic strength (40). The temperature sensitivity and instability at low ionic strength of the polAI2 DNA polymerase appear to result from a significant alteration in the tertiary structure of the enzyme. The mutant protein has a significantly lower mobility than the wild-type enzyme in discontinuous polyacrylamide gel electrophoresis of the two native proteins. Furthermore, the polAI2 enzyme sediments at a lower rate than the wild type enzyme in a sucrose velocity gradient. The decrease in electrophoretic mobility taken together with the lower sedimentation coefficient suggests that thepolAI2 mutation has produced a misfolding of the mutant protein so that it is less compact than the wild-type enzyme. This is possibly the cause of its thermal instability and rapid inactivation in low salt solutions. A striking feature of the polAI2 enzyme, even at permissive temperatures, is its decreased ability to catalyze nick translation (40). This point is illustrated in Fig. 6. Given nicked PM2 DNA, wild-type pol I catalyzes the incorporation of nucleotides at the 3'-hydroxyl end and the release of nucleotides from the 5' end of the nick in equimolar amounts at both 30" and 43". With gapped PM2 DNA, the gaps are rapidly filled in, regenerating the nicks, which then become sites for nick translation (Fig. 7). The incorporation and release of nucleotides at a nick catalyzed by the polA I2 enzyme are also equivalent. However, the rate at which nick translation proceeds is tenfold lower than that seen with the wild-type enzyme at 30", and is even further reduced at 43" (Fig. 6). With DNA that contains gaps, the mutant enzyme rapidly fills in the gaps at nearly the same rate and to the same extent as the wild-type enzyme, then catalyzes very little further synthesis or release of nucleotides (Fig. 7). Since measurements of the polymerase and 5' + 3' exonuclease activities associated with thepolAI2 enzyme show them to be nearly normal at 30" (40), the abnormally low rate of nick translation suggests that there is a substantial defect in the coordination of polymerization and 5' -+ 3' exonuclease action. Since such coordination presumably demands a rigid spatial arrangement of the two active sites, this novel defect may be a consequence of the structural perturbation caused by the polAI2 mutation.

2. Conditionally Lethul polAexI Mutant The isolation of the temperature-sensitive, conditionally lethal mutant, E. coli polAexI, established that DNA polymerase I is essential for the 40.

D. Uyemura, and I . R. Lehman, JBC 251, 4085 (1976).

32

I. ROBERT LEHMAN 800

600

400

200

0 0

5

10

200

5

10

20

TIME (MINUTES 1

FIG.6. Action of wild-type and polA I2 DNA polymerases on nicked PM2 DNA (37). viability ofE. coli (41). Except for its conditional lethality,pofAex/ resembles otherpolA mutants in its retarded sealing of nascent DNA fragments, and in its sensitivity to methylmethane sulfonate and to ultraviolet irradiation. However, the rate of joining of nascent DNA fragments in the polAexl mutant is significantly more retarded than in nonlethal polA strains (38). In contrast to the polA12 enzyme, the pofAex1 polymerase is not particularly labile in low salt solutions. The pofAex1 protein comigrates with wild-type pol I in polyacrylamide gels containing sodium dodecyl sulfate as well as in native discontinuous, polyacrylamide gels; it also has the same sedimentation coefficient. Thus, there is no obvious structural alteration comparable to that observed with the polA12 polymerase (38). The polymerase and 3' 5' exonuclease activities of thepolAex1 enzyme do not differ significantly from those of the wild-type DNA polymerase I at either 30" or 43". In contrast, the 5' + 3' exonuclease activity is substantially reduced at both temperatures (Table I). Furthermore, the 5' + 3' exonuclease activity at 43" is significantly lower than at 30". With nicked PM2 DNA (Fig. 8) as template primer, thepolAex1 enzyme catalyzes the incorporation of nucleotides at a rate far in excess of the rate of hydrolysis of nucleotides from preexisting DNA. This contrasts sharply with the action of the wild-type polymerase, which maintains an almost perfect correspondence between nucleotide release and incorporation.

-

41. E. B. Konrad and I. R. Lehman, PNAS 71, 2048 (1974).

2. DNA POLYMERASE I OF Escherichiu coli

33

z a E K K

8

z

FIG.7. Action of wild-type andpolAI2 DNA polymerases on gapped PM2 DNA (37). The low rate of nucleotide release by the polAexl enzyme is presumably due to some nick translation that occurs at 30". The increment of nucleotide incorporation beyond the amount released must therefore be due to strand displacement; that is, polymerization in the 5' + 3' direction accompanied by unwinding of the strand preceding the enzyme molecule. The discrepancy between the rates of nucleotide incorporation and release is even greater at 43" than that at 30°, probably as a result of an increase in the rate of polymerization coupled with the decrease in 5' + 3' exonuclease activity at the elevated temperature. The polymerase activity TABLE I

DEFECTIVE5'

+

3' EXONUCLEASE ACTIVITYOF DNA POLYMERASE I ISOLATED FROM E. coli polAexl Enzyme activity (Nmol/mg protein)

W)

Polymerase

5' -+ 3' exonuclease

3' -+ 5' exonuclease

30 43 30 43

14.8 41.9 23.8 66.0

6.1 11.8 0.33 (0.19p

1.6 3.4 2.5 3.1

Temperature Enzyme polA+ polAexl

A portion (25-50%) of the apparent 5' -D 3' exonuclease activity at 43" may be attributable to 3' -D 5' exonuclease action. This estimate is based on the extent of hydrolysis observed on incubation of the nicked PM2 DNA with T4 DNA polymerase, which has 3' -P 5' but no 5' -+ 3' exonuclease activity (37).

34

1. ROBERT LEHMAN 600

I

400 300 -

I

1

30'

-

5

1

- - 430

500

0

1

-

10

20

-

0

p

I

?-!--

0

5

10

20

TIME (MINUTES)

FIG.

8. Action of wild-type and polAexl polymerases on nicked

PM2 DNA (37).

of the mutant is lower than that of the wild-type enzyme in this experiment (compare Fig. 8 and Table I). This may be a consequence of the 5' + 3' exonuclease defect of the mutant. When 5' + 3' exonuclease is reduced, the 5' terminated strand must be displaced ahead of enzyme molecule for polymerization to proceed (Fig. l), and this constraint might be expected to lower the polymerization rate. It therefore appears that, at 30°, the mutant enzyme can catalyze nick translation (at a low rate) and polymerization accompanied by strand displacement. At 43", nick translation is abolished and polymerization proceeds only with strand displacement. 3. Other PolA Mutants

Pol I has been purified to homogeneity from three otherpolA mutant strains, pofA'107 (42), polAS (43), and p0lA6 (44). In none of these instances is the mutation lethal. ThepolA'l07 enzyme is defective in the 5' + 3' exonuclease but not polymerase. Thus, it is similar to the polAexl enzyme. A direct comparison of the extent of the 5' + 3' exonuclease defect with that in thepolAexl mutant has not been made. ThepolA6 mutant enzyme shows an altered pH optimum for polymerization and a reduced binding affinity for DNA. The mutational defect 42. H. L. Heijneker, D. J. Ellers, R. H. Tjeerde, B. W. Giickman, B. van Dorp, and P. H. Pouwels, Mol. Gen. Genet. 124, 83 (1973). 43. S. W. Matson, F. N. Capaldo-Kimball, and R. A. Bambara, JBC 253, 7851 (1978). 44. W. S. Kelly and N. D. F. Grindley, Nucleic Acids Res. 3, 2971 (1976).

2. DNA POLYMERASE I OF Escherichitr cdi

35

appears to lie within the carboxyl-terminal large fragment of the enzyme because the fragment has the same pH optimum for polymerization as the intact enzyme, which is considerably more alkaline than that of the wildtype pol I. The large fragment also shows the reduced binding affinity for DNA that characterizes the intact mutant enzyme. The polA5 mutation which results in a decreased processivity of polymerization was mentioned in Section 111,C,3.

V. Research Applications

pol I has long been useful in the synthesis of defined homo- and copolymers [e.g., d(A-T), d(G-C), poly(dA)(dT)] that have served as well-defined structures for the physicochemical analysis of nucleic acids. More recently pol I has become a key reagent in the preparation of radioactive DNA probes, in molecular cloning, and in DNA sequence analysis. A. PREPARATION OF HIGHLYRADIOACTIVE DNA PROBES The capacity of pol I to catalyze nick translation has been used as an effective method for the preparation of highly labeled DNAs. Such labeled DNAs, and restriction endonuclease fragments derived from them, are then used as probes for detecting homologous sequences by measurements of reassociation kinetics or by in situ hybridization techniques. Indeed, labeled nick-translated SV40 DNA has been used to detect and quantitate DNA sequences present at the level of one SV40 DNA copy per haploid mouse genome, using only microgram quantities of cellular DNA (45).

B. MOLECULARCLONING The ability of pol I to fill in small gapped regions in duplex DNA molecules completely and efficiently has been used in the construction of recombinant DNA molecules in v i m . Using a closed circular duplex DNA molecule as a vector, the procedure involves cleaving the molecule to convert it to a linear structure, adding single-stranded homopolymeric stretches to the 3' termini with terminal transferase, adding the complementary homopolymeric sequence to the 3' termini of the DNA segment to be inserted, and annealing the segment to the linear form of the vector. At this point, gaps at the annealed juncture remain and are filled in 45. P. W. J. Rigby, M. Dieckmann, C. Rhodes, and P. Berg, J M B 113, 237 (1977).

36

I. ROBERT LEHMAN

with pol I so that covalent joining can be accomplished by DNA ligase (46 1.

C. DNA SEQUENCING Pol I is a key reagent in three methods for the sequence analysis of DNA. Each depends upon the ability of pol I to copy a particular sequence of single-stranded DNA starting from the terminus of the annealed primer. 1. Partial Ribonucleotide Substitution (47)

Pol I has the unique ability to incorporate a ribonucleotide in place of the corresponding deoxynucleotide when Mg2+ is replaced by MI?+ (48). The incorporated ribonucleotide then becomes a site of base specific cleavage of the chain by alkali, to yield fragments terminated by that particular ribonucleotide.

2. The Dideoxy dNTP Method (49) Pol I can incorporate the 2’-, 3’-dideoxy analogs of the deoxynucleoside triphosphates into a suitably annealed DNA primer, and in doing so block further chain growth (50). Using a known ratio of analog to each of the natural deoxynucleoside triphosphates results in an accumulation of chains at each point in the sequence at which the nucleotide occurs. As in the ribonucleotide insertion method, a family of fragments terminated with a specific nucleotide is obtained for further sequence analysis. 3 . The Plus-Minus Method (51) This method uses both pol I and the TCinduced DNA polymerase under conditions of limiting deoxynucleoside triphosphates. In the minus reaction, one of the four deoxynucleoside triphosphates is omitted from each of four separate reaction mixtures. Synthesis by pol I proceeds until the point in the sequence at which the missing deoxynucleoside triphosphate is required. For example, in the absence of dATP each chain is terminated at the 3‘ end before an A residue. In the plus reaction, synthe46. I. R. Lehman, Science 186, 790 (1974). 47. W. M. Barnes, J M B 119, 83 (1978). 48. P. Berg, H. Fancher, and M. Chamberlin,ln “Informational Macromolecules” (H. J. Vogel, B. Bryson, and J. 0. Lampen, eds.), p. 467. Academic Press, New York, 1963. 49. F. Sanger, S. Nicklen, and A. R. Coulson, PNAS 74, 5463 (1977). 50. M. R. Atkinson, M. P. Deutscher, A. Kornberg, A. F. Russell, and J. G . Moffatt, Biochemistry 8, 4897 (1969). 51. F. Sanger and A. R. Coulson, J M B 94, 441 (1975).

2. DNA POLYMERASE 1 OF Esrhrrichici coli

37

sis by T4 DNA polymerase, in the presence of a single triphosphate (dATP in this instance) results in all chains terminating at the 3' end with an A residue. A similar reaction is run with each of the other three deoxynucleoside triphosphates. After removal of the primer by cleavage with a restriction endonuclease, the nucleotide sequence of the chain that has been replicated can be deduced from the position of the bands in the eight reaction mixtures in an autoradiograph.

ACKNOWLEDGMENTS This work was supported in part by grants from the National Institutes of Health (GM 061%) and the National Science Foundation (PCM 74-00865).


DNA Polymerase 111 Holoenzyme CHARLES McHENRY

ARTHUR KORNBERG

I. Introduction . . . . . . . . . . . . . . . 11. DNA Polymerase I11 . . . . . . . . . . . . A. Detection and Isolation . . . . . . . . . B. Purification and Identification of Subunits . C. Nomenclature . . . . . . . . . . . . . D. Nuclease Activity . . . . . . . . . . . 111. DNA Polymerase 111 Holoenzyme . . . . . . A. Detection and Isolation . . . . . . . . . B. Purification and Subunit Structure . . . . C. Structural Genes for Holoenzyme Subunits D. Mechanistic Studies . . . . . . . . . . E. Physiological Role . . . . . . . . . . . 1V.Summary.. . . . . . . . . . . . . . . . Note Added in Proof , . , . . . . . . . . .

1.

. .

. . .

. . . .

. . . .

.

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

. . . . . . . .

. . .

. . .

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

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

. . . . . . . . .

. .

. . . .

.

.

.

. . . . .

. . . . . . .

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

39 40 40 41 41 42 43 43 43 46 47 48 49 49

Introduction

In this chapter we consider the structural, functional, genetic, and mechanistic aspects of the various forms of DNA polymcrase 111 of Escherichia coli. These forms are subassemblies of the subunits of DNA polymerase I11 holoenzyme. The holoenzyme has been implicated by both biochemical and genetic criteria as the polymerase responsible for replication of most of the E. coli chromosome. The nature of template-directed 39 THE ENZYMES, Vol. XIV Copyright 0 1981 by Academic Press. Inc. AU rights of reproduction in any form reserved ISBN 0-12-122714-6

40

CHARLES McHENRYANDARTHUR KORNBERG

DNA synthesis by DNA polymerases is reviewed in this volume Chapters 1-7, and elsewhere (I). DNA polymerases have generally been isolated using as an assay the filling of gaps created in duplex DNA by nuclease action. For years DNA polymerase I (pol I) was the only DNA polymerase recognized in E. coli, and was thought to be the polymerase solely responsible for synthesis of theE. coli chromosome. Subsequent studies (2) that led to the isolation of mutations in pol I (PolA), although erroneous in their original conclusion that pol I was dispensable for replication, sparked efforts to search for new DNA polymerases. The two enzymes found were designated DNA polymerase I1 (pol 11) (3-5) and DNA polymerase I11 (pol 111) (6). Beyond identification ofpolB as the structural gene for pol 11, little is known of the physiologic functions of this polymerase. Identification of pol I11 as the product of the dnaE (PofC)gene established it as an enzyme essential for DNA replication (7). Paradoxically, pol 111 demonstrated no unique properties in vitro that might distinguish it as a replicative enzyme. Later on, a more complex physiological form of pol 111, the DNA polymerase 111 holoenzyme, was isolated by using as an assay the in vitro replication of natural chromosomes rather than the repair of gaps in nuclease-damaged DNA. The natural chromosomes used were those of the single-stranded DNA phages M13, G4, and 4x174. The use of these phages to probe E. coli replicative functions is reviewed elsewhere (I, 8). As described in Section 111, the holoenzyme contains a pol I11 core and several auxiliary subunits that permit it to function as a natural replicative enzyme. II. DNA Polymerase 111

A. DETECTION AND ISOLATION Even with the availability of apolA mutant, the discovery of pol 111 was delayed by its lability and inhibition by even moderate levels of salt. Pol 1. A. Kornberg, “DNA Replication.” Freeman, San Francisco, 1980. 2. P. DeLucia and J. Cairns, Nature (London)224, 1164 (1969). 3. T. Kornberg and M. Gefter, BBHC 40, 1348 (1970). 4. R. Moses and C. C. Richardson, BERC 41, 1557, 1565 (1970). 5. R. Knippers, Nature (London) 228, 1050 (1970). 6. T. Kornberg and M. Gefter, PNAS 68, 761 (1971). 7. M. Gefter, Y. Hirota, T. Kornberg, J. Wechsler, and C. Barnoux, PNAS 68, 3150 (1971). 8. R. Schekman, A. Weiner, and A. Kornberg, Science 186, 987. (1974).

3.

DNA POLYMERASE 111 HOLOENZYME

41

I11 was initially detected as a peak of polymerase activity that was eluted at salt concentrations lower than those needed to elute pol I1 from phosphocellulose columns (6). After conditions were found that stabilized pol I11 and optimized its activity, the levels of pol I11 activity in extracts of polA cells approached those ofpol I inpolA+ cells. Maximal activity for pol I11 is attained at pH 7.0 in morpholinopropyl sulfonic acid buffer in the presence of minimal salt concentrations. Pol I11 is inhibited 50% by 50 mM KC1; addition of 150 mM KC1 results in complete inhibition. Ethanol (10%) stimulates pol I11 twofold and partially protects the enzyme from the inhibitory effects of salt (9). General comparisons of the properties of pol I11 with pol I and pol I1 have been reviewed elsewhere ( I ) .

B. PURIFICATION AND IDENTIFICATION OF SUBUNITS Purification attempts by two laboratories indicated that pol I11 contained at least a subunit of 140,000 daltons (10, / / ) . In these purification procedures, pol I11 was monitored by either the standard gap-filling assay (10) or by a complementation assay in which pol I11 stimulates replication by extracts of a dnaE mutant (11). Subsequently, pol I11 was purified 28,000-fold to 97% homogeneity (12). The resulting enzyme is composed of three subunits: a , E , and 8 of 140,000, 25,000, and 10,000 daltons, respectively. The three subunits chromatograph together on phosphocellulose and hydroxylapatite, and remain associated during gel filtration and electrophoresis on native acrylamide gels.

C. NOMENCLATURE DNA polymerase I11 (pol 111) is the term used for the simple gap-filling form of the enzyme originally isolated (6, 9) (Table I). The E and 8 subunits are not resolved from a using the chromatographic procedures described in any purification procedure (12). Thus, pol I11 is the simplest isolated enzyme that contains the dnaE gene product, and appears to be the catalytic core of the holoenzyme. The 140,000 dalton protein (a), which is the dnaE gene product (see Section III,C), will be referred to simply as the a subunit of the DNA polymerase I11 holoenzyme, whether or not it is found 9. T. Kornberg and M. Gefier, JBC 247, 5369 (1972). 10. D. Livingston, D. Hinkle, and C. Richardson, JBC 250, 461 (1975). 1 1 . B. Otto, F. Bonhoeffer, and H. Schaller, EJB 34, 440 (1973). 12. C. McHenry, and W. Crow, JBC 254, 1748 (1979).

42

CHARLESMcHENRYANDARTHURKORNBERG TABLE I

COMPONENTS OF DNA POLYMERASE 111 HOLOENZYME Subunits

Mass (daltons x

a

140

E

25 10 83 52 32 37

9 7

Y 6

P

Alternative designation dnaE protein, polC protein

-

dnaZ protein Factor 111, dnaX protein

Factor I, cop01 III*, dnaN protein

to exhibit any independent enzymatic activity. DNA polymerase I11 holoenzyme (or holoenzyme) is the term used for the complex of proteins that includes the core DNA polymerase 111; it is the intact soluble replicative polymerase responsible for most of the replication of the E. cofi chromosome. Intermediate forms of pol 111 that are subassemblies of pol I11 and holoenzyme auxiliary subunits will be referred to as pol I11 with a superscript (i.e., pol 111', pol III*). Individual subunits, or complexes of them, are referred to as such (i.e., p, y, 6, y . 6).

D. NUCLEASE ACTIVITY Pol I11 has both 3' + 5' and 5' 3' exonuclease activities (13). The 3' 5' activity is specific for single-stranded DNA and removes mismatched nucleotides from the 3' end of an otherwise duplex structure before continuing polymerization from the duplex 3'-OH terminus. Thus, like pol I, pol I11 has the capacity to proofread its product. The 3' -+ 5' exonuclease yields 5'-mononucleotides, but fails to degrade the 5'terminal dinucleotide. Unlike pol I, the 5' + 3' exonuclease of pol I11 is inactive on duplex DNA. It can degrade single-stranded DNA and proceed into duplex regions after degradation has begun. This indicates that pol 111 catalyzes 5' + 3' degradation processively (13). The role of this 5' .--, 3' activity in replication is not clear. It is incapable of removing duplex structures encountered during chain elongation. Possibly the 5' .--, 3' exonuclease is used for other functions, such as repair. The effect of the other holoenzyme subunits on the pol I11 nuclease activities is not known. -+

+

13. D. Livingston and C.

C. Richardson, JBC

250, 470 (1975).

3.

DNA POLYMERASE 111 HOLOENZYME

111.

DNA Polymerase 111 Hoioenzyme

A.

DETECTION AND ISOLATION

43

DNA polymerase 111 holoenzyme was discovered in the investigation of the conversion of single-stranded DNA phages to the duplex replicative form (14). The general reaction scheme for these studies [for a review see Refs. (I, S ) ] are summarized in Fig. 1. The phages M13, G4, and (6x174 differ in the mechanisms of primer formation. M13 DNA, coated with single-strand binding (SSB) protein, depends on the rifampicin-sensitive RNA polymerase; G4 uses the dnaC primase to synthesize its primer; 4x174 can use the dnaC primase only after the prepriming action of six other proteins (n, n’, n”, i, dnaB, and dnaC). Despite the diversity of priming systems, all primers are elongated by the action of the DNA polymerase I11 holoenzyme. DNA polymerase 111 alone will not substitute for holoenzyme, but holoenzyme, like pol 111, has been shown to depend on a product of thednaE (polC)gene (1.5). Early studies indicated that holoenzyme could be resolved by phosphocellulose chromatography into two components, pol III* and copol III* (14, 1.5). Pol IIP was distinguished from pol 111 by its ability to use a long, primed, single-stranded template in the presence of copol 111” and by its greater size and lability.

B. PURIFICATION AND SUBUNIT STRUCTURE The holoenzyme has been purified 7400-fold from HMS-83, apolA-poll3 strain (16). Using high-resolution, denaturing electrophoresis techniques ( I Z ) , 13 bands could be detected in this preparation, sedimenting as an 11 S entity in a glycerol gradient. Before a polypeptide can be regarded as a holoenzyme subunit more evidence than its mere presence in highly purified preparations is needed. Preliminary assignments of subunits have been based upon these criteria: (i) Purification of a subunit to homogeneity and a demonstration that it is essential for reconstituting holoenzyme activity on a natural chromosome. (ii) Physical association of a polypeptide with a known polymerase component in both the holoenzyme and a simpler subassembly of subunits; with this latter criterion, it is also necessary to show that the physical properties of the putative subunit change as 14. W. Wickner and A. Kornberg, JBC 249, 6244 (1974). 15. W. Wickner, R. Schekman, K. Geider, and A. Kornberg, PNAS 30, 1764 (1973). 16. C. McHenry and A. Kornberg, JBC 252, 6478 (1977).

44

CHARLES McHENRY AND ARTHUR KORNBERG

0-

5

SSB

ss

PRIMING

LOENZYME

DNA LIGASE

RF

FIG,1. Requirements for conversion of a single-stranded phage DNA (SS) to the duplex replicative form (RF). a result of the associations, and that the subunit comigrates with the other holoenzyme components on sizing columns and glycerol gradients. (iii) Demonstration that association of a proposed subunit to pol I11 makes the new complex more holoenzyme-like, Definitive evidence that a polypeptide is a subunit should include both the first criterion and genetic proof that the polypeptide is required for replication of the E. coli chromosome in vivo. Four polypeptides (a,/3 ,y , and 6) satisfy these rigorous criteria. Three additional ones ( E , 0, and T ) have been judged to be holoenzyme subunits by the other criteria. The composition of holoenzyme and its subunit subassemblies, based on present information, is as given in Table I. Resolution of holoenzyme by phosphocellulose chromatography generates pol III* and the /3 subunit (37,000 daltons) (16, 17). Alone, each is inactive, but together they reconstitute holoenzyme activity on G4, 4x174, and M13 templates. /3 appears to exist as a dimer upon dissociation from other holoenzyme components (17). Pol 111" can be further resolved to yield a complex of two subunits ( y . 6) by treatment with o-phenanthroline and mild heat (16). This chelator presumably removes an essential metal from pol 111. Although not yet found in pol 111, zinc is an essential component required for activity in a variety of DNA and RNA polymerases. Chelation by o-phenanthroline is likely responsible for rendering pol I11 sufficiently labile to permit its selective heat inactivation and dissociation from y . 6, thus permitting y . 6 to be purified as a distinct molecular species. The y . 6 complex has no known independent enzymatic activity, but together with pol I11 and /3 reconstitutes holoenzyme activity on phage templates (16). The y and 6 subunits have recently been resolved from each other. y was purified to 65% homogeneity from an overproducing strain that carried a plasmid with the dnaZ gene, the structural gene for y (see Section II1,C) (18). Free y was resolved from pol I11 by Blue Dextran Sepharose chromatography. The 17. K . Johanson and C. McHenry JBC 255, 10984 (1980). 18. U. Hiibscher and A. Kornberg, JBC, 255, 11698 (1980).

3. DNA POLYMERASE 111 HOLOENZYME

45

purified subunit exists as a dimer in its native state (18). 6 was purified to 50% homogeneity using, as an assay, its ability to reconstitute holoenzyme activity on a 4x174 template in the presence of pol 111, p, and y

(W.

Evidence that E and 8 subunits are holoenzyme subunits was presented in Section II,B. The polypeptide T (83,000 daltons), a seventh component of holoenzyme preparations, has also been implicated as a subunit (20). Purified as a complex of pol 111, it is termed pol 111’. Addition of T to pol I11 makes pol 111 heavier and more basic, as indicated by a higher affinity for phosphocellulose. When holoenzyme was immunoprecipitated with j3 antibody, T and a were coprecipitated in a 1 : 1 ratio (20). The T polypeptide appears to be a monomer when dissociated from holoenzyme and exhibits an ATPase activity that is dependent on single-stranded DNA (21). The functional properties of pol I11 are also altered by the addition of T. Pol 111‘, resembles holoenzyme, and differs from pol 111 by a slight capacity to use a randomly primed, long, single-stranded template in the presence of spermidine (20). Pol 111’ and holoenzyme are both stimulated by spermidine; by contrast, the low levels of pol I11 activity with these single-stranded templates are further decreased by spermidine. Pol 111’ is far less efficient than holoenzyme in this assay and cannot substitute for holoenzyme in the G4 system. Thus, the addition of T to pol I11 to form pol 111’ makes pol I11 more holoenzyme-like. The T polypeptide is not required for reconstitution of holoenzyme-like activity on single-stranded phage DNA templates (20). Perhaps some of the holoenzyme subunits, required for replication of the complex E. coli chromosome, are not needed for replication of the small phages. These simpler templates may not require all the components of the intact holoenzyme complex after it has been resolved into its constituent parts. For example, pol I11 alone is able to fill in gaps in nuclease-activated duplex DNA. Reports from the Hurwitz laboratory describe the isolation of three factors that, when added to pol 111, replicate single-stranded phage templates (22,231. Elongation Factors I and I11 anddnaZ protein probably correspond to p , 6 and y , respectively (Table I). 19. U. Hiibscher and A. Kornberg, PNAS 76, 6284 (1979). 20. C. McHenry, I n “Mechanistic Studies of DNA Replication and Genetic Recombination” (B.Alberts, ed.) ICN-UCLA Symp. Mol. Cellular Biol., Vol. 19. Academic Press, New York, 1980. 21. R. Meyer, J. Shlomai, and A. Komberg, personal communication. 22. J . Hurwitz and S. Wickner, PNAS 71, 6 (1974). 23. S. Wickner and J. Hurwitz, PNAS 73, 1053 (1976).

46

c.

CHARLESMcHENRYANDARTHURKORNBERG STRUCTURAL GENES FOR

HOLOENZYME SUBUNITS

The a, p , y , and 6 holoenzyme subunits have been identified as the products of replication (dna) genes (Table 11). Conditional lethal replication mutations in these genes have been correlated with defects in the corresponding subunit. Pol I11 and holoenzyme are known to contain the product of the dnaE (PolC) gene. Both pol I11 and holoenzyme are defective when assayed in extracts of temperature-sensitivednaE cells (7, 15). Some mutator mutations have also been mapped in the dnaE locus (24-26). Inasmuch as pol I11 contains three polypeptides (IZ),it was not known which subunit was the dnaE gene product. A 4500 base-pair segment of E. coli DNA that contains the dnaE gene has recently been cloned into the ret gene of pBR322. This plasmid complementsdnaE mutants and directs the synthesis of the Q polypeptide (27). Inserts of DNA into the Hind111 site on the cloned segment abolishes the ability of the plasmid to rescue dnaE mutants and blocks the expression of Q (27) in “maxicells” (28). Therefore, it has been concluded that dnaE is the structural gene for a . A specialized dnaE transducing phage has been constructed that can transduce several different dnaE mutants (29). Lysogens of this phage restore normal pol I11 activity to extracts. Yet, pol 111 is not amplified upon induction of these lysogens (29). Similarly, a colEldnaE hybrid plasmid from the Clarke and Carbon collection rescues dnaE mutants, but does not lead to an overproduction of pol I11 (30). These results may be due to an autoregulatory mechanism, with a! being rapidly proteolyzed if not bound to other holoenzyme components, or to the need for the E and 0 polypeptides for the expression of a activity. The structural gene for y is dnaZ. Early studies indicated that y . 6 contained a product of the dnnZ gene (16). Recent work in which y and 6 were purified independently has demonstrated that y specifically complements extracts from dnaZ cells (/8,3/). Similarly, 6 and p complements extracts from dnaX ( / 9 , 3 2 )and dnaN (33,341 cells, respectively, and are judged to 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34.

C. Sevastopoulos and D. Glaser, PNAS 74, 3947 (1977). E. B. Konrad, J . Bacferiol. 133, 1197 (1978). R. Hall and W. Brammer, Mol. Gen. Genet. 121, 271 (1973). C. McHenry and M. Welch, manuscript in preparation. A. Sancar, A. Hack, and W. Rupp, J . Bacferiol. 137, 692 (1979). H.Shizuya, D. Livingston, and C. C. Richardson, PNAS 71, 2614 (1974). S. Wickner, R. Wickner, and C. Raetz, BBRC 70, 389 (1976). C. Truitt and J. Walker, BBRC 61, 1036 (1974). J. Henson, H. Chu, C. Irwin, and J. Walker, Generics 92, 1041 (1979). P. Burgers, A. Kornberg, and Y. Sakakibara, PNAS, in press. Y. Sakakibara and T. Mizukami, Molec. Gen. Genet. 178, 541 (1980).

3. DNA POLYMERASE 111 HOLOENZYME

47

TABLE I1

CHARACTERISTICS OF DNA POLYMERASE 111 HOLOENZYME MUTANTS MutanP Characteristics

dnaE

dnaZ ~

Subunit Fast stop in DNA synthesis Growth of phages 4x174 h

T7 Role in repair

~

dnaX

dnaN

~~~~

a

Y

+

+

s

P

-

-

N.D. N.D. N.D. N.D.

-

+ -

+

+

+

N.D.

+ -

+ N.D.

N.D., not determined.

be the products of these genes. Thus, these four subunits, which were first identified by biochemical in v i m assays to be holoenzyme subunits, have now been shown by genetic criteria to be required for chromosomal replication.

D. MECHANISTIC STUDIES The specific roles of all the holoenzyme subunits in the replication process are not yet clear. However, preliminary work suggests a possible cycling of certain of the subunits during the replication process (35). Holoenzyme, in the presence of ATP, can form an isolatable initiation complex upon incubation with a primed single-stranded DNA template. Upon formation of this initiation complex, elongation proceeds in the presence of an antibody directed against copol III* (p) preparations. Furthermore, it has been suggested that when isolated separately, elongation Factor I11 (8) and dnaZ protein (y) can transfer elongation Factor I Cp) to the DNA template in an ATP- or dATP-requiring process (36), but the requirement for Factor I @) in further elongation remains undetermined. In later experiments, purified antibody prepared against homogeneous j3 has been shown to inhibit only reactions for which holoenzyme is required (17). Furthermore, as in the early experiments with copol III", the p antibody blocks the formation of an isolatable initiation complex, but not the subsequent elongation (17). Thus, j3 antibody can be used as a tool to 35. W. Wickner and A. Kornberg, PNAS 70, 3679 (1973). 36. S. Wickner, PNAS 73, 3511 (1976).

CHARLES McHENRYANDARTHUR KORNBERG

48

TABLE 111 PROCESSIVITY OF POL Processivity and effects Processivity, residues Spermidine effect On incorporation On processivity SSB" effect On incorporation On processivity

111 AND HOLOENZYME

Pol 111

Holoenzyme

10 to 15

>5000

Inhibition Two- to fourfold decrease

Stimulation Twofold increase

Strong inhibition N.D.b

Stimulation Fivefold increase

Single-strand binding protein N.D.. not determined

block reinitiation so that the processivity of holoenzyme can be determined. Holoenzyme can replicate most of the G4 genome (>SO00 nucleotides; without dissociating from it (37). Other experiments have indicated the processivity of pol I11 to be only 10-15 nucleotides (Table 111). Spermidine and single-strand binding protein, both of which stimulate holoenzyme and inhibit pol 111, increase the processivity of holoenzyme on single-stranded DNA by three- to fivefold (Table 111) (37).

E. PHYSIOLOGICAL ROLE The fact that four subunits of the holoenzyme are encoded by genes in which conditional lethal replication mutations have been found testifies to their participation in DNA replication. All of these mutants promptly stop DNA synthesis at the nonpermissive temperature (Table 11). Where it has been checked, cells with these mutations do not support the growth of phages that require E. coli polymerase functions (e.g., 4x174 and A), but do support the growth of phage T7, which codes for its own polymerase (Table 11). Combining genetic and biochemical evidence, it appears likely that the holoenzyme is responsible for most of the replicative DNA synthesis in E. coli. It is considered likely that the 10 to 20 molecules of the holoenzyme in an E. coli cell are sufficient to sustain the in vivo chain growth of about 1,000 nucleotides per second (I). However, on phage templates in vitro, holoenzyme can neither remove primers nor fill in the 37. P. Fay, K. Johanson, C. McHenry, and R. Bambara, JBC 256, 976 (1981).

3. DNA POLYMERASE I11 HOLOENZYME

49

short gap preceding the 5' terminus of these primers (see Fig. 1) (38).This is analogous to the synthesis of the lagging strand of the E. coli chromosome in which the nascent Okazaki fragments must be connected. Pol I performs this function in vitro and probably does so in vivo as well ( I ) . The dnaE gene product has also been judged essential for one pathway of excision-repair of UV irradiation damage of DNA (39), but the role of other holoenzyme mutations in this pathway has not yet been examined. IV.

Summary

DNA polymerase I11 holoenzyme is a complex, multisubunit enzyme responsible for most of the replicative synthesis in E. coli. It contains a core (pol 111) that can repair short gaps created by nuclease in duplex DNA. Pol I11 contains three subunits: a (dnaE protein), E and 8 . For efficient replication of the long, single-stranded templates that resemble structures encountered at natural replication forks, pol I11 requires the auxiliary subunits p , y , and 6 , encoded by dnaN, dnaZ, and dnaX genes, respectively. Processivity in elongation by the holoenzyme exceeds 5000 residues, compared to a value of only 10 for pol 111. The /3 subunit is required for initiation of replication but antibody directed against it does not block elongation. The intact holoenzyme complex contains at least one other component, a DNA-dependent ATPase, named T . The T polypeptide is not required for reconstitution of holoenzyme activity on singlestranded templates, but alters some properties of pol I11 to make it resemble the holoenzyme. Major uncertainties about the organization and functions of the subunits of the holoenzyme account for our limited understanding of the mechanism of its action. Note Added in Roof

New information is available about the following subunits: /3, 7,and 6 . The /3 subunit is part of the holoenzyme elongation complex in which all /3 antigenic determinants are buried after the initiation complex starts to function (40). The 7 subunit when added to pol I11 (presumably forming pol 111') increases its processivity from about 10 to 50 ( 4 / ) . Spermidine (4 mM 38. 39. 40. 41.

H. Tabak, J . Griffith, K. Geider, H. Schaller, and A. Kornberg,JEC 249,3049 (1974). D. Youngs and K . Smith, Nature New Biol. 244, 240 (1973). K. Johnson and C. McHenry, manuscript in preparation. P. Fay, K. Johanson, C. McHenry, and R . Bambara, manuscript in preparation.

50

CHARLES McHENRY AND ARTHUR KORNBERG

stimulates pol 111’ and increases its processivity to 100, whereas pol I11 is inhibited under these conditions. Pol IIIwhas a processivity of 50 which is increased to 150 by SSB, a protein which inhibits pol I11 and pol 111’ (41). Thus, there is a gradient of processivity among the forms of pol I11 reflecting their structural complexity. The T subunit, a DNA-dependent NTPase, hydrolyzes both AT and dATP to the corresponding diphosphates ; GTP and dGTP are hydrolyzed at 10-20% the rate of ATP. Pyrimidine deoxynucleoside triphosphates are hydrolyzed very slowly (42). Oligo(dA),, is 80% effective as G4 DNA in stimulating this reaction (42). The 6 subunits is a provisional designation for an activity required to reconstitute holoenzyme activity in the replication of single-stranded G4 DNA (43). This activity was previously supplied as an impurity in dnuG primase preparations. Thus, 4 may be the eighth holoenzyme subunit.

42. R. Meyer, D. Rein, and C. McHenry, manuscript in preparation 43. P. Burgers and A. Kornberg, unpublished results.

T-Phage DNA Polym erases I. ROBERT LEHMAN

I. Introduction . . . . . . . . . 11. T4 DNA Polymerase . . . . . . A. Purification and Properties . . B. Reactions Catalyzed . . . . . C. Role of T4 DNA Polymerase in 111. TS DNA Polymerase . . . . . . A. Purification and Properties . . B . Reactions Catalyzed . . . . . C. Role of T5 DNA Polymerase in IV. T7 DNA Polymerase . . . . . . A. Purification and Properties . . B. Reactions Catalyzed . . . . . C. Role of T7 DNA Polymerase in

I.

. . . . . . . . . . . . . . . . . . Vivo . . . . . . . . . . . . . . . . . Vivo . . . . . . . . . . . . . . . . . Vivo . . . .

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

. . . . . .

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

.

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . . . . . . . . .

51 52 52 54 57 60 60 60 61 62 62 63 64

Introduction

Infection of Esclzerichia coli with the T series of bacteriophages (T1 through T7) results in the prompt termination of bacterial DNA replication and, after a brief interval, the onset of bacteriophage DNA replication. Each of these phages with the exception of T1 and T3 is known to induce a new DNA polymerase that is required for synthesis of the bacteriophagespecific DNA. This chapter is concerned with the T4-, T5-, and T7induced DNA polymerases, the enzymes that have been examined in 51 THE ENZYMES. Vol. XIV Copyright 0 19RI by Academic Press, Inc. All rights of reproduction in any form reserved

ISBN 0-12-122114-6

52

I. ROBERT LEHMAN

greatest detail and about which most is known. T4- and T7-induced polymerases very likely exist as part of multienzyme DNA replication complexes in vivo, and indeed act in association with accessory proteins (single-stranded DNA binding protein, primase, etc.) in duplex DNA replication in vitro. A similar situation may well hold for T5-induced DNA polymerase. Because of their close association with other proteins, the properties of the T-phage polymerases within such replication complexes may differ significantly from those of the isolated enzymes.

II. 14 DNA Polymerase

T4 DNA polymerase was the first of the phage-induced polymerases to be isolated in homogeneous form (I 1. It was also the first DNA polymerase to be identified with a specific genetic locus (T4 gene 43) (2, 3). A. PURIFICATION AND PROPERTIES T4 DNA polymerase was first purified from extracts of E. coli B infected with T4 phage bearing an umber (nonsense) mutation in gene 44 (I, 4). In such extracts the specific activity is five- to tenfold higher than in extracts from wild-type T4-infected cells. Nearly homogeneous polymerase is obtained after chromatography on phosphocellulose, DEAE-cellulose, and hydroxyalpatite ; 650 g of infected cells yield approximately 1 mg of pure enzyme. Alberts and colleagues (5) have developed an alternative purification procedure in which extracts enriched in polymerase as well as other replication proteins (the products of genes 44, 45, and 62) are prepared from cells infected with T4 bearing the regA mutation, along with mutations in genes 42 and 30. Using DNA-cellulose, hydroxylapatite, norleucineSepharose, and DEAE-cellulose chromatography, approximately 9 mg of homogeneous polymerase are obtained from 300 g of infected cells. 1. M. Goulian, 2. J. Lucas, and A. Kornberg, JBC 243, 627 (1968). 2. A. de Waard, A. V. Paul, and I. R. Lehman, P N A S 54, 1241 (1%5). 3. H. R. Warner and J. E. Barnes Virology 28, 100 (1966). 4. R. H. Epstein, A. Bolle, C. M. Steinberg, E. Kellenberger, E. B. De La Tour, R. Chevalley, R. S. Edgar, M. Susman, D. Denhardt, and A. Lielausis, C S H S Q B 27, 375 ( 1963).

5. C. F. Morris, H. Hama-Inaba, D. Mace, N. K. Senha, and B. Alberts, JBC 254, 6787 (1979).

53

4. T-PHAGE DNA POLYMERASES TABLE I

COMPARISON OF AMINOACIDCOMPOSITION OF T4 DNA POLYMERASE AND E . coli DNA POLYMERASE I" Amino acid Lysine Histidine Arginine Half-cystine Aspartic acid, asparagine Threonine Serine Glutamic acid, glutamine Proline Glycine Alanine Valine Methionine Isoleucine Leucine Tyrosine Phenylalanine Tryptophan

T4 DNA Polymerase Moles/ 1 14,000 g 85 16 46 15 113 29 65 113 40 61 55 55 35 86 59 41 12 12

E . coli DNA Polymerase I Moles/109,000 g 61 19 48 3 88 53

40 126 53 63 102 61 24 55 112 33 25 9

From Goulian er a / . ( I ) and Jovin et a / . (6).

Like DNA polymerase I (pol I), TCinduced DNA polymerase consists of a single polypeptide. However, its molecular weight, 114,000, is somewhat higher than that of the E. coli enzyme (MW 109,000).Its amino acid composition (Table I) also shows several significant differences from pol I. In particular, the T4 enzyme has 15 half-cystine residues compared to the E coli enzyme, which has only 3. This point is of interest in view of the great sensitivity of the phage-induced enzyme, and the relative insensitivity of pol I to sulfhydryl inactivation by, for example, N-ethylmaleimide (6). T4 polymerase displays a relatively broad pH optimum ranging from pH 8 to 9; at pHs 7.5 and 9.7, approximately 50% of optimal activity is observed. Maximal polymerase activity requires 6 mM M$+; Mn2+at an optimal concentration of 0.1 mM gives a rate approximately one-fourth of that with 6 mM M$+. There is no detectable activity in the absence of a divalent cation (Table 11). 6. T.

M. Jovin, P. T. Englund, and L. Bertsch, JBC 244, 2996 (1969).

54

I. ROBERT LEHMAN TABLE I1

COMPARISON OF PHAGE T4,T5 AND T7-INDUCED

Functions Polymerization: 5‘ + 3’ Exonuclease 3‘ 5’ single strands Exonuclease 3’ .+ 5‘ duplex DNA Template-primer Primed single strands Nicked duplex Activity pH optimum Inhibition by NEM Effect of 200 mM NaCl compared to 50 mM set at 100 Structure Homogeneity Molecular weight Number of subunits Genetic locus

DNA

POLYMERASES“

T4

T5

T7

+

+ + +

+ +

+

+

+

-

+

4

-

+

8-9

8-9

7.6-7.8

5

400

< 10

+

Yes 114,OOO 1 gene 43

+

-

+

Yes

Yes

%,OW

%,ooo

1

2 gene 5 , trxA

gene rs53

Adapted from A. Kornberg, “DNA Replication” Freeman, San Fracisco, 1980.

B. REACTIONS CATALYZED 1. Polymerization

Like all other DNA polymerases, T4 polymerase catalyzes the polymerization of mononucleotide units from deoxynucleoside 5‘triphosphates onto the 3’-hydroxyl terminus of a primer at the direction of a single-stranded DNA template. Thus, fully duplex DNA cannot serve as a template-primer. However, DNA made partially single-stranded by digestion from the 3’ termini with E. coli exonuclease I11 (7) becomes an effective template-primer for the T4 polymerase, and one which under optimal conditions for the enzyme permits a rate of polymerization that approaches the in vivo value (Fig. 1). Single-stranded DNA can also serve as a template-primer for T4 DNA polymerase. The product is a duplex structure in which the newly synthesized strand is covalently linked from its 5’ terminus to the 3‘ end of the template ( I ) . Formation of such a structure by T4 polymerase is plausibly explained by a model in which the single-stranded DNA loops back upon 7. C. C. Richardson and A. Kornberg, JBC 244, 29% (1%4).

55

4. T-PHAGE DNA POLYMERASES SINGLE STRANDED TEMPLATE-PRIMER

EXONUCLEASE 111 TREATED DUPLEX D N A

’1

3 - 5‘ EXONUCLEASE

5u1uwLLIIlLLUvL

5 -

3’

3 T4 POLYMERASE

FIG. 1. Action of single-stranded DNA and exonuclease 111-treated duplex DNA as template-primers for T4 DNA polymerase [adapted from Ref ( I ) . See text for details]. itself, the loop being stabilized by the annealing between regions along the same strand. Any unpaired nucleotides at the 3’ terminus are hydrolyzed by the 3‘ + 5’ exonuclease of the polymerase (see Section II,B,2) until a base-paired terminus is reached (Fig. 1). The template-primer thus formed is analogous to the exonuclease 111-digested duplex and can be replicated up to the 5‘ terminus of the template. Unlike pol I, the T4 polymerase is unable to use a DNA duplex that contains a phosphodiester break (a nick) as template-primer, presumably because of its inability to displace the 5‘-terminated strand at the nick. However, addition of the T4 gene 32 protein (the phage-induced singlestranded DNA binding protein) (4, 8) facilitates strand displacement and allows the T4 polymerase to replicate the nicked duplex. This effect is, however, observed only at low ionic strengths (9). When supplemented with accessory proteins, which include the products of genes 44, 62, and 45, in addition to gene 32, T4 DNA polymerase can initiate replication at a nick at physiological ionic strengths. DNA synthesis begins by covalent addition onto the 3‘-hydroxyl end and continues with strand displacement. 8. B. Alberts and L. Frey, Nature (London) 227, 1313 (1970). 9. N. G. Nossal, JBC 249, 5668 (1974).

56

I . ROBERT LEHMAN

Replication under these conditions is highly processive and proceeds at or near the in vivo rate of 800 nucleotides polymerized per second (10). When supplemented with the products of genes 41 and 61, these proteins can catalyze the initiation of short RNA primers on single-stranded circular DNA templates. The primers can be extended to form long DNA duplexes (10).

2. 3' + 5' Exonuclease T4 DNA polymerase lacks 5' 3' exonuclease activity but contains an extremely active 3' + 5' exonuclease that shows a strong specificity for single-stranded DNA; its turnover number is some 250-fold greater than 5' exonuclease associated with pol I and some 3-fold greater than the 3' the turnover number for polymerase action ( I ) . The products of hydrolysis are deoxynucleoside 5'-monophosphate and a single dinucleotide, derived from the 5' terminus of the polynucleotide. Thus, like E. coli exonuclease I, the 3' + 5' exonuclease of T4 polymerase is incapable of cleaving a dinucleotide (I/). The polymerase and 3' + 5' exonuclease activities associated with the 114,000 dalton DNA polymerase protein cannot be separated by, for example, proteolytic cleavage. However, nonSense mutants in gene 43 have been isolated that cause premature polypeptide chain termination, yielding a protein with a molecular weight approximately 80% that of the native protein (12). The mutant protein retains its exonuclease but not its polymerase activity. 5' exonuclease is The rate of hydrolysis of polynucleotides by the 3' dependent on chain length even at saturating levels of polynucleotide termini. This results from the ability of T4 polymerase to bind to internal nucleotides as well as to the 3'-hydroxyl terminus of a polynucleotide. Binding of the enzyme to internal sites does not result in hydrolysis and as a consequence the internal nucleotides act as inhibitors. Indeed, an increase in chain length of tenfold (from 300 to 3000 residues) results in a decrease in the rate of hydrolysis of approximately 100-fold ( / I ) .

-

-

3. Interaction of Polymerase and 3' -+ 5' Exonuclease Activities In the presence of the complementary deoxynucleoside triphosphates, a template, and a primer, T4 DNA polymerase may act either as a polymerase or exonuclease, depending upon the mode of association of the template and primer. Polymerization occurs when all three of the 10. C. C. Liu, R. L. Burk, U. Hibner, .I. Barry, and B. Alberts, CSHSQE 43,469 (1978). 11. W. M. Huang and I. R. Lehman, JBC 247, 3139 (1972). 12. N. G. Nossal and M. S. Hershfield, JBC 246, 5414 (1971).

4. T-PHAGE DNA POLY MERASES

57

following are available to the enzyme: (i) a polynucleotide template, (ii) a 3'-hydroxyLterminated primer at least one residue shorter than the template, and (iii) the appropriate deoxynucleoside triphosphate (or triphosphates) complementary to the template. In the presence of all of these components the enzyme acts as a polymerase; in the absence of any one, it functions as an exonuclease. Consider a structure in which the 3'-hydroxyl terminus of one of the two strands of a DNA duplex protrudes beyond the 5' terminus of the opposing strand (see Fig. 1). Hydrolysis of the protruding nucleotides by 3' + 5' exonuclease action will proceed until the 3' end occupies a site at least one residue interior to the 5' terminus. At this point all three of the components required for polymerase action are present and the missing nucleotide can be restored. Because fraying at the ends of the duplex creates a transient single strand, the enzyme can once again act as an exonuclease removing a residue from the 3' terminus. The cycle can be repeated again with the enzyme alternating as polymerase or exonuclease in a process that consumes deoxynucleoside triphosphates without net synthesis of DNA.

C. ROLEOF T4 DNA POLYMERASE in Vivo A functional T4 DNA polymerase is essential for the initiation and maintenance of viral DNA replication. Indeed, T4-induced polymerase was the first such enzyme to be directly linked with chromosomal replication ( 2 , 3 ) .It is now clear that T4 polymerase functions in T4 DNA replication as part of a replication complex that, in addition to the polymerase (the gene 43 product), contains six accessory proteins that permit the initiation of new DNA strands by RNA primers, and the replication of duplex DNA at rates near those observed in vivo (10). As previously noted, a chain-terminating (umber) mutant in T4 gene 43 induces the synthesis of a shortened polypeptide that lacks polymerase but retains exonuclease activity. By examining the immunological cross reactivity and molecular weights of the peptides generated following infection of E. coli with a series of amber mutants, a map of the T4 polymerase gene has been constructed (13). The physical map (Fig. 2) is in good agreement with the genetic map determined on the basis of recombination frequencies. From an analysis of the size and cross reactivity of the amber peptides it is clear that the T4 polymerase gene is translated in a counterclockwise direction along the circular genetic map of phage T4 (4). The results of temperature shift experiments with temperature-sensitive mutants in gene 43 have led to the suggestion that the production of a 13. W. M. Huang and I. R. Lehman, JBC 247, 7663 (1972).

58

I. ROBERT LEHMAN GENE 43 GENETIC El92 PHYSICAL MAP

I. I

1

:.'.I

I 100

I

.:I 80

' .

.

')'It

I

I

I

I

40

60

MW

.

.I

'

1

J

-

I 20

I

1

0

10-3

FIG. 2. Comparison between genetic and physical maps of T4 gene 43. The physical map was constructed on the basis of the gene 43 amber peptides as determined by their migration in poiyacrylamide gels in the presence of sodium dodecyl sulfate; the genetic map is that of E. F. Allen, I. Albrecht, and J . W. Drake, Genetics 65, 187 (1970). [From Huang and Lehman ( I - ? ) ] .

functional T4 polymerase, including its associated 3' + 5' exonuclease, involves the energy-dependent conversion of an inactive precursor to active enzymes (14). Thus, when E. coli is infected at 43" with the temperature-sensitive rsL.53 allele of the T4 DNA polymerase gene, T4 DNA polymerase and 3' + 5' exonuclease activities are undetectable in extracts assayed at 30". However, on shifting the culture from 43" to 30" both polymerase and exonuclease activity appear immediately and increase rapidly, even in the presence of sufficiently high concentrations of chloramphenicol, to completely block protein synthesis. Thus, during infection at 43" an inactive polymerase is synthesized that is converted to active enzyme on shifting to 30", a process that does not require protein synthesis. The appearance of active enzyme upon shifting to 30" does, however, require oxidative metabolism since it does not occur in the presence of 2,4-dinitrophenol, an inhibitor of oxidative phosphorylation (15) (Fig. 3). The nature of the energy-dependent conversion of inactive to active T4 tsL.56 DNA polymerase remains an intriguing but unsolved problem. Several, but not all, temperature-sensitive gene 43 mutants cause a large (in one instance up to 2000-fold) increase in reversion frequency of T4 rZZ mutants involving both transitions and transversions; that is, gene 43 acts as a mutator gene (16). Other temperature-sensitive gene 43 mutants produce a decrease in reversion frequency of rZl mutations, especially AT + GC transitions. These mutants, therefore, have the pro14. M. N. Swartz, H. Nakamura, and I. R. Lehman, Virology 47, 338 (1972). 15. J. Thorner, W. M. Huang, and I. R. Lehman, Virology 68, 338 (1975). 16. J. F. Speyer, J. D. Karam, and A. B. Leony, CSHSQB 31, 693 (1966).

4. T-PHAGE DNA POLYMERASES

59

I--

-430-

DNP

A-A-

01 0

' '

10

20

30

40

I

50

I

1

60

70

MINUTES AFTER INFECTION

FIG.3. Time course of appearance of T4tsLS6 DNA polymerase after infection at 43", followed by a shift to 30" in the presence of chloramphenicol (CAM) and in the presence and absence of 2,4 dinitrophenol (DNP). [From Thorner et a / . (IS)].

perties of antimutators. In vitro measurement of exonuclease-polymerase ratios of the purified mutant enzymes have shown the polymerase isolated from cells infected with a mutator mutant (rsL.76) to have a low exonuclease-polymerase ratio relative to the wild-type enzyme, and the polymerase from the antimutator (rsL144) mutant to have a correspondingly high exonuclease-polymerase ratio (17, 18). Inasmuch as the 3' + 5' exocuclease activity of pol I serves a proofreading function by excising mismatched nucleotides during DNA replication in vitro (19), mutation rates seen with these phages reflect, at least in part, the relative polymerase and 3' + 5' exonuclease activities of T4 DNA polymerase during DNA replication in vivo. On the other hand, the DNA polymerase induced by another mutator mutant, T4 rsLB8, shows no defect in the proofreading activity of its associated 3' + 5' exonuclease (20). In this instance, then, the mutator phenotype appears to result from a decrease in the accuracy of nucleotide selection during polymerization. As previously noted, T4 DNA polymerase functions in DNA replication in vivo as part of 17. N. Muzyczka, R. L. Poland, and M. J. Bessman, JBC 247, 7116 (1972). 18. M. J. Bessman, N. Muzyczka, M. F. Goodman, and R. L. Schnaar, J M B 88, 409 (1974). 19. D. Brutlag and A. Kornberg, JBC 247, 241 (1972). 20. F. D. Gillin and N . G. Nossal, JBC 251, 5219, 5225, (1976).

60

1. ROBERT LEHMAN

a multienzyme complex. Its association with accessory proteins within such a complex has in fact been shown to influence the fidelity with which it catalyzes nucleotide polymerization (10).

111.

T5 DNA Polymerase

A. PURIFICATION AND PROPERTIES T5-induced DNA polymerase has been purified to apparent homogeneity by DEAE-cellulose and phosphocellulose chromatography followed by either filtration through Sephadex G-100 or hydroxylapatite chromatography (21, 22). As judged by equilibrium sedimentation and polyacrylamide gel electrophoresis, the enzyme consists of a single polypeptide of molecular weight 96,000 (Z/, 22). Like T4-induced polymerase, the T5 enzyme requires a thiol (2-mercaptoethanol or dithiothreitol) for maximal activity, and is completely inhibited by N-ethylmaleimide. It is however strongly stimulated by 0.2 M monovalent cation, in particular, NI&+ and Na+. At this ionic strength T4 DNA polymerase retains only,about 5% of its activity. T5 polymerase has a pH optimum of 8.5; it is completely dependent upon added Mg2+(Table 11).

B. REACTIONSCATALYZED 1. Polymerization

T5 DNA polymerase can effectively use single-stranded DNA as a template, with either the 3’-hydroxyl end looped back or an oligonucleotide annealed to the chain to serve as a primer in a manner analogous to T4 polymerase (21, 22). Unlike the T4 enzyme, T5 DNA polymerase is able to initiate replication at a nick in duplex DNA even in the absence of accessory proteins (23). Replication at a nick is accompanied by displacement of the 5’-terminated strand, and is strongly influenced by temperature. At 25”, replication of denatured T7 DNA is two- to threefold greater than nicked duplex T7 DNA; at 37” it is nearly tenfold greater. The basis for the difference in behavior of the two types of template-primer is not known. In studies using the homopolymer pair poIy(dA)300.d(T),o as a 21. C. W. M. Orr, S. T. Herriott, and M. J. Bessman, JBC 240, 4652 (1965). 22. R. K . Fujimura and B. C. Roop, JBC 251, 2168 (1976). 23. S . K . Das and R. K. Fukimura, JBC 254, 1227 (1979).

4. T-PHAGE DNA POLY MERASES

61

template-primer, the T5 DNA polymerase is highly processive, with 150160 nucleotides added with each association and dissociation of the enzyme from the template-primer (24). A similar analysis of T4 DNA polymerase gave a processivity value of 12. Thus, replication by T5 DNA polymerase is intrinsically more processive than T4 polymerase. However, as noted previously, association of T4 polymerase with accessory replication proteins can substantially increase its processivity.

2. 3' -+ 5 ' Exonuclease T5 DNA polymerse has a 3' + 5' but not 5' + 3' exonuclease activity. However, it is noteworthy that bacteriophage T5 induces a 5' + 3' exonuclease separate from the polymerase that is essential for T5 DNA replication (25). The 3' + 5' exonuclease associated with T5 polymerase attacks both native and denatured DNA; however, the rate of degradation of native DNA is only one-fifth that of denatured DNA (26). The products of hydrolysis are deoxynucleoside 5'-monophosphates and a small proportion of dinucleotides. Possibly, the latter derive from the 5' termini of the polynucleotide. The optimal conditions for 3' + 5' exonuclease activity (e.g., pH, Mg 2+, thiol reagent, and ionic strength) are the same as those for polymerase action. Like T4 DNApolymerase,TS polymerase can promote the alternate insertion and hydrolysis of nucleotides at the 3' termini of a template-primer, leading to the conversion of deoxynucleoside triphosphates to their corresponding monophosphates without net synthesis of DNA. Although no accessory proteins have been identified, it is almost certain that like T4, replication of T5 DNA will require a multienzyme complex. Indeed, amber mutants of T5 have been isolated that represent at least six different genes that are essential for DNA replication. Several other mutations are known that affect the rate and extent of T5 DNA replication (27). Within such a complex, important features of the polymerase and exonuclease (template preference, processivity, and the balance between nuclease and polymerase) may be significantly altered. C. ROLEO F T5 DNA P O L Y M E R A S Vivo E~ Temperature-sensitive mutants in phage T5 that map in a specific genetic locus (fs.73)fail to synthesize T5 DNA at restrictive temperatures in 24. 25. 26. 27.

S. K. Das and R. K. Fujimura, JBC 252, 8700, 8708 (1977). G. D. Frenkel and C. C. Richardson, JBC 246, 4839, 4848 (1971). S. K. Das and R. K. Fujimura, J. Virol. 20, 70 (1976). H. E . Hendrickson and D. J. McCorquodale, J . Virol. 9, 981 (1972).

62

I. ROBERT LEHMAN

as assayed in vitro. Thus, T5 polymerase is essential for replication of the T5 chromosome (2, 27). The mutant polymerase has been purified to near homogeneity. Examination of its polymerase activity verified the thermolability first observed in partially purified preparation of the enzyme (28). The mutant enzyme is one-fifth as active at 43" as it is at 30", compared with the wild-type polymerase, which is fivefold more active at the higher temperature. In contrast, the 3' + 5' exonuclease activities of the mutant and wild-type enzymes are approximately equivalent at 43". The mutational defect therefore appears to be specifically in the polymerase activity of the T5induced enzyme.

vivo, and induce an abnormally thermolabile T5 DNA polymerase

IV. 17 DNA Polymerase

A. PURIFICATION AND PROPERTIES The DNA polymerase induced upon infection of E. coli with phage T7 has been isolated in nearly homogeneous form by chromatographic procedures similar to those employed for the isolation of the T4- and T5-induced enzymes (e.g., phosphocellulose, DEAE-Sephadex, and hydroxylapatite) (29, 30). The enzyme consists of two subunits: One, encoded by T7 gene 5 , has a molecular weight of 84,000; the other, which has a molecular weight of 12,000, is the bacterial protein, thioredoxin ( 3 / ) . Both subunits are essential for polymerase activity (32,33).Like the other T-phage DNA polymerases, the T7 enzyme has an absolute requirement for Mg 2f and is stimulated by 2-mercaptoethanol. Optimal activity is observed at pH 7.6-7.8 in phosphate buffer (Table 11). The two subunits of T7 polymerase can be separated by dialysis of the enzyme against 6 M guanidine-HC1followed by gel filtration (30).Alternatively, the gene 5 protein, free of the thioredoxin subunit, can be isolated from extracts of an E. coli thioredoxin (trxAf mutant infected with wildtype T7 (34). The larger, phage-specified, subunit retains the singlestranded-DNA-specific 3' + 5' exonuclease activity of the native en28. 29. 30. 31. 32. 33. 34.

R. K. Fujirnura and B. C. Roop, Biochemistry 15, 4403 (1976). P. Grippo and C. C. Richardson, JBC 246, 6867 (1971). S . Adler and P. Modrich, JBC 254, 11605 (1979). T. C. Laurent, E. C. Moore, and P. Reichard, JBC 239, 3436 (1964). P. Modrich and C. C. Richardson, JBC 250, 5515 (1975). D. F. Mark and C. C. Richardson, PNAS 73, 780 (1976). K. Hori, D. F. Mark, and C. C. Richardson, JBC 354, 11591 (1979).

4. T-PHAGE DNA POLYMERASES

63

zyme, but lacks polymerase and double-stranded DNA-specific 3' -+ 5' exonuclease (see Section IV, B,2). No polymerase-associated activities are detectable in the small, thioredoxin, subunit. Reconstitution of the native enzyme can be achieved by incubating the two subunits at 0" for an appropriate period, at molar ratios of thoredoxidgene 5 protein ranging from 12 to 150 depending upon the conditions and the concentration of subunits. The reconstituted enzyme is essentially indistinguishable from the native T7 polymerase in its polymerase and exonuclease activities (30, 34).

B. REACTIONSCATALYZED 1. Polymerization

Either single-stranded DNA or duplex DNA that has been made partially single-stranded by exonuclease action can serve as an effective template-primer for the T7 DNA polymerase; fully duplex DNA is inert. However, when complemented with the T7 gene 4 protein, T7 polymerase can promote the replication of duplex DNA at a nick. Under these conditions, T7 DNA polymerase catalyzes the polymerization of deoxynucleotides, while the gene 4 protein facilitates the unwinding of the duplex coupled to the hydrolysis of ATP or other nucleoside triphosphate (35,36). Moreover, in the presence of ribonucleoside triphosphates and a single-stranded DNA binding protein, the gene 4 protein can catalyze the synthesis of tetranucleotides on the displaced single strand resulting from polymerase action. The extension of the tetranucleotide primers by T7 polymerase then permits synthesis along the displaced strand. Thus, T7 polymerase acting together with the gene 4 protein and a single-stranded DNA binding protein is capable of generating a replication fork in which leading strand synthesis gives rise to a displaced single strand, which then permits lagging-strand synthesis by the generation of oligoribonucleotide primers, and their subsequent extension by deoxynucleotide polymerization (3, 36). 2. 3'

+

5 ' Exonuckases

T7 DNA polymerase has two separate 3' 4 5' exonuclease activities: One is active on single-stranded DNA and the other is active on duplex DNA (29, 30, 34). As previously noted, the active site for the single35. C. C. Richardson. L.. J. Romano, R. Kolodner, J . E. LeClerq, F. Tamanoi, M. J . Engles, F. B. Dean, and D. S. Richardson, CSHSQB 43, 427 (1978). 36. G . Hillenbrand, G . Morelli, E . Lanka, and E. Scherzinger, CSHSQB 43, 449 (1978).

64

I . ROBERT LEHMAN

stranded DNA-specific exonuclease resides entirely within the gene-5coded subunit (30,34); 3’ --* 5’ hydrolysis of duplex DNA like nucleotide polymerization requires the interaction of the gene 5 protein with thioredoxin. The specific activity of the double-stranded DNA-specific exonuclease is approximately twice that of the single-stranded DNAspecific exonuclease. The products of hydrolysis of both single-stranded and duplex T7 DNA are >98% deoxynucleoside 5’-monophosphates (SO). As judged by chromatographic analysis and sensitivity to 5’-nucleotidase, 5’-terminal dinucleotides are not formed. In contrast to the polymerase activity that is unable to initiate polymerization at a nick in duplex DNA in the absence of gene 4 protein, the double-stranded DNA-specific exonuclease can initiate hydrolysis at such nicks, as well as at the 3’-hydroxyl termini at the ends of duplex DNA molecules (30, 34). Under conditions of DNA synthesis, i.e., in the presence of the four deoxynucleoside triphosphates, the double-stranded DNA-specific 3‘ + 5’ exonuclease is inhibited, presumably reflecting the inaccessibility of the 3’ terminal nucleotides of the primer to exonuclease action as a consequence of nucleotide polymerization at the primer terminus (30,34). Similar effects have been noted with the T4 and T5 DNA polymerases. Surprisingly, the addition of one or more deoxynucleoside triphosphates to the gene 5 protein results in inhibition of the single-stranded DNAspecific exonuclease, despite the virtual absence of polymerase activity (30, 34). Similar inhibition has been noted upon addition of ribonucleoside triphosphates. Thus, the inhibition of exonuclease action may not simply reflect competition between synthesis and hydrolysis at the primer terminus. This finding further suggests that the phage-encoded gene 5 protein not only has a site for the 3’-hydroxyl terminus of a single-stranded polynucleotide, but a site for nucleoside triphosphates as well. Although there is no 5‘ 4 3’ exonuclease activity associated with T7 polymerase, gene 6, which is directly adjacent to gene 5 on the T7 chromosome, induces such an exonucleolytic activity (37-39). Like the analogous exonuclease induced by T5 infection, the 5‘ --* 3’ exonuclease specified by the gene 6 protein is essential for T7 DNA replication (39).

C.

ROLE OF

T7 DNA POLYMERASE in Vivo

T7 mutants defective in gene 5 synthesize an altered DNA polymerase and are unable to replicate T7 DNA in vivo (29, 3 2 , 3 9 ) .Similarly, whenE. 37. C. Ken and P. D. Sadowski, JBC 247, 305 (1972). 38. K. Shinozaki and T. Okazaki, Nucleic Acids Res. 5, 4245 (1978). 39. F. W. Studier, Science 176, 367 (1972).

4.

T-PHAGE DNA POLYMERASES

65

coli cells that carry a mutation in the structural gene for thioredoxin are infected with phage T7, neither T7 DNA replication nor active T7 DNA polymerase can be detected (32, 40, 41). Clearly, then, T7 DNA polymerase, a dimer composed of the phage-coded gene 5 protein and the host-specified thioredoxin, plays an essential role in T7 DNA replication. As noted previously, the homogeneous T7 DNA polymerase, while unable to polymerize nucleotides on a duplex DNA template, is able to do so when supplemented with a single-stranded DNA binding protein and the T7 gene 4 protein, an enzyme endowed with helix unwinding and ribooligonucleotide synthetic capabilities.

40. M. J. Chamberlin, J . Virol. 14, 509 (1974). W.Chase, and C. C. Richardson, Mol. G e n . Gene?. 155, 145 (1977).

41. D. F. Mark, I.


Cellular and Viral-Induced Eukaryotic Polymeruses A. WEISSBACH

I. Introduction and Perspective . . . . . . . . . 11. DNA Polymerase a . . . . . . . . . . . , . A. Purification and Properties. . . . . . . . . B. Biological Role . . . . . . . . . . . . . . 111. DNA Polymerase p . . . . . . . . . . . . . A. Purification and Properties . . . . . . . . . B. Biological Role . . . . . . . . . . . . . . IV. DNA Polymerase y . . . . . . . . . . . . . A. Purification and Properties . . . . . . . . . B. Biological Role . . . . . . . . . . . . . . V. Herpes Simplex Virus-Induced DNA Polymerase A. Purification and Properties . . . . . . . . . B. Biological Role . . . . . . . . . . . . . . VI. Vaccinia Virus-Induced DNA Polymerase . . . A. Purification and Properties . . . . . . . . . B. Biological Role . . . . . . . . . . . . . . VII. Conclusion . . . . . . . . . . . . . . . . .

1.

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

. . .

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

. . . . . . , . . . . . . . . . . . . . .

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

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

. .

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

, . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

67 69 69 73 73 74 76 76 77 79 80 80 83 83 84 85

86

Introduction and Perspective

A nomenclature for the known eukaryotic DNA polymerases was proposed in 1975 (/). This classification, which recognized three major 1. A. Weissbach, D. Baltimore, F. J. Bollum, R. C. Gallo, and D. Korn, Scicizcc 190, 401 (1975).

67 THE ENZYMES, Vol. XIV Copyright @ 1981 by Academic Press. Inc. All rights of reproduction in any form reserved

68

A. WEISSBACH

classes of cellular DNA polymerases--cY, /3, and y-has proved to be applicable to a wide range of species in the Animal Kingdom (2). In addition, the nomenclature scheme recognized the existence of a mitochondria1DNA polymerase and new viral-induced DNA polymerases that are produced in infected animal cells. This chapter limits itself to consideration of enzymes that have been well characterized, i.e., DNA polymerases a , P , and y from mammalian cells and the herpes simplex virus and vaccinia virus-induced DNA polymerases. A number of other reported DNA polymerase activities ( 2 4 ) , less well identified, are necessarily omitted. The three cellular DNA polymerases were named in the order of their discovery: DNA polymerase a was first isolated from calf thymus and characterized by Bollum (5, 6), and his work was an important starting point in the study of eukaryotic DNA polymerases. The enzyme has been identified in many species as a predominant activity, and has been purified from rat, murine, calf thymus, and human cells. In 1971 Weissbach et al. (7) reported a new, low molecular weight DNA polymerase in HeLa cell nuclei at the same time that Baril et al. (8)reported an enzyme with similar properties in rat liver nuclei. This enzyme has been designated as ppolymerase. Further identification and characterization of P-polymerase in calf thymus, rat liver, KB cells, and chick embryos followed shortly thereafter (9-11). In general, P-polymerase represents about 5% of the total DNA polymerase in growing, cultured cells, but is a major component of low DNA polymerase activity in quiescent cells. DNA polymerase y , first reported in 1972 by Fridlender et ul. (12), is a widely distributed enzyme (13) that shows a marked ability to copy ribohomopolymers, but a total inability to use natural RNA as a template. It thus has no relationship to the retrovirus reverse transcriptases. At first thought to be a minor activity in the cell, the DNA polymerase y level in 2. A. Weissbach, Annu. Rev. Biochem. 46, 25 (1977). 3. J . A. Kantor, X. H. Lee, J. G . Chirikjian, and W. G . Feller, Science 204, 511 (1979). 4. B. A. Brennessel, D. P. Buhrer, and A. Gottlieb,Anal. Biochem. 87, 411 (1978). 5. F. J. Bollurn, JBC 235, 2399 (1960). 6. M. Yoneda and F. J. Bollurn, JEC 240, 3385 (1965). 7. A. Weissbach, A. Schlabach, B. Fridlender, and A. Bolden,Notlrre New B i d . 231, 167 ( 197 1).

8. E. F. Baril, 0. E. Brown, M. D. Jenkins, and J. Laszlo, Binrhemistry 10, 1981 (1971). 9. L. M. S . Chang and F. J. Bollurn. JBC 246, 5835 (1971). 10. M. E. Haines, A . M. Holrnes, and I. R. Johnston, FEES (Fed.Eur. Biochem. SOC.) Lett. 17, 63 (1971). 11. H. Berger, Jr., R. C. Huang, and J. L. Irvin, JBC 246, 7275 (1971). 12. B. Fridlender, M. Fry, A. Bolden, and A . Weissbach, PNAS 69, 452 (1972). 13. A. Weissbach, Cell S, 101 (1975).

5. CELLULAR AND VIRAL-INDUCED DNA POLY MERASES

69

growing cultured cells is, in fact, equal to that of thep-polymerase, and in at least one case the y-polymerase is the major polymerase in the organism. Table I summarizes some relevant characteristics of the three cellular DNA polymerases obtained from various sources and also lists two wellcharacterized viral-induced DNA polymerases. It should be emphasized here that the properties, size, and behavior of the a-, P - , and y-polymerases can differ from those shown in Table I, depending on the source of the enzyme. The specific patterns shown by inhibitors, however, seem to be invariant. Each of these enzymes is considered in detail in the following sections.

II. DNA Polymerase a

DNA polymerase a has been extensively purified from calf thymus (14, 1 5 ) , human cells (16), murine cells (171, and others (2). Because of the extensive heterogeneity of the enzyme in various species, isolation of a-polymerase in a pure form has been difficult. Nevertheless, a nearhomogeneous preparation of a-polymerase has been obtained from human cells by Fisher and Korn (161, and from mouse myeloma cells by Chen et al. (17). A. PURIFICATION AND PROPERTIES Table I1 summarizes the purification of a-polymerase from cultured human KB cells as described by Fisher and Korn (16). In this procedure all buffers contained 1 rnM P-mercaptoethanol and 1 m M EDTA and, after fraction V, 20% glycerol. A protease inhibitor, p-toluenesulfonyl fluoride, is present when the cells are broken. The second DEAE-cellulose step, which offers little purification per se, is apparently important for subsequent steps. The purified protein from human cells has a specific activity of 206,000 unitdmg, a unit being defined as the incorporation of 1 nrnol of dTMP in DNMhour at 37”. The enzyme, as isolated, exists either as a monomer of 140,000 daltons or as dimers of 265,000-280,000 daltons. It has an isoelectric point of 5.0-5.2 and can be resolved in denaturing polyacrylamide gel 14. 15. 16. 17.

F. J. Bollum, Progr. Nucleic Acid Res. Mol. Biol. 15, 109 (1975). K. McKune and A. M. Holmes, Nucleic Acids Res. 6, 3341 (1979). P. A. Fisher and D. Korn, JBC 252, 6528 (1977). Y. C. Chen, E. W. Bohn, S. R. Planck, and S. H. Wilson, JBC 254, 11678 (1979).

TABLE I

EUKARYOTIC DNA POLYMERASES DNA polymerases

Source Human cells

Major cellular location Nucleus, cytoplasm

Molecular weight (X l t 3 )

Subunits (kilodaltons)

156

76, 66

(KB) Y

Novikoff hepatoma Chick embryo

Herpes simplexinduced Vacciniainduced

Infected HeLa cells Infected HeLa cells

Nucleus

3 I"

Nucleus, mitochondria

180

47

Nucleus

144

74,29

Cytoplasm

115 ~

a

~

~~~

~~~~~~~~

DNA polymerase /3 from other cells has a reported molecular weight of 40,OOO-45,OOO.

Inhibitors Aphidicolin, N-ethylmaleimide, Ara-ATP Dideoxynucleoside triphosphates, iodoacetate Dideoxynucleoside triphosphates, N-ethylmakimide Phosphonoacetate, Ara-ATP, N-ethylmaleimide Phosphonoacetate, N-ethylmaleimide

TABLE I1 PURIFICATION OF

Step Crude extract pH 5.5 precipitation

Ultracentrifugation First DEAE-cellulose

2

Second DEAE-cellulose Phosphocellulose Hydroxylapatite DNA-ceUulose Gel electrophoresis'

Fraction I I1 (resolubilized precipitate) 11' (supernatant) 111 (supernatant) 111' (pellet) IV (adsorbed) IV' (flow-through) V VI VII VIII IX

DNA POLYMERASE a FROM KB CELLS" Velum$ (ml)

Proteid (mg)

Activit9 (units)

9.9 2.2

43 19

880 890

2.8

9

2.9

9 x 10-1

1.1

4 x 10-1 4 x 10-2 1 x 10-2 I x 1 r 3 1.6 x 10-4

3.8 x lo-' 6.6 x lo-* 6.5 x lo-*

60 790 50 490 100 280 240 130

33

Specific activity (unitdmg)

Yield (%)

20 47

(100) 108

88

97

550

67

700 6,000 13,000 33,000 206,000

32 28

I5 4

" As described by Fisher and Korn (16). Reaction mixes contained in 2 5 0 ~ 1 10 , mM Tris, pH 9.2,20 mM mercaptoethanol, bovine serum albumin 200 pglml, 10 m M MgCb, activated salmon sperm DNA, 800 pglrnl, dATP, dCTP, dGTP, and dTTP, 50 g C L M each, and PHIdlTP at a final specific activity of 0.04 Ci/mmol, and enzyme. A unit is the amount of enzyme that catalyzes the incorporation of 1 nmol of labeled dTMP into an acid-insoluble product in 1 hour at 37". Quantities are expressed per gram wet weight of KB Cells. Aliquots, 400 p l , of fraction VIII were used for nondenatunng gel electrophoresis. The protein value was derived by densitometry of a stained gel. Recovery of DNA polymerase activity by elution of slices of parallel unstained gels varied between 50 and 95%. The specific activity value is based on units of loaded activity.

72

A. WEISSBACH

electrophoresis into two subunits of 76,000 and 66,000 daltons. The purified enzyme has a half-life at 0" of 14 months if stored in a concentrated form in the presence of sucrose and potassium phosphate. Optimal reaction conditions include a pH between 7.5-8.5, and Mg'+ at 4-8 mM. Salt concentrations above 50 mM are inhibitory, with about 50% of the activity lost at 100 mM KCl. a-Polymerase is markedly inhibited by Caz+ and Li+. a-Polymerase is most reactive with duplex DNA templates containing gapped regions with available 3'-OH termini (activated DNA). A surprising property of the purified enzyme is its inability to catalyze the synthesis of long DNA chains. It is only slightly processive, synthesizing an 11 2 5 nucleotide length before coming off the template (18, 19). DNA polymerase a does not act at nicks or in short gaps below 20 nucleotides in length, and does not utilize a blunt-ended DNA template. The enzyme binds to single-stranded DNA that contains 3'-OH ends and can catalyze synthesis of hairpin molecules from such templates (20, 21). Synthetic are copied at 20% the rate of DNA templates such as (dA), . (dT)12--IB activated DNA, whereas the corresponding synthetic RNA template, (A), . dTlz, copies at only 3% the rate of activated DNA. However, murine DNA polymerase a copies (dT), . rA2, faster than any other template (22).Spermidine has been found to increase the apparent K , for M. Purified calf DNA (20). The K , for dNTPs is in the range of 1-4 x thymus DNA polymerase a catalyzes both pyrophosphorolysis and pyrophosphate exchange (14). As isolated, purified human a-polymerase has no detectable nuclease activity. There have been reports that bone marrow and calf thymus contain an a-polymerase-like enzyme that contains a 3' + 5' exonuclease, and has been called DNA polymerase 8 (23). Whether this represents a new enzyme or an association of a cellular 3' + 5' exonuclease (24) with the a-polymerase is still unclear. Chen et al. (17) obtained two nearhomogeneous preparations of a-polymerase from mouse myeloma. These 18. P. A. Fisher, T. S-F. Wang, and D. Korn,JBC 254, 6128 (1979). 19. K. McKune and A. M. Holmes, BBRC 90, 864 (1979). 20. P. A. Fisher and D . Korn, JBC 254, 11033 (1979). 21. P. A. Fisher and D. Korn,JBC 254, 11040 (1979). 22. S. H. Wilson, A. Matsukage, E. W. Bohn, Y. C. Chen, and M. Sivarajan, Nucleic Acids RPS.4, 3981 (1977). 23. M. Y. W. Tsang-Lee, C. K. Tan, A. G . So, and K. M. Downey,Biochemistry 19,20% (1980). 24. G. Villani, S. Spadari. S. Boiteux, M. Defais, P. Caillet-Fauquet, and M. Radman, Biochimie 60, 1145 (1978).

5 . CELLULAR AND VIRAL-INDUCED DNA POLYMERASES

73

large molecular weight enzymes (MW = 190,000) contain subunits of 47,000 and 54,000 daltons. One of the a-polymerase species contains both 3' + 5' and 5' + 3' exonucleases associated with it. a-Polymerase has also been isolated in highly purified form from calf thymus (15) and regenerating rat liver; in both cases a number of subunits ranging from 50,000 daltons to 70,000 daltons seem to be associated with a catalytic polypeptide whose molecular weight is about 150,000. Thus, a common denominator in many of these studies is the heterogeneity of DNA polymerase a, a feature which may have important implications in the control and function of this enzyme. B.

BIOLOGICAL ROLE

It is generally agreed that DNA polymerase a has a key role in the replication of nuclear DNA and in the synthesis of the DNA of the viruses SV40, polyoma, and adenovirus (14, 25). Understanding the role of a-polymerase has been aided by the availability of specific inhibitors such as aphidicolin, or the arabinose-containing nucleotides such as ara-ATP. The use of these inhibitors supports the concept that DNA polymerase a is the major replicative polymerase in mammalian cells (26-30). It represents 90-95% of the total DNA polymerase activity of cultured growing mammalian cells and drops to low levels in cells that have ceased nuclear DNA synthesis. 111.

DNA Polymerase /3

DNA polymerase /3 is the smallest of the known eukaryotic DNA polymerases and shows remarkable chemical stability under various conditions. As a result, and although it represents only about 5% of the total DNA polymerase in growing, cultured mammalian cells, it was the first eukaryotic DNA polymerase to be isolated in a homogeneous state. This has been accomplished from calf thymus (3I ), human KB cells (32), mouse 25. 26. 27. 28. 29. 30. 31. 32.

A. Weissbach, ABB 198, 386 (1979). H. J. Edenberg, S. Anderson, and M. L. DeParnphilis, JBC 253, 3273 (1978). M. A. Waqar, M. J . Evan, and J. A. Huberman, Nucleic Acids Res. 5, 1933 (1978). M. Ohashi, T. Taguchi, and S . Ikegami, BBRC 82, 1084 (1978). E. Wist and H. Prydz, Nucleic Acids Res. 6, 1583 (1979). E. Wist, BBA 562, 62 (1979). L. M. S. Chang, JBC 248, 3789 (1973). T. S-F. Wang, W. D. Sedwick, and D. Korn, JEC 250, 7040 (1975).

74

A. WEISSBACH

myeloma (33),Novikoff hepatoma (34) and chick embryos (35). The purification of DNA polymerase /3 from the latter is summarized in Section II1,A. A. PURIFICATION AND PROPERTIES The procedure used by Stalker et al. (34) is shown in Table 111, and yields a homogeneous enzyme after a 200,000-fold purification with a remarkable apparent yield of 46%. The starting material, Novikoff hepatoma, is an ascites tumor with a generation time of 12 hours when maintained in rats, so relatively large quantities of cells can be obtained conveniently. The purification relies on the sequential use of three chromatographic separations on DEAE-Sephadex, phosphocellulose, and hydroxylapatite. The final step in the purification procedure uses singlestranded DNA cellulose as an affinity column, which in a 25-fold enrichment step provides the pure enzyme. The enzyme is stable at 4" during the isolation procedure, and stabilization of the enzyme during purification is facilitated by the use of 10% glycerol in the elution buffers and, at the final step, by having bovine serum albumin (1 mg/ml) present. With this method, 200 pg of purified DNA polymerase P are obtained per kilogram of cells. The enzyme in whole cells is stable at - 20" for months, and the purified enzyme has been stored for 1 year at - 196" without loss of activity. The enzyme, as isolated, has a molecular weight of 31,000 although the calf thymus (31), KB cell (32), and chick (35) and mouse P-polymerases (33) have been reported to have molecular weights of 44,000, 43,000 and 40,000 daltons, respectively. Purified DNA polymerase /3 has no detectable nuclease activity. It shows an alkaline isoelectric point (8.51, a pH optimum of 8.4-9.2 and a K, for deoxynucleoside triphosphates of 7-8 p M . For maximal synthesis Mg2+ at 5-10 mM is required; MI?+ (1 mM) can also be used. The enzyme is stimulated twofold by 50 mM NaCl or by 100-200 mM KC1. The latter salt levels severely inhibit a-polymerase. Phosphate and pyrophosphate are inhibitory to /3-polymerase and should be avoided in reaction mixes. Neither the mouse nor human enzyme can catalyze pyrophosphate exchange, pyrophosphorolysis, or dNTP turnover (33, 36). 33. K. Tanabe, E. W. Bohn, and S. H. Wilson, Biochemistry 18, 3401 (1979). 34. D. M. Stalker, D. W. Mosbaugh, and R. R. Meyer, Biochemistry IS, 3114 (1976). 35. M. Yamaguchi, K. Tanabe, Y. N . Taguchi, M . Nishizawa, T. Takahashi, and A. Matsukage, JBC 255, 9942 (1980). 36. T. S-F. Wang, W. D. Sedwick, and D. Korn, JBC 249, 841 (1974).

75


TABLE I11

PURIFICATION OF NOVIKOFF HEPATOMA DNA POLYMERASE P" Protein (mg)

Total unitsh

Specific (unitdmg)

Purification (-fold)

Yield

Fraction 1. Cell extract 11. Ammonium sulfate 111. DEAE-Sephadex IV. Phosphocellulose V. Hydroxylapatite VI. DNA-Cellulose

14,500 4,610 8,866 81.2 1.34 0.031

3,880 3,780 9,820 4,390 3,150 1,800

0.268 0.820 11.3

1.0 3.06 42.2 202 8,770 217,000

100 97.4 253

54.1

2,350 58,100'

(%)

113

81.2 46.4

' I From Stalker et al. (35).The reaction mixtures contained the following components in a final volume of 125 111: 25 mM Tris-HCI, pH 8.4; 5 mM 2mercaptoethanol; 7 mM magnesium acetate; 0.5 mM EDTA; 0.015 m M each of dATP, dCTP, dGTP, and PHIdTTP (specific activity 975 rnCi/mmol); 50 m M NaCI; 15% ( w h ) glycerol; 250 Fg/ml activated DNA; and 0.01-0.3 units of DNA polymerase fraction. Incubations were carried out for 1 hour at 37" and acid-insoluble radioactivity was determined. When incorporation was not linear for 1 hour, the data were extrapolated from a 30-minute incubation. * A unit is defined as the incorporation of 1 nmol of total nucleotide into DNA per hour at 37". ' With several different preparations, the specific activity varied from 32,000 to 62,000 unitslmg.

A reported characteristic of DNA polymerase /3 is its relative insensitivity to urea, acetone, and alcohol (14). The enzyme is stabilized by glycols and stimulated by spermidine (up to 10 mM) (34). Another general property of the P-polymerases is their relative resistance to N-ethylmaleimide (NEM), which is a powerful inhibitor of DNA polymerases a and y . At 4 mM, NEM shows a 28% inhibition of DNA polymerase @ from Novikoff hepatoma, a value that is slightly higher than previously reported for the human enzyme (37). The NEM partial inhibition is not unexpected since p-hydroxymercuribenzoate inhibits @-polymeraseat concentrations above 50 p M (14, 34). An important characteristic of P-polymerase is its ability to copy a synthetic ribohomopolymer such as (A), . dTlz as well as the corresponding deoxyribohomopolymer (dA), . dTlz or activated DNA (34).This is in contrast to a-polymerase, which utilizes the deoxyribohomopolymer (dA), . dT,,_,, eight times better than (A), . dTlz, which is, in fact, copied at only 3% the rate of activated DNA (18). Rat DNA polymerase 0 has been reported to have a uniquely high requirement for primers when 37. K. W. Knopf, M. Yamada, and A. Weissbach, Biochemistry 15, 4540 (1976).

76

A. WEISSBACH

copying poly(A) templates, and can thus be distinguished from y-polymerase or oncornavirus reverse transcriptase (38). Steady-state kinetic measurements suggest an ordered BiBi mechanism for polymerization and a scheme depicting two DNA binding sites on the enzyme has been advanced (33).Although the specific activity of the purified Novikoff hepatoma DNA polymerase p prepared by Stalkeret al. (S4)is 58,000 with activated DNA as a template, Ono et al. (38)reported rat ascites hepatoma DNA polymerase preparations with a specific activity of lo6 units/mg on an (A), * dTlz-ls template.

B. BIOLOGICAL ROLE The level of DNA polymerase L,3 in quiescent or growing cells or during the cell cycle has been reported to be relatively constant, leading to the suggestion that it may be involved in DNA repair synthesis (39). Hiibscher et al. (40) have shown that &polymerase can participate in the repair of UV-damaged DNA in neuronal nuclei, an organelle in which DNA polymerase p is the only detectable polymerase activity. The further role of this enzyme in other types of DNA synthesis is unknown at the present time.

IV.

DNA Polymerase y

DNA polymerase y exists in at least two forms and is found in the nucleus, cytoplasm, and mitochondria (41-43). Like a-polymerase, DNA polymerase y readily undergoes reversible aggregations that, in v i m at least, are salt-dependent. It comprises about 5% of the total DNA polymerase activity in the growing, cultured mammalian cell, and therefore is about equal to the &polymerase level. In developing chick embryos, y-polymerase represents 45% of the total DNA polymerase activity and is present in larger amounts than either 0- or P-polymerases (44). It is clear that the so-called “mitochondrial” DNA polymerase is one species 38. K. Ono, A. Ohashi, K. Tanabe, A. Matsukage, M. Nishizawa, and T. Takahashi, Nucleic Acids Res. 7, 715 (1979). 39. G. Pedrali Noy, L. Dalpra’, M. A. Pedrini, G. Ciarrocchi, E. Gidotto, F.Nuzzo, and A. Falaschi, Nucleic Acids Res. 1, 1183 (1974). 40. U. Hiibscher, C. C. Kuenzle, and S. Spadari, PNAS 76, 2316 (1979). 41. S. Spadari and A. Weissbach, JBC 249, 5809 (1974). 42. G. Pedrali Noy and A. Weissbach, BBA 477, 70 (1977). 43. A. Bolden, G. Pedrali Noy, and A. Weissbach, JBC 252, 3351 (1977). 44. M. Yamaguchi, A. Matsukage, and T. Takahashi, JBC 255, 7002 (1980).

5 . CELLULAR AND VIRAL-INDUCED DNA POLY MERASES

77

of the y-polymerase class, and that the nuclear species of DNA polymerase y can be distinguished from it (43). Despite extensive efforts, the enzyme has not been prepared in pure form from mammalian tissues (45), perhaps due, in part, to its heterogeneity: but it has been purified to near-homogeneity from chick embryos (44). A.

PURIFICATION AND PROPERTIES

An outline of the purification of the y-polymerase from chick embryos as described by Yamaguchi et al. (44) is shown in Table IV. In this procedure, frozen 11-day-old embryos are minced and sonicated in a buffer containing 0.5 M KCl, 10% glycerol, and eventually, 0.5% Triton X-100. The purification scheme uses two phosphocellulose column chromatographic steps, a Sephadex G-200 gel filtration, and hydroxylapatite adsorption chromatography. Following the second phosphocellulose column, the enzymatic activity separates into two gel components, a 180,000- and a 280,000-dalton species, during gel filtration on a Sephadex G-200 column, and each species is further purified separately. The final separation step, which gives a 1000-foldenrichment, involves affinity chromatography on a double-stranded DNA cellulose column, and can be compared to the single-stranded DNA cellulose columns used in the purification of a- and P-polymerases. Attempts to purify DNA polymerase y by d n i t y chromatography on poly(rA)-Sepharose columns leads to inactivation of the enzyme. The purified enzyme can be stored at -80" but loses 50% of its activity in one freeze-thaw cycle. The purified enzyme sediments at 7.5 S and the molecular weight is estimated to be 180,000. SDS polyacrylamide gel electrophoresis shows a prominent polypeptide at 47,000 daltons, so the native enzyme appears to be a tetramer of this subunit. Based on this, the specific activity of the purified enzyme is calculated to be 660,000 unitslmg on a poly(A) template. With (A), dTlz-ls as a primer-template, the K , value for dTTP is about 1 p M, the optimal pH 8.5-9.0, and the optimal KCI concentration is 220 p M. In the presence of increasing levels of potassium phosphate, the optimal KCI concentration drops proportionately (45); and Mn2+ at 0.5-0.6 mM is fivefold more effective than the optimum Mg2+concentration of 12 mM. The structure of native DNA polymerase y in mammalian cells will probably differ somewhat from the avian enzyme. Rat liver DNA polymerase y can be obtained as a 4 S species (60,000 daltons) (43), whereas the smallest species of the chick native enzyme sediments at 7.5 45. K-W. Knopf, M. Yamada, and

A. Weissbach, Biochemistry

15, 4540 (1976).

TABLE IV

PURIFICATION O F DNA POLYMERASE y FROM CHICK

Step Crude extract First phosphocellulose and ammonium sulfate fractionation Second phosphocellulose Sephadex G-200 Hydroxylapatite Double-stranded DNA cellulose

Fraction I

I1

EMBRYOS~

Protein

Activityb units

(mg)

(%)

0,38 3s

80 4.8 4.1 x 10-' 1.0 x lo-'

I11 IV- 1 IV-2 v-1 v-2 VI- 1

9.0 x

4.1(14) 5.0(17) 3.6(12)

(VI-1-dT VI-2

(1.3 x 10-7 1.3 x lCV

(4 4.5( 15)

9.0 x 1C2 8.4 x 10-3 8.1 x 10-3

specific activity (unitdmg)

2x77) 8.4(28) 9.0(30)

56 84 100 490 620 400.000

Purification (-fold) 1 9.2 150

220 260 1,300 1,600 1,100,000 (1,500,000) 920,000

From Yamaguchi et al. (44). The assay mixture contained in 25pl,50 mM Tns, pH 8.5, 1 mM dithiothreitol, 0.5 mM MnC&.,80 pglml poly (rA) 16 pg/mI dT,,-,,, 0.1 mM [3H]dTTP(60cpdpmol), 15% glycerol, 400 pg/ml bovine serum albumin, 100-120 mM KCI, 20-40 mM potassium phosphate (pH 8.5) and enzyme. A unit is the amount of enzyme that catalyzes the polymerization of 1 nmol of dTMP in 60 minutes.

* Quantities are expressed per gram wet weight of

11-day-old chick embryos.

' Fractions in the peak of DNA polymerase activity (see Fig. 5A).

S. CELLULAR A N D VIRAL-INDUCED DNA POLY MERASES

79

S (150,000-180,000 daltons). By contrast, HeLa cell DNA polymerase y can be separated into two species on phosphocellulose chromatography, both with the same or similar apparent molecular weight of 110,000 (do), and human lymphoblast DNA polymerase y has a reported molecular weight of 120,000 (46). The interspecies difference is further illustrated by the report that sea urchin DNA polymerase y has a sedimentation value of 3.3 s (47). A salient feature of y-polymerases is their ability to copy ribohomopolymers at a rate greater than activated DNA. Under proper conditions the HeLa cell DNA polymerase y will utilize (A), * dT12-18five to ten times more efficiently than activated DNA (44). This is in contrast to the template characteristics of a-polymerase, which utilizes this synthetic template at 3% the rate at which it uses activated DNA. In addition, y-polymerase is active at potassium phosphate concentrations (50 mM) that are inhibitory to DNA polymerase /3(14,45), an enzyme that is known to copy (A), . dTlz-18 at about the same efficiency it copies activated

DNA. B. BIOLOGICAL ROLE Of the three cellular DNA polymerases, only DNA polymerase y is capable of synthesizing continuous long DNA chains in a processive manner (48). DNA polymerase a , by comparison, is highly discontinuous, polymerizing 10-15 nucleotides at a time and then leaving the template (18,49).,&Polymerase also shows discontinuous synthesis when copying a poly(A) template (49). The ability of the y-polymerase to carry out a processive and continuous synthesis of a DNA chain may explain its known physiological roles. One of the forms of the enzyme is responsible for mitochondrial DNA synthesis (43, 50, 51) and another, the nuclear y-polymerase, is involved in the replication of adenovirus DNA (52, 53). The synthesis of adenovirus DNA and mitochondria1 DNA share a strand-displacement step in their replication process; this has led to the 46. M. Robert-Guroff, A. W. Schrecker, B. J . Brinkman, and R. C. Gallo, Biochemistry 16, 2866 (1977).

47. A. Habara, H. Nagano, and Y. Mano, BBA 561, 17 (1979). 48. M. Yamaguchi, A. Matsukage, and T. Takahashi, Nature (London) 285, 45 (1980). 49. A. Matsukage, M. Nihizawa, T. Takahashi, and T. Hozumi, J . Eiochern. (Tokyo),in press (1980). 50. U. Hubscher, C. C. Kuenzle, and S . Spadari, PNAS 76, 2316 (1979). 51. W. timmemann, S-M. Chen, A. Bolden, and A. Weissbach,JBC 255, 11847 (1980). 52. P. C. Van der Vliet and M. M. Kwant, Nature (London) 276, 532 (1978). 53. H. Krokan, P. SchaEer, and M. L. DePamphilis, Biochemistry, 18, 4431 (1979).

80

A. WEISSBACH

suggestion that y-polymerase has a unique role in strand-displacement syntheses (52, 25). Since both mitochondria1 DNA and adenovirus DNA are synthesized in a continuous mode without the apparent formation of short intermediates, such as Okazaki fragments, the processive character demonstrated by y-polymerase in vitro is also reflected in vivo . However, it is apparent that the basic physiological role of DNA polymerase y in the nucleus of the cell remains unknown. V.

Herpes Simplex Virus-Induced

DNA

Polymerase

The recognition in 1963 that herpes simplex virus (HSV) induced a new DNA polymerase in infected cells (54, 5 5 ) followed shortly after the discovery of DNA polymerase a , and predates the identification of DNA polymerases p and y . The HSV-induced DNA polymerase is therefore one of the earliest eukaryotic DNA polymerases studied. The altered properties of the enzyme were recognized by Keir et al. ( 5 3 , and the enzyme was partially purified and characterized by Weissbach er a!. (56). Highly purified, near-homogeneous preparations of the HSV polymerase have been prepared from HSV-1-infected HEp-2 cells (57) and from African green monkey cells (58). Purification of the viral-induced enzyme is facilitated by the large amounts of virus that are produced in the infected cell (59). Thus, the amount of HSV-1 DNA polymerase in HSV-1-infected HeLa cells can rise to four times the combined level of all the host cell DNA polymerases. A. PURIFICATION AND PROPERTIES As described by Knopf (58), African green monkey cells (RC-37; Italdiagnostic Products) grown in monolayers were infected at 5 pfdcell with HSV-1 (Angelotti) that had previously been passed through RC-37 five times. Six hours after infection the cells were collected, disrupted by sonication in 0.25 M potassium phosphate, pH 7.5, containing 0.5% Triton X-100. All the subsequent purification steps shown in Table V were per54. H. M. Keir, J. Hay, J. M. Momson, and J. Subak-Shape, Nature (London)210, 369 (1966). 55. H. M. Keir, J. Subak-Shape, W. I. H . Shedden, D. H. Watson, and P. Wildy, Virology 30, 154 (1966). 56. A. Weissbach, S-C. L. Hong, J. Aucker, and R. Muller, JBC 248, 6270 (1973). 57. K . L. Powell and D. J . M. hrifoy, J . Vlrol. 24, 616 (1977). 58. K. Knopf, EJB 98, 231 (1979). 59. M. Yamada, G. Brun, and A. Weissbach, J . Virol. 26, 281 (1978).

TABLE V PURIFICATION OF

HSV-1-DNA POLYMERASE FROM INFECTED RC-37 CELLP ~~

Purification

Volume (ml)

Total protein (mg)

Total activity (units)

Specific activity (unitdmg protein)

Purification

(%)

Cell extract dialysate DEAE-cellulose Phosphocellulose DNA-cellulose DNA-cellulose peak (fraction 37)

370 575 247 30 0.9

1191.4 217.8 28.5 1.38 0.033

125,280 150,480 95,168 3 1,570 1,575

105.2 690.9 3339.2 22876.8 47727.3

1 6.6 31.7 217.5 453.7

100 120 76 25

~~

Total recovery

~

From Knoff (58). Reaction mixtures contained in lOOpl50 mM Tns-HCI (pH KO), 7.5 mM MgCl,, 100 m M ammonium sulfate, 5 0 p g bovine serum albumin, 0.5 mM dithiothreitol, 0.1 mM each of dATP, dCTP, dGTP, and PHld'lTP (0.4 Ci/mmol), and 25 pg of activated salmon sperm DNA prepared as described by Pedrali Noy and Weissbach (42).A unit is the amount of enzyme that catalyzes the polymerization of 1 nmol of nucleotide in 60 minutes under standard assay conditions. a

82

A. WEISSBACH

formed with buffers containing 0.5 mM dithiothreitol and 1 mM phenylmethylsulfonyl fluoride. The purification is relatively simple and involves three chromatographic separations on DEAE-cellulose, phosphocellulose, and double-stranded DNA cellulose, which yield a highly purified preparation after only a 450-fold purification. Using similar steps with DEAE-cellulose, phosphocellulose, and single-stranded DNA-cellulose separations, Powell and Purifoy (2 years prior to Knopf's report) purified the HSV-induced polymerase from HEp-2 cells almost 1700-fold with almost a 50% recovery (57). The purified enzyme, stored in 50 mM TrisHC1, 1 mM EDTA in 50% glycerol is stable at - 20 or -70". As isolated by Knopf, the purified enzyme shows a major polypeptide of 144,000 daltons on SDS polyacrylamide gel electrophoresis, which is in agreement with the 150,000-dalton species found by Powell and Purifoy (57). The enzyme isolated from RC-37 cells also shows the presence of two other polypeptides of 74,000 and 29,000 daltons, which were not observed by Powell and Purifoy and which may represent impurities. A prominent feature of HSV-DNA polymerase, and one that facilitates its identification, is its activity at high salt concentrations. The presence of 150 mM KCI or 100 mM (N&)2S04 leads to a two- to threefold enhancement of the enzymatic activity, whereas the cellular a-polymerase is inhibited nearly 90% at these salt concentrations. The HSV-1-induced DNA polymerase has a pH optimum of 8-8.5, and a M$+ optimum of 3 mM (in the presence of activated DNA template). Dithiothreitol ( 5 mM) also stimulates the enzyme threefold. The enzyme is inhibited by Zn2+, N-ethylmaleimide, and the pyrophosphate analogs, phosphonoacetic acid, or phosphoformate (60-62). Inorganic pyrophosphate does not inhibit the enzyme, which is able to catalyze pyrophosphate exchange into dNTPs. Aphidicolin, a powerful inhibitor of DNA polymerase a , also inhibits the HSV-induced DNA polymerase as well as the vaccinia-induced DNA polymerase described in Section VI (63). It has been observed that any inhibitor of DNA polymerase a also inhibits the HSV-induced DNA polymerase and the vaccinia-induced DNA polymerase, and vice versa (21). Since there is no known relationship or structural similarity between these viral-induced enzymes and DNA polymerase a , it will be of consid60. A. Bolden, J. Aucker, and A. Weissbach, J . Virol. 16, 1584 (1975). 61. S . Leinbach, J. M. Reno, L. Lee, A . F. Isbell, and J. A. Baezi, Biochtrnistry 15, 426 (1976). 62. B. Eriksson, A. L&rsmn,E. He!gstrand, N. G. Johansson, and B. Oberg, BBA607,53 (1980). 63. G . Pedrali Noy and S . Spadari, J . Virology 36, 457 (1980).


83

erable interest to elucidate the active sites of these enzymes and compare them. The HSV-DNA polymerase contains a 3' + 5' exonuclease activity that copurifies with the enzyme and is apparently an intrinsic activity. This is in contrast to the purified host-cell DNA polymerases, which are devoid of nuclease activity in their most purified form, although preparations of DNA polymerase (Y with nuclease activity have been reported (22, 2 3 ) . Whether the exonuclease serves as a "proof-reading'' activity, as has been postulated for E. coli DNA polymerase I (64) and T4-DNA polymerase ( 6 3 , remains to be determined.

B. BIOLOGICAL ROLE Herpes virus contains a relatively large genome of about 10' daltons. A genome of this size would be expected to code for 100-150 proteins, and it would not be surprising if one of these proteins might be a new DNA polymerase. Genetic evidence for this exists since certain viral DNA negative mutations are located at the chromosomal site that determines the DNA polymerase expression (66, 67). It thus appears self-evident that HSV-induced DNA polymerase is required for synthesis of the viral DNA. In addition, almost all other members of the herpes group seem to induce a new DNA polymerase in host cells after infection (25).

VI.

Vaccinia Virus-Induced DNA Polymerase

The pox viruses, of which vaccinia virus is a member, are among the largest viruses and contain DNA genomes of 1.2-2 x 10' daltons. Jungwirth and Joklik (68) and Magee and Miller (69) suggested in the 1960's that vaccinia virus could induce a new DNA polymerase in infected cells. This viral-induced DNA polymerase was partially purified by Berns er al. (70), and later clearly separated from the host DNA polymerases by 64. M. P. Deutscher and A. Kornberg, JBC 244, 3019 (1969). 65. M. S. Hershfield and N . G. Nossal, JBC 247, 3393 (1972). 66. P. Chartrand, C. S . Crumpacker, P. S . Schaffer, and N. M. Wilkie, Virology 103,311 (1980). 67. L . E. Schnipper and C. S. Crumpacker, PNAS 77, 2270 (1980). 68. C. Jungwirth and W. K. Joklik, Virology 27, 80 (1965). 69. W. E. Magee and 0. V. Miller, Virology 31, 64 (1967). 70. K. I. Berns, C. Silverman, and A. Weissbach, J . Virol. 4, I5 (1969).

84

A. WEISSBACH TABLE VI

PURIFICATION OF

VACCINIAVIRUSDNA POLYMERASE"

Fraction

Activity (units x lo-'])

Protein (mg)

Specific activity (unitdmg)

55 28 6.6 3.9 2.8 1.7

1,495 268 10.9 1.3 0.52 0.089

36 104 610 2,800 5,400 19,000

I. Extract' 11. DEAE-cellulose'

111. IV. V. VI.

DNA-agarose Phosphocellulose Hydroxylapatite Glycerol gradient

From Challberg and Englund (72). A unit is the amount of enzyme that catalyzes the incorporation of 1 nmol of total nucleotide into an acid insoluble form in 30 minutes at 37". ' Activity in Fractions I and I1 includes both vaccinia and host polymerases.

Citarellaet al. (71). It has been purified to near homogeneity from infected HeLa cells by Challberg and Englund (72). A. PURIFICATION AND PROPERTIES Because of the relatively large amount of viral-induced DNA polymerase formed in the infected cell ( 7 / ) , Challberg and Englund (72) were able to isolate 100 p g of purified enzyme from 27 g of vacciniainfected HeLa cells. Vaccinia-infected HeLa cells, obtained 53 hours after infection, and stored at -2W, were broken by Dounce homogenization in 10 volumes of 10 mM NaC1, 2 mM Tris, pH 7.6, 0.1 mM benzamidine. The lysate was clarified by centrifugation at 15,OOOg, and the supernatant fluid containing 13% glycerol and 4 m M diisopropyl fluorophosphate (DFP) was incubated 1 hour at 0" and applied to a DEAE-cellulose column. The outline of the further purification procedure is shown in Table VI, and is unique in that the DNA affinity column step is performed before the phosphocellulose and hydroxylapatite steps. Elution of the enzyme activity in each chromatographic separation utilizes buffers containing 10% glycerol and yields about a 50% recovery of enzymatic activity in each step. The final step of the preparation yields an enzymatic activity that is at least 500-fold purified from the crude cytoplasmic fraction and is 95% homogeneous. 71. R. V. Citarella, R. Muller, A. Schlabach, and A. Weissbach, J. Virol. 10, 721 (1972). 72. M. D. Challberg and P. T. Englund, JBC 254, 7812 (1979).

5. CELLULAR AND VIRAL-INDUCED DNA POLYMERASES

85

The vaccinia DNA polymerase activity in the infected cells is stable for one month at -20” and is stable for 24 hours at 0” in the cytoplasmic extract (fraction I). The most purified preparations (fractions IV and VI) are stable for months at -20”. In the absence of protease inhibitors, such as DFP and benzamidine, proteolysis of the enzyme during purification occurs even at the phosphocellulose step. Native vaccinia-DNA polymerase is a single polypeptide with a molecular weight of 110,000-1 15,000. It is maximally active in the presence of 5 mM MgC1, and shows a pH optimum in 50 mM potassium phosphate, at 8-9. Its activity in Tris-HC1 at the same pH is 10% that shown in potassium phosphate buffers. The enzyme requires the presence of SH groups and is inhibited by 10 mM N-ethylamaleimide or 30 pM p chloromercuribenzoate. In contrast to the herpes simplex-induced DNA polymerase, the vaccinia-DNA polymerase is inhibited by salt (50% at 200 mM NaCl). The vaccinia DNA polymerase shows maximal activity in an activated DNA template, but will neither nick-translate nor strand-displace a nicked 4x174 DNA template. The enzyme seems sensitive to the secondary structure of the template since in copying 4x174 templates it pauses at regions that contain potential hairpin structures (73). As previously reported (71), the purified polymerase contains a strong exonuclease activity that is apparently part of the DNA polymerase polypeptide since both the polymerase and nuclease activity show the same kinetics of heat inactivation at 45”. The intrinsic nuclease activity is a 3’ + 5’ exonuclease that produces 5’-mononucleotides. Although the pH optimum of the exonuclease, 8-9, is similar to the pH optimum of the polymerase activity, the nuclease activity is twice as active in Mn2’ (50 pM MnC1,) as in the optimum MgCl, concentration (10 mM). In addition, the exonuclease is twice as active in Tris-HC1 as in potassium phosphate and is inhibited 50% by 50 mM NaCl. The polymerase-associated exonuclease hydrolyzes single-stranded DNA somewhat faster than the equivalent duplex DNA. This preference for single-stranded DNA increases as the size of the DNA piece becomes smaller.

B. BIOLOGICAL ROLE The vaccinia-induced DNA polymerase is assumed to be required for the synthesis of the viral DNA, although this remains unproved. Since the genetic loci for this enzyme on the vaccinia chromosome has not been 73. M. D. Challberg and P. T. Englund, J B t 254, 7XLU (IYIY). 74. A. Kornberg, “DNA Replication.” Freeman, San Francisco, 1980.

86

A. WEISSBACH

determined, genetic analysis of the components of DNA replication, as was done for the herpes virus, remains to be investigated. VII.

Conclusion

The mechanism(s) of DNA replication in the cell’s nucleus remain unknown. Further understanding of the physiological role of each of the cellular DNA polymerases will parallel the unraveling of the complex events that accompany and control the synthesis of nuclear DNA. The smaller viral chromosomes, which should be more vulnerable to genetic manipulation and analysis, would seem to offer a promising avenue of research, in parallel perhaps, to the extraordinary detail emerging from the studies of E. coli and its phages (74). The present lack of knowledge portends that our perception of the types and nature of eukaryotic DNA polymerases, as well as their roles, may change within the next few years.

Reverse Transcriptase INDER M. VERMA

I . Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Purification and Properties . . . . . . . . . . . . . . . . . . . .

A. Purification . . . . . . . . . . . B. Enzymatic Activities of Virions , . C. Physical Properties . . . . . . . 111. Biosynthesis of Reverse Transcriptase IV. Mechanism of Reverse Transcription . V. Applications to Molecular Biology . . A. Synthesis of Complementary DNA B. Synthesis of Double-Stranded DNA C. End Labeling . . . . . . . . . . D. Other Utilities . . . . . . . . .

1.

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

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

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

87 88 88 89 92 94 95 99 99 100 101 103

Introduction

Reverse transcriptase can faithfully transcribe RNA into complementary DNA (cDNA). It was first found in the purified virions of murine leukemia virus (MLV) by Baltimore (f)and Rous sarcoma virus (RSV) by Temin and Mizutani (2). The viral RNA acted as the template to direct the incorporation of deoxyribonucleoside triphosphates. The enzyme plays a central role during the life cycle of a retrovirus (3).Temperature-sensitive mutants with a lesion in the reverse transcriptase are unable to establish 1. D. Baltimore, Nature (London)226, 1209-1211 (1970). 2. H. Temin and S. Mizutani, Narure (London)226, 1211-1213 (1970) 3. I . M . Verma, RBA 473, 1-37 (1977). 87 THE ENZYMES, Vol. XIV Copynght 0 1981 by Academic Press. Inc. All rights of reproduciion in a n y form reserved ISBN 0-12-122714-6

88

INDER M. VERMA

infections (4 1. Reverse transcriptase manifests at least three enzymatic activities: (i) Conversion of RNA into DNA (RNA-dependent DNA polymerase); (ii) conversion of single-stranded DNA to double-stranded DNA (DNA-dependent DNA polymerase); and (iii) selective removal of RNA moiety from an RNA-DNA hybrid (RNase H) (3). It has been suggested that reverse transcriptase may also contain DNA endonuclease (5) and swivelase (“unwinding-like”) activities (6). In addition, reverse transcriptase also contains a “tRNA-primer binding site” (7). Due to its multiple activities, the enzyme reverse transcriptase has been alternatively referred to in the literature as RNA-dependent DNA polymerase, RNA-directed DNA polymerase and DNA polymerase of retrovirus (EC 2.7.7.7). Several review articles have been written on reverse transcriptase over the last few years (for general introduction, Temin and Baltimore (Sf, Temin (91, and Stephenson (10); for detailed review on purification, properties, etc., Green and Gerard ( I / ) , Sarngadharen er af. (12), Verma (3), and Gerard and Grandgenett (13). This chapter is limited to a brief review of the purification of the enzyme and its enzymatic activities, mechanism of reverse transcription, and research applications to molecular biology.

11.

A.

Purification and Propeflier

PURIFICATION

Reverse transcriptase resides in the core of the virion (14, 15) and can be easily solubilized by nonionic detergents like Nonidet P40 (16-19) or 4. M. Linial and W. S. Mason, Virology 53, 258-273 (1973). 5. D. P. Grandgenett, A. C. Vora, and R. D. Schiff, Virology 89, 119-132 (1978). 6. M. S . Collett, J. P. Leis, and A. J. Faras, J . Virol. 26, 498-509 (1978). 7. A. Panet, W.H. Haseltine, D. Baltimore, G. Peters, F. Harada and J. E. Dahlberg, PNAS 72, 2535-2539 (1975). 8. H. M. Temin and D. Baltimore, Advan. Virus Res. 17, 129-186 (1972). 9. H. M. Temin, Annu. Rev. Genet. 8, 155-177 (1974). 10. J. R. Stephenson (ed.) “Molecular Biology of RNA Tumor Viruses.” Academic Press, New York, 1980. 11. M. Green and G. F. Gerard, Progr. Nucleic Acid Res. Mol. Biol. 14, 187 (1974). 12. M. G. Sarngadharan, H. S . Allaudeen, and R. C. Gallo, Merhods Cancer Res. 12, 3-47 (1976). 13. G. F. Gerard and D. P. Grandgenett, in ”Molecular Biology of RNA Tumor Viruses (J. R. Stephenson, ed.), pp. 345” Academic Press, New York, 1980. 14. J. Tooze (ed.). “Molecular Biology ofTumor Viruses.” Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1973.

6. REVERSE TRANSCRIPTASE

89

Triton X- 100 (20-22). The detergent-lysed virions manifest all three major activities associated with purified reverse transcriptase. The solubilized lysate is generally fractionated by one or a combination of ion-exchange chromatography, affinity chromatography, velocity sedimentation, and gel filtration. The details of these procedures have been described elsewhere (3, 13). Most of the studies have been carried out on reverse transcriptase obtained from avian retroviruses [avian myeloblastosis virus ( A M V ) ] , Rous sarcoma virus, and murine leukemia viruses. In our experience, the following protocol yields homogeneous preparations of reverse transcriptase relatively fast and in fairly high yields. Purified virions are lysed with nonionic detergent followed by centrifugation at 15,000g for 30 min. The supernatant is adsorbed onto a poly(C)agarose (23) or heparin-Sepharose column (24) and the enzyme eluted with a salt gradient. The enzymatic activity is monitored by utilizing poly(C) . oligo(dG)12-18 as template-primer and radiolabeled dGTP as substrate. The peak fractions of enzymatic activity are pooled and chromatographed on a phosphocellulose column. Material eluting from the phosphocellulose column is highly pure; however, it does contain traces of DNase activity. If the enzyme is to be used for synthesis of cDNA, the peak fraction of enzymatic activity obtained from the phosphocellulose column is further purified by velocity centrifugation on a glycerol gradient (17). The yield of the enzymatic activity from the starting material is about 50-60%. The enzyme should be stored at either -70” in 15% glycerol or -20” in 50% glycerol. B. ENZYMATIC ACTIVITIES OF VIRIONS

Purified reverse transcriptase exhibits both synthetic (DNA polymerase) and degradative (RNase H) activities. These and related enzyme activities found in purified virions are as listed below. 15. R. C. Nowinski, N . H. Sarkar, and E. Fleissner, Merhods Cancer Res. 8,237 (1973). 16. D. L. Kacian, K. F. Watson, A . Burny, and S. Spiegelman, EEA 246,365-383 (1971). 17. I. M . Verma and D. Baltimore, “Methods in Enzymology” Vol. 29, pp. 125-131, 1973. 18. D. P. Grandgenett, G. F. Gerrard, and M. Green, PNAS 70, 230-234 (1973). 19. A. J. Faras, J. M. Taylor, J. P. McDonnell, W. E. Levinson, and J . M. Bishop, Eiochetnisfry 11, 2334-2342 (1972). 20. J. Hurwitz and J. P. Leis, J . Virol. 9, 116-129 (1972). 21. L. H. Wang and P. H. Duesberg,J. Virol. 12, 1512-1521 (1973). 22. J. W. AbreU and R. C. Gallo,J. Virol. 12, 431 (1973). 23. S. L. Marcus, M. J. Modak, and L. F. Cavalieri, J . Virol. 14, 853 (1974). 24. M. Golomb and D. P. Grandgenett, JBC 254, 1606-1613 (1979).

90

INDER M. VERMA

1. DNA Polymerase

Reverse transcriptase can utilize both polyribonucleotides and polydeoxyribonucleotides as templates to direct the synthesis of complementary polydeoxyribonucleotides (25). The two activities are virtually inseparable and apppear to contain a common active site. The enzyme can efficiently transcribe both the homopolymers and heteropolymers. Like other known DNA polymerases, reverse transcriptase also requires a preformed primer to initiate DNA synthesis (26). Although both ribo- and deoxyribo-oligomers can serve as primers, the deoxyribo-oligomers are considerably more efficient primers (3). The primer for initiation of viral DNA synthesis is transfer RNA (27-29), which presumably provides the 3’-OH end to form a phosphodiester bond with the substrate. The direction of synthesis is from 5‘ to 3’ (2.5).The reverse transcriptase appears to be a zinc metalloenzyme (30, 31) and its divalent ion requirements have been tabulated elsewhere (3). Unlike the bacterial DNA polymerases, but like the eukaryotic DNA polyrnerases, reverse transcriptase lacks the 3‘-5’ exonuclease activity, which has been postulated to guarantee a high degree of fidelity during transcription (32). Not surprisingly, reverse transcriptase has been reported to catalyze the incorporation of an exceptionally large number of incorrectly paired bases (33) when homopolymeric templates are used. However, it is not known if the relatively high degree of mistakes is also introduced if heteropolymeric templates are used. 2 . Ribonuclease H The RNase H activity associated with purified reverse transcriptase specifically degrades the RNA moiety of an RNA-DNA hybrid (34). The degradation is not dependent on the concurrent synthesis of complemen25. D. Smoler, I. Molineux, and D. Baltimore, JBC 246, 7697 (1971). 26. I. M. Verma, N. L. Meuth, E. Bromfeld, K. F. Manly, and D. Baltimore, Narure New Biol. 233, 131-134 (1971). 27. R. C . Sawyer, F. Harada, and J. E. Dahlberg, J. Virol. 13, 1302-131 1 (1974). 28. A. J. Faras, J. E. Dahlberg, R. C. Sawyer, F. Harada, J. M. Taylor, W. E. Levinson, J. M. Bishop, and H. M. Goodman,./. Virol. 13, 1134 (1974). 29. G. Peters, F. Harada, J. E. Dahlberg, A. Panet, W. Haseltine, and D. Baltim0re.J. Virol. 21, 1031-1041 (1977). 30. B. J. Poiesz, N. Battula, and L. A. Loeb, BBRC 56, 959 (1974). 31. D. S . Auld, M. Kawaguchi, D. M. Livingston, and B. L. Vallee BBRC 62, 296 (1975). 32. A. Kornberg, “DNA Synthesis.” Freeman, San Francisco, 1974. 33. N. Battula and L. A. Loeb, JBC 251, 982 (1976). 34. K. Molling, D. P. Bolognesi, W. B. Auer, W. Biisen, H. W. Plassmann, and P. Hausen, Nature New Biol. 234, 240-243 (1971).

6. REVERSE TRANSCRIPTASE

91

tary DNA (cDNA) since preformed hybrids are susceptible to RNase H activity (35),which acts as an exoribonuclease and requires free ends (36). In contrast, the cellular RNase H acts as an endoribonuclease (37). Ribonuclease H associated with purified reverse transcriptase from either AMV or MLV cleaves at the 3’ end of the 3’-5‘phosphodiester bond to yield products containing 5’-phosphate and 3’-OH ends (35, 36, 38). The products generated during RNase H activity are 4 to 20 nucleotides long. No mononucleotides are detected in the reaction product. RNase H associated with AMV reverse transcriptase is a processive exonuclease (38, 39), while the RNase H activity associated with the purified MLV reverse transcriptase has been variously reported to act in a random (38) or processive manner (40). Although RNase H and DNA polymerase activities reside on the same polypeptide ( / 8 ) , they appear to have different functional sites. For instance, (a) DNA polymerase activity is more heat-labile than the corresponding RNase H activity (36, 4/); (b) RNase H activity is selectively inhibited by NaF (42); (c) digestion of reverse transcriptase with chymotrypsin leads to inactivation of DNA polymerase activity 8 to 10 times faster than the corresponding RNase H activity (43);(d) DNA polymerase activity purified from AMV is inactivated eight times faster than the corresponding RNase H activity by N-ethylmaleimide (44);and (e) differential sensitivity to low pH (441, sodium pyrophosphate (45), pyridoxal phosphate (46), etc. 3. DNA Endonuclease A DNA endonuclease has been reported in the purified virions and purified AMV reverse transcriptase. The virion-associated DNA endonuclease has a molecular weight of 32,000 and is referred to a ~ 3 ( 5 )2. It ~ has been purified from both AMV and ASV core structures and appears to 35. D. Baltimore and D. Smoler, JBC 247, 7282-7287 (1972). 36. J. Leis, I. Berkower, and J . Hurwitz, In “DNA Synthesis in Vitro” (R.D. Wells and R. B. Inman, eds.), pp. 287-308. University Park Press, Baltimore, Maryland, 1973. 37. W. Keller and R . Crouch. P N A S 69, 3360-3364 (1972). 38. 1. M. Verma,J. Virol. 15, 843-854 (1975). 39. D. P. Grandgenett and M. Green, JBC 249, 5148-5152 (1974). 40. K. Molling, J . Virol. 18, 418 (1976). 41. 1. M. Verma, W. S. Mason, S. D. Drost, and D. Baltimore, Nature (London) 251, 27-31 (1974). 42. L. Brewer and R . D. Wells, J . Virol. 14, 1494-1502 (1974). 43. M. T. Lai and I. M. Verma, J . Virol. 25, 652-663 (1978). 44. M . Gorecki and A. Panet, Biochemisfry 17, 2438-2442 (1978). 45. J. C. Meyers and S. Spiegelman, P N A S 75, 5329-5333 (1978). 46. M. J. Modak, BERC 71, 180-187 (1976).

~

92

INDER M. VERMA

be structurally related to the large, but not the small, subunit of AMV reverse transcriptase (47). Both the virion-associated ~32”’’endonuclease and the reverse transcriptase-associated DNA endonuclease can nick supercoiled DNA in the presence of Mn2+without any site specificity (47). However, ~ 3 2 ” endonuclease ” in the presence of Mg2+,nicks supercoiled E. coli ColEl plasmid DNA preferentially near the EcoRI cleavage site (13, 48). The purified reverse transcriptase can be treated with chymotrypsin to generate a 32,000 dalton fragment that exhibits DNA endonuclease activity (49). The DNA endonuclease activity appears to be more heat-stable than either the DNA polymerase or RNase H activities. No similar DNA endonuclease activity has been found to be associated with mammalian retrovirus purified reverse transcriptase, but an endonuclease activity has been demonstrated in MLV virions (13). Studies with temperature-sensitive mutants with lesions in reverse transcriptase may provide direct evidence of the viral origin of the DNA endonuclease activity. 4. DNA Swivelase

DNA swivelase removes superhelical turns from form I DNA, converting it to a series of relaxed covalently closed structures. These relaxed structures with various topological winding numbers can be separated by agarose gel electrophoresis. In the purified virions of ASV, DNA swivelase activity has been observed after treatment with nonionic detergent. Similarly, Collett el al. (6) have reported the presence of “unwinding-like’’ activity associated with purified AMV reverse transcriptase. However, it has not been shown unequivocally that the DNA swivelase activity is viral-coded protein. Several other enzymatic activities have been variously reported to be associated with purified retroviruses (8);however, this chapter is confined to viral-coded proteins only.

C. PHYSICAL PROPERTIES The physical structure of purified reverse transcriptase from avian, feline, and murine retroviruses has been studied extensively. Most of the structural work has been done on the purified reverse transcriptase obtained from AMV, ASV, M-MLV, and R-MLV (3, 13).Table I summarizes the physical and biochemical properties of purified AMV and M-MLV 47. R . D. S c h E and D. P. Grandgenett, . I Virol. . 28, 279-291 (1978). 48. R. D. Schiff and D. P. Grandgenett, .I.Virol. 36, 889-893 (1980). 49. D. P. Grandgenett, M. Golomb, and A . C. Vora, J . Virol 33, 264-271 (1980).

93

6. REVERSE TRANSCRIPTASE TABLE I

COMPARISON OF PROPERTIES OF M-MULV REVERSETRANSCRIFTASE, ISOLATED a A N D ap FORMS OF AMV REVERSETRANSCRIFTASE Properties

MuLV

a

ffp

Molecular weight Number of subunits rIl2 of DNA polymerase (min, at 45") f l , * of DNA polymerase with template (min at 45") Binding affinity of primer tRNA (liter/mole) Mode of action of RNAase H

84,000 1 6.5 6.5

65,000 1 7.5 7.5

170,000 2 7.0 15.5

k 3 , Fig. 1) for both pMB9 and X DNA (55). The mechanism was independent of whether the substrate was linear or a covalently closed circle. The kinetic study of Halford et al. (55) evaluated the rate constants k , and k 2 in the scheme h

i!

Form I --&Form I1 -LForm 111

Although the authors found a two-step mechanism under various ionic and temperature conditions, the absolute values of k and k 2 were highly dependent on the conditions and ranged from 0.045 to 1.05 min-' for k l , and 0.020 to 0.42 for k 2 . Interestingly however, the ratio of k , to k 2 under the various conditions employed was approximately two to one in all cases. The authors suggest that this 2: 1 ratio is statistical in nature and due to the double strandedness of the DNA substrate. In generating Form I1 DNA from Form I DNA, cleavage can occur in one of two strands. However, in converting Form I1 into Form 111 DNA, cleavage must occur in the complementary strand (5.5). Thus, as suggested by Halford et al., binding and cleavage in both steps of the two-step reaction occur at equal rates after the statistical factor is accounted for. From the above discussions concerning a one-step versus a two-step mechanism for EcoRI endonuclease, it is apparent that the actual mechanism observed, and thus the relative values of k 3 and k P 5(Fig. l), is highly dependent on the reaction conditions, the reaction temperature, the nature and concentration of the substrate, and the substrate to enzyme ratio. Studies similar to these have also been carried out for other restriction endonucleases. A two-step mechanism has been demonstrated for Hpa I1 (79, 80), andHindIII (55). However, Sail (55) and MnoI (80)show double-

,

80. B . R. Baumstark, R. J. Roberts, and U. L. RajBhandary, JBC 254, 8943 (1979).

172

R. D. WELLS, R. D. KLEIN, A N D C . K. SINGLETON

strand scission during one binding event. For BumHI, a two-step mechanism has been interpreted in one instance ( / 5 ) ,whereas a one-step mechanism has been proposed under a different set of conditions (55). Thus the Bum HI data again points to the importance of the effect of reaction conditions on the mechanism of restriction endonucleases.

IV.

bnic Strength and Solvent Effects on Enzyme Specificity

A. EFFECTONEcoRI The influence of a variety of reaction conditions on the rates and specificities of restriction endonuclease reactions has been modestly studied. Some of the variables that have been examined include ionic strength, divalent metal ion (magnesium, manganese, cobalt, zinc, etc.), pH, solvent effects (glycerol, dimethylformamide, dimethyl sulfoxide), and enzyme concentrations. One of the most interesting observations is the ability of some enzymes to recognize a subset of their canonical recognition sequences as a function of one or more of these factors. As with most other topics covered in this chapter, the most extensive work in this area has been performed with EcoRI. Polisky et al. (81) were the first to demonstrate that the substrate specificity for EcoRI could be reduced from the canonical hexanucleotide (G-A-A-T-T-C) to the tetranucleotide (N-A-A-T-T-N). The reduction in site specificity occurred when the ionic strength of the reaction medium was kept low, the magnesium chloride concentration was reduced from 5 to 2 mM, and the pH was raised to approximately 8.5. The enzymatic activity that was responsible for cleavage at the N-A-A-T-T-N site was termed EcoRI". It was also pointed out by these workers that the different subset sites (N-A-A-T-T-N) were cleaved at substantially different rates, presumably dependent on differences in the sequence surrounding the AATT core. Other studies (82-84) have elaborated on this basic observation. Addition of managanese chloride to the reaction medium, or replacement of magnesium chloride with manganese chloride, also relaxes the specificity. A more thorough examination (84) of the types of conditions that enhance the EcoRI* activity demonstrated that 10 mM Tris buffer (pH 8.8) plus 81. B. Polisky, P. Greene, D. E. Garfin, B. J. McCarthy, H. M. Goodman, and H. W. Boyer, PNAS 72, 3310 (1975). 82. M. Hsu and P. Berg, Biochemistry 17, 131 (1978). 83. C. M. Clarke and B. S. Hartley, BJ 177,49 (1979). 84. T. I. Tikhonenko, E. V. Karamov, B. A. Zavizion, and B. S. Naroditsky, Gene 4, 195 (1978).

10. TYPE I1 RESTRICTION ENZYMES

173

2 mM managanese chloride was optimum. In addition, the presence of 40 to 50% glycerol, or several organic solvents in concentrations of 1 to 6%, enhanced the relaxation of specificity. A more complete evaluation of the role of divalent metal ions on the EcoRI and EcoRI* activity has been performed (Malcolm et al., personal communication). Cobalt and zinc, which have an ionic radius to charge ratio similar to magnesium, were found to act as cofactors for EcoRI. Malcolmet af. found that a general decrease in sodium chloride concentration and/or an increase in pH caused a stimulation of the EcoRI" activity. The salt concentration or pH value at which EcoRI" activity appeared depended on the cation under study. The relative order of the ability of metal ions to increase EcoRI* activity under these conditions was manganese > magnesium > zinc > cobalt. A detailed evaluation of restriction sites that are recognized by EcoRI* has been recently reported (85). Using the conditions reported previously (81, 82), Woodbury et al. attempted to gain further insight into the nature and selectivity of EcoRI* activity by carefully evaluating the rates of cleavage on 6x174 replicative-form DNA. This genome contains no EcoRI sites but does contain a number of EcoRI* sites. From kinetic analysis of the cleavage reactions it was possible to generate a hierarchy of the double-stranded recognition sequences: G-A-A-T-T-C, the canonical sequence, was most reactive. G-G-A-T-T-Twas the next most reactive sequence (G-G-A-T-T-A and G-G-A-T-T-G were not cleaved), followed by A-A-A-T-T-T and G-A-A-T-T-N (where N= A,T). N-A-A-T-T-N' (N and N' are unsepcified) were cleaved at a lesser rate, and sites having only the central A-T dimer were cleaved the slowest. Studies were also performed in the presence of managanese chloride at moderate salt concentrations and at near neutral pH values; the same heirarchy of cleavage sites was observed. Woodbury and co-workers utilized the cleavage specificity heirarchy to define possible contact sites between various functional groups of the bases within the recognition site and the EcoRI protein molecule. It should be noted that even under standard EcoRI digestion conditions (Table I) using only modest amounts of enzyme, someEcoR1" sites can be recognized as well as the canonical recognition site. Bishop demonstrated that under standard digestion conditions, G-A-A-T-T-A was recognized by EcoRI (86). However, instead of double-strand scission occurrhg at this site, only single-strand scission (specific nicking) occurred. The nick generated in this manner was found on the same strand in all molecules. 85. C. P. Woodbury, 0. Hagenbuchle, and P. H. von Hippel, JBC 255, 11534 (1980). 86. I. 0. Bishop, J M B 128, 545 (1979).

174

R . D. WELLS, R . D. KLEIN, ANDC. K. SINGLETON

B. OTHERENZYMES Heininger et al. (87) showed that the Bsu endonuclease, which cleaves in the middle of the tetranucleotide G-G-C-C, decreased its substrate specificity at high nuclease concentrations. Conditions that enhanced the recognition of the -G-C- sequence were high pH (approximately 8 . 3 , low ionic strength, and high glycerol content. Glycerol, (12%) was found to be particularly effective in generating additional cleavage products. In addition, sodium chloride enhanced the recognition of G-G-C-C by Bsu but inhibited the activity that recognizes the -G-C- sequence. The amount of Bsu that was required to show the first sign of degradation at the -G-Csequence depended strongly on the reaction medium. Considering all factors, the conditions that were most conducive to the formation of additional restriction fragments were 25 mM Tris buffer (pH 8.51, 10 mM magnesium chloride, and 25% glycerol. Under these conditions as little as a twofold excess of Bsu was sufficient to generate additional restriction fragments, whereas under the standard Bsu reaction conditions, reduced specificity was first observed with a 20- to 40-fold excess of the endonuclease. Single-stranded viral DNA was also cleaved by Bsu when these reaction conditions were used (see Section IX). Furthermore, Heininger et af. (87) found that HueIII, which also recognizes G-G-C-C, did not show a reduction in specificity (to -G-C-) under the high pH and glycerol conditions. This is additional evidence that the mechanism of recognition and cleavage may be different for different enzymes, even among isoschizomers (88) which cleave between the same base pairs. The activity of BurnHI can also be influenced by the presence of hydrophobic reagents such as glycerol, dimethyl sulfoxide, ethylene glycol, ethanol, and dioxane (89). The cleavage specificity ofBamHI in the presence of these reagents was distinct from that under normal reaction conditions. Cleavage did not occur at sites that are subsets of the BarnHI recognition sequence (G-G-A-T-C-C). Instead, cleavage occurred at sites completely unrelated to the canonical recognition site. Several lines of evidence indicated that this activity was not due to contaminating endonucleases, and thus, the activity was thought to be an intrinsic property of BarnHI (89). To the best of our knowledge, this is the first report of reagents that actually change the recognition site of a restriction endonuclease instead of simply relaxing the specificity to a subset of the canonical 87. K. Heininger, W. Horz, and H. G. Zachau, Gene 1, 191 (1977). 88. Isoschizomers are defined as enzymes which recognize and cleave within the same sequence. The precise site of cleavage is not necessarily the same. For example, SmaI (CCCkGG) and XmaI (CkCGGG) are considered to be isoschizomers. 89. J. George, R. W. Blakesley, and J . G . Chirikjian, JBC 255,6521 (1980).


175

site ( 8 9 ~ )Heininger . et nl. may have observed a similar phenomena with Bsu (87). Another study reported on the alteration of specificity of several restriction endonucleases in the presence of organic solvents (90). The enzymes studied were EcoRI, XbaI, SalI, HhaI, PstI, BamHI, and SstI. However, the enzymes used in this investigation were not characterized as to their degree of purity. Hence, it is not possible to draw rigorous conclusions regarding the modification of site recognition by the enzymes in question; the observed cleavages could be due to contaminating activities. The specificity of BstI (G-&-G-A-T-C-C)is reduced to a BstI* site (NL-G-A-T-C-N) in the presence of large amounts of enzyme (>100 U/pg A DNA) (91). This reduced specificity was enhanced in the presence of glycerol as well as under other conditions that also promoteEcoR1" activity (91). It is not surprising that the kinetics and specificity of restriction endonucleases are altered by the presence of various reagents. The history of nucleic acid enzymology clearly indicates that a variety of other enzymes are influenced in a similar fashion. These include the E. coli and M. luteus DNA polymerases' capacity to incorporate ribonucleotides in place to of deoxyribonucleotides (92, 93), to carry out de novo reactions (W), utilize ribo- versus deoxyribo- templates and primers ( 9 3 , and to incorporate the wrong nucleotides (96). Furthermore, the type of divalent cation used in reactions of terminal transferase (97), E. coli RNA polymerase (98), and RNA ligase (991, influences the substrate specificity of each of these enzymes. Many other cases have also been documented. By selectively altering the specificity of restriction endonucleases it may be possible to increase their utility as reagents, as well as to provide new model systems for the study of DNA-protein interactions. The effect of incubation temperature has not been evaluated for many of the endonucleases (see Table I). However, it has been shown (27) that 89a. J . George and J. G. Chirikjian (personal communication) have recently found that BfiniHl actually cleaves subsites (G-G-A-N-C-C, G-G-N-T-C-C, G-A-A-T-C-C) of its canon-

ical recognition sequence in the presence of the hydrophobic reagents. 90. E. Malyguine, P. Vannier, and P. Yot, Gene 8, 163 (1980). 91. M. C. Catherine and B. S. Hartley, BJ 177, 49 (1979). 92. P. Berg, H. Fancher, and M. Chamberlin, "Symposium on Informational Macromolecules," p. 467. Academic Press, New York, 1963. 93. J. H. van de Sande, P. C. Loewen, and H. G. Khorana, JBC 247, 6140 (1972). 94. J. F. Burd and R. D. Wells, J M B 53, 435 (1970). 95. T. M. Tamblyn and R. D. Wells, Biochemisfry 14, 1412 (1975). %. N. Battula and L. A. Loeb, JBC 249, 4086 (1974). 97. K. Kato, J. M. Goncalves, G. E. Houts, and F. J. Bollum, JBC 242, 2780 (1967). 98. J. S . Krakow, G. Rhodes, and T. M. Jovin, In "RNA Polymerase" (R. Losick and M. Chamberlin, eds.), p. 127. Cold Spring Harbor Lab., Cold Spring Harbor, New York, 1976. 99. D. M. Hinton, J. A. Baez, and R. I. Gumport, Biochemisfry 17, 5091 (1978).

176

R . D. WELLS, R . D. KLEIN, ANDC. K. SINGLETON

Hue111 has remarkable heat stability, being stable at 70" for more than 1 hour with no decrease in activity. The rate of the Hue111 reaction increases approximately threefold from 37" to approximately 75". Even at this high temperature, the canonical -G-G-C-C- is faithfully recognized by Hue111 (27). Temperature may affect other enzymes differently, and thus may provide new insights concerning the mechanism of recognition and specificity. V.

Synthetic Oligonucleotides as Substrates

Studies by Greene et al. (75) demonstrated that the self-complementary octanucleotide d(pT-G-A-A-T-T-C-A)can serve as substrate for EcoRI endonuclease. Cleavage of the octamer required that it be base-paired. The optimum reaction temperature (15") was slightly below the T , of the duplex octamer (17-19"). Although the turnover number for EcoRI with this substrate was comparable to that with other substrates (Table 11), the value of the K , was about 200-fold higher. This discrepancy may reflect the fact that a significant proportion of the octanucleotide may have been single-stranded under the conditions employed. d(pT-G-A-A-T-T-U-A) was found not to be a substrate for EcoRI (75). Dwyer-Hallquist and Agarwal (personal communication) utilized four self-complementary octanucleotides that contain the recognition site for HpaI to investigate the interaction of this enzyme with the recognition site. The nonderivatized oligomer was d(G-G-T-T-A-A-C-C). The other three oligomers contained base analogs: 5-bromodeoxyuridine in place of the second thymidine, uridine in place of this residue, and inosine in place of the second guanosine. The kinetic data obtained using two of the oligomers is given in Table 11. The uridine- and inosine-containing oligomers did not serve as substrates for Hpa I. The bromodeoxyuridinecontaining oligomer did act as a substrate, albeit at a reduced efficiency as compared to the nonderivatized oligomer. Both oligomers had to be in the duplex form for cleavage to occur. The conclusions drawn by these workers were that some nonproductive binding of the bromodeoxyuridinecontaining oligomer to IJpaI was occurring, and that Hpa1 requires the presence of the 5-methyl group of the thymidine adjacent to its cleavage site for binding and cleavage. Synthetic duplex oligonucleotides ranging in length from 6 to 13 base pairs were used by Baumstark er al. in studies with HpaII and MnoI endonucleases (80). Both enzymes recognize the same four base-pair sequence (C-C-G-G). Both were shown to be inactive against single-strand substrates (Section IX). The data obtained in this study demonstrated that

177


HpaII and MnoI, although possessing the same recognition sequence, differ in their interaction with, and cleavage of, the duplex substrates used (80). When the duplex 5' d(pG-A-A-C-C-G-G-A-G-A)

I

I

I

I

3' d(T-T -G-G-C-C

I

I

I

I

-T -C-Tp)

I

3' 5'

was employed as substrate, cleavage by HpaII occurred primarily in the upper strand. This is suggestive of a two-step mechanism for this enzyme as previously discussed. In contrast, MnoI cleaved both the upper and lower strands with equal efficiency. Finally, by using various combinations of the oligomers available to these workers, it was demonstrated that both HpaII and MnoI require at least one base preceding the 5' terminus of the recognition site for strand scission to occur. Goppelt et nl. (100) utilized several oligodeoxynucleotides to investigate specific and nonspecific interactions between EcoRI and DNA. The interaction between oligomers (some possessing the EcoRI recognition site and others not) and EcoRI was followed by circular dichroism. Each single-strand oligomer bound to EcoRI in the absence of Mg2+ ions with K ~ 107 = M - 1 . Complex formation with both specific (EcoRI sitepossessing) and nonspecific oligomers gave rise to a decrease of the ellipticity of the protein in the range of the absorption of the peptide bond. Such a change in ellipticity suggested a conformational change was occurring upon complexation with DNA (100). The dependence of complex formation on ionic strength was interpreted by Goppelt ef a!. (100) to correspond to an involvement of two ion-pair bonds between DNA and enzyme. The importance of electrostatic interactions in complex formation between EcoRI and DNA was also supported by nucleic acidcellulose affinity column studies carried out by these authors (100). Several studies on the use of oligonucleotides and dinucleotides as inhibitors of restriction endonuclease activity on high molecular weight substrates have been carried out (Section VII).

VI.

Substituted DNAs

A.

STUDIES WITH

BASE ANALOGS

Early work employing DNA substrates with modified bases to investigate restriction endonuclease-DNA interactions was carried out with 100. M.Goppelt, A. Pingoud, G . Maass, H. Mayer, H. Koster, and R. Frank, EJB 104, 101 (1980).

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R. D. WELLS, R . D. KLEIN, AND C. K. SINGLETON

EcoRI. These studies have been reviewed (69a); the results are summarized here. The presence or absence of the 2-amino group of dG, a minor groove constituent, has no effect on binding and cleavage by EcoRI at its recognition site (G-A-A-T-T-C) (101).5-Hydroxymethylcytosine was also shown not to affect the restriction of DNA by EcoRI, suggesting that the 5-H of cytosine is unimportant in recognition and subsequent cleavage by this enzyme (102). These workers demonstrated, however, that glucosylation at this position rendered the DNA resistant toEcoRI cleavage, probably because of steric hindrance (102, 103). Berkner and Folk found that the V,,, and K , ofEcoRI were unaltered when thymidine was replaced with uridine (103). However, 5-hydroxymethyluridine substitution lowered the V,,, by 20-fold with no effect on K,, suggesting a stearic or electronic influence of the 5-hydroxymethyl moiety on EcoRI cleavage (103). Berkner and Folk investigated the effects of substituted pyrimidines in DNA on the activity of several restriction endonucleases (104). The studies involved endonucleases that contain A-T base pairs within their recognition site (HpaI, HindII, HindIII, EcoRI, BamHI, and HAEII, Table I) and those that contain only G-C base pairs within the recognition site (Hpa 11, and HhaI). DNA that contains glucosylated 5-hydroxymethylcytosine was resistant to cleavage by each of these enzymes, consistent with findings previously discussed (102, 103). These results suggest that a large group (glucosyl) situated within the major groove interferes with restriction by a number of enzymes (104). DNA that contained 5-hydroxymethyluridine (in place of thymidine) allowed discrimination between the two groups of enzymes that were studied. Endonucleases that contain A-T base pairs within their recognition site cleaved this substituted DNA at much diminished rates. However, no effect was observed on the rate of cleavage by HpaII and HhaI (104). Somewhat more interesting were the results Berkner and Folk obtained with DNA in which thymidine was replaced by uridine (104). No effect was seen on the cleavage rate of EcoRI and BamHI with uridinesubstituted DNA. HpaI, HindII, and HindIII, however, all exhibited a much reduced rate of cleavage with this substrate. All five of these enzymes contain A-T base pairs within their recognition sites, yet it appears that only HpaI, HindII, and HindIII utilize the 5-methyl group of 101. 102. 103. 104.

P. Modrich and R. A. Rubin, JEC 252, 7273 (1977). D. A . Kaplan and D. P. Nierlich, JBC 250, 2395 (1975). K. L. Berkner and W. R. Folk, JEC 252, 3185 (1977). K. L. Berkner and W. R. Folk, JBC 254, 2551 (1979).


179

thymidine in the interaction with their sites of recognition, whereas EcoRI and BamHI do not. Several studies have been carried out on the interaction between restriction endonucleases and 5-bromodeoxyuridine-substitutedDNAs. Digestion of DNA that contained 5-bromodeoxyuridine (instead of thymidine) by EroRI, HaeII, HpaI, HpaII, BamHI, and Hind111 was slower than digestion of unsubstituted DNA (47, 104, 10-5). The most sensitive endonucleases with regard to initial velocity were Barn H I , HindII, and HpaI (102). The effects of 5-bromodeoxyuridine may be related to its alteration of the pyrimidine ring electronic properties. Such alterations include the possible influence of stacking, ionization, and tautomeric characteristics of the base (104). Results with HpaII (104) and SmaI (47), neither of which contains thymidine in its recognition site, indicate that the effect of 5-bromodeoxyuridine of altering the cleavage reaction lies outside the recognition site per s e . Marchionni and Roufa found that while the overall rate of cleavage for SmcrI on substituted and unsubstituted DNAs was the same, one of the three SrnaI sites of the DNA under investigation was highly resistant to SrnaI digestion with substituted DNA (47). Mention should be made of work by Petruska and Horn (105) who found that, unlike all other restriction endonucleases examined thus far, MboI activity was enhanced fivefold on 5-bromodeoxyuridine-substituted DNA. This unusual finding was interpreted in a manner that depended on the distance of the nearest thymidine residue from the actual cleavage site (105).

Finally, even though the presence of 5-bromodeoxyuridine in the DNA substrate reduces the rate of cleavage of most restriction endonucleases that have been examined (with the exception of MboI), the reduced rate forEroRI does not mask an earlier interesting finding. Thomas and Davis observed that EroRI exhibited differential rates of cleavage at the five EcoRI sites of A DNA (106). The difference in frequency of cleavage between the most preferred site and the least preferred site was about tenfold. Similar results were obtained by Forsblum et al. in noting a preference by EcoRI for the various recognition sites in adenovirus DNA (107). Marchianni and Roufa also observed the preference for certain sites in A DNA by EcoRI as seen by Thomas and Davis, even though the A 105. J. Petruska and D. Horn, BBRC 96, 1317 (1980). 106. M. Thomas and R . W. Davis, J M B 91, 315 (1975). 107. S . Forsblum, R. Rigler, M. Ehrenberg, U. Pettersson, and L. Philipson, Nucleic Acids Res. 3, 3255 (1976).

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R. D. WELLS, R . D. KLEIN, AND C. K. SINGLETON

DNA contained 5-bromodeoxyuridine in place of thymidine and the overall rate of cleavage by EcoRI was about one-half that of unsubstituted A DNA (47). Thus, although cleavage was affected by the presence of 5-bromodeoxyuridine, preferential recognition of certain sites was not.

B. METHYLATEDDNAs Many investigations have focused on the interaction of restriction endonucleases with methylated DNAs. Such studies are important in determining the biological relationship between modification and restriction. However, findings on these interactions can also infer mechanistic details in a general way. Cleavage or lack of cleavage at a methylated site by a particular restriction endonuclease indicates whether or not the methylated residue is important in the interaction between the enzyme and its recognition site. Inferences of steric hindrance and/or electronic effects can be made from the results of studies on methylated DNA-restriction endonuclease interactions. The results of a number of such studies are summarized in Table I11 [see (108-1 / 8 ) ] , A fascinating new application of restriction endonucleases involves utilizing isoschizomers, one of which is inhibited by methylation of its recognition site while the other is not, to investigate the distribution and function of methylated bases in eukaryotic DNAs (119, 109, 120). For example, HpaII cannot cleave at its recognition site, C-C-G-G, when the internal C is methylated at the 5 position. Yet, MspI can cleave regardless of the presence or absence of 5-methylcytosine at this position (Table 111). Thus, comparison of the bands produced by MspI and those produced by Hpa I1 indicate the location of 5-methylcytosines within the sequence C-C-G-G. Other pairs of isoschizomers that can be employed in a similar 108. M. B. Mann and H. 0. Smith, Nucleic Acids Res. 4, 4211 (1977). 109. C. Waalwijk and R. A. Flavell, Nucleic Acids Res. 5, 3231 (1978). 110. T. W. Sneider, Nucleic Acids Res. 8, 3829 (1980). 111. P. H. Roy and H. 0. Smith, J M B 81, 427 (1973). 112. A. C. P. Lui, B. C. McBride, G. F. Vovis, and M. Smith, Nucleic Acids Res. 6, 1 (1979). 113. B. Dreiseikelmann, R. Eichenlaub, and W. Wackernagel, BBA 562, 418 (1979). 114. R. E. Streeck, Gene, in press (1980). 115. S. Lacks and B. Greenberg, JBC 250, 4060 (1975). 116. A. P. Bird and E. M. Southern, J M B 118, 27 (1978). 117. A. Dugaiczyk, J. Hedgepeth, H. W. Boyer, and H. M. Goodman, Biochemistry 13, 503 (1974). 118. P. H. Roy and H. 0. Smith, J M B 81, 445 (1973). 119. F. Gautier, H. Biinemann, and L. Grotjahn, EJB 80, 175 (1977). 120. A. Razin and A. D. R i g s , Science 210, 4470 (1980).

18 1

10. TYPE I1 RESTRICTION ENZYMES TABLE I11

RESTRICTIONENDONUCLEASES SHOWN METHYLATED DNA9 Enzyme

Site cleaved

HpaII Hap I1

(C)CGGb C(C)GG GGC(C) G(A)TC

Msp I HneIII Fnu EI MboI Dpn I Sau 3A1 BurnHI BglII Dpn 11 TuqI A I Hha I HaeII EcoRI Bsp I Hind11 Hind111 180

-

G(A)TC' G(A)TC GG(A)TCC AG( A)TCT

-

T(C)GA -

-

Site not cleaved

TO CLEAVE

Reference

C(C)GG C(C)GG (C)CGG GG(C)C G(A)TC GAT(C)"." G(A)TC

-

CPy(C)GPuG G(C)GC RG(C)GCY GA( A)TTC GG(CXCy= GTPyPu(A)C (A)AGCTT

The methylated residues, either 5-methylcytidine or bmethyladenosine, are enclosed in parentheses. Dashes designate not determined. * Half-methylation of the duplex recognition site. Methylation is required for cleavage. Nicking occurs in unmethylated strand. The position of the methylated C is uncertain.

manner to localize methylated residues at other sites can be inferred from Table 111. This approach is a very powerful technique, since information on distribution, tissue specificity, and function of methylated bases in eukaryotic DNAs can be obtained (120). VII.

A.

Inhibitor Studies

REAGENTSTHATMODIFY PROTEINS

Nath and Azzolina (personal communication) investigated the effect of sulfhydryl group inhibitors on the activity of several restriction endonucleases. BamHI was shown to be completely inactivated by 5 , S-dithio-

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R. D. WELLS, R. D. KLEIN, A N D C . K. SINGLETON

bis(2-nitrobenzoic acid) (DTNB) and p-mercuribenzoate. The latter reagent also completely inactivated Pvu I. DTNB partially inhibited HindIII, PvuI, AvaI, SmaI, PstI, and SstII. p-Mercuribenzoate also demonstrated partial inhibition of AvaI, SmaI, and PstI. No inhibition with either reagent was found with SalI, BglII, and HpaI. Similarly, EcoRI (100) and BglI (61) were shown to be insensitive to sulfhydryl-inhibiting reagents. Thus, the importance of sulfhydryl groups for restriction endonuclease activity appears to be highly dependent upon the particular endonuclease. Methyl acetimidate, at concentrations of 3 to 33 mM was found by Woodhead and Malcolm to inactivate EcoRI endonuclease (72). This compound is specific for primary amino groups (1211, and the inhibition suggests that lysine residues may be important in the interaction of EcoRI with DNA (72). Lysine residues, as well as arginines, have also been implicated in the interaction of BglI with DNA. Modification of the BglI lysine residues by pyridoxal 5'-phosphate resulted in diminished binding and catalysis for this enzyme (61 1. 2,3-Butanedione modification of arginine residues of BglI inhibited the enzyme activity but did not affect the binding properties ofBglI (61). However, no study was performed of the influence of these modifications on the capacity of the modified enzyme to recognize noncanonical DNA sites, nor was there an attempt to determine which amino acid residues within the protein were modified. B. POLYNUCLEOTIDES AS INHIBITORS Woodhead and Malcolm (72) studied the ability of various DNAs to protect EcoRI from methyl acetimidate inhibition. Protection was shown to be independent of the presence or absence of Mg ions. Their investigation demonstrated that EcoRI will bind to DNA that does not contain the EcoRI recognition site. The kinetic analysis of the data led these authors to conclude that nonspecific binding ofEcoRI is 15,000 times weaker than binding to the recognition site (72). Although the finding that EcoRI endonuclease binds nonspecifically to DNA is in agreement with the studies on oligonucleotides (100; see Section V) and other studies discussed below, there is disagreement as to the relative strength of specific versus nonspecific binding. Studies on the nonspecific DNA binding properties of EcoRI have also been carried out by Langowski et al. (73). This work demonstrated that d(G-G-A-A-T-T-C-C), eukaryotic DNA, synthetic alternating copolymers (both duplex and single-stranded) and duplex homopolymers, and tRNA were competitive inhibitors of EcoRI cleavage of pBR322 DNA. Only 121. M. J . Hunter and M. L. Ludwig, JACS 84, 3491 (1962),


183

poly(dG) . poly(dC) was found not to inhibit cleavage. The K, values for the various inhibitors ranged from to lo-" M (M nucleotides). This competitive inhibition was interpreted as representing nonspecific binding of EcoRI to DNA and RNA. By comparing results of inhibition between poly(dAT) and d(G-G-A-A-T-T-C-C), the authors concluded that nonspecific binding is about two orders of magnitude weaker than specific binding of DNA by EcoRI (73). The reason(s) for the discrepancy in the strength of nonspecific binding versus specific binding found in this work and in the studies by Woodhead and Malcolm (72) are not apparent at this time. Similar inhibition studies to those above have demonstrated that Bum HI, like EcoRI, possesses nonspecific DNA binding properties (76). Hinschet al. found that BamHI activity on pJC80 DNA was competitively inhibited by nonsubstrate polynucleotides with K, values from > to lop6 M ( M base pairs) (76). Inhibition was seen with polyribo- and polydeoxyribonucleotides. The amount of inhibition shown by a given DNA was dependent on base composition, base pairing, and helix conformation. The nonspecific DNA binding property demonstrated for EcoRI and Bum HI is a typical property for a protein that recognizes DNA at a given target site. Models of DNA-protein interaction for DNA binding proteins generally lead to the conclusion that these proteins possess an appreciable affinity for nontarget sequences leading to nonspecific binding that is important in the overall binding mechanism ( I , 122, 123). Finally, inhibition of Barn HI activity by dinucleotides has been investigated (76, 1244). The results of these two studies are conflicting, and there appears to be no good correlation between the amount of inhibition by a particular dinucleotide and its presence or absence from the BamHI recognition site. Thus, it may be that the enzyme binds the dinucleotides by a mechanism different from that with a true substrate.

VIII.

Influence of Drugs and Other Ligands on Cleavage Specificities

Several investigations have focused on the ability of region-specific DNA binding ligands to preferentially inhibit cleavage at certain restric122. P. H. von Hippel, In "Biological Regulation and Development, Volume I: Gene Expression" (R.F. Goldberger, ed.), p. 279. F'lenum, New York, 1979. 123. T. M. Jovin, Annu. Rev. Biuchem. 45, 889. 124. Y.H. Lee, D. Clanton, and J . G . Chirikjian, FP 38, 294 (1979).

184

R . D. WELLS, R. D. KLEIN, AND C. K. SINGLETON

tion sites. A number of ligands, such as carcinogens, dyes, and antibiotics, are known to bind specifically to DNA. Studies were undertaken in an effort to enhance the specificity of restriction enzyme cleavages, as well as to further understand the mode of interaction of these ligands with DNA. In 1976, Kania and Fanning (125) showed that the binding of 6,4’-diamidino-2-phenylindole(DAPI) to lambda and adenovirus DNAs caused a preferential inhibition ofEcoRI cleavage at some sites but not at others. Also, Nosikov el al. (126) found that distamycin A and actinomycin D preferentially inhibited some of the sites of the following enzymes: EcoRI, EcoRII, HindII, HindIII, HpaI, and HpaII. Distamycin A and DAPI preferentially complexes with A-T-rich regions, whereas actinomycin binds to G-C pairs. van de Sande and his associates (quoted in 1) have studied the capacity of a variety of drugs to inhibit the cleavage at some specific sites by DNA restriction enzymes. The drugs that demonstrated the largest effects were olivomycin and Hoechst 33258. At an appropriate drug/DNA ratio, the cleavage of one EcoRI site on lambda DNA was completely inhibited whereas cleavages at other sites were unaffected. Studies were also performed on HpaI cleavage of PM2 DNA and on HhaI reaction with MI3 replicative form DNA. Due to the relatively low concentration of drug necessary to cause these specific inhibitions, these workers suggested that these findings must be due to differences in the sequences surrounding the canonical recognition sites. A similar approach employed a broader range of drugs and antibiotics (127). The restriction enzymes studied were EcoR1, HindIII and BarnHI. An interesting observation from this work was that the restriction sites located in the center of lambda DNA were most sensitive to the inhibitory action of various intercalators, whereas these interior recognition sites were the least preferred when the intercalators were absent. Thus, the intercalators enhanced the discrimination shown by EcoRI against the interior sites. The binding of cis-dichlorodiammineplatinum(I1)to pSMI plasmid DNA causes a preferential inhibition of PstI cleavage at one of the four PstI sites. This inhibition is believed to be due to the presence of four G-C base pairs that neighbor the Pst1 site (128). 125. J. Kania and T. G. Fanning, EJB 67, 367 (1976). 126. V. V. Nosikov, E. A. Braga, A. V. Karlishev, A. L. Zhuze, and 0. L. Polyanovsky, Nucleic Acids Res. 3, 2293 (1976). 127. K. Nath, Absrr. / l r h Intern. Congr. Biochem. 01-5-HSI (1979). 128. G . L. Cohen, J. A. Ledner, W. R. Bauer, H. M. Ushay, C. Caravana, and S. J. Lippard, JACS 102, 2487 (1980).


IX.

185

Cleavage of Single-Stranded DNA Substrates by Certain Restriction Endonucleases

In the early 1970s, it was believed that restriction enzymes only recognized sites on duplex DNA with twofold rotational symmetry. Later it was realized that HueIII specifically cleaved single-stranded DNAs from 6x174, M13, and fl viruses. This was an apparent violation of the “twofold rotational symmetry” notion. This observation was of substantial interest from both the standpoint of the enzymology of restriction endonucleases and of the study of properties of single-stranded viral genomes in solution. Further studies have demonstrated that HueIII plus several other enzymes apparently recognize folded-back duplex regions within the single-stranded viral genomes. The subject of single-stranded DNA cleavage has been recently reviewed (129). The enzymes that have been tested for their capacity to cleave singlestranded DNA substrates are listed in Table IV [See (130-/33)], with HueIII as probably the most thoroughly studied. A substantial body of data (27 and reviewed in 129) reveals that HueIII cleaves duplex DNA approximately 16 times faster than single-stranded viral DNA that contains potential short duplex regions. Although not yet proven, it is also possible that a b u m fide single-stranded G-G-C-C site may be cleaved, though at an even slower rate. One point of discrepancy in Table IV concerns the ability of HpaII to cleave single-stranded DNA. Two studies involving the interaction of chemically synthesized oligonucleotides and single-stranded viral DNAs with HpuII seem to validate the notion that single-stranded substrates are not cleaved, or are cleaved more slowly, by this enzyme (133, 134). Since single-stranded viral DNA has extensive duplex structure (reviewed in I ) , it is curious that such DNA is not a substrate for all restriction endonucleases that cleave replicative form DNA. Even though Hue111 and HhuI recognition sites are 100% G-C, high G-C content in the site is neither a necessary nor sufficient criterion for cleavage. MboI, MboII and HinI, whose sites are less than 50% G-C, cleave single129. s. K . Neuendorfand R. D. Wells, (1980). In “Gene Amplification and Analysis, Vol. I: Restriction Endonucleases” (J. G. Chirikjian, ed.), p. 101. Elsevier, North Holland, Amsterdam, 1981. 130. R. W. Blakesley and R. D. Wells, Nature (London)257, 421 (1975). 131. K . Hoviuchi and N . D. Zinder, PNAS 72, 2555 (1975). 132. 0. J . Yo0 and K . L. Agarwal,JBC 255, 10559 (1980). 133. B . R. Baumstark, R. J . Roberts, and U. L. RajBhandary, JBC 254,8943 (1979). 134. J . L. Hines and K. L. Agarwal, personal communication.

186

R . D. WELLS, R. D. KLEIN, A N D C . K. SINGLETON TABLE IV

RESTRICTION ENDONUCLEASES TESTED ON 4x174,M13 OR F1 VIRALDNAs Enzyme Enzymes that cleave HaeIII Hha I SfaI Hin fI MboI MboII Msp I Bsp RI Enzymes that do not cleave" Ah1 Bum HI EcoRI HaeII Hind11 Hind111 Hpa I HpaIIr Mno I Psr I SalI Sma I

Recognition site

Reference

GGCC GCGC GGCC GANTC GATC GAAGA CCGG GGCC

(/29-131, 6.0 (27, 65, 130) (130) (27, 65) (27) (65) (132) (33 )

AGCT GGATCC GAATTC PuGCGCPy GTPyPuAC AAGCTT GTTAAC CCGG CCGG CTGCAG GTCGAC CCCGGG

(130, 6 5 ) (27) (130, 7S)* (130)

(/.TI), 68) (130) (27)'' (130, 6 s . 131, 27, J33)b (133)b (27) (27) (2 7)

It has been shown that EcoRI (72, 73. 1 0 0 ) and BarnHI (76) bind to singlestranded DNA but d o not cleave. Binding to single-stranded DNA may be a general property of restriction endonucleases. * Tested on synthetic oligodeoxyribonucleotides. The discrepancy of results with HpaII is discussed in the text. K. L. Agarwal, personal communication.

stranded DNA. On the other hand, HpaII, MnoI, and SmaI, whose recognition sequences contain 100% G-C, do not cleave single-stranded DNA. In addition, sites for HaeII, which are a subset of those cleaved by HhaI, are not cleaved in 4x174 viral DNA (Table I V ) . Thus, subtleties in DNA structure or enzyme-substrate specificities, or both, determine if a particular enzyme recognition site in single-stranded DNA is cleaved. Another study (132) indicates that MspI can cleave single-stranded synthetic deoxyoligonucleotides at a slow rate with cleavage dependent on low temperatures and the surrounding nucleotide sequences. Similarly,

187


MspI cleaves 4x174 viral DNA at a 27-fold slower rate than the cleavage of duplex 4x174 DNA. Koncz et al. (33) have shown that BspRI endonuclease specifically cleaves 4x174 viral DNA when a 100-fold excess of enzyme is present. BspRI is an isoschizomer of HaeIII, both recognizing G-G-C-C. These workers found that the canonical G-G-C-C sites were cleaved in the viral DNA like they were in the replicative form.

X.

Cleavage of DNA-RNA Hybrids

There has been substantial interest in the possibility of using DNA restriction endonucleases for the specific cleavage of DNA-RNA hybrids. Whereas several laboratories have attempted to evaluate this possibility, the presence of contaminating ribonucleases in the restriction endonuclease preparations gave rise to complications. If restriction endonucleases could specifically cleave both the DNA and RNA strands of DNA-RNA hybrids, this technique would provide a valuable tool for studying gene expression and for RNA sequence studies. Molloy and Symons (135) describe the cleavage of a DNA-RNA hybrid, prepared from cucumber mosaic virus RNA and its copy DNA, by the following restriction enzymes: EcoRI, HindII, SalI, MspI, HhaI, AluI, TaqI, andHaeII1. These investigators showed that the DNA strand of the hybrid was cleaved, at least in some cases, in the same place as for duplex DNA. This cleavage required 20- to 50-fold higher enzyme levels than those required for cleavage of duplex DNA. It was not certain if the RNA strand was cleaved. Alternatively, it is possible that the RNA was randomly degraded to small oligonucleotides since the appropriate determinations were not performed to rule out this possibility. As previously described, two of these enzymes (HhaI and HueIII) cleaved certain single-stranded viral DNAs. However, four of the enzymes (EcoRI, HindII, ScilI, Alrr I) that do cleave the DNA-RNA hybrids do not cleave viral single-stranded DNA. Thus, even though these authors have not studied the fate of the RNA strand, it is likely that DNA-RNA hybrids are, in fact, cleaved. This approach is likely to be used expediently in further studies. Brown (136) has reported that HaeIII will cleave a duplex DNA 5' 3'

. .

. .dG-dG-rC-rC. . . 3'

.

.dC-dC-dG-dG.

..

5'

135. P. L. Molloy and R. H . Symons, Nucleic Acids Res. 8, 2939 (1980). 136. N . L. Brown, FEBS (Fed. Eirr. Eiochern. Soc.) Lett. 93, 10 (1979).

188

R. D. WELLS, R. D. KLEIN, AND C. K. SlNGLETON

that contains a site partially substituted with two ribocytidine residues. However, no kinetic data were provided. XI.

lnsolubilized Restriction Enzymes

To the best of our knowledge, only one paper (137) has appeared that describes the covalent linking of restriction enzymes to an insoluble matrix. These workers used cyanogen bromide to couple Barn HI and EcoRI to Sepharose. Over 90% of the activities were found in an insoluble form, and the insoluble enzymes specifically cleaved lambda, adenovirus, and SV40 DNAs in the same manner as the soluble enzymes. In addition, EcoRI" activity was observed. The thermal stability of both enzymes was increased by at least lo", and the preparations could be lyophilized to dryness without loss of activity. In addition to BamHI and EcoRI, HpaI and TaqI have been carried through this procedure. The use of insolubilized restriction endonucleases has several advantages over standard techniques: the coupled resin can be employed in DNA cleavage reactions in a manner analogous to column chromatographic techniques, reactions can be terminated by rapidly pelleting the resin from solution by centrifugation, and finally, the recovered enzymes can be reused several times. It is quite likely that this technique will receive more attention in further studies. XII.

Crystallization of Restriction Endonucleases

Crystals of EcoRI have been obtained by two different groups. Both groups have obtained crystals of the endonuclease itself, and of the endonuclease complexed with an oligonucleotide that possesses the EcoRI recognition site (138, 139 and Rosenberg, personal communication). The unit cell of the EcoRI crystals contains four enzyme monomers per asymmetric unit (138, 139). At high enzyme concentrations in solution, the EcoRI subunit associates as a tetramer (70), in good agreement with the crystallographic data. The data for the Eco RI-oligomer complex crystals (G-A-A-T-T-C (139) and C-G-C-G-A-A-T-T-C-G-C-G(J. M. Rosenberg, personal communication)) suggests that the unit cell in both instances possesses four enzyme monomers plus two duplex DNA fragments in an 137. Y. H. Lee, R. W. Blakesley, L. A. Smith, and J. G . Chirikjian, Nucleic Acids Res. 5, 679 (1978). 138. J. M. Rosenberg, R. E. Dickerson, P. J . Greene, and H. W. Boyer, J M B 122, 241 (1978). 139. T.-S. Young, P. Modrich, E. Jay, and S.-H. !Grn,JMB, 145, 607 (1981).


189

asymmetric unit. Thus, one DNA duplex is assumed to be associated with one enzyme dimer, the active form ofEcoR1 in solution (3). More detailed crystallographic data should prove to be highly enlightening with regard to understanding the interaction between EcoRI endonuclease and its recognition site. XIII.

Genes for Restriction Endonueleases

Only a small number of restriction and modification genes have been cloned to date. These include the genes for HhaII, EcoRI, EcoRII, PstI, and the methylase genes ofBspI, EcoRII, PstI andBsu. Attempts to clone several other restriction endonuclease genes have been futile. It is likely, however, that new approaches to cloning restriction enzyme genes will be made, since such clones will provide useful tools for studying the regulation and expression of these genes as well as being of practical value as a source of the enzymes. The restriction and modification genes of the HhaII system were the first to be cloned. Smith and his associates (20) cloned a three kilobasepair segment of DNA from Haemophilus haemolyticus using the plasmid pBR322 as a vector. Transformed cells carrying the appropriate recombinant plasmid expressed both theHha I1 restriction function as well as the modification function. E. coli cells harboring this plasmid were used for the partial purification of HhaII. Further studies from Smith's laboratory (140, and Hausler and Smith, unpublished) have generated another recombinant plasmid that contains a 2.2 kilobase-pair segment of Haemophilus haemolyricus DNA possessing both the restriction endonuclease and modification methylase genes. The DNA sequence of this insert has been compared to that of the EcoRI gene; no regions of extensive homology were observed (Hausler, Smith and Modrich, personal communication). Modrich and his associates and Boyer et al. have independently cloned 2.2 kilobase-pair segments of DNA that are required for the expression of the EcoRI restriction and modification phenotypes. These DNA segments were sequenced independently using different sequencing procedures. The DNA sequences obtained by these two laboratories are in complete agreement (14/-/43). Since the amino and carboxyl terminal sequences of 140. B. Hausler, FP 39, 946 (1980). 141. A. K. Newman, R. A. Rubin, S.-H. Kim, P. Modrich,JBC, 256, 2131 (1981). 142. R . A. Rubin, P. Modrich, and T. C. Vanaman, JBC, 256, 2140 (1981). 143. P. J. Greene, M. Gupta, H. W. Boyer, W. E. Brown, and J. M. Rosenberg,JBC, 256, 2143 (1981).

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R . D. WELLS, R . D. KLEIN, AND C. K. SINGLETON

the restriction and modification enzymes have been partially determined (142), these investigators could unequivocally establish the reading frame of the DNA sequence. The DNA sequence indicated that the EcoRI endonuclease consists of 277 amino acid residues, whereas the methylase gene is 326 amino acid residues in length. The genes for the two proteins do not overlap and, in fact, are separated by a 29 nucleotide intercistronic region. Because both the restriction endonuclease and modification enzymes have the same recognition site (G-A-A-T-T-C), it was of interest to compare the amino acid sequences of the two proteins. Both groups concluded that there is a general lack of homology between the two polypeptides, suggesting different evolutionary origins for the two proteins. Moreover, both groups have concluded that, based on circular dichroism measurements and theoretical structural predictions, the two enzymes differ markedly in their secondary structure. Thus, the mechanism of interaction of these two enzymes with the same DNA recognition sequence remains an enigma. A few other restriction endonuclease genes have been cloned with virtually no characterization in terms of restriction mapping or DNA sequence analysis. The genes for EcoRII endonuclease and methylase have been cloned and exist together on a 5.8 megadalton DNA fragment (144). The modification methylase gene of Bacillus sphaericus R has been cloned inE. coli using a pBR322 vector. The gene is carried by a 2.5 kb restriction fragment (145). Though recombinant clones do not exhibit BspI restriction endonuclease activity, the level of methylase activity was found to be higher than in the parental strain (143). Also, the modification and restriction systems of Bsu 12451 and Bsu 124711 have been cloned into Bacillus subtilis 168 (146). The genes for PstI restriction and modification enzymes have also been cloned using the plasmid pBR322 as a vector. The smallest in a recombinant plasmid encoding both the modification and endonuclease genes was approximately 4,000 bp in length (147). When this plasmid was introduced intoE. coli minicells, two new proteins of 33,000 and 35,000 daltons were identified. The 35,000 dalton protein comigrated with the prominent component of a partially purified PstI endonuclease preparation.

144. 145. 146. 147,

V. G.Kosykh, Y. I. Buryanov, and A. A. Bayev, Mol. Gen. Genet. 178,717 (1980). E. Szomolanyi, A. Kiss, and P. Venetianer, Gene 10, 219 (1980). T. Shibata. S. Ikawa, Y.Komatsu, T. Ando, and H. Saito, J E 139, 308 (1979). R. Y. Walder, J. L. Hartley, J. E. Donelson, and J. A. Walder, PNAS 78, 1503 (1980).

10. TYPE 11 RESTRICTION ENZYMES

XIV.

191

Conclusions

Restriction endonucleases have been widely used for gene cloning and mapping, for investigations on gene expression in both prokaryotic and eukaryotic systems, and as models for the study of protein-DNA interactions. It is expected that this usage will be expanded in both academic and industrial directions. Most restriction endonucleases have been meagerly characterized in biochemical terms. Interestingly, however, the characterizations that have been carried out show that these enzymes, although grouped together as site-specific DNA cleavage enzymes, exhibit remarkable diversity. This diversity is evident in both the physical and kinetic properties of each enzyme, as well as whether a particular endonuclease can interact with and cleave methylated DNAs, base-analog containing DNAs, single-stranded DNA's, DNA-RNA hybrids, and noncanonical duplex DNA. Such diversity draws attention to the rich source of biochemical and enzymological knowledge that can be obtained by employing this class of enzymes as both biochemical reagents and subjects of investigation. ACKNOWLEDGMENTS This work was supported by the National Institutes of Health (CA 20279) and the National Science Foundation (PCM 15033). C. K. Singleton was supported, in part, by a fellowship from the American Cancer Society (PF-1904). We also wish to express our appreciation to our friends and colleagues who freely shared their unpublished results with us. Due to the extremely rapid pace of progress in this field, this chapter would be far less complete without their help. We thank R. Roberts for checking the accuracy of Table I.


Endonucleases Spec@ for Single-Strunded Poly nucleotides I. ROBERT LEHMAN

I. Introduction . . . . . . . . . . . . . . . . . . . . . . 11. Neurospora crassa Endonuclease . . . . . . . . . . . . A. Physical Properties . . . . . . , . . . . . . , . . . B. Reaction Catalyzed . . . . . . . . . . . . . . . . . C. Biological Role . . . . . . . . . . . . . . , . . . . 111. Nuclease S l . . . . . . . . . . . . . . . . . . . . . . IV. Mung Bean Endonuclease . . . . . . . . . . . . . . . . A. Physical Properties . . . . . . . . . . . . . . . . B. Reactions Catalyzed . . . . . . . . . . . . . . . . . V. Research Application of Single-Strand-Specifc Endonucleases I

1.

. . . . ,

. . . . .

. . . . . . . . . .

. . ,

. . . . . . .

.

. . .

. .

.

. . .

193 194 194 196 197 198 199 199 200 201

Introduction

Endonucleases with a high degree of specificity for polynucleotides that lack an ordered structure have been isolated from a variety of fungi, including Neurospora crassa ( I , 2 ) , Aspergillus oryzae (3, 4 ) , Penicillium cit1. 2. 3. 4.

S. Linn and 1. R. Lehman, JBC 240, 1287 (1965). S . Linn, and I. R. Lehman, JBC 240, 1294 (1965). T. Ando, BBA 114, 158 (1966). V. M.Vogt, EJB 33, 192 (1973). 193 THE ENZYMES,Vol. XIV Copyright 0 1981 by Academic Press, Inc. AU rights of reproduction in any form reserved ISBN 0-12-122714-6

194

I . ROBERT LEHMAN

rinum P (5, 6), and Ustilago maydis (7, 8 ) . Similar enzymes have been purified from mung bean sprouts (9, 1 0 ) and wheat seedlings ( I I ) . Because of their ability to discriminate between single- and double-stranded polynucleotides, these enzymes have become exceedingly useful as reagents for the analysis of nucleic acid structure. Two of these nucleases, from N. crassa and U . maydis, appear to be required for DNA repair and recombination in viva A summary of the properties of the six singlestranded polynucleotide specific endonucleases is given in Table I. The enzymes to be considered in detail are those from N. crassa, Aspergillus oryzae, and mung bean.

11,

Neurospora crassa Endonuclease

A. PHYSICAL PROPERTIES The N. crassa endonuclease was the first of the single-strand-specific endonucleases to be discovered ( I , 2). Though purified extensively from both conidia and stationary phase mycelia (approximately 10,000 fold), the enzyme has not yet been obtained in homogeneous form. As judged by sedimentation in sucrose density gradients, it has a sedimentation coefficient, s20,w, of 3.7 and a molecular weight (assuming a globular structure) of 55,000 (12). The important feature of the enzyme, and that which has made it a useful reagent, is its high degree of selectivity for single-stranded DNA or single-stranded regions within duplex DNA. Although the 10,000-fold purified enzyme preparation attacks native DNA at 2% the rate of denatured DNA, it is clear that hydrolysis of native DNA is due largely to the activity of a contaminating nuclease, which can be selectively inactivated by incubation at 55”, or by brief exposure to thiols. After such treatment, native DNA is attacked at less than 0.1% the rate found with denatured DNA. The partially purified enzyme is active under a variety of conditions including temperatures ranging from 25” to 65”, pHs ranging from 6 to 9 in 5. M. Fujimoto, A. Kuninaka, and H. Yoshino, Agr. Biol. Chem. 38, 777 (1974). 6. M. Fujimoto, A. Kuninaka, and H. Yoshino, Agr. Biol. Chern. 38, 785 (1974). 7. W. K . Holloman, and R. Holliday, JBC 248, 8107 (1973). 8. W. K. Holloman, JBC 248, 8114 (1973). 9. D. Kowalski, W. D. Kroeker, and M. Laskowski, Sr. Biochemistry, 15, 4457 (1976). 10. W. D. Kroeker, D. Kowalski, and M. Laskowski, Sr. Biochemistry. 15, 4463 (1976). 11. W. D. Kroeker and J. L. Fairley, JBC 250, 3773 (1975). 12. S. Linn, “Methods in Enzymology,” Vol. XII, Part A, p. 247, 1967.

TABLE I

PROPERTIES OF SINGLE-STRANDED POLYNUCLEOTIDE-SPECIFIC

Enzyme source

Degree of purity (%)

Molecular weight 55,000

Ratio DNasel RNase

Final products of hydrolysis

PH optimum

>90% Nucleoside

7.0-9.0

N . crassa (1. 2)"

Unknown

(1. maydis

Unknown

42,000

3

>90% Nucleoside 5'-phosphates

8.0

>90

32,000

5

>90% Nucleoside

4.0-4.3

1

A . oropzae (-3, 4 ) P. citrinum (5, 6 ) Mung bean (9. 1 0 ) Wheat seedlings

5.0

Zn" co2+ Mg'+ Mg2+ Ca2+ co2+ Zn" Zn" coz+ Zn2+

5.0

Zn2+

4.8-5.5

Zn2+required for stabilization

5'-phosphates

(7. 8 )

>90

Unknown

0.7

>90

39,000

1.2

>90

43,000

2

5'-phosphates >90% Nucleoside 5'-phosphates >90% Nucleoside 5'-phosphates >90% Nucleoside 5'-phosphates

(11)

Numbers in parentheses are reference numbers.

Divalent cations

ENDONUCLEASES

Inhibitors

Comments

2-Mercaptoethanol. Preferential attack at G potassium and dG residues phosphate, ATP 2- Me rc aptoethano I, Preferential attack at G potassium and dG residues phosphate, ATP

Potassium phosphate, NaF

Possesses 3'-nucleotidase activity; cold labile Possesses 3'-nucleotidase activity Possesses 3'-nucleotidase activity ; preferential attack at A and dA residues

196

I. ROBERT LEHMAN

the presence of a variety of divalent cations, and at ionic strengths ranging from 0.03 to 0.20. Although the enzyme shows some activity in the absence of added divalent cations (20% of maximal), it is strongly inhibited (95%) by EDTA at 0.1 mM. This inhibition cannot be overcome by an excess of Mg'+, but is specifically reversed by Coz+.The enzyme is also inhibited to some extent by potassium phosphate (80% inhibition at 33 mM) and ATP (50% inhibition at 0.5 r n M ) (13). The N. crussu endonuclease hydrolyzes RNA at approximately the same rate as DNA. The same enzyme is responsible for both activities. Thus, the two cochromatograph on hydroxylapatite, have the same pH and temperature optima, show similar rates of heat inactivation, respond in the same way to EDTA and to divalent cations, and have similar specificities for single-stranded polynucieotide substrates.

B. REACTION CATALYZED The N. crussa endonuclease hydrolyzes single-stranded DNA and RNA to a mixture of mono- and oiigonucleotides. As judged by the formation of acid-soluble products, the hydrolysis of single-stranded DNA proceeds in two phases: an initially rapid phase that yields a digest composed of a mixture of 5'-mononucleotides (approximately 30%) in which dGMP predominates, and oligonucleotides of various lengths ranging from dinucleotides to larger than pentanucleotides. At the end of the second phase, mononucleotides comprise approximately 45% of the digest and the remainder is mainly in the form of di- and trinucleotides. More prolonged incubation yields greater than 90% mononucleotides. The N. crussu nuclease can very effectively remove single-stranded regions in double-stranded DNA. This point is illustrated in Fig. 1. In the experiment shown, duplex bacteriophage T7 DNA was treated withE. coli exonuclease I11 to remove from 25 to 5000 nucleotides from the 3' termini of the duplex. Treatment of these DNAs with the N. c r a m endonuclease resulted in an amount of material made acid-soluble that was comparable to that removed initially by exonuclease 111. The value of the slope (0.86) is consistent with the extent of conversion to acid-soluble nucleotide expected under the conditions of the experiment. The N. crussu enzyme is thus able to remove single-stranded stretches in double-stranded DNA to within ten nucleotides of the hydrogen-bonded nucleotides; the value of ten represents the experimental error in the analysis. 13. M. J. Fraser, R. Tjeerde, and K. Matsumoto, Can. J . Biochem. 54, 971 (1976).

I I.

197

SINGLE-STRAND-SPECIFIC ENDONUCLEASES W -1

rn

14

I

I

I

1

I

I

I

3

slope = 0.89

0.1

-

-

-n 0

a

I

0

0

2

1

I

I

I

I

4 6 8 10 12 14 PERCENT MADE ACID SOLUBLE BY EXONUCLEASE 111

16

FIG. 1. Effect of pretreatment of ”P-labeled native T7 DNA with exonuclease 111 on its susceptibility to the N. crussa nuclease [from Linn and Lehman ( 2 ) ] . The abscissa gives the level of made acid-soluble after exonuclease 111 treatment; the ordinate gives the additional made acid-soluble by subsequent treatment with the N. crassu nuclease.

C. BIOLOGICAL ROLE Recent studies by Fraser and his colleagues have suggested that the N. endonuclease represents the end product of a series of proteolytic processing steps beginning with an inactive pronuclease (13, 14). Ronuclease can be purified from fresh mycelia. This form of the enzyme, which has a molecular weight of 88,000 based on sucrose density gradient sedimentation, can be activated by treatment with trypsin or by permitting the mycelial extracts to “age.” In the latter case activation is presumably a consequence of the action of endogenous proteases. The active enzyme, which has a molecular weight of 61,000, shows both endonuclease activity specific for single-stranded DNA and duplex DNA-specific exonuclease activity. Treatment of this form of the enzyme with trypsin or endogenous proteases results in its conversion to the single-strand-specific endonuclease that is devoid of exonuclease activity and has a molecular weight of

crassa

14. S. Kwoong and M. J. Fraser, Can. J . Biochern. 56, 370 (1978).

198

I. ROBERT LEHMAN

55,000 ( I , 2). It therefore appears that the single-strand-specific endonuclease and the double-strand-specific exonuclease activities are associated with a single polypeptide. This polypeptide is released into the culture medium from mycelia during the exponential phase of growth (15). The endo-exonuclease may function in recombination and DNA repair in vivo. In fact, in several respects it resembles therecBC nuclease ofE. coli, an enzyme that is required for normal recombination in this organism (16). Moreover, endo-exonuclease-deficientmutants of N. crussa (uvs-3 and nuh-4) have been isolated that are sensitive to ultraviolet light and to radiomimetic agents, and show an abnormally low frequency of mitotic recombination (17). In these two instances the level of inactive endoexonuclease precursor is higher than the wild type; thus the two mutations may result in the inability of the precursor to undergo conversion to the active form of the enzyme (18). A single-stranded DNA-specific endonuclease with properties identical to the Neurospora enzyme has been purified from the smut fungus Ustilago may& (7). It is also present at reduced levels in an ultravioletsensitive and recombination-defective mutant (8).

111.

Nucleare 51

Nuclease S1 was first identified by Ando (3) in preparations of “Takadiastase” from Aspergillus oryzae and purified some 1000-fold. It has subsequently been purified to near homogeneitv by Vogt (4). The purified protein has a sedimentation coefficient, S20,w of 3.3 and a molecular weight of 32,000. Like the N. crassa endonuclease, nuclease S1 appears to be a metalloenzyme. Dialysis of the enzyme against 1 mM EDTA results in its inactivation. Activity can be largely restored by the addition of either Co2+ or Zn*+. In contrast to the N . crassa enzyme, which has a broad pH optimum in the range of 7 to 9, nuclease S1 shows optimal activity at pH 4 . 6 4 . 3 . It is essentially inactive at pHs higher than 6.0. The enzyme is relatively insensitive to ionic strength; its activity at 0.4 M NaCl is 55% that at 0.1 M, the ionic strength at which it is optimally active. An exception is sodium phosphate, which at pH 4.6 inhibits the enzyme at concentrations as low 15. 16. 17. 18.

M. J. Fraser, Nucleic Acids Res. 6, 231 (1979). P. J. Goldrnark and S. Linn, JBC 247, 1849 (1972). E. Kafer and M. Fraser, M o t . Gen. Genet. 169, 117 (1979). T. Y.-K. Chow and M. J. Fraser, Can. 1. Eiochem. 57, 889 (1979).

11.

SINGLE-STRAND-SPECIFIC ENDONUCLEASES

199

as 10 mM. Although the rate of hydrolysis of denatured DNA is not greatly diminished at high ionic strengths, the extent of hydrolysis is reduced, presumably because of the tendency of DNA to renature under these conditions. Like the N. crassa endonuclease, nuclease Sl hydrolyzes both RNA and DNA; however, it is approximately fivefold more active on DNA than RNA. The products after extensive hydrolysis are nucleoside 5 ' monophosphates. The purified enzyme is highly specific for singlestranded DNA. Thus, under conditions where 96% of denatured DNA from bacteriophage lambda is rendered completely acid-soluble, less than one phosphodiester bond scission is introduced per ten molecules of native lambda DNA. On the other hand, both S1 and N. crassa endonucleases are able to recognize and cleave partially denatured regions in duplex DNA generated by the superhelicity of covalently closed duplex DNAs (19, 20) or by mismatches produced by mutational alterations in a DNA duplex (21). These enzymes can also cleave short single-stranded regions in duplex DNA (2, 22, 23) as well as the nonhydrogen-bonded loops in tRNA (24).

IV.

Mung Bean Endonucleate

A.

PHYSICAL PROPERTIES (3)

The mung bean nuclease has ,een obtainec in near homogeneous form. It has a molecular weight of 39,000 as determined by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate. Sucrose gradient sedimentation and Sephadex G-100 filtration yield values of 43,000 and 42,000, respectively. The enzyme contains one sulfhydryl group and three disulfide bonds. Approximately 70% of the enzyme molecules contain a peptide bond cleavage at a single site in the protein. Thus, reduction of the enzyme with 2-mercaptoethanol prior to gel electrophoresis in the presence of sodium dodecyl sulfate generates, in addition to the intact 39,000 dalton polypeptide two polypeptides of MW 25,000 and 15,000, 19. 20. 21. 22. 23. 24.

P. Beard, J . F. Morrow, and P. Berg, J . Virol. 12, 1303 (1973). A. C. Kato, K. Bartok, M. J . Fraser, and D. T. Denhart, BBA 308, 68 (1973) T. E. Shenk, C. Rhodes, P. W. J . Rigby, and P. Berg, PNAS 72, 989 (1975). J . E. Gerrnond, V. M. Vogt, and B. Hirt, EJB 43, 591 (1974). K. Shishido and T. Ando, BBA 390, 125 (1975). H. Tenehouse and M. J. Fraser, Can. J . Biochom. 51, 569 (1973).

200

I. ROBERT LEHMAN

which are presumably linked covalently by a disulfide bond. No difference in enzymatic activity can be detected between the intact and nicked forms of the enzyme. The mung bean endonuclease appears to be a glycoprotein containing 29% carbohydrate by weight. The mung bean enzyme is rapidly inactivated at pH 5, the pH at which it is optimally active. It can, however, be stabilized by the addition of 0.1 mM Zn2+and 1 mM cysteine or other thiols (glutathione or dithiothreitol). Dialysis of the enzyme at pH 5 results in complete loss in activity. However, activity can be restored by the addition of Zn2+and cysteine. In contrast, dialysis of the mung bean nuclease at pH 5 in the presence of 1 mM EDTA results in a loss of enzymatic activity that cannot be restored by the addition of Zn2+and cysteine; other divalent cations (Co2+,Mg2+, Mn2+,Caz+)are equally ineffective. Presumably a metal ion is removed by the EDTA dialysis that results in irreversible inactivation of the enzyme. B. REACTIONS CATALYZED (4) The mung bean endonuclease catalyzes the hydrolysis of singlestranded polyribo- and polydeoxynucleotides at approximately equivalent rates, to produce 5'-phosphoryl-terminatedmono- and oligonucleotides. It possesses an intrinsic 3'-nucleotidase which also acts on both mono- and oligonucleotides. Comparable endonuclease and 3'-nucleotidase activities are associated with a nuclease isolated from wheat seedlings (I I). Although showing a strong (up to 1000-fold) preference for polynucleotides lacking ordered structure, the mung bean endonuclease is less specific in this regard than the N. crassa or S1 nucleases. Thus, the mung bean endonuclease catalyzes as many as 50 double-strand cleavages in native T7 DNA at levels of enzyme that are required to convert denatured T7 DNA to >90% acid-soluble material. Under conditions that tend to destabilize the DNA duplex (i.e., lower ionic strength and increased temperature, 30" versus 22") native T7 DNA can be degraded completely. An analysis of the products generated under these conditions during the early phases of hydrolysis of native T7 DNA suggests that following an initially small number of endonucleolytic cleavages (possibly at very A-T rich regions), hydrolysis occurs preferentially from the ends of the duplex, generating a mixture of mono-, di-, and trinucleotides. Such a mode of hydrolysis is consistent with the degree of specificity of the mung bean nuclease for single-stranded polynucleotides. Thus, short single-stranded stretches that are formed transiently at the ends of the duplex, particularly under conditions of low ionic strength and elevated temperature, may be cleaved by the enzyme. Reiteration of this process should then lead to complete degradation of the DNA.

11.

SINGLE-STRAND-SPECIFIC ENDONUCLEASES

V.

Research Applications of Single-Strand-Specific Endonucleases

20 1

Because of their high degree of specificity for denatured regions in duplex DNA, all three of the enzymes described have served as useful probes for the identification and, if necessary, elimination of such regions. However, because of its very high degree of specificity for single-stranded as opposed to duplex DNA, its ease of preparation from readily available sources, and its relatively high specific activity the S1 endonuclease has become the enzyme of choice in the structural analysis of DNA and DNA-RNA hybrids. For example, the S1 enzyme has been widely used in DNA and DNA-RNA annealing experiments as a general reagent for the selective removal of nonhybridized, and hence single-stranded polynucleotide. The enzyme has also been utilized in mapping the location of small deletions in viral chromosomes. To cite a specific example, heteroduplex molecules formed from the complementary strands of a deletion mutant and wild-type SV40 DNA contain a single-stranded loop susceptible to the action of S1 nuclease at a position corresponding to the deletion. Incubation of the heteroduplex with S1 nuclease results in hydrolysis of the molecule at the loop, leaving fragments whose length corresponds to the position of the deletion within the SV40 molecule. In fact, by means of this technique S1 nuclease has been used to locate a deletion as short as 32 base pairs in the SV40 genome (21). S1 nuclease has been used widely for the analysis of spliced mRNAs generated as a consequence of intervening sequences in eukaryotic genes (25). The procedure consists of hybridization of unlabeled mRNAs to "P-labeled single-stranded DNA of high specific radioactivity. If the mRNAs are spliced, RNA-DNA hybrids flanked by single-stranded DNA, will result, together with loops of nonhybridized single-stranded DNA, at splice points in the mRNA. When these structures are treated with S 1 endonuclease, the single-stranded DNA is hydrolyzed, resulting in a fully duplex structure with discontinuities in the DNA at the splice points. These duplexes can be resolved and their size determined by electrophoresis through agarose gels.

25. A. J. Berk and P. A. Sharp, PNAS 75, 1274 (1978).


Exodeoxyribonucleases of Escherichia coli BERNARD WEISS

1. General Properties . 11. Specific Exonucleases A. Exonuclease I . . B. Exonuclease 111 .

. . . . C. Exonuclease IVA and IVB . D. Exonuclease VII . . . . . . E. Exonuclease VIII . . . . .

1.

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

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

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

. . . . . . .

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

203 206 206 211 225 226 229

General Properties

Deoxyribonucleases (DNases) catalyze the hydrolysis of phosphodiester bonds in polydeoxyribonucleotides. They are divided into two general classes: exonucleases and endonucleases. As classically defined ( I ), exonucleases hydrolyze linear chains stepwise from the 3' or 5' ends, releasing mononucleotides predominantly, whereas endonucleases attack chains more or less randomly at internal loci to release large fragments initially. Natural phenomena were not so easily categorized, however. DNases were soon discovered that attacked the ends of DNA molecules to release oligonucleotides as well as mononucleotides (Z), or to release 1. M. Laskowski, Ann. N . Y. Acad. Sci. 81, 776 (1959). 2. M. P. Deutscher and A. Kornberg, JBC 244, 3029 (1969).

203 T H E ENZYMES, Vol. XIV Copyright 0 1981 by Academic Press, Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

204

BERNARD WElSS

dinucleotides almost exclusively ( 3 ) , or to release high molecular weight fragments initially (4). Accordingly, the terms are redefined as follows: An endonuclease can attack a covalently closed circular strand, but an exonuclease cannot because it requires a chain terminus. If an exonuclease preparation is contaminated with a small amount of an endonuclease, it is capable of rapidly degrading circular DNA molecules. Therefore, the following additional or alternative criteria are frequently used: (i) An exonuclease releases mononucleotides or small oligonucleotides early in the course of a digestion. (ii) An exonuclease usually displays directionality, degrading a strand 3' + 5' or 5' + 3'. (iii) The activity of an exonuclease is often affected by the concentration of free substrate termini in a reaction mixture; it may be enhanced, for example, by shearing the DNA substrate or by partially predigesting it with an endonuclease, or it may be blocked by terminal substituents such as 3'-phosphornonoesters or acetyl groups. It should be noted, however, that the terms exonuclease and endonuclease are merely descriptive and do not represent mutually exclusive categories. Some enzymes, such as exonucleases 111 and V of E. coli, possess both exo- and endonucleolytic activities, i.e., they can at times hydrolyze circular molecules and are capable of initiating hydrolysis from the ends of linear strands. Exonuclease activity is usually assayed by measuring the conversion of an acid-insoluble polynucleotide substrate into acid-soluble mono- or oligonucleotides. This digestion may be followed by measurement of radioactivity (if a radiolabeled substrate is used) or of Azso.The products may be readily identified by chromatography or by electrophoresis. If the substrate is a x2P-labeledDNA, the average chain length of the products may be determined from the proportion of radioactivity released into a Norit-nonadsorbable (i.e., base-free) form by E. coli alkaline phosphatase (a phosphomonoesterase). The 3'- and 5'-mononucleotide products may be distinguished by their susceptibility to 3'- or 5'-nucleotidase. Directionality of attack may be established through techniques introduced by Lehman (S), using terminally labeled substrates. Thus, the DNA may be uniquely labeled at its 3' end with the aid of radiolabeled nucleoside triphosphates and a DNA polymerase, or it may be labeled at its 5' end with the aid of "P-labeled ATP and polynucleotide kinase. The substrate is then incubated with the exonuclease to see if the terminal label is released early or late during the course of digestion. Exonucleases can also be characterized with respect to their tendency 3. D. M . Trilling and H. V. Aposhian, PNAS 60, 214 (1968). 4. E. A. Friedman and H. 0. Smith, JBC 247, 2846 (1972). 5. I. R. Lehrnan, JBC 235, 1479 (1960).

12. E. coli EXONUCLEASES

205

to act in either a processive or in a distributive (random) manner (6). The termprocessive (used in the sense of “going forward”) describes the behavior of a nuclease molecule that binds to a polynucleotide and continues to degrade it without becoming unbound, whereas a distributive enzyme catalyzes a single cleavage first on one chain and then on another at random. Some enzymes are intermediate in behavior. Three techniques have been used to determine processivity . The first two depend on a high molecular ratio of substrate to enzyme in the reaction mixtures. In the first (6), one starts with a homogeneous substrate preparation and then examines the length distribution of molecules in a partial digest. A distributive enzyme yields partially digested molecules with a Gaussian length distribution. A processive enzyme, however, yields a mixture that appears to consist entirely of completely digested and completely intact molecules; partially digested molecules are rare because they are equal in number to the enzyme molecules to which they are bound. By the second method, Thomas and Olivera (7) measured the relative rates of release of nucleotides from the 3‘ and 5’ ends of a substrate, e.g., from [3H]d(pT)~-[32P]d(pT)z used alone or annealed to poly(dA). Because a processive exonuclease tends to degrade a strand from one end to the other before proceeding to a second strand, it appears to release labeled material from both ends of the substrate simultaneously. In the third method, exemplified by the studies of Wu et al. (a), one observes the effect of the addition of a second substrate to an ongoing reaction; a processive enzyme remains bound to the first substrate and continues for a while to degrade it at the previous rate. The interpretation of experiments on processivity and directionality are often dependent on a knowledge of the molecular ratio betweeen substrate and enzyme. At a high substrate-to-enzyme ratio, a processive 3‘ + 5’ exonuclease appears to degrade DNA from 3’ and 5‘ ends simultaneously, as described above. If higher levels of enzyme are used, enough to saturate the ends of the chains, the enzyme appears to be distributive by the first two methods cited above. Because it is rare in most other areas of enzymology to use stoichiometric amounts of enzyme, many investigators do not concern themselves with these considerations. When dealing with exonucleases, however, it is quite possible to inadvertently employ stoichiometric amounts of enzyme in a reaction mixture for the following reasons: (i) Because the substrate can be uniformly, and hence highly, radioactively labeled, it is tempting to use small amounts of it; (ii) the 6. N . G. Nossal and M. F. Singer, JBC 243, 913 (1968). 7. K. R . Thomas and B. M. Olivera, JBC 253, 424 (1978). 8. R. Wu, G. Ruben, B. Siegel, E. Jay, P. Spielman, and C. D. Tu,Eiochemisrry 15, 734 ( 1976).

206

BERNARD WElSS

conversion of a significant percentage of a high molecular weight substrate into an acid-soluble form requires many catalytic events per molecule, necessitating much enzyme; (iii) a very long DNA strand may contain only one binding site for the enzyme, e.g., a 3’-hydroxyl terminus. The following are a few additional observations bearing on the properties of the exonucleases to be described in the sections to follow. (i) With few exceptions, the activity of an exonuclease is a function of the concentration of chain termini (rather than that of DNA-phosphorus), therefore, the K , should be reported in those terms. (ii) Specificity for singlestranded substrates has been customarily determined with denatured DNA, but single-stranded DNA undergoes transient intrastrand basepairing, making it susceptible to double-strand-specific enzymes; a homopolymer substrate is more appropriate as a standard of comparison. (iii) Most, but not all DNases, require bivalent cations (usually Mg2+)for optimum activity, but it is not generally appreciated that the optimum amount is not only a function of the nature of the enzyme but also of the substrate concentration; for that reason, optimum cation concentrations are not described here. II. Specific Exonucleares

Table I outlines some of the distinguishing characteristics of each of the exonucleases of E. coli. They all cleave phosphodiester bonds so as to produce 5’-rather than 3’-phosphomonoesters. In cases where directionality of an enzyme is known, it is always at least 3‘ + 5 ’ . Some of the enzymes have, in addition, a 5‘ + 3’ activity. The exonucleases of E. coli were covered in a review of bacterial DNases by Lehman in 1971 (9). Since then, exonucleases VII and VIII have been discovered, and there have been many new findings with respect to exonucleases 111 and v. The sections below describe only exonucleases I, 111, IV, VJI, and VIII. Separate chapters in this volume deal with exonuclease V (see Linn, Chapter 13 this volume) and with the DNA polymerases (see Section I, Chapters 1-3, this volume). A. EXONUCLEASE I 1. Introduction

Exonuclease I, discovered by Lehman (51, is a 3’ + 5‘ exonuclease that is specific for single-stranded DNA. It attacks a DNA strand processively 9. I. R. Lehman, “The Enzymes,” 3rd ed., Vol. 4, p. 251, 1971.

207

12. E. c d i EXONUCLEASES TABLE I

EXONUCLEASES O F E .coli Enzyme Exonuclease I

Gene sbcB

(Exonuclease 11) Exonuclease 111

Exonucleases IVA and IVB Exonuclease V (recBC DNase)

Characteristics” Single-strand specific, processive; leaves 5’-terminal dinucleotide intact 5’ exonuclease activity of Former name for 3‘ DNA polymerase I (see below) Double-strand-specific; associated activities: DNA-3’-phosphatase, AP endonuclease, and RNase H Single-strand-specific; degrades oligonucleotides completely; not well characterized* ATP-dependent; both 3’ + 5’ and 5’ --+ 3’; double-strand specific, processive; inactive at a nick; releases large oligonucleotides initially; also a single-strand specific endonuclease; a recombination enzyme Former name for 5’ + 3‘ exonuclease activity of DNA polymerase I (see below) Single-strand specific, processive; EDTAresistant; both 3’ -+ 5’ and 5‘ + 3’; oligonucleotide products Double-strand specific; not well characterizedb; product of the cryptic Rac prophage of E. coli K12; detected only in sbcA mutants 3‘ 5‘ activity that is single-strand specific; 5’ 3’ activity: double-strand specific; monoand oligonucleotide products; RNase H activity; repair functions 3’ 5’ only; single-strand specific 3’ + 5’ activity: single-strand specific; does not 3’ activity: singleattack dinucleotides; 5’ strand specific but can attack a duplex after initiating hydrolysis at a single-stranded end --+

xthA

recBC

(Exonuclease VI) Exonuclease VII

xse

Exonuclease VIII

recE

DNA polymerase I

polA

DNA polymerase I1 DNA polymerase Ill

polB p0lC

-+

-

-+

--+

‘I

Unless otherwise noted, directionality is 3‘ + 5’ and products are 5’-mononucleotides.

* Directionality is unknown.

(7), releasing 5‘-mononucleotides,and degrading it down to its 5’-terminal dinucleotide, which it leaves intact ( 1 0 ) (Fig. 1).

2. P~r$cationand Properties Purification methods have relied on the tendency of the enzyme to bind to DNA. In earlier procedures, the enzyme was coprecipitated with DNA 10. 1. R. Lehman and A. L. Nussbaum, JBC 239, 2668 (1964).

208

BERNARD WEISS

FIG. 1. The exonuclease I reaction: processive 3’ 5’ hydrolysis of 5’mononucleotides from single-stranded DNA, sparing the 5‘-terminal dinucleotide. n = number of nucleotides in the original chain. --f

when crude extracts of E. coli B were treated with protamine (5) or with streptomycin ( l o ) , and was then further purified by chromatography. In one method, it was obtained as a by-product of the purification of DNA polymerase I (10). Subsequently, the enzyme has been purified about 1200-fold to apparent homogeneity with the aid of affinity chromatography on denatured DNA-cellulose (1 1 ). The gene for exonuclease I was incorporated into multicopy bacterial plasmids (ColE 1 derivatives) by molecular cloning technology, yielding a strain that overproduced the enzyme 15-fold (12); further amplification may be possible with other cloning vectors. Such strains are preferred sources for the enzyme. Estimates of 70,000 and of 72,000 for the molecular weight of the exonuclease I protomer are based on electrophoresis in SDSpolyacrylamide gels (11, 13). An earlier value of 43,000 obtained by Sephadex gel filtration (see 15) was erroneously low, probably because of retardation of the protein under conditions of low ionic strength (13). The Lehman and Nussbaum method of purification, from E. coli B ( l o ) ,yields predominantly a monomeric protein; that is, the molecular weight of the enzyme as determined under nondenaturing conditions, by gel filtration and velocity sedimentation, agrees with that obtained by SDS-gel electrophoresis for the enzyme that has been denatured and reduced. Another method of purification, from E. coli K12 (/I), yields predominantly a different form of the enzyme, probably a dimer. Although its electrophoretic mobility in SDS is similar to that of the Lehman and Nussbaum preparation, it migrates more slowly than the latter under nondenaturing conditions. Both forms of the enzyme are active ( / I ) . 3 . The Reaction Catalyzed The enzyme is usually assayed by measuring the degradation of radiolabeled, heat-denatured DNA to acid-soluble products (10). Because the glucosylated DNA of bacteriophage T4 is resistant to most other 1 1 . R. K . Ray, R. Reuben, I. Molineux, and M. Gefter, JBC 249, 5379 (1974). 12. D. Vapnek, N. K. Alton, C. L. Bassett, and S. R. Kushner, PNAS 73, 3492 (1976). 13. V. MacKay and S. Linn, BBA 349, 131 (1974).

12. E . coli EXONUCLEASES

209

enzymes (14),its use as a substrate enhances the specificity of the assay (15).The enzyme requires a pH of 9.5 and the presence of Mg2+ for optimum activity. Studies with mutants indicate that with this assay at least 90% of DNase activity measured in crude extracts ofE. coli is due to exonuclease I(15). Exonuclease I has been characterized extensively (5, 10). It has no measurable activity on double-stranded DNA or RNA. An apparent exception is its ability to degrade almost completely some duplexes composed of strands of homopolymers or of an alternating copolymer, such as poly(dA) . poly(dT) or poly(dA-dT). The probable explanation is that the spontaneous breakage and reformation of hydrogen bonds in such molecules causes the creeping or slippage of one strand over another so that a single-stranded 3' end is always presented to the enzyme during digestion. The enzyme, however, is unable to digest poly(dG) * poly(dC), which has a more highly ordered structure. If a duplex substrate contains an unpaired 3' end, the enzyme digests the single-stranded region to within 6 to 8 nucleotides of the base-paired region (16). Exonuclease I attacks a DNA chain only at a free 3'-hydroxyl terminus. It is blocked by 3'-phosphoryl or 3'-acetyl end groups, and its action on strands with 3'-hydroxyl termini is inhibited in the presence of strands with 3'-phosphoryl termini. Oligonucleotides with chain length as low as 3 to 6 are also attacked, but with an estimated I(, about lo6higher than that for denatured DNA. Dinucleotides are resistant to cleavage. 4. Biological Role Exonuclease I is specified by the sbcB locus at 43.6 min (near his) on the genetic map of E. coli (17). The locus is near a major integration site for bacteriophage P2. Consequently, sbcB is among the genes lost by P2 eduction, a deletion of chromosomal genes accompanying the spontaneous curing of P2 lysogens. The fact that the eductants have no growth defect in nutrient media indicates that exonuclease I is dispensible. The proximity of the gene to a prophage attachment site, as well as its superfluous nature, raises the evolutionary possibility of a phage origin for the gene. Mutants lacking exonuclease I have no demonstrable defects in replication, recombination, or repair (15,18). On the contrary, the normal pres14. C. C. Richardson, JBC 241, 2084 (1966). 15. D. M. Yajko, M. C. Valentine, and B. Weiss, JMB 85, 323 (1974). 16. D. Brutlag and A. Kornberg, JBC 247, 241 (1972). 17. B. J . Bachmann and K . B. Low, Microbid. Rev. 44, 1 (1980). '18. S . R . Kushner, H. Nagaishi, A . Templin, and A . J. Clark, P N A S 68, 824(1971); S. R. Kushner, H. Nagaishi, and A. J. Clark, PNAS 69, 1366 (1972).

2 10

BERNARD WEISS

ence of the enzyme in wild-type E. coli has a deleterious effect on a recombination pathway. As reviewed by Clark (19), E. coli has several pathways for general (homologous) genetic recombination. One requires exonuclease V (the recBC gene product) and another requires the recF gene. recBC mutants are recombination-deficient, but this deficiency can be reversed by an sbcB mutation, provided the recF gene is functional (“sbc” is an acronym for “suppressor of recBC”). Presumably the sbcB gene product destroys a DNA intermediate in the recF pathway of recombination, so that the recF pathway does not normally operate in wild-type E. coli. There is a set of exonuclease I-deficient mutants, previously designated xonA (15), that do not suppress the deficiency of recBC mutants in assays for genetic recombination. They do, however, suppress to a variable extent their sensitivity to mitomycin C and to ultraviolet irradiation. Some xonA mutants produce a temperature-sensitive exonuclease I and are, therefore, known to be altered in the structural gene for that enzyme. The xonA and sbcB mutations belong to the same complementation group and are about equally contransducible with the his operon ofE. coli. While it is still possible that the sbcB mutations might be polar mutations affecting separate genes for exonuclease I and for the suppression of recombination deficiency, it is more likely that the xonA mutants are merely leaky sbcB mutants. For that reason, sbcB is the current designation for the exonuclease I gene (17). +

5 . Research Applications Exonuclease I has been used in a sensitive assay for single-strandspecific endonuclease activity. The assay is based on the principle that a single endonucleolytic cleavage in a circular DNA strand renders it almost completely degradable by exonuclease I (20). It can also be used to remove traces of single-stranded DNA in preparations of double-stranded DNA, but for this purpose it is more practical to use the readily obtainable endonucleases from Aspergillus oryzae ( S 1 nuclease) or from Pseudomonas BAL-31. Exonuclease I has enabled the measurement of the average length of single-stranded 3‘ tails in duplex molecules ( 2 / ) ,and its unique ability to degrade the glucosylated DNAs of the T-even phages (after DNA denaturation) has facilitated their chemical analysis (22). 19. 20. 21. 22.

A. J. Clark,Annu. Rev. Genet. 7, 67 (1973). P. Sadowski and J. Hurwitz, JBC 244, 6192 (1969). V. MacKay and S . Linn, JEC 249, 4286 (1974). I. R. Lehrnan and E. A. Pratt, JEC 235, 3254 (1960).


21 1

Because of its inability to attack a 5'-terminal dinucleotide, exonuclease I has remained an invaluable tool for studying the 5' ends of DNA. Limit digests produced by exonuclease I have been used to isolate and to identify the 5'-terminal dinucleotides in native DNAs (23) as well as those at the new termini created by site-specific endonucleases (24). In a similar manner, the DNA-5'-adenylate intermediate in the DNA ligase reaction was characterized after isolation of terminal deoxydinucleotides covalently bound to AMP (25). The digestion of high molecular weight DNA by exonuclease I is seldom more than 90% complete (10). For digestion up to the terminal dinucleotide, therefore, the enzyme is usually used in conjunction with an endonuclease, such as pancreatic DNase I. The endonuclease bypasses any .factitious 3'-phosphoryl termini and intrastrand duplex regions, and it increases the concentration of substrate termini.

B. EXONUCLEASE I11 1. Introduction

a. Description. Exonuclease I11 is a monomeric protein (MW = 28,000) with four catalytic activities (Fig. Z), namely (it a 3' 5' exonuclease specific for bihelical DNA (exodeoxyribonuclease activity) and (ii) for the RNA strand in an RNA-DNA hybrid duplex (RNase H activity); (iii) it hydrolyzes 3'-terminal phosphomonoesters fiom DNA (DNA-3'phosphatase activity); and (iv) it cleaves DNA endonucleolytically at an apurinic or apyrimidinic site (AP endonuclease activity) creating base-free deoxyribose 5-phosphate end groups. 6 . History. In 1964 Richardson and his colleagues (26) described a DNA phosphatase-exonuclease that was present in partially purified preparations of DNA polymerase I. It was discovered as a protein that enhanced the ability of calf thymus DNA to serve as a primer for the polymerase; it did so presumably by removing inhibiting 3'-phosphoryl end groups from the DNA. The phosphatase and exonuclease activities were copurified and shown to be inactivated by heat at the same rate, suggesting that they were properties of the same enzyme. In addition, the enzyme was shown to degrade a mixed copolymer of ribo- and 23. B. Weiss and C. C. Richardson, J M B 23, 405 (1967). 24. T.J. Kelly and H. 0. Srnith,JMB 51, 393 (1970). 25. I. R. Lehrnan, Srience 186, 790 (1974). 26. C. C. Richardson and A. Kornberg, JBC 239, 242 (1964); C. C. Richardson, I. R. Lehrnan, and A. Kornberg, ibid., p. 251.

212

BERNARD WEISS (a) exodeoxyri bonuclease :

; - I

;

-

_-

_"Ill - *

Ill

I

-.

'

*-

(b) RNase H

(c) DNA-3'-phosphatase

---m ---m (dl A P endonuclease

--

-m - - ---- m--

---P

sQ3--- 5: P

P

P

---P

PJ o $ p J - - -

FIG.2. The reactions catalyzed by exonuclease 111. deoxyribonucleotides, a finding from which it was later assumed that it had RNase H activity (27). The RNase H activity was directly demonstrated in 1972 by Keller and Crouch (28). The laboratories of W.Verly and D. Goldthwait were largely responsible for the initial discovery that E. coli extracts were capable of enzymatic cleavage at apurinic or apyrimidinic sites in DNA. Such sites are places where a base has been removed from DNA by hydrolysis of the glycosylic bond that joins it to a deoxyribose residue, leaving the sugarphosphate backbone intact. Depurination, which can occur spontaneously, is accelerated at elevated temperatures and at low pH. In these early experiments, the preparation of a partially depurinated substrate was facilitated by prior treatment of the DNA with methyl methanesulfonate, an alkylating agent; its major adducts are methylated bases (e.g., 7-methylguanine and 3-methyladenine) with unstable glycosylic bonds. 27. 1. Berkower, J . M. Leis, and J. Hurwitz,JBC 248, 5914 (1973). 28. W. Keller and R. Crouch, P N A S 69, 3360 (1972).


213

In 1969 Friedberg and Goldthwait (29) described “endonuclease 11,” an activity specific for alkylated DNA, but they did not know which of the many alkylation-produced lesions were specifically recognized by the enzyme. Verly and Paquette (30) reported an “endonuclease for apurinic sites” in extracts ofE. coli. When Hadi and Goldthwait ( 3 / ) found that endonuclease I1 preparations cleaved DNA that had been partially depurinated by acid, Paquette et al. (32) suggested that the two enzymes might be the same. They found that their AP endonuclease could be purified by the method used for endonuclease I1 and that it catalyzed cleavage only at the apurinic (alkali-labile) sites in an alkylated substrate, i.e., the DNA could be cleaved by the enzyme only to the extent that it could be cleaved by NaOH. That this endonucleolytic activity was catalyzed by exonuclease I11 was unsuspected until later work with bacterial mutants. Milcarek and Weiss (33) isolated a set of mutants deficient in exonuclease 111, as measured by the exonuclease and DNA-3’-phosphatase assays. Yajko and Weiss isolated another set deficient in “endonuclease 11” activity (W), as measured by an assay for the breakdown of alkylated DNA. When Marcus Rhoades discovered by chance that an “endonuclease 11” mutant was deficient in exonuclease I11 activity, all the mutants were analyzed and found to be deficient in both activities (34, 35). If one activity was unusually thermolabile in a mutant, so was the other, indicating that they were specified by a common structural gene. The mutations were also corevertible and cotransducible, and the multiple activities were specified by cloned DNA fragment about 3 kilobaae pairs in length (36). The endonuclease, exonuclease, and phosphatase activities were also copurified with a homogeneous monomeric protein (37). c. Nomenclature. From about 1973 to 1978 there was much apparent controversy and confusion in the literature. It stemmed more from conflicting terminology than from conflicting data. The term “endonuclease 11” was initially defined as an enzymatic activity purified by a standard 29. E. C. Friedberg and D. A. Goldthwait, PNAS 62, 934 (1969). 30. W. G . Verly and Y. Paquette, Can. J . Biochem. -50, 217 (1972). 31. S-M.Hadi and D. A. Goldthwait, Biochemisrry 10, 4986 (1971). 32. Y. Paquette, P. Crine, and W. G . Verly, Can. J . Biochem. 50, 1 1 9 9 (1972). 33. C. Milcarek and B. Weiss, J M B 68, 303 (1972). 34. D. M. Yajko and B. Weiss, PNAS 72, 688 (1975). 35. B. Weiss, S . G . Rogers, and A. F. Taylor, In. “DNA Repair Mechanisms” (P. C. Hanawalt, E. C. Friedberg, and C . F. Fox, eds.), p. 191. Academic Press, New York, 1978. 36. S . G . Rogers and B. Weiss, Gene, 11, 187 (1980). 37. B. Weiss, JBC 251, 1896 (1976).

214

BERNARD WEISS

scheme and detected by a standard assay (38); the equation for its reaction could not be written. The assay employed alkylated DNA and probably measured cleavage at apurinic sites (32) due mainly to exonuclease I11 (35). Later, the purification scheme (39) and assay (40) were changed, and “endonuclease 11” was used to refer to an enzyme that cleaved DNA at alkylated residues rather than at depurinated sites, and was clearly separable from exonuclease I11 (40, 41). Hence, there was confusion about the identity of exonuclease 111, endonuclease 11, and Verly’s enzyme for apurinic sites. When the existence of endonuclease I1 (as newly defined) could not be confirmed by its discoverers (42), the term was dropped. Meanwhile, Verly and Rassart (43) had purified to homogeneity the main AP endonuclease of E. coli. Its physical and enzymatic properties were not distinguishable from those of exonuclease I11 (44), which is responsible for about 85% of the AP endonuclease activity in crude extracts (35). Nevertheless, Gossard and Verly (44) referred to it as “endonuclease VI,” mainly to separate it from the controversy that surrounded “endonuclease 11” rather than to imply that it was distinct from exonuclease 111. For simplicity, we should adhere to the principle of one name per enzyme, and it is therefore urged that the term “endonuclease VI” be dropped in favor of the more familiar designation “exonuclease 111.” “Endonuclease VI activity” should therefore be referred to as “the AP endonuclease activity of exonuclease 111.” We will thus follow the convention by which the terms “exonuclease 11” and “exonuclease VI” were abandoned. d. Similar Enzymes. Although Hemophilus injuenzae is distantly related to E. coli, it possesses an enzyme that is similar to E. coli exonuclease 111 with respect to its physical properties and its multiple catalytic activities (45, 46). Streptococcus pneumoniue also contains a phosphatase-exonuclease similar to exonuclease 111, but AP endonuclease and RNase H activities have not been reported for it (47). 38. E. C. Friedberg, S-M.Hadi, and D. A. Goldthwait, JEC 244, 5879 (1969). 39. S-M. Hadi, D. Kirtikar, and D. A. Goldthwait, Biochemistry 12, 2747 (1973). 40. D. M. Kirtikar, G . R. Cathcart, and D. A. Goldthwait, PNAS 73, 4324 (1976). 41. D. M. Kirtikar, G. R. Cathcart, J. G. White, I. Ukstins, and D. A. Goldthwait, Biochemistry 16, 5625 (1977). 42. D. M. Kirtikar, G . R. Cathcart, J. G. White, I. Ukstins, and D. A. Goldthwait, Biochemistry 17, 4578 (1978). 43. W. G. Verly and E. Rassart, JEC 250, 8214 (1975). 44. F. Gossard and W. G . Verly, EJB 82, 321 (1978). 45. J. K. Gunther and S. H. Goodgal, JBC 245, 5341 (1970). 46. J. Clements, S. G. Rogers, and B. Weiss, JEC 253, 2990 (1978). 47. S. Lacks and H. Greenberg, JEC 242, 3108 (1967).

215

12. E. coli EXONUCLEASES TABLE I1

MOLECULARPROPERTIES OF EXONUCLEASE 111" Property Stokes radius, a Sedimentation coefficient, s20,w Frictional ratio,$!!,,, Molecular weight, native enzyme'' Molecular weight, reduced and denatured enzyme'

Value 2.29 nm 2.92 S 1.15

27,400 28,500

From Weiss (37). From sedimentation and diffusion coefficients with an assumed value for the partial specific volume. ' Determined by SDS gel electrophoresis. "

2. Purijicution and Properties Exonuclease 111 has been purified by ordinary methods of solvent or salt fractionation followed by ion exchange chromatography ( 2 6 , 3 7 , 4 3 , 4 8 ) In . one scheme (481, it was obtained as a by-product in the large-scale preparation of DNA polymerase I, from which it was separated by gel filtration chromatography. Two methods yielded preparations that were practically homogeneous by SDS gel electrophoresis (37, 43). Starting with wild-type cells, a 1600-fold purification was required. Currently, the enzyme is most easily purified from a strain that bears the gene for exonuclease 111 on a multicopy plasmid that is thermoinducible for DNA replication (36). Such cells can be induced to overproduce the enzyme 50- to 100-fold and to yield about 1 to 2 mg of pure (>98% homogeneous) enzyme per gram of cell paste after only a 15- to 30-fold purification (49). The physical parameters of exonuclease I11 (37) are summarized in Table 11. According to this data, the enzyme is a globular monomeric protein of MW = 28,000. Verly and Rassart (43) obtained values of 32,000 to 33,000 for the molecular weight of their endonuclease for apurinic sites, based on SDS gel electrophoresis and gel filtration chromatography; these differences are probably within experimental error. 3 . The Reactions Cutcilyzed u . Generul Properties. Rates for each of the four enzymatic reactions have been determined at equivalent unsaturating concentrations of each 48. T. M. Jovin, P. T. Englund, and L. L. Bertsch, JBC 244, 2996 (1969). 49. S. G. Rogers and B . Weiss, "Methods in Enzymology," Vol. 65, p. 201, 1980.

2 16

BERNARD WEISS

substrate and under optimum conditions for each assay (SO). These relative rates of phosphodiester bond cleavage are as follows: exodeoxyribonuclease, 100; AP endonuclease, 37; DNA-3'-phosphatase, 22; and RNase H [on poly(rA) . poly(dT)], 12. The pH optimum for the endo- and exonucleolytic activities is between 7.6 and 8.5 in Tris . HCl buffer (26,37); a lower pH optimum (6.8-7.4) has been reported for the phosphatase activity in phosphate buffer (26). The enzyme is partially active in the absence of added divalent cations, but this activity can be inhibited by EDTA, suggesting that the purified enzyme contains bound metal ions. Mg2+or Mn2+are required for optimum activity; they are about equally effective and can reverse the inhibition caused by EDTA. Ca2+can substitute for Mg2+in the AP endonuclease reaction (SO) but not in the exonuclease or phosphatase reactions. The exonuclease activity normally seen in the absence of added divalent cations is inhibited by the Ca'+. If Ca'+ is used in the AP endonuclease reaction, therefore, one can minimize the exonucleolytic degradation that would normally occur after the endonucleolytic cleavage (50). A similar effect has been reported for citrate in the presence of Mg2+(51). Zn2+ inhibits exonuclease I11 even in the presence of Mg'+ (>90% at lop4M ) (26 ) . Exonuclease I11 is inhibited by the sulfhydryl reagent p-chloromercuribenzoate (50-90% at M). Although dispensible during the standard assay, 2-mercaptoethanol stabilized the enzyme during prolonged incubations (26). b. Exodeoxyribonuclease Reaction. Exonuclease I11 has a 3' + 5' exodeoxyribonuclease activity specific for double-stranded DNA. It releases 5'-mononucleotides from the 3' end of DNA molecules so that partially digested duplexes have projecting 5' tails. The standard assay for the enzyme measures the exonucleolytic release of acid-soluble radioactive material (i.e., nucleotides) from radiolabeled linear DNA duplexes. The enzyme is highly specific for bihelical substrates; it has no detectable activity on single-stranded homopolymers. Because of intrastrand hydrogen-bonding, single-stranded DNA is also a substrate for the enzyme, but it is degraded at only + t o 3 the rate of duplex DNA (33). The glucosyl residues of bacteriophage T4 DNA render it largely resistant to the exonucleolytic activity of exonuclease I11 ( I # ) , whereas it remains a good source of substrate for the endonuclease and phosphatase reactions (33, 34). 50. S. G . Rogers and B. Weiss, unpublished data. 51. S. Ljungquist, B. Nyberg, and T. Lindahl,FEBS (Fed. Eur. Biochem. Soc.)Lerr. 57, 169 (1975).


2 17

The action of exonuclease III at a 3' terminus is not blocked by a phosphomonoester (which it can hydrolyze through its phosphatase activity), by one or two paired or mismatched ribonucleotides (52),or by up to 3 mispaired deoxyribonucleotides (16). The enzyme can initiate hydrolysis at a single-strand break, but not at the low temperatures and moderate salt concentrations that favor base-stacking across the nick (53). Under standard assay conditions (37), one molecule of the enzyme cleaves 150 phosphodiester bonds per minute. As the digestion proceeds, the duplex portion of the substrate becomes progressively shorter until there is an insufficient number of base pairs to hold the strands together. After 40 to 50% digestion of a duplex DNA preparation, the rate of hydrolysis slows abruptly because the remaining DNA is mostly singlestranded (26). When the substrates are poly(dA) . poly(dT) or poly(dAdT), the kinetics of degradation are different from those seen with naturally occurring DNAs. These synthetic polymers can maintain duplex structures even after 50% hydrolysis because their strands can slip or creep over one another through the transient breakage and reformation of hydrogen bonds. The exonuclease reaction therefore continues almost unabated until almost 100% of the substrate is digested. At 25" a strand can be degraded up to its 5'-terminal dinucleotide; at higher temperatures the limit products are longer 5'-terminal oligonucleotides (-52),probably because of the reduced stability of oligonucleotide duplexes. The mode and extent of degradation are influenced greatly by the incubation temperature. At 5" it is limited; at 23" it is processive; and at 37" it is largely distributive. Thus, at 5", in the presence of 270 mM NaCI, an enzyme molecule removes only 6 nucleotides from a 3' terminus and remains bound to the DNA in a complex that can be isolated by gel filtration chromatography: the addition of more enzyme leads to the removal of 6 more nucleotides by each newly bound enzyme molecule (53). The results suggest that enzyme molecules are bound in fixed positions and have a limited reach, or that their translocation is restricted under these conditions. Donelson and Wu (-5.3) found that the release of mononucleotides was not influenced by the length of the single-stranded 5' tail, and concluded therefore that the binding was not to the very end of that strand. When the temperature of their reaction mixtures or of their isolated enzyme-DNA complexes was shifted to 223", there was unlimited digestion of the polynucleotide substrates. A striking feature of exonuclease 111 is that at 23-28", in the presence of enough enzyme to saturate the DNA termini, the digestion of a homogeR. Roychoudhury and R. Wu, JBC 252, 4786 (1977). 53. J . E. Donelson and R. Wu,JBC 247, 2661 (1972).

52.

2 18

BERNARD WEISS

neous preparation of linear duplexes proceeds synchronously (? 5%) for the first 250 nucleotides (8). At 23" the enzyme is processive for at least 100 nucleotides as determined by an experiment in which the ongoing degradation of radiolabeled DNA by a limiting amount of enzyme was not impeded by the subsequent addition of a large excess of unlabeled substrate (8). At 37", however, the enzyme appears to work in a distributive or nonprocessive fashion; Thomas and Olivera ( 7 ) , using substrates with uniquely labeled 3' and 5' termini, found that exonuclease I11 removed less than 50 nucleotides before dissociating from a chain terminus. Exonuclease I11 may act in a halting or skipping fashion on DNA that is tightly bound to protein. Riley and Weintraub (54) found that when exonuclease I11 digests the 140-base pair DNA of the nucleosome core, it does so in a 3' -+5' direction on each strand, removing 10 nucleotides at a time. It was not determined if they were released as decanucleotides or as mononucleotides. The reaction required larger amounts of enzyme than for the digestion of free DNA, and it appeared to be nonprocessive, or else processive for 510 bases at a time. c. RNuse H Activity. Exonuclease I11 catalyzes the selective degradation of the RNA strand in an RNA-DNA hybrid duplex; the RNA strand is degraded at 10,000 times the rate of the DNA strand (35). 5'ribomononucleotides are released in what is presumably a 3' + 5' exonucleolytic reaction. This activity is distinct from that of the enzyme called RNase H of E. coli, which is an endonuclease, and from the RNase H activity associated with the 5' + 3' exonucleolytic activity of DNA polymerase I. RNase H activity is detected by the release of acid-soluble material (mononucleotides) from a radiolabeled ribohornopolymer in the presence of the complementary unlabeled deoxypolynucleotide, e.g., . poly(dT). There is no measurable activity of exonufrom p~ly([~H]rA) clease I11 on single- or double-stranded RNA molecules. At the time of its discovery, exonuclease I11 was found to be capable of degrading a mixed copolymer containing about one ribonucleotide for every eight deoxyribonucleotides (26). Without direct demonstration, exonuclease I11 was assumed to have RNase H activity (27) and was therefore used by Keller and Crouch (28) as an experimental control in studies of other enzymes with RNase H activity. These latter studies demonstrated that exonuclease I11 could attack poly(rA) poly(dT), break the phosphodiester bonds between ribonucleotides, and release 5 ' mononucleotides exclusively; however, the authors employed excessive amounts of an enzyme preparation of unstated purity and found that the DNA strand was also completely degraded. Roychoudhury and Wu (52) 54. D. Riley and H. Weintraub, Cell 13, 281 (1978).


2 19

found that the enzyme catalyzes the consecutive cleavage of two ribonucleotides from the 3’ end of a DNA duplex, thus suggesting a 3’ + 5 ‘ exonucleolytic mechanism. Finally, it was demonstrated that the RNase H activity copurifies with the exonuclease, and that the highly purified enzyme preferentially degrades the RNA strand in a hybrid duplex (35). This unusual specificity gives us some insight into how exonuclease 111 and perhaps some other double-strand-specific enzymes recognize their substrates. The simplest way for an enzyme to exercise specificity for a DNA-containing duplex is to recognize a deoxyribose (or even just a 2‘-hydrogen) on one strand and to attack the other strand. Thus, presented with an RNA-DNA hybrid duplex, exonuclease 111 binds to the DNA strand and degrades the RNA strand, but not vice versa (55). d. DNA--?’-phosphatuse Activity (26). 3’-terminal phosphomonoesters are hydrolyzed from DNA by exonuclease 111 to yield 3’-hydroxyl end groups and inorganic phosphate. The DNA is then subject to exonucleolytic degradation by the enzyme. The rate of hydrolysis of 3’phosphomonoesters from duplex DNA is about twice that for heatdenatured DNA. The enzyme has no measurable activity on 3‘phosphoryl-terminated RNA or on d(T-T-Tp), but it attacks a mixed polymer terminated at its 3’ end by ribonucleoside 3’-phosphate residues. Originally the activity was measured with a substrate consisting of uniformly labeled [32P]DNA into which 3’-phosphoryl termini were introduced through partial endonucleolytic digestion with micrococcal nuclease. The assay measured the release of radioactive material in an acidsoluble, Norit-nonadsorbable (base-free) form, i.e., as 34Pi. With crude extracts, this assay is subject to about 15% interference by other enzymes, presumably by other exonucleases coupled with 5’-nucleotidases. An alternative assay, employing a substrate uniquely labeled at its 3‘ termini, is >99% specific for exonuclease I11 in crude extracts (33). e . A P Endonucfease Reaction. Exonuclease I11 cleaves DNA molecules 5‘ to an apurinic or apyrimidinic site. At the newly created termini are nucleoside 3’-hydroxyl end groups and base-free deoxyribose 5-phosphate residues (35). Apurinic sites have been shown to be substrates for the enzyme even after their reaction with NaBH4 @), a treatment that reduces the free aldehyde groups on the base-free sugars. The enzyme has no measurable endonucleolytic activity on free DNAs lacking AP sites. It does, however, make endonucleolytic cleavages in chromatin 55. Curiously, this mechanism was presented initially only for the purpose of simplifying a figure (Ref. 37 and Fig. 3, Section II,B,3,f). It was later taken seriously, and its chief prediction, namely, the sparing of the DNA strand in the hybrid duplex, was tested and verified.

220

BERNARD WElSS

DNA at a point where the 5' end of a strand projects beyond the nucleosome core (54, 5 6 ) . The reaction is demonstrated most specifically by the cleavage of partially depurinated supercoiled DNA. 4x174 RFI VHIDNA (covalently closed circular duplexes) is treated with acid to introduce about 2 apunnic sites per molecule. The assay substrate is rapidly renaturable, but nicking at AP sites by exonuclease I11 permits the strands to remain separated after a mild denaturing treatment and so to be adsorbed to nictrocellulose filters (46). Other assays have measured the degradation of partially depurinated linear duplexes, either by the release of large fragments from DNA entrapped in a polyacrylamide gel (38)or by the release of acidsoluble material (4.7); they are less specific because they measure exonucleolytic degradation as well. A P sites have been introduced into these substrates by treatment of the DNA with alkylating agents or acid (43) or by the treatment of uracil-containing DNA with uracil-DNA glycosylase (46). Concomitant exonucleolytic degradation has been reduced by using glycosylated T4 phage DNA, citrate, or Ca'+ in the reaction mixture (see a , this section). f. Reaction Mechanisms. The relatively low molecular weight of exonuclease I11 suggests that a single active site may be responsible for all of its catalytic functions. According to the common-site model (Fig. 3), the enzyme contains three regions. One region recognizes adeoxyribose on the strand opposite the one that is cleaved, thus conferring a specificity for DNA-DNA or DNA-RNA duplexes. A second region recognizes and cleaves phosphoester bonds. A third region recognizes a topological feature common to all of the substrates, namely, a space created by a missing or displaced base. In the endonuclease substrate, the space is at an AP site. In the phosphatase substrate, the space is created by the missing terminal nucleoside. In the exonuclease substrates, an interstrand space is created by the transient unwinding that occurs commonly at ends of duplexes because the terminal bases are less constrained by stacking forces than is the rest of the DNA. Although alternatives are not precluded, the following observations are consistent with the common-site model: (i) Unlike other enzymes that work at 3' ends (such as DNA polymerases or exonuclease I), the enzyme does not require a free 3'-hydroxyl group, but attacks 3'-phosphorylterminated DNA; it therefore probably recognizes some other topological feature. (ii) Similarly, the enzyme does not recognize the chemical group 56. Perhaps, during the binding of DNA to histones, some nucleoside residues become unpaired and are rotated away from the central axis of the double helix to leave spaces that resemble AP sites (see Section II,B,3,D.

22 1


( A ) Enzyme:

(C)3' - Phosphatase

3 recognition sites

(B)Endonuclease

(D) Exonuclease

FIG. 3. The common-site model for exonuclease 111. The enzyme (shaded area) is shown attacking duplexes, cleaving phosphoester bonds at the arrows. The enzyme (A) is pictured as having three regions. One region recognizes and cleaves phosphoester bonds; a second region recognizes the duplex structure by recognizing a deoxyribose on the strand opposite the one that is cleaved. The thud region recognizes a space created by (B) a missing base, (C) a missing 3'-terminal nucleoside, or (D) terminal unwinding. From Weiss (-37), with permission. unique to an AP site, i.e., a free aldehyde; it cleaves at an AP site even if the aldehyde is reduced (44). (iii) The enzyme does not care whether a purine or pyrimidine is missing at the AP site (351,so it probably recognizes the space itself rather than some feature of the unpaired base. (iv) Although the exonuclease is specific for duplex substrates, it removes from one to three mispaired nucleotides from the 3' terminus (16). (v) The enzyme does not attack DNA at single-strand breaks at 5" in 70 mM NaCl ( 5 3 , conditions that favor the stacking of bases across the nick. (vi) In all the exonuclease I11 mutants that have been studied (12 of independent origin), the exonuclease, phosphatase, and endonuclease activities are similarly affected (33-35, 57). (vii) All the reactions produce new 3'57. B.

(1976).

J. White, S. J. Hochhauser, N. M.Cintron, and B. Weiss,J.

Bacteriol. 126, 1082

222

BERNARD WEISS

hydroxyl termini and result from cleavages 5' to the recognized moiety whether it be a terminal nucleotide, a terminal phosphate, or a base-free sugar (35). The model, therefore, correctly predicted that the endonucleolytic cleavage would be 5' to the AP site. An alternative model that had been first considered stated that an enzyme with two catalytic sites had evolved to mediate sequential steps in an excision-repair pathway. According to that model, however, endonucleolytic cleavage should have been 3' to the AP site so that subsequent 3' + 5' exonuclease activity could remove the lesion (i.e., the base-free sugar). In its most general form, the common-site model implies the cleavage of unpaired nucleotides that lie adjacent to paired regions. The following molecules, however, are not cleaved endonucleolytically although they contain such regions: (i) circular single-stranded 4x174 DNA, which has both relatively stable and transient bihelical regions (58); (ii) heteroduplexes of mutant A phage DNAs that contain unpaired regions ( 5 0 ) ; (iii) covalently closed circles of the alternating copolymer poly(dA-dT) (59); and (iv) duplexes with long, single-stranded 3' ends (16). From the terminal unwinding hypothesis, we might also expect that di- and trinucleotides would be released, in addition to mononucleotides, during exonucleolytic digestion, but such products have not been found (26). g. Biological Role. Exonuclease I11 is an abundant and very active enzyme in crude extracts of E. coli; there are about 3500 molecules per cell. Its structural gene, xthA, is located at 38 min on the chromosomal map (17,57) and is expressed constitutively. Exonuclease I11 production is not affected by exposure of the cells to mitomycin C (an alkylating agent) or by mutations in the gene for dUTPase (58), both of which result in the production of AP sites. From the study of xth mutants, we know that under the standard assay conditions exonuclease I11 accounts for about 85-90'36 of the AP endonuclease activity in crude extracts ofE. coli (34,35, 60). Within the cell, its exonuclease activity may be second only to that of exonuclease V, an ATP-requiring enzyme, although the relative activity of the latter under physiological conditions is uncertain (see Linn, Chapter 13, this volume). Most of the residual AP endonuclease activity in xth mutants appears to be due to endonuclease IV (60, 61). In crude extracts of Hemophilus influenzae, exonuclease I11 is also the major AP endonuclease and ATP-independent exonuclease, and this AP endonuclease activity far exceeds the restriction endonuclease activity (46). 58. A. F. Taylor and B. Weiss, unpublished data. 59. P. Modrich and I. R. Lehman, JBC 245, 3626 (1970). 60. S. Ljungquist, T. Lindahl, and P. Howard-Flanders,J . Bacteriol. 126, 646 (1976). 61. S. Ijungquist, JBC 252, 2808 (1977).


223

Tight xth mutants have the following abnormalities: (i) a slightly increased sensitivity to alkylating agents (34, 41, 62); (ii) an increased level of genetic recombination, or hyper-Rec phenotype (63);and (iii) an inability to tolerate a dut (dUTPase) mutation (35). These defects can be explained by their loss of AP endonuclease activity. The mutants are normal with respect to their growth rate, their sensitivity to ultraviolet irradiation, and their mutation frequency (331. The hypothesis that the exonuclease activity of exonuclease I11 was important in DNA repair was attractive because its properties were ideal for this purpose. The enzyme degrades only one strand in a given region, leaving an intact template opposite a 3‘-hydroxyl primer terminus. Because of its ability to remove mismatched nucleotides, it was implicated in the “proofreading” of newly synthesized DNA (16). There is no evidence for this assumption; xrh mutants do not have high mutation rates. The enzyme is clearly nonessential; a deletion mutant is viable (57). It is nevertheless possible that the exonuclease activity of the enzyme may have an important function for which other exonucleases can substitute; however, xth mutations have at least been transduced into a polAl mutant (deficient in the 3’ + 5‘ exonuclease activity of DNA polymerase I) and into an sbcB-recBC mutant (deficient in both exonucleases I and V) without noticeable effect (57). The effects of enzyme overproduction are also unremarkable. A strain bearing a multicopy xth plasmid grew normally despite a constitutive 30-fold overproduction of the enzyme (36). Moreover, a thermoinducible plasmid yielding a 120-fold overproduction could be isolated mostly intact after induction (36), suggesting that even at these levels of intracellular exonuclease activity, there is no significant degradation of the plasmid DNA. These findings suggest that there may be few, if any, free 3‘ ends in intracellular DNA; such termini may normally exist only within replication or topoisomerase complexes. The DNA-3’-phosphatase function of the enzyme was thought to be of possible importance (26) because 3’-phosphoryl-terminatedDNA is not a substrate for the DNA polymerases of E. coli, and because exonuclease I11 is responsible for >99% of this phosphatase activity in cells under normal growth conditions (33, 64). In E. coli, however, no enzymes are currently known to produce 3’-phosphoryl end groups in DNA. Similarly, 62. The apparent marked sensitivity of one mutant (Ref. 3 4 ) was due in part to a second uncharacterized mutation that was not cotransducible withxthA (B. Weiss, unpublished). A deletion mutant was not very sensitive to methyl methane sulfonate (Ref. 41). 63. J. Zieg, V. F. Maples, and S. R. Kushner, J . Barterid. 134, 958 (1978). 64. Alkaline phosphatase, which can also catalyze this reaction, is normally repressed except during growth in low-phosphate media.

224

BERNARD WElSS

it is hard to think of a use for the RNase H function. It would not etlectively degrade an RNA Drimer in DNA synthesis because it attacks the wrong end of the strand, and it should not selectively remove misincorporated ribonucleotides because its DNase activity is greater. Exonuclease I11 does, however, appear to be important in the repair of DNA that contains uracil residues or AP sites (see Friedberg, et nl., Chapter 13, this volume). AP sites can occur not only from spontaneous depurination but also from the action of glycosylases that specifically hydrolyze unusual bases, such as uracil, from DNA. AP sites are repaired through the sequential action of AP endonucleases, DNA polymerase I (via its 5‘ -+ 3’ exonuclease and its polymerase activities), and DNA ligase. Thus, a nonlethal dur (dUTPase) mutation, which enhances the misincorporation of uracil into DNA, becomes conditionally lethal in the presence of nonlethal mutations affecting exonuclease 111(35),DNA polymerase I (65),or DNA ligase ( 6 5 ) . The dut-xth double mutant is inviable, probably because of the accumulation of lethal AP sites. An ung (uracil DNA glycosylase) mutation restores its viability, presumably by blocking the removal of uracil and hence the formation of AP sites (35).While these results indicate that exonuclease 111 is essential for the efficient repair of AP sites in DNA, and while they imply that it is the AP endonuclease activity of the enzyme that is essential, they do not rule out the possibility that it is the exonuclease activity that is important (66). The basis for the hyper-Rec phenotype ofxth mutants is unknown. Enhanced recombination is not seen in assays that measure only the frequency of incorporation of foreign DNA during transduction, conjugation, or transformation. It has been seen only in an assay for recombination between two distant but homologous chromosomal regions (63). The hyper-Rec phenotype might be due to decreased degradation of recombination intermediates (as in the case of exonuclease I mutations) or to the production of recombinogenic lesions (perhaps AP sites) (67). 4. Research Applications

DNA that has been partially digested with exonuclease I11 is an ideal primer for most DNA polymerases and has been widely used for this 65. B-K. Tye and I. R. Lehman, J M B 117, 293 (1977). 66. One theory, for example, is that the exonuclease exerts an “anti-ligase” function; it would create a gap to prevent premature sealing of the incision site by DNA ligase, thereby giving DNA polymerase I more time to work. There is, however, no evidence that DNA ligase would catalyze a reaction involving a base-free sugar. 67. By analogy, dut mutants, which generate AP sites transiently, are also hyper-Rec. See S. J. Hochhauser and B. Weiss, J . Bacteriol. 134, 157 119781.


225

purpose. The enzyme can be used to create gaps in DNA at specific sites, such as restriction enzyme cleavage sites. These gaps can be repaired with triphosphates that contain base analogs or they can be treated with a mutagen, such as bisulfite (681, that is specific for single-stranded regions. The result is localized mutagenesis of a specific region of a genome. If the gaps are repaired by DNA polymerase and labeled nucleoside triphosphates, the region can be subjected to sequence analysis. Wu and his colleagues described methods for controlled synchronous exonucleolytic degradation that can be used to determine the nucleotide sequence for a limited distance at the 3' ends of DNA (8, 53). Exonuclease I11 has also been used in conjunction with a single-strand-specific endonuclease (S 1 nuclease) to generate small deletions at a restriction site in a cloned gene (69).

The AP endonuclease activity of the enzyme can be used to detect AP sites in DNA molecules and to distinguish them from sites of ribonucleotide misincorporation, which are also alkali-labile. In conjunction with other DNases, exonuclease I11 can also be used to identify, and to isolate, specific protein binding sites in a DNA molecule; the bound protein partially protects such sites from nucleolytic degradation (54).

c.

EXONUCLEASES IVA AND IVB

1. Introduction

Exonucleases IVA and IVB catalyze the degradation of oligonucleotides to nucleoside 5'-monophosphates. Our knowledge of them is limited to the information in one publication, by Jorgensen and Koerner (70), which was based on work with relatively impure fractions.

2 . Purijication and Properties Extracts of E. coli B were subjected to ammonium sulfate fractionation followed by DEAE-cellulose chromatography, and two separately eluting peaks of DNase activity were found. Although their catalytic properties were indistinguishable, they were still eluted at different salt concentrations when rechromatographed on DEAE-cellulose. By this method, 68. D. Shortle and D. Nathans, PNAS 75, 2170 (1978). 69. S . Sakonju, D. F. Bogenhagen, and D. D. Brown, Cell 19, 13 (1980). 70. S . E. Jorgensen and J. F. Koerner, JBC 241, 3090 (1966).

226

BERNARD WEISS

exonuclease IVA was purified 120-fold and exonuclease IVB was purified about 30-fold. 3. Catalytic Properties The exonucleases have a broad pH optimum, between 8.0 and 9.5. They require Mg2+,and are distinguished from exonuclease I by a greater heat stability and an ability to degrade dinucleotides. They are assumed to be exonucleases because early in the course of digestion, they release mononucleotides exclusively. The preparation of exonuclease IVB had an RNase activity (50% of DNase) that may have been due to contamination from a closely eluting RNase peak. The DNase activities were assayed by measuring the release of acidsoluble, ultraviolet-absorbing material from an oligonucleotide mixture. The substrate was prepared by digesting salmon sperm DNA with pancreatic DNase (an endonuclease) to the point of 20-25% acid solubility. Because this susbstrate preparation was degraded at about 20 times the rate of either native or heat-denatured salmon sperm DNA, it was concluded that exonucleases IVA and IVB were specific for oligonucleotides. This conclusion is unwarranted because the substrates were compared at widely different concentrations. The concentration of nucleotide phosphorus was the same in each reaction, and therefore, the short-chain oligonucleotides had, of course, a much higher concentration of 3' termini than did the native or denatured DNAs (71 1. Because of uncertainty about enzyme purity and about substrate specificity, therefore, we do not know if exonucleases IVA and IVB are unique, if they are other known enzymes, or if they are enzyme mixtures. D. EXONUCLEASE VII 1. Introduction Exonuclease VII (72) attacks single-stranded DNA processively from both the 3' and 5' ends, releasing large oligonucleotides that are then further degraded to smaller ones (Fig. 4). The enzyme has been thoroughly reviewed by Chase and Vales (73) and therefore is described briefly here. 71. It was perhaps for this reason that Jorgensen and Koerner noted, in a control experiment, that their preparation of authentic exonuclease I was more active on the oligonucleotide preparation than on denatured DNA. 72. J. W. Chase and C. C. Richardson, JBC 249, 4545, 4553 (1974).

227


FIG.4. The exonuclease VII reaction. Processive attack at both ends of the singlestranded DNA releases large oligonucleotides that are further degraded by enzyme molecules released from other chains.

2 . Purification and Properties The purification procedure (72), which has recently been improved (73), uses as an enzyme source a mutant that is deficient in both exonuclease I and DNA polymerase I. Streptomycin sulfate is used to coprecipitate the enzyme with cellular DNA, following which the enzyme is further purified by acetone fractionation and column chromatography. Tne native enzyme has a molecular weight of 88,000 as determined from gel filtration and velocity sedimentation data. An unusually high frictional coefficient of 3.07 indicates an asymmetric shape. If the molecule is rodshaped, it will cover a 100-nucleotide length of DNA, which is the approximate size of its initial degradation products. 3 . The Reactions Catalyzed

Exonuclease VII is highly specific for either single-stranded DNA or for the single-stranded ends of duplexes. Like exonuclease I (see Section II,A) it can digest poly(dA-dT), but at a slower rate than single-stranded DNA. The degradation, which is both 3’ 5’ and 5‘ + 3‘, is not affected by the presence or absence of phosphornonoesters at either terminus. Degradation is processive as evidenced by the lack of a significant fraction of partially digested DNA molecules in a partial digest. The initial products appear to be oligonucleotides of chain length 2100 nucleotides. These are further digested so that the limit products are 2 to 25 or more ---$

73. J. W. Chase and L. D. Vales, in “Gene Amplification and Analysis, Vol. 11: Analysis of Nucleic Acid Structure by Enzymatic Methods” (J. G . Chirikjian and T. S. Papas, eds.), in press. Elsevier-North Holland, Amsterdam, 1981.

228

BERNARD WEISS

nucleotides in length. No mononucleotides are produced. Of special interest is the enzyme's ability to excise thymine dimers from a duplex substrate that contains nicks near the dimers, introduced for example by an endonuclease of Micrococcus luteus. This reaction may be analogous to its ability to remove a small displaced or unpaired region from either end of a synthetic duplex, such as d(C)s-d(T)m * poly(dA) or d(Ts-d-(C)a. poMdA). The standard assay for the enzyme measures the conversion of sheared radiolabeled, denatured DNA to an acid-soluble form in the presence of EDTA. Exonuclease VII is unique among the exonucleases ofE. coli in that it is not inhibited by EDTA. The enzyme has a sharp pH optimum at 7.9 and has a 5- to 10-fold higher activity in phosphate as compared to Tris buffer.

4. Biological Role The structural gene for exonuclease VII, designated xseA, has been located with the aid of mutants that produce a thermolabile enzyme. It is at 53 min on the genetic map ofE. coli (17, 7 4 ) . Deletion mutants are fully viable, indicating that the gene is nonessential (75).xseA mutants have a slightly increased sensitivity to ultraviolet irradiation, are sensitive to nalidixic acid, and have a hyper-Rec phenotype (increased recombination frequency) (74). The defects of the exonuclease VII mutants may be difficult to observe because of redundant roles played by other enzymes. Therefore, an xseA mutation has been combined with apolAex mutation (defective in the 5' 3'-exonuclease of DNA polymerase I) and with recBC (exonuclease V) mutations. An xseA-polAex double mutant is more hyper-Rec and more temperature-sensitive than either alone, and although the .rseA mutation does not affect the rate of excision of thymine dimers from a wild-type strain, it markedly reduces the rate of such excision in a recBC-polAex multiple mutant (76-78). --f

5. Reseurch Applications (79) Exonuclease VII has been used in conjunction with other singlestrand-specific enzymes to determine the extent and location of single74. J. W. Chase and C. C. Richardson, J . Bacteriol. 129, 934 (1977). 75. L. D. Vales, J. W. Chase, and J. B. Murphy,J. Bocferiol. 139, 320 (1979). 76. J. W. Chase and W. Masker, J . Bncreriol. 130, 667 (1977). 77. W. E. Masker and J. W. Chase, in, "DNA Repair Mechanisms" (P. C. Hanawalt, E. C. Friedberg, and C. F. Fox, eds.), p, 261. Academic Press, N e w York, 1978. 78. J. W. Chase, W. E. Masker, and J. B. Murphy, J . Bacteriol. 137, 234 (1979). 79. For a more detailed discussion of the techniques mentioned in this section, see Ref. 73.

12. E. roli EXONUCLEASES

229

stranded regions in DNA molecules. Endonuclease S1 of Aspergillus oryzae digests away all single-stranded regions, whether they be terminally or internally located. Exonuclease I removes only a terminal 3' tail. Exonuclease VII removes both 3' and 5' tails. These enzymatic reagents have been used in various combinations to determine the structure of exonuclease V digestion intermediates that have long, single-stranded tails (80), to measure the length of inverted repeats and intervening sequences in a DNA after intrastrand annealing ( 8 / ) , to analyse DNA-mRNA hybrids for regions in the DNA that are not represented in the mRNA due to splicing (82), and to isolate material between inverted repeats, such as DNA cloned by the poly(dA) poly(dT) tailing technique (83). E.

EXONUCLEASE VIII

1. Ziitroduction

Exonuclease VIII (84, 85') is relatively specific for double-stranded DNA. Its reaction products and mode of attack have not been described. The enzyme is specified by the recE gene on an integrated cryptic prophage (the Rac prophage) carried by some lines ofE. coli K12. Enzyme production is normally repressed in wild-type cells; it is expressed at higher levels in sbcA mutants. The discovery of exonuclease VIII and of its evolutionary origin arose from three separate lines of investigation that yielded the following findings. (i) A recBC (exonuclease V) mutant is recombination-deficient, but if it is also rac -, it becomes recombination-proficient during mating at the time it receives a functional rac (recombination activation) locus from an Hfr donor cell (86). This phenomenon resembles zygotic induction, i.e., the derepression of prophage genes upon conjugational transfer into a repressor-free (nonlysogenic) recipient. (ii) recBC mutants can also become recombination proficient through ansbcA mutation (87) located in or near the rac (integrated Rac prophage) locus (88), and such mutants possess a new DNase activity that, unlike exonuclease V, does not require 80. V. MacKay and S. Linn, JBC 249, 4286 (1974). 81. P. Nisen, R. Medford, J. Mansour, M. Purucker, A. Skalka, and L. Shapiro, PNAS 76, 6240 (1979).

82. 83. 84. 85. 86. 87. 88.

A. J. Berk and P. A. Sharp, PNAS 75, 1274 (1978). S. P. Goff, and P. Berg, PNAS 75, 1763 (1978). S. R. Kushner, H . Nagaishi, and A. J. Clark, PNAS 71, 3593 (1974). J. R . Gillen, A. E. Karu, H. Nagaishi, and A. J. Clark,JMB 133, 27 (1977). K. B . Low, Mol. Gen. Genet. 122, 119 (1973). A. Templin, S. R. Kushner, and A. J. Clark, Genetics 72, 205 (1972). R . G. Lloyd and S. D. Barbour, Mol. Gen. Genet. 134, 249 (1974).

230

BERNARD WEISS

ATP (89).(iii) ex0 (previously called red) mutants of bacteriophage A lack an exonuclease required for phage-mediated recombination. After growth on rac+ cells, however, derivatives can be isolated that are recombination-proficient and have regained a DNase activity. These ArecE (or Arev, or A-reverse) phages have acquired by recombination a new region of DNA, not homologous with'h, but homologous with therac region of the chromosome (90-92). Exonuclease VIII is believed, therefore, to be a relative of the exonuclease of bacteriophage A, specified by A-like cryptic prophage. 2. Purijication and Properties The enzyme has been purified by conventional methods from an sbcA mutant of E. coli K12 (84) and from cells infected with ArecE (85). The phage-infected cells yielded about 12 times more enzyme than the sbcA mutants. The enzymes prepared from both sources were physically similar and were both precipitated by antiserum to the ArecE exonuclease. The enzyme appears to consist of a single polypeptide with a molecular weight of 1.4 x lo5, as determined by sedimentation analysis in a glycerol gradient and by sodium dodecyl sulfate gel electrophoresis (85, 93). Surprisingly, this value is quite different from that for A exonuclease (MW = S2,000),and antibody to the latter failed to inhibit exonuclease VIII significantly. Therefore, there must be considerable evolutionary divergence between these enzymes. It has been suggested that exonuclease VIII might be a gene fusion product, thus accounting for its higher molecular weight (85).

3 . Catalytic Properties The enzyme is assayed by the degradation of radiolabeled bacterial DNA to an acid-soluble form (84). The purified enzyme has a marked specificity for double-stranded versus single-stranded DNA (40-fold) (84, 85). Unlike exonuclease V (the recBC enzyme), which also functions in recombination, exonuclease VIII is ATP-independent and cannot degrade the glycosylated DNA of bacteriophage T4 (89). It requires Mg2' and has a pH optimum between 8.0 and 9.0. Questions of identity and origin have dominated investigations of this enzyme, leaving even the most basic 89. S. D. Barbour, H . Nagaishi, A . Templin, and A. J. Clark, PNAS 67, 128 (1970). 90. M. M. Gottesman, M. E. Gottesman, S. Gottesman, and M. Gellert, J M B 88, 471 (1974).

91. K. Kaiser and N. E Murray, Mol. Gen. Genet. 175, 159 (1979). 92. R. Evap, N. R. Seeley, and P. L. Kuempel, Mol. Gen. Gener. 175, 245 (1979). 93. A value of 1.22 x lo5 was separately obtained by the latter method [Ref. (SS)].


23 1

questions of mechanism unexplored. Thus, we do not know the nature of the reaction products or direction of attack. The only evidence that it is an exonuclease stems from its inability to degrade circular single-stranded DNA molecules (84); circular duplexes were not tried despite the enzyme's specificity for double-stranded DNA. It is assumed that the enzyme will prove to be similar to A exonuclease (W), i.e., a 5' -+ 3' processive exonuclease that releases 5'-mononucleotides and is unable to initiate degradation at a single-strand break; because of the physical differences between these enzymes, however, this assumption may not be warranted. 4. Biological Role

As discussed above, the enzyme can substitute for A exonuclease in A-mediated general recombination. It is also part of the recE pathway, an alternative pathway for homologous recombination in uninfected E. coli (19). TherecE pathway is manifest in mutants for the major recombination pathway (recBC mutants) that regain recombination proficiency concomitant with enhanced expression of the structural gene for exonuclease VIII (recE). This apparent derepression can occur through mutation in a control locus (sbcA) or through zygotic induction during conjugation. Neither loss of the prophage nor mutation in the sbcA locus significantly affects the frequency of genetic recombination in recB +C+cells. The theoretical role of exonucleases in recombination has been reviewed by Radding (95). By restriction endonuclease analysis, Kaiser and Murray (91) have studied the structure of several XrecE phages and of the rac region of the chromosome. Their results indicate that the Rac prophage undergoes excision from the bacterial chromosome and recombines with phage A to form ArecE. Two regions of the Rac prophage, amounting to no more than a few percent of its length, share homology with X DNA. The two phages are, therefore, distantly related.

94. J . W. Little, I. R . Lehman, and A . D. Kaiser, JBC 242, 672 (1967); J. W. Little, h i d . , p. 679; D. M . Carter and C. M. Radding, JBC 246, 2502 (1971); Y. Masamune, R . A. Fleischman, and C . C. Richardson, ibid.. p. 2680. 95. C. M. Radding, Annu. Rev. Biochem. 47, 847 (1978).


recBC-like Enzymes: Exonuclease V Deoxyribonucleases KAREN M. TELANDER MUSKAVITCH STUART LINN

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . 11. Biological Role . . . . . . . . . . . . . . . . . . . . . . . . .

111.

IV.

V. VI.

A. Recombination and Repair . . . . . . . . . . . . . . . . . B. Restriction . . . . . . . . . . . . . . . . . . . . . . . . . C. Viability . . . . . . . . . . . . . . . . . . . . . . . . . . D. Replication . . . . . . . . . . . . . . . . . . . . . . . . . Purification and Biophysical Properties . . . . . . . . . . . . . The Reactions Catalyzed . . . . . . . . . . . . . . . . . . . . A. ATP-Dependent Double-Stranded DNA Exonuclease . . . . . B. Single-Stranded DNA Exonuclease . . . . . . . . . . . . . C. Single-Stranded DNA Endonuclease . . . . . . . . . . . . . D. DNA-Dependent ATPase . . . . . . . . . . . . . . . . . . E. ATP-Dependent Double-Stranded DNA Unwinding . . . . . . Models for the Mechanism of Action of the ATP-Dependent DoubleStranded DNA Exonuclease . . . . . . . . . . . . . . . . . . Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . .

.

.

. .

. . . .

234 235 235 237 237 237 238 240 240 244 245 245 246 247 250

233 THE ENZYMES,Vol. XIV Copyright 0 1981 by Academic Press. Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

234 1.

KAREN. TELANDER MUSKAVITCH AND STUART LINN

Introduction

The first report of an exonuclease V (exoV) activity, an ATP-dependent double-stranded DNase, was made by Tsuda and Strauss in 1964 ( I ) .They utilized the bacterium Micrococcus lysodiekticus, which is now known as M. luteus. Since then similar activity has been observed in nine additional bacterial species: Bacillus laterosporus (2 1, Mycobacterium smegamatis ( 3 ) , Escherichia coli (4-7), Diplococcus pneumoniae ( 8 ) , Haemophilus influenzae (9), Bacillus subtilis (10, I / ) , Bacillus cereus (12), Pseudomonas aeruginosa ( 13), and Alcaligenes faecalis (14). This widespread occurrence suggests that exoV enzymes are ubiquitous in bacteria. In eukaryotes, the only purified ATP-dependent double-stranded DNase was discovered by Gafurov et al. (15, 16) in sea urchin embryos (Strongylocentrotus intermedius). However, work on yeast mitochondria1 mutagenesis suggests the presence of an ATP-dependent DNase (17), and Neurospora crassa has an enzyme with a combination of nuclease activities resembling bacterial exoV (18). Thus, exoV-like enzymes may be present in all living organisms, prokaryotic or eukaryotic. The bacterial enzymes constitute a distinct class with a number of characteristics in common: (1) An ATP-dependent double-stranded DNA exonuclease that degrades DNA to short oligonucleotides, (2) a DNAdependent ATPase that hydrolyzes ATP to ADP and Pi, (3) a requirement for a divalent cation and an alkaline pH optimum, (4) single-stranded DNA exonuclease activity, and ( 5 ) a large multisubunit structure. Further Y. Tsuda and B. S. Strauss, Biochemistry 3, 1678 (1964). M. Anai, Seikagaku 39, 167 (1967). F. Winder and M. P. Coughlan, EBA 134, 215 (1967). G. Buttin and M. Wright, CSHSQE 33, 259 (I%@. M. Wright and G. Buttin, Bull. SOC. Chim. Bioi. 51, 1373 (1%9). M. Oishi, PNAS 64, 1292 (1969). S. Barbour and A. J. Clark, PNAS 65, 955 (1970). G. Vovis and G. Buttin, BEA 224, 29 (1970). E. A. Friedman and H. 0. Smith, JEC 247, 2846 (1972). 10. A. V. Chestukhin, M. F. Shemyakin, N. A. Kalinina, and A. A. Prozorov, FEBS (Fed. Eur. Biochem. Soc.) Lett. 24, 121 (1972). 11. S. Ohi and N. Sueoka, JEC 248, 7336 (1973). 12. G. Banfalvi, F. Antoni, and S. Csuzi, Stud. Eiophys. Berlin 50, 89 (1975). 13. R. V. Miller and A. J. Clark, J. Eacteriol. 127, 794 (1976). 14. J. D. C . Rosamond and M. R. Lunt, EJ 163, 485 (1977). 15. N. N. Gafurov and V. A. Rasskazov, Dokl. Akad. Nauk SSSR 219, 1495 (1974). 16. Y. M. Gafurov, L. L. Terentev, and V. A. Rasskazov, Biochemistry ( U S S R )44, 996 (1979). 17. R. N. Bastos and H. R. Mahler, JEC 249, 6617 (1974). 18. T. Y. K. Chow and M. J. Fraser, Can. J . Biochern. 57, 889 (1979). 1. 2. 3. 4. 5. 6. 7. 8. 9.

13. recBC-LIKE ENZYMES

235

work on some exoV enzymes has revealed associated single-stranded DNA endonuclease and double-stranded DNA unwinding activities. These similarities led Wilcox and Smith to propose a common nomenclature for these enzymes (191, in which the bacterial source abbreviations are those used for restriction endonucleases, and the enzyme class is noted as exoV. Thus Eco exoV, Hind exoV, etc., are used here. The exoV enzymes belong to a larger group of ATP-dependent DNases and DNA-dependent ATPases, which also includes such enzymes as type-I restriction endonucleases, helicases, and gyrases. This larger group has been reviewed by Whitehead (20). The approach of this review is a comparative one. Emphasis is placed on enzymes about which we have the most information: Eco exoV, Hind exoV, Bsu exoV, and Mlu exoV.

It.

Biological Role

A.

RECOMBINATION AND REPAIR

The observation of decreased or undetectable exoV activity in recombination and repair mutants of four bacteria has sparked a great deal of interest in these enzymes. Mutants in the recB or recC genes of E. coli form fewer recombinants in Hfr crosses than do wild-type strains (21-241, are more sensitive to X-rays, ultraviolet light, and mitomycin C (21, 2-3, 24), and lack exoV activity (5-7). Similarly, an H. irtfuenzae mutant selected for sensitivity to methylmethane sulfonate is deficient in recombinant formation following transformation and lacks Hind exoV activity (251, and a D . pneumoniae mutant isolated by sensitivity to y-rays produces transformants at 15% of the wild-type frequency, is UV sensitive, and lacks exoV activity (26). ExoV activity is also missing in therecE5 mutant of B. subtilis (27), but is only decreased five- to tenfold in another recstrain that is a poor recipient for transformation and transduction (10). 19. K. W. Wilcox and H . 0. Smith,JBC 251, 6122 (1976). 20. E. P. Whitehead, i n . "Macrmolecules in the Functioning Cell" (F. Salvatore, G. Marino, and P. Volpe, eds.), p. 159. Plenum, New York, 1979. 21. P. Howard-Flanders and R . P. Boyce, Radicrt. Res., Sicppl. 6 , 156 (1966). 22. B. Low, PNAS 60, 160 (1968). 23. N . S. Willetts and D. W. Mount, J . Bacreriol. 100, 923 (1969). 24. A. J. Clark, A m i i . Rev. Genet. 7 , 67 (1973). 25. M . L. Greth and M. R. Chevallier, BBRC 54, 1 (1973). 26. G. Vovis and G . Buttin, BBA 224, 42 (1970). 27. J. Doly, E. Sasarman, and C. Anagnostopoulos, Murcit. Res. 22, 15 (1974).

236

KAREN TELANDER MUSKAVITCH AND STUART LlNN

Recombination and repair lesions in bacteria that lack exoV are not absolute, so the deficiency must be carefully quantitated with well-defined strains. For E. coli recB and recC mutants, the frequency of recombinants following Hfr crosses is decreased no more than 100-fold below that of the wild type (23), and in A phage-prophage crosses it is decreased no more than twofold (28).Postreplication repair is almost normal in recB mutants (29), and recB and/or recC mutants are only slightly more sensitive than wild-type E . coli to ultraviolet light (23). Finally, work by Stahl and Stahl on A phage chi mutants indicates that these genes are essential to A phage-phage recombination, at least under special conditions (30). A thorough investigation of the H . injuenzae mutants has also been made. The results demonstrate the importance of relocating an allele from its original highly mutated background and the difficulty of quantitating transformation frequencies. When the mutant allele, add-18, originally isolated by Greth and Chevallier (2.51, is moved to an unmutagenized strain, the ability of the strain to form transformants, quantitated as transformants per competent cell, is decreased only twofold relative to the wild type. However, if quantitated as transformants per colony-forming unit, there is a sevenfold decrease, and if quantitated as transformants per microgram of DNA bound, there is a 17-fold reduction (31). Other mutants 1ackingHind exoV activity give similar results, and since the frequency of transformants per competent cell is almost the same as for wild-type cells, one might conclude that exoV is not involved in recombination. However, a small effect appears reproducible. These mutants are also slightly more sensitive to y-rays, UV light, methylmethane sulfonate, and mitomycin C, and these phenotypes have been used to group the mutant alleles into three complementation groups ( 3 2 ) . While the three complementation groups for loss of Hind exoV activity only suggest that this enzyme may have three subunits (32), in E. coli and B . subtilis direct identification of the exoV enzyme as the product of at least one rec gene has been possible. Tomizawa and Ogawa (33) have shown that a temperature-sensitive recB mutation produces a temperature-sensitive Eco exoV enzyme; however, a temperaturesensitive recC mutation has no effect on the enzyme. Further work by 28. P. Lin, E. Bardwell, and P. Howard-Flanders,PNAS 74, 291 (1977). 29. K. C. Smith and D. H. C. Meun, J M B 51, 459 (1970). 30. F. W. Stahl and M. M . Stahl,Generics 86, 715 (1977). 3 1. K . W. Wilcox and H . 0. Smith, J . Bacreriol. 122, 443 ( 197s). 32. J . Kooistra, G . D. Small, J . K . Setlow, and R. Shapanaka, J . Bacterial. 126, 31 (1976). 33. J . 1. Tomizawa and H. Ogawa, Nfiritre New Eiol. 239, 14 (1972).

13. wcBC-LIKE ENZYMES

237

Kushner revealed that of the DNase activities only the double-stranded DNA exonuclease activity of Eco exoV is made temperature sensitive by the temperature-sensitive recB allele (34). In B. subtifis, Doly and Anagnostopoulos (35) discovered that although a recES mutant has no exoV activity, the inactive enzyme can be purified and has an altered subunit. It should be noted that the five subunits of Bsu exoV identified by these workers may be the products of proteolysis (36).

B. RESTRICTION Studies of mutants in the E. coli recB and recC genes indicate that exoV is responsible for completing the degradation of foreign DNA after it has been acted upon by a host-restriction endonuclease (37).

C. VIABILITY It has been observed, both in E. coli and H . injuenzae, that mutants lacking exoV activity have a slower growth rate and reduced viability when compared to the wild type (32, 38). Typically, less than one-half of the cells present in a liquid culture are able to form colonies on agar. Studies of E. coli mutants indicate that reduced viability occurs because some of the cells cannot divide and others can divide only a limited number of times. The reason why mutants sometimes fail to divide is unknown (39). Attempts to clone the E. coli recB and recC genes have been unsuccessful, suggesting that the level of exoV is an important factor in determining the viability of the bacterium, perhaps in an area different from repair and recombination.

D. REPLICATION Involvement of Eco exoV in replication could explain the decreased viability of mutants. Although data on this point are only suggestive, studies of spheroplast lysates and toluene-treated E. coli indicate that 34. 35. 36. 37. 38. 39.

S. R . Kushner, J . Bacteriol. 120, 1219 (1974). J. Doly and C. Anagnostopoulos, EJB 71, 309 (1976). M. F. Shemyakin, A . A. Grepachevsky, and A. V. Chestukhin, EJB 98, 417 (1979). V. F. Simmon and S. Lederberg, 1.Bacteriol. 112, 161 (1972). F. Capaldo-Kimball and S. D. Barbour, 1.Bacteriol. 106, 204 (1971). F. N . Capaldo and S. D. Barbour, J . Bacleriol. 115, 928 (1973).

238

KAREN TELANDER MUSKAVITCH A N D STUART LINN

exoV is essential to ATP-dependent DNA synthesis observed under these conditions (40,41). In addition, Hendler and co-workers (42) have isolated a complex of E. coli exoV and DNA polymerase I that carries out ATPdependent DNA synthesis, and tentatively report the isolation of similar complexes with DNA polymerases I1 and 111. However, since recB and recC mutants do replicate, the Eco exoV activity involved in replication must still exist in these mutants, or is replaceable. 111.

Purification and Biophysical Properties

Some properties of various exoV enzymes, including the sea urchin (S. intermedius) enzyme, and their relative purity are summarized in Table I. Homogeneous enzymes have been obtained from E. coli, H. influenzae, and B. subtilis. The subunit structure of the Eco and Bsu exoV enzymes varies with the purification technique. In the less purified Eco exoV preparation of Oishi and co-workers (see 44,45), an additional smaller subunit is detected when compared with the preparation of Goldmark and Linn (43). The larger subunit fraction identified by Lieberman and Oishi may contain both of the large subunits reported by Goldmark and Linn, because the only characterization done by the former group was glycerol gradient sedimentation. The smaller subunit may be purified away from an enzyme core in the Goldmark and Linn preparation, or may be fortuitously present in the less purified Nobrega et a/. preparation (44). Two observations suggest that the small subunit may be a true subunit ofEco exoV: (1) It is required for recovery of activity after high salt treatment and separation of the two subunit fractions (45), and (2) Hind exoV has three subunits, two large and one small (46). The difference between the two B. subtilis preparations would seem to be due to proteolysis. Shemyakin et al. (36) add phenylmethylsulfonyl fluoride to the lysis buffer and report purification of a stable enzyme preparation with about an 8% yield. In contrast, Doly and Anagnostopoulos (35)report that the Bsu exoV is unstable at later stages of their purification and obtain a 1.7% yield. Ohi and Sueoka (11) also report 40. G. W. Brazill, R. Hall, and J . D. Gross, Nature (London)New Biol. 233, 281 (1971). 41. D. J. Harper, P. L. Chen, and P. L. Carl, BBA 474, 363 (1977). 42. R. W. Hendfer, M. Pereira, and R. Scharff, PNAS 72, 2099 (19751. 43. P. J. Goldmark, and S. Linn, JBC 247, 1849 (1972). 44. F. G . Nobrega, F. H. Rola, M. Pasetto-Nobrega, and M. Oishi, PNAS 69, 15 (1972). 45. R . P. Lieberman and M. Oishi, PNAS 71, 4816 (1974). 46. K. W. Wilcox, M. Orlosky, E. A. Friedman, and H . 0. Smith, FP 34, 515 (1975).

239

13. rerBC-LIKE ENZYMES TABLE I

PURIFICATION AND PROPERTIES OF

Source of enzyme

E. coli (43)

Purification (-fold)

Specific activity" (U/rng)

17,000

57,000

s

~

EXOV ENZYMES ~Molecular , ~

(S)

weight

12

270,000

E. coli (44, 4 5 )

3,500

21,600

12.4

350,000

H. influenme (9.46)

2,000

28,000

12

290,000

B. subtifis (35)

350

237

B . subtilis (.16)

5 ,000

$4,000

2,300 I50 100 743 4,200 97,000 820 200

67,000 54,500 8,700 28,000 2,450 91,OOO 1,900 4,700

M . luteus (47) B. laterosporus (48) M. smegmatis (49) D. pneumotiiae (8) B. cereus (50) P. aeruginosa (13) A. fueculis (14) S . inrermedius ( 1 6 )

-

270,000

300,000

-

300,000

-

450,000

Subunit molecular weight

140,OOO 128,000 170,000 60,000 115,000 107,000 68,000 81,000 70,000 62,000 52,500 42,500 155,OOO 140,000

-

-

" All values have been adjusted so that 1 unit is the amount of enzyme needed to render acid-soluble 1 nmol of duplex DNA-nucleotides in 30 rnin.

instability and a poor yield during purification in the absence of a protease inhibitory. Several generalizations can be made from Table I about bacterial exoV enzymes. They are around 300,000 daltons and are composed of two nonidentical, large subunits. In some cases a third smaller subunit may be part of the enzyme. The high level of purification required to obtain homogeneous enzyme indicates that relatively few enzyme molecules are present in the cell. 47. M . Anai, T. Hirahashi, and Y. Takagi, JBC 245, 767 (1970). 48. M. Anai, M. Mihara, M. Yamanaka, T. Shibata, and Y. Takagi,J. Biochem. (Tokyo) 78, 105 (1975). 49. F. G. Winder and M. F. Lavin, BBA 247, 542 ( 197 I ). 50. G . Banfalvi, S. Csuzi, A. Ohlbaum, and F. Antoni, ABB 14, 53 (1979).

240 IV.

KAREN TELANDER MUSKAVITCH AND STUART LINN

The Reactions Catalyzed

A. ATP-DEPENDENT DOUBLE-STRANDED DNA EXONUCLEASE ATP-dependent double-stranded DNA exonuclease activity defines an exonuclease V enzyme (51). In-depth characterization of this activity has been done for Eco exoV, Hind exoV, Bsu exoV, and Mlu exoV. A terminus is required on the double-stranded DNA substrate. The enzymes are active on linear duplex DNA from many sources (If, 3 6 , 4 3 , 47, 5 / , 52), and phosphomonoester groups are not required for Eco exoV activity (53). Covalently closed circular double-stranded DNAs are not substrates (36, 43, 51, S2, 54), nor are nicked duplex circles regardless of whether 3’- or 5’-phosphomonoesters are present at the nick (52-54). If a single-stranded gap of greater than 5 nucleotides is introduced into a duplex circular DNA, Eco exoV is able to degrade the DNA (53, 5 3 , having presumably opened the DNA with its single-stranded DNA endonuclease activity. It has been reported that Hind exoV is not active on gapped circular DNA, but these experiments were done under conditions where the single-stranded DNA endonuclease is not active (52). The degradation of linear, but not circular, duplex DNA by Eco exoV is an exploitable property, although its utilization is limited by the difficulty of purifying the enzyme. The enzyme can be used to assay restriction endonucleases (56) and to eliminate cellular DNA in plasmid preparations. Both Eco exoV andHind exoV are inhibited by cross-links within DNA in a manner that suggests the enzyme degrades the DNA from a terminus until it encounters a cross-link, and then stalls (57, 58). Eco exoV is inactive on duplex RNA but has limited activity on RNA-DNA hybrids, presumably degrading DNA termini up to a hybrid region (53).The treatment of circular or linear duplex DNAs with UV light or X-rays does not change their respective failure or success as substrates forEco exoV (53). ATP is a required cofactor for the double-stranded DNase of all exoV enzymes. Nucleoside triphosphate specificities for all the bacterial en51. 52. 53. 54. 55. 56. 57. 58.

M. Wright, G . Buttin, and J. Hurwitz,JBC 246, 6543 (1971). E. A. Friedman and H. 0. Smith, JBC 247, 2859 (1972). A. E. Karu, V. MacKay, P. J. Goldmark, and S . Linn, JBC 248, 4874 (1973). Y. Takagi, K. Matsubara, and M. Anai, BBA 269, 347 (1972). A. Prell and W. Wackernagel, EJB 105, 109’( 1980). D. Lackey and S. Linn, “Methods in Enzymology,” Vol. 65, p. 26, 1980. A. E. Karu and S. Linn, PNAS 69, 2855 (1972). M. Orlosky and H. 0. Smith,JBC 251,6117 (1976).

13. recBC- LIKE ENZYMES

24 1

zymes have been investigated: Eco exoV (43, 51, 591, Hind exoV (9),Bsu exoV ( l o ) ,Mlu exoV (60), Bla exoV (48), Msm exoV (49), Dpn exoV (8), Bce exoV ( 5 0 , Pae exoV (13), and Afa exoV (14). ATP is generally the most effective, and dATP nearly as effective. The efficacy of other riboand deoxytriphosphates varies with the enzyme. During DNA degradation the ATP is hydrolyzed to ADP and Pi and, as might be expected, all pure di- and monophosphates, as well as P-y ATP analogues, are ineffective. Interestingly, many ATP residues are hydrolyzed for each phosphodiester bond cleaved in the DNA. The optimum ATP concentration is usually between 10-s and M . In most cases, higher concentrations of ATP inhibit the double-stranded DNA exonuclease, but not the other activities of the enzymes. A divalent cation is required by all the exoV enzymes. Mg2+is generally best with an optimal concentration of around 10 mM. Mn*+is Less effective and has a lower optimal concentration. In some cases other divalent cations can stimulate the ATP-dependent double-stranded DNA exonuclease ( 3 , 4 7 ) but usually, when they are added in the presence of MgC12, they are inhibitory (3, 8, 9, 47, 50, 6 1 ) . As more fully discussed below, changes in the species or concentration of the divalent cation can have dramatic effects on the activities of the exoV enzymes (36, 61, 62). The pH optimum for all of the enzymes is around 9, and some have interesting ionic strength dependences. NaCl can inhibit Eco exoV if present at greater than 40 mM ( 5 / ) ,and there is evidence that K+, but not Na+, enhances the ability to degrade more than one DNA duplex (63). Hind exoV is 60% more active in 0.5 M Tris-HC1 than in 0.1 M , but is 50% inhibited by 0.125 M NaCl(9). In the case of Msm exoV, the identity of the salt is not important, but an ionic strength of approximately 0.3 M is required for maximum activity (49). Various factors that can inhibit the ATP-dependent double-stranded DNA exonuclease have been studied. In B. subtifis, an inhibitor was detected in crude extracts and later identified as small DNA fragments (48). Proteins that bind DNA can also decrease the production of acid-soluble oligonucleotides. Eschierichia cofi single-stranded DNA-binding protein inhibits Eco exoV (64), and low molecular weight, basic, relatively thermostable proteins from B. subtifis affect the Bsu exoV (65). Many E. coli 59. D. C. Eichler and I. R. Lehman, JBC 252, 499 (1977). 60. M. Anai, T. Hirahashi, M. Yamanaka, and Y. Takagi, JBC 245,775 (1970). 61. J. Rosamond, K. M. Telander, and S. Linn, JBC 254, 8646 (1979). 62. K . W. Wilcox and H. 0. Smith, JBC 251, 6127 (1976). 63. U. Hermanns and W. Wackernagel, EJB 76, 425 (1977). 64. V. MacKay and S. Linn, JBC 251, 3716 (1976). 65. A . V. Chestukhin, V. I. Fedchenkov, and M. F. Shemyakin, M d . B i d . 13,499 ( 1979).

242


bacteriophages are also known to produce proteins that inhibit Eco exoV (6669). There has been only limited work on the effects of chemical inhibitors, the most interesting being the inhibition by pyridoxal phosphate of Eco exoV (70). The product of the ATP-dependent exonuclease activity is a mixture of acid-soluble 5'-phosphomonoester oligonucleotides. The average length of the oligonucleotides varies between 3 and 6 nucleotides depending upon the enzyme (9, 13, 36, 43, 47, 49, 71). When the time course of product formation is investigated using either sedimentation velocity analysis or viscometry , it is observed that the acid-soluble oligonucleotides are present from the very beginning of the reaction. When present in excess, undegraded substrate DNA molecules persist until the end of the reaction (8, 9, 43, 47, 49, 53, 71, 72). These data indicate that the ATP-dependent double-stranded DNase is exonucleolytic and processive, completely degrading one DNA molecule to oligonucleotides before beginning to degrade another. Studies with cross-linked duplex linear DNAs support this mechanism (13, 57, 58). The sedimentation velocity analyses indicate that intermediates in the double-stranded DNA exonuclease reaction do not tend to accumulate. However, they may be present in Dpn exoV and Bla exoV reactions (8, 71), and can be observed at short times in Hind exoV reactions (62, 73). Changes in the concentrations of ATP, Mgz+and salt can greatly increase the accumulation of intermediates in the processive exoV degradation. For Mfu exoV, low concentrations of ATP, which limit its availability, result in the production of sedimentable DNA fragments (54), but no further work has been done on the fragments. To increase the intermediate yield for the E. coli, H. influenzae, and 3. subtilis enzymes, the concentration of ATP is increased to 1-5 mM (36, 53, 62) which, as noted previously, inhibits the production of acid-soluble oligonucleotides. Additional modifications of the reaction conditions can further favor the production of intermediates: for Eco exoV the ionic strength is increased slightly (53); for Hind exoV both the concentration of Mg2+and the ionic strength are 66. Y. Sakaki, J . Virol. 14, 1611 (1974). 67. A. E. Karu, Y. Sakaki, H. Echols, and S. Linn, JBC 250, 7377 (1975). 68. R. Pacumbaba and M. S. Center, J . Virol. 16, 1200 (1975). 69. F. VanVliet, M . Couturier, J. DeLafonteyne, and E. Jedlicki, Mol. Gen. Genef. 164, 109 (1978). 70. M. Anai, T. Fujiyoshi, J. Nakayama, and Y. Takagi, JBC 254, 10853 (1979). 71. M . Anai, M. Yamanaka, T. Shibata, T. Mihara, T. Nishimoto, and Y . Tagaki, J. Biochem. (Tokyo) 78, 115 (1975). 72. P. J. Goldmark and S. Linn,PNAS 67, 434 (1970). 73. E. A. Friedman and H. 0. Smith, N ~ f u r eNew Biol. 241, 54 (1973).

243

13. recBC- LIKE ENZYMES TABLE I1

CHARACTERISTICS OF REACTION INTERMEDIATES“ Characteristics Tailed fragments Length of duplex region (base pairs) Length of tails (nucleotides) Range Mean Termini of tails Single-stranded fragments Length (nucleotides)

Eco exoV (75)

Hind exoV (74, 6 3 )

300-27,OOO

1000-36,000b and -2000b

800-5,OOO

51.5,00Ob

3,200 5’ and 3‘

4,200 3’

100-500

5 21 ,OOOb

T7 phage DNA, the substrate in these experiments, is 39,000 base pairs long. Estimated from data reported in micrometers assuming 2800 base pairs or 2900 nucleotides per micrometer for duplex and single-stranded DNA, respectively.

decreased (62, 73); forBsu exoV the pH is decreased (36). The amounts of Eco exoV and Hind exoV are also increased to give a more synchronous reaction. Biochemical (53, 62, 73, 74) and electron micrographic (62, 7 4 ) characterization of the Eco exoV and Hind exoV intermediates reveal that they are of two classes: (1) Duplex DNA fragments with terminal singlestranded tails, and (2) large single-stranded fragments. Table I1 summarizes the details of their properties. While similarities between the intermediates produced by Eco exoV and Hind exoV are evident, several important differences exist. Hind exoV produces only 3’ tails, whereas Eco exoV produces about equal numbers of 3‘ and 5’ tails. Only rarely do intermediates of the Hind exoV reaction have two tails, but it is not uncommon to observe a single-stranded tail on each end of a duplex Eco exoV intermediate. This latter difference may reflect the fact that one enzyme per terminus was added in the Hind exoV study, whereas three to four per terminus were added in the Eco exoV study. Hind exoV produces a distinct class of duplex intermediates about 2000 base pairs long, possibly with single-stranded tails, which accumulate during the course of the reaction. No similar, distinct class is observed for Eco exoV. Further studies of Eco exoV intermediates have been carried out using a very gentle technique to spread the molecules for electron microscopy (75). The presence of terminal single-stranded loops associated with one 74. V. MacKay and S. Linn, JBC 249, 4286 (1974). 75. K . M. T. Muskavitch and S . Linn, in, “Mechanistic Studies of DNA Replication and Genetic Recombination” (B. Alberts and F. C. Fox, eds.), ICN-UCLA Symp. Mol. Cell. Biol., Vol. 19, p. 901, 1980.

244


or two single-stranded tails is revealed, as is the rare occurrence of pairs of single-stranded loops located within otherwise duplex regions. The relevance of all these data to the mechanism of the double-stranded DNase is discussed in Section V. Unfortunately, extensive characterization of the intermediates produced by Bsrr exoV has not yet been done, but sedimentation velocity analysis (36) of the intermediates suggests that they may be similar to the intermediates produced by Eco exoV and Hind exo V. Along with partially degraded DNA molecules, complexes between enzyme and DNA may be regarded as reaction intermediates. Whereas no one has succeeded in detecting Eco exoV-DNA complexes, complexes have been detected with the enzymes from H. itzjkenzue (19), M . linteus (761, and M . sntegrnatk (77). The optimum ATP and Mgz+concentrations for complex stability vary. When testing with SV40 DNA, Hind exoV binds only to the linear duplex form of DNA and only at termini. Mlu exoV binds best to blunt duplex DNA such as T7 DNA, and more poorly to molecules with single-stranded tails, such as hDNA or T7 DNA following limited exonuclease 111 digestion. The binding of Bsu exoV to duplex DNA is unique; it appears to bind all over the DNA and cause a nonspecific pairing of duplexes, the extent of which depends upon the concentration of enzyme (78). ATP is not required.

B. SINGLE-STRANDED DNA EXONUCLEASE With the exception of Pric exoV, all of the known bacterial exoV enzymes have associated single-stranded DNA exonuclease (8, 9, I I , 13, 14, 35, 36, 42,47-50, 72). ATP dependence varies from none for one preparation ofBsrr exoV (11) to total forEco exoV (721, Hind exoV (9), Mll4 exoV (47),and Afii exoV (14). The rate of degradation of single-stranded DNA is always less than that for duplex DNA, but the degree varies between a twofold decrease for Eco exoV (43) and a 100-fold decrease for Blri exoV (48). [This difference probably explains why molecules with singlestranded tails are degraded more slowly than duplex molecules with blunt ends (55, 62).] Reaction conditions for maximum activity are usually the same as for the duplex exonuclease. The single-stranded DNA exonuclease activity ofEco exoV is not inhibited by high concentrations of ATP, but Afrr exoV has the same ATP dependence for both of its exonuclease 76. B . Van Dorp, M . T. E . Ceulen, H. L. Heijneker, and P. H. Pouwels, BBA 299, 65 (1973). 77. F. G . Winder and P. A. Sastry,FEBS (Fed. Eitr. Biochetn. Soc.) L e u . 17, 27 (1971). 78. S. Ohi, D. Bastia, and N . Sueoka, Notidre (Lotidoti) 248, 586 (1974).

13. rrcBC- LIKE ENZYMES

245

activities. Studies of Eco exoV indicate that the single-stranded DNA exonuclease is processive and produces acid-soluble oligonucleotides (72).

C. SINGLE-STRANDED DNA ENDONUCLEASE Only three species of exoV have been tested for the presence of singlestranded DNA endonuclease activity. It is present in Eco exoV (72) and Hind exoV (52),but absent in Prie exoV under a variety of reaction conditions (13). It is important that the enzyme be tested under more than one set of reaction conditions because this activity is not maximally active under conditions optimal for the ATP-dependent double-stranded DNA exonuclease. The Hind exoV single-stranded endonuclease is active only at Tris-HC1 concentrations less than 0.25 M , and so was originally overlooked (32).Eco exoV single-stranded DNA exonuclease has a pH optimum of 7 and is stimulated sevenfold by ATP (72). The optimum ATP concentration is about 2 mM and inhibition is not observed at high ATP concentrations (4.3). It can be inhibited by E. c d i single-stranded DNA binding protein (SSB) ((54) and CaC1,(61 1. With fd single-stranded circular DNA in the absence of ATP, the endonuclease acts randomly (72), but recent work by Prell and Wackernagle (55) suggests that this is not the case for single-stranded gaps located within duplex DNA. D. DNA-DEPENDENT ATPASE All exoV enzymes have a DNA-dependent ATPase activity that hydrolyzes ATP to produce ADP and inorganic phosphate (8, 11, 13, 14, 43, 48-51, 60, 79). The optimal conditions for the ATPase are generally those for the exonuclease activities. A DNA cofactor is required: Any DNA that can be hydrolyzed by the enzyme suffices, but single-stranded DNA is less effective than double-stranded DNA (11, 43, 50). An exchange reaction between ATP and ADP that does not require DNA has been observed for Mlu exoV (80). Normally, the hydrolysis of ATP is coupled to the degradation of DNA in such a way that the number of ATPs hydrolyzed per phosphodiester bond cleaved can be determined. This number is greater than one and varies between approximately 3 and 40 for different enzymes (8, 11, 13, 14, 43, 48-50, hU, 79). It can also vary with reaction conditions (14, 4 3 ) . Several examples of the uncoupling of ATPase from DNase are known. Eco 79. H . 0. Smith and E. A. Friedman, JBC 247, 2854 (1972). 80. M . Anai and Y. Takagi, JBC 246, 6389 (1971).

246


exoV, Hind exoV, and Pue exoV form a complex with cross-linked, linear DNA that (although unable to degrade the DNA) continues to hydrolyze ATP (13, 57, 58). A similar phenomenon is observed for Eco exoV in the presence of an RNA-DNA hybrid (53).SSB inhibits the DNase activity of Eco exoV but not the ATPase such that more ATP molecules are hydrolyzed for each phosphodiester bond cleaved (64). High levels of ATP inhibit the double-stranded DNA exonuclease of Eco exoV and Hind exoV but do not so affect the ATPase ( 4 3 , 5 3 , 5 9 , 6 2 ,79). The effect can be enhanced for Hind exoV if the concentration of Mg2+ is decreased (62). For Eco exoV the most dramatic uncoupling is seen in the presence of Caz+ (81). The DNase activities can all be completely inhibited without significantly affecting the ATPase (61). It should be noted that not everything that inhibits the double-stranded DNA exonuclease activity is also an uncoupler. Pyridoxal phosphate (70) and a variety of divalent metallic salts (61) inhibit both DNase and ATPase. E. ATP-DEPENDENT DOUBLE-STRANDED DNA UNWINDING The ability of exoV to catalyze unwinding is suggested by the nature of the double-stranded DNA exonuclease reaction intermediates and by the lack of any obvious strict coupling between ATPase and DNase. Intermediates with single-stranded regions are formed by Hind exoV if the concentration of Mg2+is lowered (62). In a similar way, unwinding is favored over DNase by Eco exoV in the presence of SSB (64)or high levels of ATP (53,59, 74). In the presence of Ca2+,only ATP-dependent unwinding and associated ATP hydrolysis are catalyzed by Eco exoV (61,811. Under nearly physiological conditions, which include 1 mM MgC12, 1 mM CaCl,, and 5 mM ATP, linear duplex T7 DNA appears to be unchanged by Eco exoV, although ATP is hydrolyzed. If SSB is also present in such a reaction “forks” (two single-stranded tails at the same terminus) are formed at the ends of duplex DNA and grow larger with time until only single-stranded DNA remains. This single-stranded DNA is about one-third unit length because of a very low level of nicking activity that is still retained by Eco exoV. The exact amount of nicking vanes with the reaction conditions (61). If SSB is added after the reaction has begun but just shortly before it is terminated, small denatured regions, or “bubbles,” are observed within the duplex. Thus, with Ca2+present, Eco exoV apparently moves along in the interior of the DNA double helix, transiently denaturing the duplex in its vicinity as it travels down the molecule. When present, SSB prevent 81. J. Rosamond, B. Endlich, K. M. Telander, and S. Linn, CSHSQB 43, 1049 (1979).

13. rPcBC-LIKE ENZYMES

247

reannealing of the strands unwound by the enzyme, and single-stranded DNA accumulates. It has been suggested that Hind exoV moves within a DNA double helix in a similar manner ( 6 2 ) . Recent electron microscopic studies by Taylor and Smith indicate that the unwinding by Eco exoV in the presence of Ca2+proceeds by way of single-stranded loops associated with two single-stranded tails (8.2,83 ) in a manner reminiscent of the double-stranded DNA exonuclease (75). However, under the unwinding conditions internal paired single-stranded loops (twin loops) are observed more frequently (see below).

V.

Models for the Mechanism of Action of the ATP-Dependent Double-Stranded DNA Exonuclease

Only the H. itiflirenzue and E. coli enzymes have been studied in sufficient depth to formulate detailed models for their action on doublestranded DNA. Wilcox and Smith have proposed the model shown in Fig. 1 for Hind exoV ( 6 2 ) . Following binding to a terminus,Hind exoV begins to travel along the interior of the double helix if ATP is present. At approximately 6000 base pair intervals, the enzyme begins to hydrolyze one strand, forming single-stranded fragments. After continuing in this mode for approximately 4000 base pairs, the enzyme nicks the previously undegraded strand, releasing a duplex of roughly 2000 base pairs with a 3’ single-stranded tail about 4000 nucleotides long. The enzyme remains bound to the original DNA molecule and reverts to its “tunneling” mode, again transiently denaturing the double helix as it moves through it until, after about 2000 base pairs, it begins again to degrade the same strand. Hiri d exoV continues this processive fragmentation until it completes the formation of intermediates from one molecule. It can then begin either to degrade another intact duplex, or to degrade an intermediate to acidsoluble oligonucleotides. The former is the preferred course. Figure 2 depicts a mechanism for Eco exoV recently proposed by Muskavitch and Linn (75). It is based upon an earlier model (74) but is more consistant with further electron microscopic observations made by them (75) and by Taylor and Smith (8.2,83). It is proposed to apply to reactions with or without Ca2+.As shown, the enzyme binds to a duplex terminus (Fig. 2A) and begins to unwind the DNA double helix, forming one strand 82. A. Taylor and G. R . Smith,iri. “Mechanistic Studies of DNA Replication and Genetic Recombination” 1B.Alberts and F. C. Fox, eds.), ICN-UCLA Symp. Mol. Cell. Biol., Vol. 19, p. 909, 1980. 83. A. Taylor, and G . R . Smith, CeII, 22, 447 (1980).

248


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FIG.2. Model for the degradation of duplex DNA by Eco exoV (75). The structures in the circle represent true reaction intermediates; those on the periphery are formed by disruption of the intermediates during observation procedures (e.g., electron microscopy).

13. recBC-LIKE ENZYMES

249

into a loop and displacing the other as a tail (Fig. 2B). Both 3’- and 5’-terminated tails are formed with equal frequency. If SSB is present when such a loop-plus-tail structure is disrupted (during electron microscopic sample preparation, for example), renaturation (Fig. 2B‘) is prevented and a fork is formed (Fig. 2B”). When the loop is several thousand nucleotides in length it is hydrolyzed, beginning at its terminus to form single-stranded fragments several hundred nucleotides long. During this hydrolysis, a loop and two tails are observed at a terminus (Fig. 2C), or, if disrupted, either a fork (Fig. 2C”) or a single-stranded tail (Fig. 2C’), depending upon whether SSB is present. If the strand composing the loop is not cleaved but is partially threaded past the enzyme, it can reanneal with the complementary region of the tail. The DNA still held by exoV in the loop, and that complementary to it in the tail, then form a pair of single-stranded loops (twin-loops), previously called “rabbit ears” (Fig. 2,RE). This process may occur at times when loop DNA is being threaded through the enzyme before it is cut off as a single-stranded fragment. SSB in the reaction does not prevent the formation of twin loops. Complete hydrolysis of the loop to single-stranded fragments produces a molecule with a single-stranded tail and the enzyme bound at the single strand : double-strand junction (Fig. 2D). Since the enzyme is observed to be essentially processive, hydrolysis of the tail must occur in a manner that allows the enzyme to remain associated with the same duplex DNA molecule. This could be accomplished by hydrolysis of the tail from its junction with the duplex toward its terminus as depicted in Fig. 2E. The enzyme, having completed hydrolysis of the tail, is able to reinitiate the reaction cycle (Fig. 2A). Repetition continues until the duplex molecule is completely reduced to single-stranded fragments. The released enzyme can then begin to unwind and hydrolyze another DNA molecule, or to degrade the single-stranded fragments to acid-soluble oligonucleotides. It is obvious that the proposed mechanisms for Eco exoV and Hind exoV are very similar. The one essential difference is that, unlike Eco exoV, Hind exoV appears to proceed via a two-stage process, first forming a distinct class of intermediates and then degrading them. Both act processively from the end of a linear molecule and simultaneously unwind the double helix as they move within its interior (rather than along its exterior). As previously suggested (84), ATP hydrolysis is believed to provide the energy for this combination of translocation and denaturation. The type-I restriction endonucleases represent, in contrast, a class of enzymes that use ATP hydrolysis to power translocation along the ex84. F. G. Winder, Natirre New B i d . 236, 75 (19721.

250

KAREN TELANDER MUSKAVITCH A N D STUART LINN

terior of the DNA double helix (85). The details of the manner in which Eco exoV and Hind exoV unwind the DNA may be slightly different, since Hind exoV produces only 3' tails, where Eco exoV produces both 3' and 5' tails. However, it is reported that Hind exoV also forms both loopplus-tail structures and twin-loops (8.3). The preeminence of unwinding, even in reactions that degrade DNA to oligonucleotides, and particularly under conditions believed to approximate those within the bacterium, suggests that unwinding is an important function of the enzyme in vivo. Further credence is lent to this view by work with temperature-sensitive mutants of E. coli . At the nonpermissive temperature, the mutants display the decreased viability characteristic of recBC mutants (86). Among the nuclease activities, only the doublestranded DNA exonuclease of the isolated enzyme is found to be temperature sensitive (34).It is possible that a deficiency in unwinding is the lesion in these mutants causing both decreased viability and.loss of the doublestranded DNA exonuclease. Unwinding, which creates single-stranded DNA, may also explain the role of Eco exoV in recombination. Single-stranded DNA is believed to play an important role in recA-mediated recombination (87). Since other cellular enzymes also are able to unwind DNA (88, 8 9 ) , loss of exoV activity need not result in a total inability to carry out recombination. VI.

Conclusion

ExoV enzymes are found throughout the bacterial kingdom. All have ATP-dependent double-stranded DNA exonuclease and DNA-dependent ATPase activities. Also associated with the enzyme may be ATPdependent single-stranded DNA exonuclease, ATP-stimulated singlestranded DNA endonuclease, and ATP-dependent DNA unwinding. Investigation of the mechanism of action of the enzyme indicates that DNA unwinding is fundamental to several of the enzyme's activities and may well be its most important function in the cell. The complete degradation of DNA to acid-soluble oligonucleotides may only serve a function in restriction. 85. B. P. Endlich and S. Linn. (1980), this volume, Chap. 9. 86. S. R. Kushner, J . Bacterial. 120, 1213 (1974). 87. T. Shibata, C. DasGupta, R. P. Cunningham, and C . M. Radding, PNAS 76, 1638 ( 1979). 88. M. Abdel-Monem and H. Hoffman-Berling, H E 65, 431 ( 1976). 89. M. Abdel-Monem, M . C . Chanal, and H. Hoffman-Berling, H E 79, 33 (1977).

Enzymes That Incise Damaged DNA ERROL C. FRIEDBERG THOMAS BONURA ERIC H. RADANY JACK D. LOVE

.

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . .

251

11. Enzymes That Attack Phosphodiester Bonds in DNA Following

Hydrolysis of N-Glycosylic Bonds (AP Endonucleases) . . . . . . A. DNA Glycosylase-Associated AP Endonuclease Activities . . . . B. A P Endonucleases Associated with Other Catalytic Activities . . C. AP Endonucleases Without Associated Catalytic Activities. . . . D. Concluding Comments on AP Endonucleases . . . . . . . . . . 111. Enzymes That Attack Phosphodiester Bonds in Damaged DNA with Intact N-Glycosylic Bonds . . . . . . . . . . . . . . . . . . . A. UV Endonuclease of Escherichia coli . . . . . . . . . . . . . B. UV Endonuclease Activity in Other Organisms . . . . . . . . . C. Endonuclease V of Escherichiu coli . . . . . . . . . . . . . . D. Endonuclease Activity Directed at Photoalkylated Purines in DNA Note Added in Proof . . . . . . . . . . . . . . . . . . . .

1.

252 252 263 267 273 274 274 277 277 278 279

Introduction

The excision of damaged or inappropriate nucleotides from DNA can occur by a number of different biochemical pathways, depending on both the nature of the specific base damage in question and on the particular organism under investigation (I). Evidence to date indicates that an integ1. P. C. Hanawalt, P. K. Cooper, A. K. Ganesan, and C. A. Smith,Annu. Rev. Biochem. 48, 783 ( 1979).

25 1 THE ENZYMES,Val. XIV Copyright 01981 by Academic Press, Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

252

E. FRIEDBERG, T. BONURA, E. RADANY, J. LOVE

ral component of most (if not all) pathways of excision repair of DNA is the enzyme-catalyzed hydrolysis of phosphodiester bonds by specific enzymes that we collectively designate as DNA-incising uctivities . Such enzymes fall into two major classes: ( I ) Those that attack phosphodiester bonds in DNA subsequent to the hydrolysis of the associated glycosylic bond that links a nitrogenous base to the deoxyribose-phosphate backbone: and (2) those that directly attack phosphodiester bonds in damaged DNA. The former class of enzymes is generally designated as AP (apurinicl apyrimidinic) endonucleases (2 ) because their endonuclease activity is confined to sites of base loss in DNA. Such substrate sites arise by the spontaneous hydrolysis of N-glycosylic bonds in DNA (the rate of which is markedly accelerated by certain chemical modifications of bases), or by enzyme-catalyzed hydrolysis of these bonds by DNA glycosylases (31. Most known AP endonucleases appear to be free of known DNA glycosylase activities. However, other AP endonucleases have not yet been physically separated from certain specific DNA glycosylases . Until evidence to the contrary is available, we will consider DNA glycosylaseassociated AP endonucleases as a distinct category of DNA-incising activities. In this chapter attention is focused principally on DNA-incising activities from Escherichicr coli, since this organism has been the subject of extensive genetic and biochemical investigations on the excision repair of DNA ( I ) . However, where appropriate, enzyme activities from other biological sources are considered. II. Enzymes That Attack Phorphodiester Bonds in DNA Following Hydrolysis of N-Glycosylic Bonds (AP Endonucleases)

A. DNA GLYCOSYLASE-ASSOCIATED AP ENDONUCLEASE ACTIVITIES 1. UV DNA-lticising Activity .from Bncteriophage T4-l~fected Esclierichicr coli

The earliest report of a gene product coded by phage T4 involved in the excision of pyrimidine dimers from UV-irradiated DNA, was by Takagi et 2. S . Linn, ii7 "DNA Repair Mechanisms" (P. C. Hanawalt, E . C. Friedberg, and C. F. Fox, eds.), p . 175. Academic Press, New York, 1978. 3. T. Lindahl, Progr. Nircleic Acids Res. Mol. B i d . 22, 135 (1979).

14. ENZYMES THAT INCISE DAMAGED DNA

253

in 1968 (4). These investigators reported that incubation of UVirradiated DNA with extracts of TCinfected E. coli resulted in the loss of thymine-containing pyrimidine dimers from the acid-insoluble fraction of the DNA. Subsequently it was demonstrated that extracts of TCinfected E. coli contain an activity that catalyzes the preferential nicking of UVirradiated duplex DNA (5-7). This activity was shown to be dependent on a functional denV gene (8) formerly called the v gene (9) of phage T4. The isolation of denV gene mutants that are temperature-sensitive with respect to both UV sensitivity and UV DNA-incising activity provided direct evidence that the denV locus is the structural gene for an enzyme activity required for the incision of UV-irradiated DNA (10). This activity has been variously referred to in the literature as the T4 UV endonuclease (5, 6 , I / ) ,or endonuclease V (12, 13) of phage T4. For reasons that will become evident in the course of later discussion, we suggest that it is more appropriate to refer to this activity as the T4 UV DNA-incising activity. The T4 U V DNA-incising activity has been extensively purified in a number of laboratories, (6, 7, 12, 14, 1 3 , generally using assays that directly measure hydrolysis of phosphodiester bonds in UV-irradiated DNA. An alternative assay described by Seawell et ul. (16) measures the direct binding of protein to UV-irradiated DNA. The interested reader is referred to the references previously cited for detailed descriptions of the purification protocols. It is important to note that in all published purification schemes it has been assumed that the T4 UV DNA-incising activity was an endonuclease that directly catalyzed the hydrolysis of phosphodiester bonds 5’ to pyrimidine dimer sites in DNA. However studies, described later in this chapter, suggest that the T4 UV DNA-incising id.

4. Y. Takagi, M. Sekiguchi, S. Okubo, H. Nakayama, K. Shimada, and S. Yasuda, T. Nishimoto, and H. Yoshihara, CSSSQB 33, 219 (1968). 5. E. C. Friedberg and J . J . King, BBRC 37, 646 (1969). 6. S. Yasuda and M. Sekiguchi, PNAS 67, 1839 (1970). 7. E. C. Friedberg and J . J. King, J . Bacteriol. 106, 500 (1971). 8. W. B. Wood and H. Revel, Bacteriol. Rev. 40, 847 (1976). 9. W. Harm, Virology 19, 66 (1%3). 10. K. Sato and M. Sekiguchi, J M B 102, I5 (1976). 11. K. Minton, M . Durphy, R . Taylor, and E. C. Friedberg, JBC 250, 2823 (1975). 12. S . Yasuda and M . Sekiguchi, BBA 442, 197 (1976). 13. Y. Nishida, S. Yasuda, and M . Sekiguchi, BBA 442, 208 (1976). 14. E. C. Friedberg, A. K. Ganesan, and P. C. Seawetl, “Methods in Enzymology,” Vol. 65, p. 191, 1980. 15. P. C. Seawell, E. C. Friedberg, A. K. Ganesan, and P. C. Hanawalt, in, “DNA Repair: A Laboratory Manual of Research Procedures” (E. C. Friedberg and P. C. Hanawalt, eds.), Dekker, New York, 1981. 16. P. C. Seawell, T. J. Simon, and A. K. Ganesan, Biochemisrry 19, 1685 (1980).

254


activity consists of a pyrimidine-dimer-specific DNA glycosylase plus an associated AP endonuclease. T4 UV DNA-incising activity has a calculated MW -18,000 as measured by gel filtration (1l ) .The activity has no requirement for any known cofactor and is fully active in the presence of 10 mM EDTA (6, 7, 11). In the absence of EDTA, activity is slightly stimulated in the presence of Mg2+or Mn2+;other divalent cations and monovalent ions have little effect on its activity (6, 7, I / ) . Yasuda and Sekiguchi (6) reported that the T4 UV DNA-incising activity is sensitive to inhibition in 0.4 mM p-chloromercuribenzoate. However, Friedberg el a/.(14) observed only -20% inhibition by 0.1 mM p-chloromercuriphenylsulfonic acid, and Friedberg and King (7) observed no loss of activity in the presence of 0.1 mM p-chloromercuribenzoate. Enzyme activity has a broad pH optimum between 7-9 with a maximum at pH 7.2 (6, 7, if). The activity is induced very early after infection of E. coli, reaching maximum levels about 10 minutes after infection (7, 12). The T4 UV DNA-incising activity attacks cyclobutyl dipyrimidines (pyrimidine dimers) in both duplex and single-stranded DNA (6, 7, 11, 12). Both linear and circular DNA that contain pyrimidine dimers are recognized as substrate. Dimers that contain glucosylated hydroxymethylcytosine (present in the DNA of the T-even phages) are also recognized as substrate. Single-strand incisions in duplex UV-irradiated DNA are catalyzed 5‘ to pyrimidine dimers (11, 12), leaving 3’-OH and 5’-P termini (11). Within the limits of accurate measurement, saturating amounts of enzyme activity catalyze the formation of 1 nick per pyrimidine dimer (17). When heteroduplex DNA that contains pyrimidine dimers in only one strand is treated with T4 UV DNA-incising activity, all detectable nicks occur exclusively in the dimer-containing DNA strands (18). In addition, enzymatically photoreactivated UV-irradiated DNA is no longer a substrate for the enzyme (19). DNA treated with nitrogen mustard, methyl methanesulfonate, 4-nitroquinoline-l-oxide,mitomycin C, N methyl-N’-nitro-N’-nitrosoguanidine, or ionizing radiation is not a substrate for the enzyme (13, 2 0 , 2 / ). These studies indicate a strict specificity of the enzyme activity for cyclobutyl dipyrimidines in DNA, although the possibility that the enzyme recognizes this photoproduct in RNA has not been excluded. 17. E. C. Friedberg, i n , “Molecular Mechanisms for Repair of DNA’ (P. C. Hanawalt and R. B. Setlow, eds.), p. 125. Plenum, New York, 1975. 18. T. J. Simon, C. A. Smith, and E. C. Friedberg, JBC 250, 8748 (1975). 19. A. K. Ganesan, JME 87, 103 (1974). 20. E. C. Friedberg, Mutation Res. 15, 113 (1972). 21. M. It0 and M. Sekiguchi, Jap. J . Genet. 51, 129 (1976).


255

Further investigations have demonstrated that this stringent specificity is a property of a DNA glycosylase activity and not of an endonuclease. The pyrimidine dimer-DNA glycosylase function of a UV DNA-incising activity was first demonstrated by Grossmanef 01. (22) and by Haseltineet a / . ( 2 3 ) using a preparation of enzyme from M. lufeus (see Section II,A,2). Direct evidence for such activity in T4 enzyme preparations has been provided by studies from this laboratory ( 2 4 , 2 5 ) .DNA labeled in thymine with tritium was UV-irradiated under photosensitizing conditions t o produce 17% of thymine in thymine-containing pyrimidine dimers. The DNA was incubated with saturating amounts of T4 UV DNA-incising activity dnd then reirradiated at 254 nm to photoreverse pyrimidine dimers (Fig. 1). This protocol was found to promote the release of free thymine from DNA. The generation of free thymine was shown to be strictly dependent on the presence of thymine-containing pyrimidine dimers in the substrate DNA, incubation with T4 denV gene product, and photoreversal (24, 35). These results are consistent with the notion that the thymine detected is derived from pyrimidine dimers in which N-glycosylic bonds were hydrolyzed by catalytic action of the T4 enzyme preparation. Experiments have also demonstrated that the amount of radioactivity measured in free thymine is one-half the amount lost from thyminecontaining pyrimidine dimers by photoreversal at several reirradiation fluences (Fig. 2). The amount of free thymine liberated from enzymetreated DNA irradiated with 12,000 JimZof photoreversing fluence corresponds to 60% of the thymine-containing pyrimidine dimers, or 5% of the total radioactivity. Two important conclusions can be drawn from this data. First, since no other photoproducts are known to be present at such high concentrations in UV-irradiated DNA, it is highly improbable that free thymine is derived from any source other than thymine-containing pyrimidine dimers, especially in light of the kinetic correlation between free thymine release and pyrimidine dimer photoreversal. Second, in the experiment previously quoted, in which enzyme-treated DNA was exposed to 12,000 J/m2 of photoreversing light, at least 60% of thyminecontaining pyrimidine dimers in DNA must have had an N-glycosyl bond hydrolyzed by T4 enzyme treatment, so pyrimidine dimer-DNA glycosylase activity cannot be a minor component of T4 UV DNA-incising 22. L. Grossman, S . Riazuddin, W. A. Haseltine, and C. Lindan, CSHSQE 43, 947 (1979). 23. W. A. Haseltine, L. K. Gordon, C. P. Lindan, R. H. Grafstrom, N. L. Shaper, and L. Grossman, Nufure (London) 285, 634 (1980). 24. E. H. Radany and E. C. Friedberg, Natrtre (London) 286, 181 (1980). 25. E. H . Radany, J. D. Love, and E. Friedberg,in, “Chromosome Damage and Repair” (E. Seeberg and K. Kleppe, eds.). Plenum, New York, in press.

256


PYRIMIDINE DlMER DNA GLYCOSYLASE

3' A P ENDONUCLEASE

MONOMER IZATION OF THYMINE DlMER

FIG. 1. Model for two-step incision of UV-irradiated DNA. The figure shows a diagrammatic representation of part of a single strand of DNA that contains a thymine-thymine dimer (a). The dimer is shown to be attacked successively by a pyrimidine dimer-DNA glycosylase (which specifically hydrolyzes the 5' glycosyl bond of the dimer) (b), and by an AP endonuclease (c). The latter is shown to cleave the phosphodiester backbone 3' to the apyrimidinic site. [Also shown (d), is the predicted release of free thymine following monomerization of the dimer after it has been attacked by the DNA glycosylase activity.] From Grossman et a / . ( 2 2 ) and Haseltine et a / . (23). activity. This would be expected, since genetic studies have shown that both pyrimidine dimer-DNA glycosylase activity (3)and UV DNAincising activity (5) are associated with the denV gene of phage T4. Finally, the 1 : 2 stoichiometry between radioactivity in free thymine following photoreversal, and that in thymine-containing pyrimidine dimers lost following photoreversal, indicates that only one of the N-glycosylic bonds

14.

257

ENZYMES THAT INCISE DAMAGED DNA

c

w

I

Y

-

. .

PP

‘ h b m --a v)

0

1

2

3 4 5 6 7 PHOTOREVERSAL OF FLUENCE ( kJ/m2)

8

9

FIG.2. Quantitative relationship between the amount of radioactivity measured in free thymine and that associated with the reversal of thymine-containing pyrimidine dimers. The latter parameter is calculated from the measured loss of thymine-containing dimers in ENA. (From Radany ef a / . (15); reproduced with permission of the publishers.) in the dimer is hydrolyzed by the enzyme. The previously documented evidence that nicking of UV-irradiated DNA is catalyzed 5’ with respect to pyrimidine dimers ( I ] , 12) indicates that the 5’-glycosyl bond is uniquely hydrolyzed by the enzyme preparation. Further evidence for such a proposed mechanism of action of the T4 UV DNA-incising activity stems from studies by Seawell et al. (16) who have shown that when UV-irradiated DNA is incubated with T4 enzyme preparations at low temperature, alkaline-labile (presumably apyrimidinic) sites are demonstrable. In addition, the hydrolysis of phosphodiester bonds in such a substrate is effected under near neutral pH by subsequent incubation of the DNA with exonuclease 111 of E. coli. (Exonuclease 111 contains the quantitatively major AP endonuclease activity of E. coli; see Section II,B,l). Warner et d.(26) have also provided evidence for a T4 pyrimidine 26. H. R . Warner, B. F. Demple, W. A. Deutsch, C. M. Kane, and S. Linn, PNAS, in press.

258


dimer-DNA glycosylase. They utilized the observation that DNA polymerase I ofE. coli catalyzes DNA strand elongation at a nick containing a 3' terminal nucleotide significantly more rapidly than at a nick containing a 3' terminal apurinic or apyrimidinic site. When testing the primer-template activity of duplex UV-irradiated DNA incubated with T4 UV DNA-incising activity, they observed it to be low. However, the addition to the T4 enzyme-treated DNA of an AP endonuclease that catalyzes incision of DNA on the 5' side of a site of base loss (5' AP endonuclease) restored normal priming activity for DNA polymerase I. The authors reasoned that the original incubation of UV-irradiated DNA with T4 enzyme must have generated nicks with 3' terminal AP sites, which were then excised by the subsequent addition of 5' AP endonuclease (Fig. 3). The formation of AP sites in UV-irradiated DNA by the T4 enzyme implied the catalytic hydrolysis of one or more DNA glycosyl bonds, presumably at pyrimidine dimers. Warner et al. (26) also demonstrated the photoreversal-dependent release of free thymine from UVirradiated DNA incubated with purified T4 enzyme preparations. The release of free thymine following the monomerization of thyminecontaining pyrimidine dimers in DNA provides a convenient assay for the purification of the TCpyrimidine dimer-DNA glycosylase activity. Such studies are in progress. The incision of UV-irradiated DNA by T4 enzyme preparations demonstrates that the preparations purified thus far contain AP endonuclease activity in addition to the pyrimidine dimer-DNA glycosylase. Attempts to physically separate T4 pyrimidine dimer-DNA glycosylase and AP endonuclease activities have not been successful.

2 . UV DNA-Incising Activity from Micrococcus luteus The discovery of enzyme activity from extracts of M. luteus (formerly called M. lysodeikticus) that preferentially attacks UV-irradiated DNA dates back to a report by Strauss (27) in 1962. Subsequent evidence for such an activity has been reported by a number of laboratories (4,28-35). A UV DNA-incising activity assumed to be a direct-acting endonuclease 27. B. S. Strauss, PNAS 48, 1670 (1%2). 28. A. Rorsch, C. van der Kamp, and J. Adema, BBA 80, 346 (1964). 29. R. L. Elder and R. F. Beers, J . Bacrerriol. 90, 681 (1965). 30. B. Strauss, T. Searashi, and M. Robbins, PNAS 56, 932 (1966). 31. E. Moriguchi and K . Suzuki, BBRC 24, 195 (1%6). 32. H. Nakayarna, S. Okubo, M . Sekiguchi, and Y. Takagi, BBRC 27, 217 (1%7). 33. K. Shimada, H. Nakayama, S. Okubo, M. Sekiguchi, and Y. Takagi, BBRC 27, 539 (1967). 34. S. Okubo, H. Nakayama, M. Sekiguchi, and Y. Takagi, BBRC 27, 224 (1%7). 35. W. L. Carrier and R . B . Setlow, EBA 129, 318 (1966).


259

5’ A P ENDONUCLEASE

FIG. 3. The figure demonstrates the presence of a putative 3’-acting AP endonuclease associated with the phage T4 pyrimidine dimer-DNA glycosylase. The former activity produces a 3’ terminal deoxyribose-phosphate residue (a), which is a poor primer for DNA polymerase I of E. coli. However, following fiwther incubation with a S’-AP endonuclease, the deoxyribosephosphate residue is excised (b), leaving a normal 3’ terminus. The template DNA strand is omitted for clarity. specific for pyrimidine dimers was first purified by Kaplan et a/. (36) and by Carrier and Setlow (37). In 1971 Nakayama et a/. (38)reported that two species of U V DNA-incising activity could be distinguished by isoelectric focusing and by column chromatography, and in 1977 Riazzudin and Grossman (39) reported the extensive purification of two species of U V DNA-incising activity from M. luteus. which they designated as “correndonucleases” I and 11. A mixture of the two “correndonucleases” from M. luteus was used by Grossman ef id. (22) and Haseltine et a / . (23) to explore the distribution of pyrimidine dimers in a sequenced segment of the operator-promotor region of the E. coli lac operon. The rationale of these experiments was that enzyme-catalyzed nicking of the sequenced DhA immediately 5’ to all pyrimidine dimers by a direct-acting endonuclease (as anticipated by the investigators), or by a combined 5’-DNA glycosylase plus 5’-AP endonuclease (as shown in Fig. 4) should, following denaturation of the duplex DNA, yield fragments of precisely predictable length from the “P-labeled 36. 37. 38. 39.

J. C. Kaplan, S. R. Kushner, and L. Grossman, PNAS 63, 144 (1969). W. L. Carrier and R. B. Setlow, J. Bucteriol. 102, 178 (1970). H. Nakayama, S. Okubo, and Y. Takagi, BBA 228, 67 (1971). S. Riazzudin and L. Grossman, JBC 252,6280 (1977).

260


1 FIG. 4. This figure schematically indicates the method by which Haseltine et c d (23) detected a pyrimidine dimer-DNA glycosylase activity in enzyme preparations from M . lrrterrs . The bars under the polynucleotides indicate relative lengths of 5’ radiolabeled fragments (*) following gel electrophoresis of the denatured products of incubation of UVirradiated DNA with M. hrterw “correndonucleases.” Note that the concerted action of a S’-dimer-DNA glycosylase and a 3’-AP endonuclease generates fragments, the size of which is reduced at their 3’ ends by I nucleotide after treatment with alkali due to the presence of an alkaline-labile 3’ terminal apyrimidinic site. Enzyme-catalyzed hydrolysis of phosphodiester bonds by a 5’-endonuclease, or direct hydrolysis of phosphodiester bonds acting immediately S to the dimers, would not result in the alkaline-dependent size reduction of DNA fragments observed.

-

5’ ends (Fig. 4). Surprisingly, these investigators observed by agarose gel electrophoresis that the labeled DNA fragment lengths were -1 nucleotide longer than expected, suggesting that phosphodiester bond hydrolysis catalyzed by the UV DNA-incising activity was between the dimerized pyrimidines rather than on their 5‘ side. The predicted size distribution of denatured DNA fragments resulting from incubation of the UV-irradiated DNA with enzyme was obtained if prior to gel electrophoresis the DNA was subjected to treatment in strong alkali. This result suggested that alkaline-labile sites (presumably sites of depyrimidi-


26 1

nation) were present 5‘ to the sites of phosphodiester bond hydrolysis (Fig. 4). Based on these findings, these investigators proposed the twostep model for the incision of DNA that contains pyrimidine dimers, shown in Fig. 1. The previously reported isolation of two “correndonuclease” activities may reflect the purification of two AP endonucleases from M. luteus, each of which was contaminated with a single pyrimidine dimer-DNA glycosylase. In 1981 Pierre and Laval (40,4/) reported the purification and characterization of two chromatographically distinct AP endonucleases from M. lureus (AP endonucleases A and B), one with a PI of 4.8 and the other with a PI of 8.8 (see Section 11,C,2). These values correspond very closely with the PIS of 4.7 and 8.7 originally reported for “correndonucleases” I and 11. However, Pierre and Laval (40,411 reported that both AP endonucleases catalyze phosphodiester bond breakage 5‘ to sites of base loss. This observation is inconsistent with the results of the gel electrophoresis of DNA reported by Grossman el al. (22) and Haseltine et al. (23 ), which showed alkaline-labile sites exclusively at the 3’ end of DNA fragments, indicating the action of 3‘-AP endonuclease. Hopefully further studies on the AP endonucleases of M. luteus will resolve this apparent contradiction. Haseltine et (11. (23) purified the M. luteus pyrimidine dimer-DNA glycosylase further and achieved a substantial reduction in AP endonuclease activity. Although the most purified fraction still contains detectable AP endonuclease, it is uncertain whether or not the M. luteus pyrimidine dimer-DNA glycosylase contains a physically associated AP endonuclease activity. In this respect it is of interest to note that Tomilinet al. (42) isolated only a single “UV endonuclease” activity fromM. luteus that also contained AP endonuclease activity. The kinetics of heat inactivation of both the “UV endonuclease” and AP endonuclease activities were identical. Furthermore, the two catalytic activities were inactivated at the same concentrations of cyanide and proflavin. Tomilin et al. (42) also reported that the UV DNA-incising activity of their M. luteus fraction was competed for by the addition of duplex DNA that contained apurinic sites to enzyme reactions with UV-irradiated DNA. While these experiments do not provide definitive evidence that the AP endonuclease and UV DNAincising activity are resident in the same molecule, they indicate that the 40. J. Pierre, and J. Laval, Biochemistry, in press. 41. J. Pierre, and J. Laval, Biochemistry, in press. 42. N. V. Tomilin, E. B. Raveltchuk, and T. V. Mosevitskaya, Eur. J . Biochem. 69, 26s (1976).

262


AP endonuclease activity in question is required for UV DNA-incising activity. The precise nature of the physical relationship between pyrimidine dimer-DNA glycosylase and AP endonuclease activities in the T4 and M. Iuteus enzyme preparations is obviously an important area for further detailed investigation. (See Note Added in Proof, p. 279.) 3 . Endonuclease III of Escherichia c d i

Endonuclease 111 of E. coli is another enzyme activity in which AP endonuclease and DNA glycosylase functions have not been physically separated. This activity was originally described by Radman (43) who was searching for endonucleases involved in the correction of mismatched base pairs in DNA and used heavily UV-irradiated DNA as a substrate. These studies led to the isolation of a DNA incising activity that attacks UV-irradiated, but not unirradiated, DNA. However, in contrast to UV DNA-incising activities specific for pyrimidine dimers in DNA, extensive photoreactivation of UV-irradiated DNA did not eliminate substrate sites value for endonuclease 111. The enzyme activity was shown to have a szo,w of 2.6, with no cofactor requirement. The pH optimum was -7.0 and activity was inhibited in the presence of tRNA or 1.0 M NaCl. Independent investigations by Gates and Linn (44)led to the isolation of a DNA incising activity with properties indistinguishable from those of endonuclease 111. These investigators purified the enzyme approximately 1600-fold, but not to physical homogeneity. The purified enzyme was clearly shown to recognize a photoproduct in DNA other than thyminecontaining pyrimidine dimers. Thus, PM2 DNA irradiated under standard conditions contained -85 thymine-containing pyrimidine dimers and one enzyme-sensitive site per molecule. However, when dimers were introduced into DNA by irradiation at 309 nm in the presence of a photosensitizer (a protocol that results in far fewer nondimer photoproducts in DNA), the resulting substrate contained -95 thymine-containing pyrimidine dimers per PM2 DNA molecule, but was not nicked by preparations of endonuclease 111. Gates and Linn examined the activity of endonuclease I11 on a variety of DNA substrates and demonstrated nicking on heat- and acid-treated DNA and X-irradiated DNA. Both substrates share in common the presence of sites of base loss. Whereas the enzyme was shown to attack duplex DNA that contained only apurinic sites, alkali-stable lesions in UV-irradiated and Os0,-treated DNA were also attacked by the enzyme preparation. A reasonable explanation for this observation stems from 43. M. Radman, JBC 251, 1438 (1976). 44. F. T. Gates, Ill. and S. Linn, JBC 252, 2802 (1977).


263

studies by Linn and his colleagues (45, 46) that demonstrate DNA glycosylase activity in purified preparations of endonuclease I11 that excises, 5,6-dihydrothymine and 5,6-dihydroxydihydrothyminefrom DNA as the free base. These products had previously been shown by Cerutti (47) to be present in UV-irradiated or X-irradiated DNA, as well as in DNA treated with OsO,. Attempts to physically separate the DNA glycosylase and AP endonuclease activities have not as yet been successful. Endonuclease I11 is believed to catalyze incision of duplex DNA 3‘ to sites of base loss. This conclusion is based on experiments similar to those previously described (see Section II,A,l and Fig. 3), i.e., apurinic DNA treated with endonuclease 111 is a poor primer-template for E. coli DNA polymerase I. The DNA becomes a much better substrate for polymerization when either endonuclease IV or endonuclease VI (both of which are believed to be 5’-acting AP endonucleases, see Sections II,B, 1 and II,C, 1) is added to the reaction prior to the polymerization reaction. The interpretation offered is that endonuclease I11 leaves a 3’-depurinated terminus that is not favored by DNA polymerase I as a primer for DNA synthesis. The excision of the 3’-deoxyribose-phosphateresidue by the 5’-AP endonuclease action of endonuclease IV or AP endonuclease function of exonuclease I11 leaves a normal 3‘ terminus that is more effectively utilized by E. coli DNA polymerase I.

B. AP ENDONUCLEASES ASSOCIATED WITH OTHER CATALYTIC ACTIVITIES I . AP Endonuclease Activity of Eronuclease III from Escherichin coli The quantitatively major AP endonuclease activity of E. coli has been referred to in the literature as endonuclease 11, endonuclease VI, or as the AP endonuclease function of exonuclease 111 (see Chapter 12 by B. Weiss, this volume). Endonuclease I1 was discovered by Friedberg and Goldthwait (48-50) and defined as an activity that catalyzes the incision of 45. B. Demple and S. Linn,Nature (London) 287, 203 (1980). 46. S. Linn, B . Demple, D. W. Mosbaugh, H. R. Warner, and W. A. Deutsch, in “Chromosome Damage and Repair” (E. Seeberg and K. Kleppe, eds.), Plenum Press, New York, in press. 47. P. Cerutti, Nuturwissenschuften 61, 51 (1974). 48. E. C. Friedberg and D. A. Goldthwait, CSHSQB 33, 271 (1968). 49. E. C. Friedberg and D. A. Goldthwait, PNAS 62, 934 (1969). 50. E. C. Friedberg, S-M. Hadi, and D. A. Goldthwait, JBC 244, 5879 (1%9).

264

E. FRIEDBERG, T. BONURA, E . RADANY, J. LOVE

DNA alkylated with methyl methanesulfonate. Subsequently, Hadi and Goldthwait (51) demonstrated that the activity also attacked depurinated DNA. Independent studies by Verly and his colleagues (52,53)resulted in the isolation and purification of an AP endonuclease originally thought to be distinct from endonuclease I1 sinceall substrate sites in alkylated DNA attacked by this AP endonuclease were shown to be alkaline-labile, i.e., sites of depurination arising from the spontaneous hydrolysis of alkylated purines. In contrast, endonuclease I1 was shown to attack both alkali-labile and alkali-stable sites in alkylated DNA. The discovery of a specific 3-methyladenine-DNA glycosylase in extracts of E. coli (54) provides an explanation for this confusion. Endonuclease I1 of E. coli as originally defined was presumably a mixture of 3-methyladenine-DNA glycosylase and AP endonuclease activities. However, prior to this and other retrospective clarifications about the identity of endonuclease I1 and the major AP endonuclease of E. coli, Verly designated the latter enzyme as endonuclease VI (55 1. We recommend that the terms endonuclease I1 and endonuclease VI be dropped, because the AP endonuclease activity appears to be clearly associated with the previously described enzyme, exonuclease 111. Milcarek and Weiss (56) and Yajko and Weiss (57) isolated a series of mutants of E. coli defective in both the 3’-exonuclease and the associated 3’-phosphatase functions of exonuclease 111. Of the mutants characterized genetically, all mapped to the xthA locus, the structural gene coding for exonuclease 111, which is situated between the pfkB and pncA loci of E. coli at approximately 38.5 min on the E. coli genetic map. Studies with preparations of exonuclease 111 purified to 98% homogeneity demonstrated the presence of AP endonuclease activity that could not be separated from exonuclease and phosphatase activities by electrophoresis, sedimentation, or gel filtration (58). All xthA mutants were defective in exonuclease I11 activity and were also shown to be defective in AP endonuclease activity (58). Exonuclease I11 as purified by Weiss in terms of its exonucleasephosphatase activity is a monomeric protein of MW -28,000. The enzyme

-

51. S-M. Hadi and D. A . Goldthwait, Biochemistry 10, 4986 (1971). 52. W. G. Verly and Y. Paquette, Con. J . Biochem. 50, 217 (1972). 53. Y. Paquette, P. Crine, and W. G . Verly, Can. J . Biochem. 50, 1199 (1972). 54. S . Riazzudin and T. Lindahl, Biochemistry 17, 21 10 (1978). 55. W. G. Verly,in, “DNA Repair Mechanisms” (P. C. Hanawalt, E. C. Friedberg, and C. F. Fox, eds.), p. 187. Academic Press, New York, 1978. 56. C. Milcarek and B . Weiss, J M B 68, 303 (1972). 57. D. M. Yajko and B . Weiss, PNAS 72, 688 (1975). 58. B. Weiss, JEC 251, 1896 (1976).

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has also been purified about 8000-fold by Verly and Rassart (59) as a monomeric protein of similar molecular weight. The AP endonuclease activity is optimal at pH 8.5 and is maximally stimulated by Mg2+ at 20 mM at pH 8.0 (60). In the absence of M&+, activity is present, but is completely inhibited in the presence of EDTA at concentrations > 1 mM. If the enzyme is dialyzed against 10 mM EDTA to remove divalent cations, and then against buffer without EDTA, no enzyme activity is detected (60). Activity can be restored by the addition of Mg2+,but not by Ca2+,Mn'+, Zn2+,or Cu2+. These observations suggest that the enzyme may be a metallo-protein containing bound Mg'+. A number of different experimental approaches have yielded results consistent with the conclusion that the enzyme catalyzes the hydrolysis of phosphodiester bonds 5' to apurinic sites in duplex DNA, leaving 3'-OH and 5'-Ptermini (60). The AP endonuclease activity of exonuclease I11 of E. coli does not degrade native DNA. It is generally assumed that the enzyme attacks sites of pyrimidine or purine loss in duplex DNA with equal facility, however quantitative comparisons of this parameter have not been documented. The enzyme is at least as active on DNA that contains AP sites reduced with sodium borohydride as on the unreduced substrate, indicating that the aldehyde function of C, in deoxyribose is not required for enzyme action. Gossard and Verly (60) suggested that the mechanism of action of the enzyme is probably distinct from the @elimination reaction catalyzed by alkali at sites of base loss in DNA. This conclusion is supported by the observation that alkaline-catalyzed /3-elimination most frequently results in hydrolysis of phosphodiester bonds on the 3' side of sites of base loss (611, and has an apparent requirement for the aldehyde function at C1 of the deoxyribose residue, since reduction of C, to the alcohol prevents the reaction. Hydrolysis of apurinic DNA 5' to sites of base loss would necessitate excision of the deoxyribose-phosphate moiety in the 5' --* 3' direction during the excision repair in vivo of base damage to DNA. Such a biochemical pathway provides no obvious function for the 3' exonuclease function of exonuclease 111. Gossard and Verly (60) suggested that the exonuclease may remove one or more nucleotides from the 3' end of sites of phosphodiester bond hydrolysis, thus precluding ligation of enzymecatalyzed phosphodiester bond breaks. However such a function for exonuclease 111 has not been directly demonstrated, nor is there evidence 59. W. G. Verly and E. Rassart, JBC 250, 8214 (1975). 60. F. Gossard and W. G . Verly, Eitr. J . Biochem. 82, 321 (1978). 61. C. R. Bayley, K . W. Brarnrner, and A. D. Jones,JCS., 1903 (1961).

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that a 5’-Pterminus associated with an apurinic site can be enzymatically ligated to an adjacent 3’-OH end. An intriguing model reconciling the multiple catalytic activities of exonuclease I11 has been suggested by Weiss (58, and Chapter 12, this volume). He proposes that the enzyme recognizes “spaces” in DNA duplexes that provide substrate sites for each of its 3 catalytic domains: The exonuclease recognizes a “space” created by the unwinding of a terminal base pair due to the lowered base-stacking intcractions operating at the ends of DNA duplexes; the AP endonuclease recognizes the “space” created by loss of a base; and the 3’-phosphatase recognizes the “space” corresponding to a “missing” 3’-terminal nucleoside residue. The exact biological role of the AP endonuclease activity of exonuclease I11 is not clear. Surprisingly, mutants of E. coli (including deletion mutants) defective in exonuclease I11 (xth-) are not particularly sensitive to treatment with alkylating agents such as methyl methanesulfonate (3). This may reflect the ability of other AP endonucleases to assume the essential function(s) of this enzyme during the repair of sites of base loss. Alternatively, repair of such sites may occur by the action of DNA purine insertase activity. In contrast,E. coli mutants defective in bothxth and duf genes are only conditionally viable (62). One possible explanation for this difference is that in the absence of AP endonuclease activity, the apyrimidinic sites created by incorporation of dUMP into DNA, and the subsequent excision of uracil by ura-DNA glycosylase, are lethal lesions. This apparent contradiction suggests that the function(s) that operates in the repair of apurinic sites in the absence of AP endonuclease activity is not able to deal effectively with apyrimidinic sites. Such a conclusion is consistent with the apparent failure to discover DNA pyrimidine insertase activity in any biological source, but is equally consistent with the notion that some AP endonucleases do not attack apurinic and apyrimidinic sites in DNA with equal efficiency. 2. Exonuclease III-Like AP Endonucleuse Activity from Other Organisms Haemophilus injuenzae is another biological source from which an AP endonuclease with associated 3’-exonuclease and 3’-phosphatase activities has been isolated. Clements et al. (63) partially purified such an enzyme with a MW -30,000. Divalent cation is required for maximal 62. B. Weiss, S. G. Rogers, and A. F. Taylor, in “DNA Repair Mechanisms” (P. C. Hanawalt, E. C. Friedberg, and C. F. Fox, eds.), p. 191. Academic Press, New York, 1978. 63. J . E. Clements, S. G . Rogers, and B . Weiss, JBC 253, 2990 (1978).

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267

activity, Mg2+and Mn2+being equally effective at optimal concentrations of 5 mM. Like the AP endonuclease activity ofE. coli, activity is detected in the absence of divalent cation, and the enzyme is totally inactivated in the presence of 1 mM EDTA. AP endonuclease and exonuclease activities cannot be resolved by electrophoresis, sedimentation, or gel filtration. In addition, a mutant of H . injuenzue (corn-lU) known to be defective in exonuclease activity, is also defective in AP endonuclease activity. Like the AP endonuclease activity of exonuclease I11 of E. coli, the AP endonuclease of H. itzfluenzue catalyzes the hydrolysis of phosphodiester bonds 5' to sites of base loss in DNA, creating 3'-OH and 5'-P termini. C. AP ENDONUCLEASES WITHOUTASSOCIATED CATALYTIC ACTIVITIES

I . Etidonuclemse IV qf Escherichicr coli The AP endonuclease activity of exonuclease I11 is quantitatively the major AP endonuclease detectable in crude extracts of wild-type E. coli. In extracts ofxth- mutants defective in exonuclease 111, about 10% of the normal level of AP endonuclease active on duplex AP DNA is detectable, essentially all of which can be accounted for by an activity designated as endonuclease IV (641. Endonuclease IV has been purified about 3000-fold by Ljungquist (641, but this preparation is not physically homogeneous. The enzyme has a relative S value of 3.4 with a Stokes radius of 25 A andf/fo = I . 18. These results are consistent with a globular protein of MW -30,000. The activity has a broad pH optimum between 8.0-8.5 and has no requirement for known cofactors. Activity is sensitive to inhibition by sulfhydryl blocking agents. Normal levels of activity are expressed in the presence of EDTA and there is no detectable stimulation or inhibition of the enzyme activity when EDTA in the standard reaction is replaced by MgCl, (0.5-10 mM) or CaCl, (1.0 mM). Higher concentrations of CaCI, (5.0 mM) are inhibitory. Endonuclease IV activity is unusually resistant to NaCl and retains 50% maximal activity in 0.56 M salt. The enzyme is also quite heat resistant: heating to 60" for 5 minutes in the presence of 0.2 M NaCl results in no detectable loss of activity. Activity is not affected by tRNA at concentrations between 20-500 pglml. Endonuclease IV does not attack covalently closed circular PM2 DNA but does catalyze the stoichiometric nicking of such DNA when it con64. S. Ljungquist, JBC 252, 2808 (1977).

268


tains apurinic sites. The enzyme also degrades heavily UV-irradiated or X-irradiated DNA, however all substrate sites in such DNA are satisfactorily accounted for as sites of base loss. The polarity of DNA incision relative to sites of base loss, and the chemistry of the termini created, have not been reported. However, Linn and his colleagues (26, 46) have observed that the addition of endonuclease IV to reactions of apurinic DNA with endonuclease I11 enhances the priming activity of the product for DNA synthesis by E. coli DNA polymerase I (see Section II,A,3). This result suggests that like the AP endonuclease activity of exonuclease 111, endonuclease IV catalyzes hydrolysis of phosphodiester bonds 5’ to sites of base loss in DNA. 2 . AP Eiidotiuclecises of Micrococcus luteus Two AP endonucleases have been extensively purified and characterized from M . luteus and are referred to as AP endonucleases A and B (4U, 4 1 ) . As judged by polyacrylamide gel electrophoresis, each is a monomeric protein of MW -35,000 and neither protein has detectable nonspecific endonuclease, exonuclease, phosphatase, 3-methyladenine-DNA glycosylase, or uracil-DNA glycosylase activity. The two AP endonuclease activities are distinguishable by a different affinity for phosphocellulose and by markedly different isoelectric points (4.8 and 8.8 for enzymes A and B, respectively). In addition, AP endonuclease A has a half-life of 4 minutes at 45”, whereas AP endonuclease B has a half-life of 10 minutes at the same temperature. Activity A has a narrow pH optimum around 7.5, whereas activity B has a broad pH optimum between 6.5-8.0. Neither requires divalent cations for activity, although both are stimulated by Mg’+, are inhibited by Ca2+ or Mn”, and are inhibited 90% in the presence of 0.4 mM EDTA. A number of distinct experimental approaches indicate that both activities catalyze the hydrolysis of phosphodiester bonds 5’ to sites of base loss in duplex DNA (however see discussion in Section 11,A,2), leaving 3’-OH and 5’-P termini. Whether or not AP endonucleases A and B represent independently coded gene products is an important question that remains to be resolved. Conceivably one is a posttranslationally modified form of the other, or arises as an artifact of proteolysis in cell-free preparations of M. luteus. It would be of distinct interest to examine the antigenic relatedness of these two enzymes. AP endonuclease activities with properties similar to those of endonuclease IV ofE. coli have been purified and characterized to varying degrees from a number of other biological sources, including calf thymus, calf liver, plants, mouse epidermal cells, S . cerevisiae, B. subtilis, B.


269

stenrothermophilits, and human placenta. A summary of the essential features of these enzymes (65-77) is presented in Table I.

3. Etidotzucleiise VII of Escherichicr coli

The AP endonucleases thus far discussed have been characterized in terms of their activity on duplex DNA that contains sites of base loss. Studies in this laboratory have led to the identification of an AP endonuclease that catalyzes the degradation of single-stranded polydeoxyribonucleotides or DNA that contains depyrimidinated sites, but not of intact single-stranded polymers or DNA. This activity is present in extracts of both wild type and xth- strains of E. coli and has been designated as endonuclease VII of E. coli (78-79~). The standard assay of endonuclease VII activity measures the degradation to acid-soluble products of poly(dU) . [3H](dT)200that contain depyrimidinated sites produced by the excision of uracil from the polymer by preincubation with purified ura-DNA glycosylase. The detection of endonuclease VII in crude extracts of wild-type E. coli by this assay is complicated by the presence of two contaminating activities. One is due to low molecular weight basic proteins and polyamines that are known to promote p-elimination reactions at sites of base loss (3) and result in the selective degradation of the depyrimidinated polymer. Unlike endonuclease VII, however, these substances do not bind to DEAE-cellulose and are readily separated from the enzyme by column chromatography. The 65. L. Thibodeau and W. G. Verly, FEBS (Fed. Eur. Biochem. Soc.) Lett. 69, 1 (1976). 66. L. Thibodeau and W. G. Verly, JBC 252, 3304 (1977). 67. V. Bibor and W. G . Verly, JBC 253, 850 (1977). 68. T. Inoue and T. Kada,JBC 253, 8559 (1978). 69. P. R. Armel and S. S. Wallace, Nucleic Acids Res. 9, 3347 (1978). 70. A. B . Futcher and A. R. Morgan, Can. J. Biocllem. 57, 932 (1979). 71. S . Ljungquist and T. Lindahl, JBC 249, 1530 (1974). 72. S. Ljungquist, B. Nyberg, and T. Lindahl,FEBS (Fed. Eur. Biochem. S o c . ) L e t t . 57, 169 ( 1975). 73. J. P. Kuebler and D. A. Goldthwait, Biochemistry 16, 1370 (1977). 74. G. Ludwig and H. W. Thielmann, Nucleic Acids Res. 6, 2901 (1979). 75. T. P. Brent, Niicleic Acids Res. 4, 2445 (1977). 76. U . Kuhnlein, E. E. Penhoet, and S. Linn, P N A S 73, 1169 (1976). 77. W. S. Linsley, E. E. Penhoet, and S. Linn,JBC 252, 1235 (1977). 78. E. C. Friedberg, T. Bonura, R. Cone, R. Simmons, and C. Anderson, in "DNA Repair Mechanisms" (P. C. Hanawalt, E. C. Friedberg, and C. F. Fox, eds.), p. 163. Academic Press, New York, 1978. 79. E. C . Friedberg, C. T. M. Anderson, T. Bonura, R. Cone, E. H. Radany, and R. J. Reynolds, Progr. Nucleic Acid Res. M o l . Biol., 26, 197 (1981). 79a. T. Bonura, R. Schultz and E. C. Friedberg, in preparation.

TABLE I

AP ENDONUCLEASES FROM A VARIETY OF BIOLOGICAL SOURCES

Source

Molecular weight

Cofactor requirement

Plmsoeolirs rnti(tifl0ru.r

40,000

None

R . stearothertno~kilrrs

28.000

None

56,000

None

Stimulation or inhibition of activity Mg2+or Mn2+stimulate about Cfold; inhibited in presence of NaCl Stimulated by monovalent cations: inhibited by divalent cations Stimulated by 50 mM NaCl, and by Mg2+,Mn2+or Ca2+:inhibited in presence of 500 mM NaCl

General comments

Reference

Temperature optimum at 60" -

S. cerei7isine

(Endonuclease D) (Endonuclease E)

MgZ+ None None

S . rereifsine

Calf thymus Calf liver Mouse epidermis

35,000 28 ,000 3 1 .OW

Human lymphocytes Human skin fibroblasts

Human placenta

Mgz+ Mg2+ None MgZ+

27,000-31,000

None

Inhibited in presence of 400 mM NaCl Inhibited by 1 mM EDTA Inhibited by Ca2+,EDTA, or tRNA Stimulated by Mg2+,or KCI: inhibited by high ionic strength Inhibited by EDTA

Stimulated by Mg2+, Mn2+,Coz+, Zn2+ or 40 mM KCl: inhibited by CaZ+or EDTA

Two activities identified in extracts Temperature optimum at 40" Optimal pH at 9.5 -

Activity in XP group A and D cells has 5 to 10-fold higher K , than normal Six chromatographically separable forms identified

(76)

(77)

14.


27 1

second contaminating activity is due to exonuclease VII in extracts of E. coli. This enzyme catalyzes the exonucleolytic degradation of both the intact and the depyrimidinated polymer in the presence of EDTA (80). Exonuclease VII also binds to DEAE-cellulose, but is totally separable from endonuclease VII by successive chromatography on DEAEcellulose and phosphocellulose. Endonuclease VII has been purified free of nonspecific endonuclease and has a relative S value of 4.2 measured by sedimentation in 15-40% glycerol gradients in the presence or absence of 0.25 M KCI.A molecular weight of -55,000 has been calculated from gel filtration of the enzyme, relative to appropriate standard markers. Activity is unaffected by the presence of 5 mM EDTA in the standard reaction. In the absence of EDTA, activity is slightly stimulated by 5 mM MgC1, and by 1-2 mM CaCI,. Endonuclease VII activity is inhibited 50% in the presence of 0.15 M NaCl or in the presence of 0.58 nM (as nucleotide) tRNA. Endonuclease VII can be differentiated from previously described AP endonucleases of E. coli by a number of criteria (Table 11). Note that the only other enzymes active in the presence of EDTA are endonucleases I11 and IV. Both have been directly examined for activity against depyrimidiand found to be inactive under conditions that nated poly(dU) . [3H](dT)200 promote readily detectable degradation of depurinated duplex PM2 DNA. The substrate specificity of endonuclease VII is uncertain. In addition to the single-stranded depyrimidinated heteropolymer mentioned above, the enzyme degrades depyrimidinated poly(dU) and single-strandeddepyrimidinated PBS2 DNA, as well as single-stranded circular M 13 DNA that contains depyrimidinated sites. (The latter substrate is produced by growing phage M 13 in a strain of E. coli defective in dUTPase and in uracil-DNA glycosylase activities. The resulting phage DNA contains dUMP residues that can be converted to depyrimidinated sites by preincubation with uracil-DNA glycosylase .) Duplex form I PM2 DNA that contains apurinic sites (created by beat and acid treatment of the DNA) is not attacked by the enzyme, nor is duplex PBS2 DNA containing depyrimidinated sites a substrate for endonuclease VII. Depyrimidinated sites in single-stranded DNA that have been reduced with sodium borohydride are also not attacked by the enzyme. The reduction of the aldehyde function of the deoxyribose moiety to the alcohol stabilizes the associated phosphodiester bond against p elimination. Thus it is possible that endonuclease VII catalyzes phosphodiester bond hydrolysis by a p elimination mechanism. Single-stranded DNA substrates containing apurinic sites have not yet been examined with endonuclease VII. 80. J . W. Chase and C. C. Richardson, JBC 249, 4545 ( 1974).

TABLE I1

APURINIC/hYRMIDINIC (AP) ENDONUCLEASES OF Escherichio coli Enzyme Endonuclease 111

Endonuclease IV

Endonuclease V'

AP Endonuclease of Exonuclease 111 Endonuclease VI1

Principal properties No requirement for divalent cation: inhibited in presence of tRNA; 2.7 S: MW -27,000; pH optimum -7.0; requires duplex DNA with AP sites No requirement for divalent cation; not stimulated by Mg2+ or Ca2+:no inhibition in presence of tRNA: 3.4 S: MW -33,000; pH optimum -8.0-8.5; inhibited in presence of PCMB; requires duplex DNA with AF' sites Requires Mg2+for activity: inhibited in presence of tRNA: 2.3 S: MW -20,000: pH optimum 9.25; requires duplex DNA with AP sites Requires Mg2+for optimal activity; inhibited in presence of EDTA; 2.9 S ; MW -32,000 pH optimum -8.5 ; requires duplex DNA with AP sites No requirement for divalent cation; stimulated by Mg2+or Caz+;inhibited in presence of tRNA; 4 . 3 S ; MW -45,000: pH optimum -7.0; insensitive to PCMB; attacks single-stranded DNA and polydeoxypyrimidines with depyrimidinated sites; duplex DNA with apurinic sites is not a substrate

Associated catalytic activities

Reference

DNA glycosylase activity that recognizes 5,bsaturated thymine photoproducts None detected

(43,4446)

Acts on a variety of damaged DNAs as well as certain native DNAs 3', 5'-exonuclease (exonuclease HI), 3'-phosphatase, RNase H Not determined

(48-53 ) ( 5 5 4 U ,6 2 ) (78-790)

" Endonuclease V is not considered in the text as an AP endonuclease, but is included in the table for the sake of completeness since it has been reported to attack DNA heated at pH 5.2. From Gates and Linn (99).


273

D. CONCLUDING COMMENTS ON AP ENDONUCLEASES AP endonucleases, discovered in the late 1960s, were originally thought to constitute a specific class of endonucleases required for the repair of sites of base loss that arose spontaneously in either native DNA, or in DNA treated with chemical agents (such as certain alkylating agents) that promote increased lability of N-glycosylic bonds. The discovery of DNA glycosylases as a general class of enzymes involved in the excision of damaged or inappropriate bases (8/) provided a more obvious role for AP endonucleases in DNA repair. The findings reviewed above, that some AP endonuclease activities may be physically associated with specific DNA glycosylase activities, suggests that this class of enzymes may be fundamental to understanding the enzymology of the incision of a major fraction (if not all) of DNA damage in living cells. Thus for example, it is possibly significant that endonuclease I11 (which apparently has an associated DNA glycosylase activity specific for monoadduct thymine photoproducts with saturated 5,6 bonds) and the AP endonucleases associated with the pyrimidine dimer-DNA glycosylases of phage T4 and M. luteus all catalyze the hydrolysis of phosphodiester bonds 3’ to sites of base loss. On the other hand endonuclease IV and the AP endonuclease activity of exonuclease 111, both of which are thought to be 5’-acting AP endonucleases, are apparently not associated with DNA glycosylase activities. Conceivably 3’-acting AP endonucleases function exclusively in concert with specific DNA glycosylases, while the 5’-AP endonucleases are more general in their action. Clearly it is necessary to establish more information on the polarity of DNA incision by all known AP endonucleases and to survey them thoroughly for associated DNA glycosylase activity. It also remains to be satisfactorily demonstrated whether or not all AP endonucleases attack sites of purine and pyrimidine loss with equal facility. This potential basis for substrate specificity may provide further understanding of the biological relevance of the multiplicity of AP endonucleases detected in different organisms. In this regard it should be noted that with the exception of the AP endonuclease activity of exonuclease 111, no mutants of E. co/i have been isolated that are defective in AP endonucleases. Such strains would be of enormous value in understanding the biological role of the various AP endonucleases in this organism. Finally, in view of the apparent central role that AP endonucleases play in the excision repair of DNA, the importance of making the distinction between a direct-acting endonuclease and an AP endonuclease that acts subsequent to a physically associated or distinct DNA glycosylase, is 81. T. Lindahl, Nuture (London) 259, 64 (1976).

274


obvious. All newly discovered endonuclease activities that attack modified or damaged DNA should be very carefully investigated from this point of view.

111.

Enzymes That Attack Phosphodiester Bonds in Damaged DNA with Intact N-Glycosylic Bonds

OF Escherichiu coli A. UV ENDONUCLEASE

The phenomenon of pyrimidine dimer excision during post-UV incubation of irradiated E. coli was reported as early as 1964 (82, 83). The isolation of a series of UV-sensitive mutants defective in pyrimidine dimer excision established the involvement of at least 3 genes (uvrA, -B, and -C) in the incision of UV-irradiated DNA (I). However the isolation of the products of these genes and the direct demonstration of UV DNA-incising activity in cell-free preparations was achieved much later. Braun and Grossman (84)reported the partial purification of an endonuclease activity that selectively attacked UV-irradiated and not unirradiated DNA. This activity also has the property of binding to UVirradiated DNA that contains pyrimidine dimers . When irradiated DNA was subjected to monomerization of pyrimidine dimers by incubation with yeast photoreactivating enzyme the binding activity was lost. The activity was not detected in extracts of uvrA or uvrB mutants of E. coli, but was present at normal levels in extracts of uvrC mutants. The molecular weight of this activity was estimated at - 14,000. Subsequent studies by Seeberg and his colleagues yielded different results. These investigators prepared lysates of E. coli using lysozyme lysis of sucrose-permeabilized cells (8.5).In the absence of added ATP, extracts of wild-type and of uvrA, -B, and -C mutants showed similar nicking of either unirradiated or UV-irradiated supercoiled ColE 1 DNA. However, in the presence of 1.5 mM ATP, a fivefold increase in activity that nicked UV-irradiated DNA was observed only with extracts of wild-type strains. The addition of an extract of any one of the three uvr mutants to that prepared from either of the other two resulted in the restoration of ATPdependent nicking of UV-irradiated DNA. This observation provided the 82. 83. 84. 85.

R. B. Setlow and W. L. Carrier, PNAS 51, 226 (1964). R. P. Boyce and P. Howard-Flanders, PNAS 51, 293 (1964). A. Braun and L. Grossman, PNAS 71, 1838 (1974). E. Seeberg, J. Nissen-Meyer, and P. Strike, Nature (London) 263. 524 (1976).


275

basis for a complementation assay that has been extremely useful for the purification of the uvrA, -B, and -C gene products (86,87). The uvrB and -C gene products cochromatograph on DEAE-cellulose. The uvrA gene product does not bind to DEAE-cellulose but does bind to phosphocellulose, eluting at about 250 m M KCI. Neither the isolated uvrA gene product, nor the combined uvrB and -C gene products alone show appreciable U V endonuclease activity under standard assay conditions. However, when these fractions are combined, an ATP-dependent UV DNA-incising activity is demonstrated. This activity recognizes pyrimidine dimer sites in DNA, since substrate sites are lost when UVirradiated duplex DNA is pretreated with photoreactivating enzyme. Gel filtration measurements yielded a MW 100,000 for the uvrA gene product and a value of -70,000 for the uvrB gene product (86). The uvrA, -B, and -C genes have been cloned into bacterial plasmid vectors in a number of laboratories (87-91). Sancar et al. (88-90) developed a so-called “maxicell” procedure for the specific radiolabeling of proteins coded by infecting plasmids. Applying this procedure to cells infected with plasmids carrying either the uvrA or uvrB genes, they demonstrated monomeric gene products of MW = 114,000 and 84,000, respectively, by pol yacrylamide gel electrophoresis. Van Sluis et al. (personal communication) measured a MW = 27,000 for the product of the cloned uvrC gene expressed in minicells. The uvrA protein has been purified to apparent homogeneity and has been reported to bind to unirradiated single-stranded DNA, UV-irradiated single-stranded DNA, and UV-irradiated duplex DNA (87, 92, W).It is likely that in the latter substrate the protein binds to locally denatured regions associated with pyrimidine dimers. DNA binding activity requires the presence of Mg2+and is stimulated by either ATP or GTP. In contrast, the ATP dependence for UV DNA-incising activity by the uvrABC gene product complex cannot be replaced by GTP, ADP, or AMP (87, 92, 94).

-

86. E. Seeberg, PNAS 75, 2569 (1978). 87. E. Seeberg, in “DNA Repair Mechanisms” (P. C. Hanawalt, E. C. Friedberg, and C. F. Fox, eds.), p. 225. Academic Press, New York, 1978. 88. A. Sancar, R . P. Wharton, S. Seltzer, B . M. Kacinski, N . D. Clarke, and W. D. Rupp, J . Ma/. B i d . 148, 45 (1981). 89. A. Sancar, N . D. Clarke, J . Griswotd, W. J . Kennedy, and W. D. Rupp,J. Mol. B i d . 148, 63 (1981). 90. A. Sancar and W. D. Rupp, BBRC 90, 123 (1979). 91. H. Pannekoek, I. A. Noordermeer, C. A. Van Sluis, and P. van de Putte,J. Bncferiol. 133, 884 (1978). 92. E. Seeberg and A-L. Steinum, in “Chromosome Damage and Repair” (E. Seeberg and K. Kleppe, eds.). Plenum, New York, in press.

276

E. FRIEDBERG, T. BONURA, E. RADANY, J . LOVE

Rupp and his colleagues (9-?)reported that the rrvrA protein binds to both unirradiated single- and double-stranded DNA and that UV-irradiation increases the binding affinity of the protein for both substrates, particularly for duplex DNA. These investigators also reported that 10 mM ATP increases binding affinity for unirradiated single-stranded, but not double-stranded, DNA. The MWAprotein also catalyzes the hydrolysis of ATP in the absence of DNA (92, 94). The K , value for this reaction corresponds to that measured for the ATP requirement for endonuclease activity by the complex, suggesting that the ATPase activity may reflect the ATP requirement for nicking. The purified uvrB protein does not have any demonstrable DNA binding or ATPase activity. Very little is known about the properties of the MVVCprotein or of the composition of the IIWABCcomplex in wild-type cells. The irvrABC gene product complex is discussed in Section 111 of this chapter because there is no direct evidence that this complex functions by a DNA glycosylase/AP endonuclease mechanism, as has been demonstrated for the M. luteirs and T4 UV DNA-incising activities. However, the evidence indicating that the complex functions as a direct-acting endonuclease is still limited and indirect. The standard assay for activity on UV-irradiated DNA measures the hydrolysis of phosphodiester bonds without the use of conditions that promote p-elimination of apyrirnidinic sites in the DNA. However, this observation is not inconsistent with a two-step mechanism of DNA incision if the complex contains both PD-DNA glycosylase and AP endonuclease activities. The uvrABC protein complex has also been shown to attack DNA that contains substrate sites other than pyrimidine dimers, e.g., in DNA treated with 8-methoxypsoralen plus 360 nm light, both sites of psoralen monoadduct and of psoralen interstrand crosslink formation are attacked by the enzyme activity (94).This supports the results of in vivo studies that have shown uvrABC mutants of E. coli to be abnormally sensitive to treatment with psoralen plus long wavelength UV radiation (9.5). Such mutants are additionally sensitive to Cnitroquinoline- I-oxide, nitrogen mustard, and mitomycin C (see Ref. 1 ) suggesting that the rrvrABC complex recognizes conformational distortions in the DNA duplex rather than specific forms of base damage to DNA. Such a broad substrate specificity is more consistent with the expression of a direct-acting endonuclease than with a DNA 93. w. D. Rupp, A. Sancar, B. Kacinski, R . Wharton, S. Seltzer, J . Griswold, and N . Clarke, in “Chromosome Damage and Repair” (E. Seeberg and K. Kleppe, eds.), Plenum, New York, in press. 94. E. Seeberg, Prog. Ncic. Acids Res. Mol. Biol., in press. 95. R . S . Cole, D. Leortan, and R. R. Sinden, J M R 103, 39 (1976).


277

glycosylase activity. However, the molecular mechanism of incision of DNA that contains the various forms of damage previously mentioned need not be the same in each case. Thus it remains to be definitely established that the incision of DNA that contains pyrimidine dimers, for example, does not uniquely occur by a DNA glycosylase/AP endonuclease mechanism.

B.

uv ENDONUCLEASE ACTIVITY IN OTHERORGANISMS

There have been numerous reports in the literature of eukaryote DNAincising activity that preferentially attacks UV-irradiated relative to unirradiated DNA, however few of these activities have been purified or extensively characterized (see Ref. %). Van Lancker and Tomura (97) reported the purification to apparent homogeneity of an enzyme from rat liver with an estimated MW 15,000. Sedimentation velocity analyses in neutral and alkaline sucrose density gradients indicate that the purified enzyme catalyzes the formation of single-strand breaks in duplex UVirradiated or acetylamino fluorene-treated DNA. The enzyme is active without added MgC12, but is optimally stimulated at 5-10 mM MgC12. Waldstein et d.(98) reported a %fold purification of a UV DNA-incising activity from calf thymus that apparently recognizes pyrimidine dimers specifically. Maximum activity of the enzyme is observed between pH 7.5-8.0, and 5-15 mM MgC1, is optimal for activity. No activity is observed in the presence of 10 mM EDTA. The activity is extraordinarily labile, which has precluded significant purification thus far. However, preliminary gel filtration studies suggest that the activity is associated with a protein of high molecular weight.

-

C. ENDONUCLEASE V OF Escherichici coli

Endonuclease V is a protein that has been purified -6000-fold by Gates and Linn (99) on the basis of its ability to cleave single-stranded fd DNA. The enzyme also acts on either Form I or Form I1 duplex DNA at a rate about 10% of that on single-stranded DNA. However, duplex DNA exposed to UV radiation, heating at pH 5.2, partial denaturation and renaturation, or treatment with OsO, is degraded several times faster than is 96. E. C. Friedberg, K . H . Cook, J. Duncan, and K . Mortelman,Photochem. Phorobiol. Rev. 2, 263 ( 1977).

97. J. L. Van Lancker and T. Tomura, BBA 353, 101 (1974). 98. E. A. Waldstein, S. Peller, and R. B . Setlow, PNAS 76, 3746 (1979). 99. F. T. Gates, Ill, and S. Linn, JEC 252, 1647 (1977).

278


native DNA. A substrate particularly favored by the activity is duplex PBS2 DNA (which contains uracil instead of thymine). Mg2+is required for activity on fd, PM2, UV-irradiated PM2, or PBS2 DNA. With PBS2 or fd DNA as substrate, the enzyme has a sharp pH optimum at 9.25. When PBS2 DNA labeled with both [32P]-and [3H]uridine was incubated with endonuclease V, the acid-soluble products of degradation were shown to be oligonucleotides. No free uracil or dUMP were detected. The substrate specificity and mechanism of action of endonuclease V are unclear. Gates and Linn (99) suggested that the enzyme might recognize regions of DNA with abnormal secondary structure. Based on this premise these authors argued that at the pH optimum of 9.2-9.5, transient denatured regions may form in duplex DNA by titration of uracil or thymine residues, with consequent disruption of A-U or A-T base pairs, respectively. Surprisingly however, the activity of endonuclease V on heat denatured PBS2 DNA is less than that on the duplex form, and the pH optimum with fd DNA (which is naturally single-stranded) is also about 9.5. AT D. ENDONUCLEASE ACTIVITY DIRECTED PHOTOALKYLATED PURINESIN DNA

Livnehet al. (100) have reported the identification of an enzyme activity that recognizes a specific substrate produced by photoalkylation of DNA. Reaction of Form I PM2 DNA with isopropylalcohol in the presence of a free radical photoinitiator plus UV light at 300 nm, leads to the specific substitution of purine moieties in the DNA, yielding 842hydroxy-2-propy1)adenineand 8-(2-hydroxy-2-propyl)guanine.These investigators identified an activity in extracts of M. luteus that degrades this DNA. The activity is not completed by the simultaneous presence of single-stranded undamaged DNA or single-stranded UV- or X-irradiated DNA. However, the inclusion of duplex UV- or X-irradiated DNA results in inhibition of activity. Divalent cations are not required for the activity, which is fully active in 1 mM EDTA. ATP (0.1 mM) results in about 40% inhibition, and 0.1 mM caffeine about 30% inhibition of activity. No release of free modified purines is detected during incubation of photoalkylated DNA with M. luteus extract. In addition, when substrate is present at saturating concentrations, the addition of a tenfold excess of depurinated duplex DNA does not inhibit enzyme activity. Both of these observations argue against a DNA glycosylase/AP endonuclease mechanism of incision of the DNA. The authors thus conclude that this endonu100. Z. Livneh, D. Elad, and J. Sperling, PNAS 76, 5500 (1979).


279

clease directly attacks phosphodiester bonds in photoalkylated DNA. In speculating about the nature of the substrate sites recognized by the endonuclease, the authors point out that 8-substitution of purines by a group as small as bromine results in destabilization of the normal anti conformation of the affected nucleotide. Physical studies on such substitutions have shown that a stable configuration requires rotation around the N-glycosylic bond to assume the syn conformation. It is possible that the syn conformation assumed by any 8-substituted purines in DNA provides the molecular basis for the specific substrate recognized by this class of endonucleases. If so, one would assume that the purified enzyme might also attack DNA that contains N-acetoxy-2-acetylaminofluorene adducts, since this compound has also been shown to bind at the %position of guanine in DNA and to promote the conformation change previously discussed (101). ACKNOWLEDGMENTS Studies in the senior author’s laboratory were supported by research grants from the U.S. Public Health Service (USPHS) (CA-12428), the American Cancer Society (NP 174), and The Foundation-March of Dimes (1-6721, as well as by contract EY-76-S-03-0326 with the U S Department of Energy. E. H. Radany is a predoctoral fellow supported by USPHS training grant GM-07364, and J. D. Love is a USPHS postdoctoral fellow supported by grant CA-06441. We thank numerous colleagues for providing preprints of their work cited in this paper.

Note Added in Proof

Very recent investigations in this laboratory have provided genetic and biochemical evidence for the physical association of pyrimidine dimer-DNA glycosylase and AP endonuclease activities on the polypeptide encoded by the d m V gene of phage T4. [S. L. H. McMillan, E. H. Radany, and E. C. Friedberg, Fed. Proc. 40, 1763 (1981); S. McMillan, H. J . Edenberg, E . H. Radany, R. C. Friedberg, and E. C. Friedberg, J. Virol., in press.]

101. D. Grunberger and I. B. Weinstein,in “Biology of Radiation Carcinogenesis” (J. M. Yuhas, R. W. Tennant and J. Regan, eds.), p. 175. Raseen, New York, 1976.


Pancreatic DNase STANFORD MOORE

I. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ill. Chemical Structure . . . . . . . . . . . . . A . Sequence, . . . . . . . . . . . . . . . B . Essentiality of Specific Residues . . . . IV. Catalytic Properties . . . . . . . . . . . . . A . Roles of Divalent Metal Ions . . . . . . B . Substrate Specificity . . . . . . . . . . . V. Actin as an Inhibitor of DNase I . . . . . . VI. Research Applications . . . . . . . . . . . 11. Purification

1.

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

. . . . . . . . .

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

. . . . . . . . . .

281 282 285 285 286 288 288 290 293 295

Introduction

Bovine pancreatic deoxyribonuclease is the most thoroughly studied of the enzymes of the DNase I class (EC 3.1.21. l), defined as enzymes that cleave the substrate endonucleolytically to yield primarily S-phosphodiand oligonucleotide end products. In this chapter, the term DNase will refer to the bovine enzyme, unless otherwise specified. Bovine pancreatic DNase I can be resolved into four components of similar catalytic activity, DNases A, B, C, and D. For most purposes the mixture of the four is a suitable catalyst, subject to the degree of freedom from contaminating proteases or ribonucleases : when one of the subfractions has been studied, the letter designation will be added. The information on DNase to 1970 has been summarized by Laskowski 28 1 THE ENZYMES, Vol. XIV Copyright 0 1981 by Academic Press, Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

282

STANFORD MOORE

(I) in Volume IV of this series. This chapter will cover primarily the information that has been gathered in the subsequent decade. Comparative studies on pancreatic DNases from different species have shown that the ovine and human enzymes are closely similar to the bovine catalyst, as cited in the following section on purification. The characterization of DNases from other tissues has included the finding that the DNase I secreted by the bovine parotid gland ( 2 , 3 )is very similar but not identical, chemically (4), to the bovine pancreatic enzyme. The results provide one of the several examples of enzymes built to the same basic design via different genes in different tissues of the same species. Moving further afield in terms of biological source, the DNase in germinating barley that Brawerman and Chargaff (5) showed had some enzymatic properties in common with bovine pancreatic DNase, has been reinvestigated by ‘iao (6). By applying current techniques, he has succeeded in purifying and characterizing the enzyme from malt diastase and finds it to be remarkably homologous in all of its chemical and enzymatic properties with the mammalian enzyme. Thus, the detailed picture of the chemical structure and catalytic properties of bovine pancreatic deoxyribonuclease summarized in the following pages applies to a broader spectrum of enzymes than was envisaged at the outset of the research.

II. Purification

The starting material for the chromatographic purification of pancreatic DNase has usually been the amorphous enzyme prepared from bovine pancreas by ammonium sulfate fractionation according to the procedure of Kunitz (7). In an extension of the chromatographic experiments of Price et al. (8) on sulfoethyl-Sephadex, Salnikow et al. (9) observed four active components by chromatography on phosphocellulose (Fig. 1). Pancreatic juice submitted to initial chromatography on DEAE-cellulose gave a simi1. M . Laskowski, Sr., “The Enzymes,” 3rd. Ed., Vol. 4, p. 289, 1971. 2. W. D. Bail and W. J. Rutter, J . Exp. Z o d . 178, I (1971). 3. W. D. Ball, BBA 341, 305 (1974). 4. R. L. Lundblad, S . Hoffman, C. M. Noyes, and H. S. Kingdon,J. Dent. Res. 56, 320 (1977). 5. G . Brawerman and E. Chargaff, JBC 210, 445 (1954). 6. T.-H. Liao, Phytochernistry 16, 1469 (1977). 7. M. Kunitz, J . Gen. Physiof. 33, 349 (1950). 8 . P. A. Price, T.-Y. Liu, W. H. Stein, and S. Moore, JBC 244, 917 (1%9). 9. J. Salnikow, S. Moore, and W. H. Stein, JBC 245, 5685 (1970).

15.

283

PANCREATIC DNase

1

1

’M f”

-

> .-

u

a 0

300

Effluent

600

900 ...

(ml)

FIG. 1. Chromatography of bovine pancreatic DNase (Worthington DP grade) on phosphocellulose. Column, 2 x 75 cm: temperature, 25”; flow rate, 35 ml per hour: 4-mI fractions were collected: column equilibrated with 0.25 M sodium acetate at pH 4.7: initial eluent, 150 ml of 0.38 M sodium acetate buffer, pH 4.7; linear gradient with 400 ml each of the initial and the limit buffer, 0.7 M sodium acetate, pH 4.7: (---) absorbance at 280 nm: (0-0) enzymatic activity. From Salnikow ei nl. (Y), reproduced with permission. lar pattern of DNases on phosphocellulose. The major enzyme, DNase A, was the protein taken for detailed sequence analysis; it is a glycoprotein with a neutral carbohydrate side chain. The protein moiety of DNase B is indistinguishable from that of DNase A; the carbohydrate side chain contains sialic acid. From amino acid analyses, DNase C was characterized as being the same as A except for a proline residue substituted for a histidine. This conclusion was confirmed by peptide maps (10) interpreted in the light of the sequence studies. Liao ( 1 1 ) has shown that DNase D has the same sequence as C but contains sialic acid in the carbohydrate portion. In order to obtain the small amount of DNase D in a stable form ( / I ) , the diisopropyl fluorophosphate-treated protein fraction was rechromatographed on DEAE-cellulose with use of a CaC1, gradient, thus combining the experience of Hugli (f2)on the chromatographic removal of traces of chymotrypsin and chymotrypsinogen B and of Price et nl. (8) on the stabilization of DNase against proteolysis by Ca2+. Liao ( I / ) has summarized the differences in the four enzymes (Table I). In Fig. 1 it can be noted that the activity-to-protein ratio may be slightly lower for the sialylated DNases B and D than for A and C; this observation is borne out by the specific activities in Table I. When DNase I is used as a specific biochemical reagent in the preparation of nuclear ribonucleic acids or nuclear proteins, the purification prob10. J. Salnikow and D. Murphy, JBC 248, 1499 (1973). 11. T.-H. Liao, JBC 249, 2354 (1974). 12. T. E. Hugli, JBC 248, 1712 (1973).

284

STANFORD MOORE TABLE 1

ANALYSESOF DNASESA, B ,

c, A N D D“

Analysis for

DNase A”

DNase B”

DNase C”

DNase Dh

Mannose Galactose N - Acet ylglucosamine Sialic acid Proline Histidine Specific activity (unitsimg)

5.8 0.0 I .9 0.0 9.0 6.2 1158

4.5 I .O 3.4 1 .O 9.2 5.7 92 1

4.7 0.0 1.9 0.0 10.1 4.9 I045

4.3 I .O 3.3 0.8 9.9 5.1 837

“ From Liao ( / I ) ,

reproduced with permission.

’ Constituents expressed as residues per molecule

lem is a special one. The separation of DNases A , B, C, and D from one another is not necessary, but freedom from traces of ribonucleases or proteases is essential. Wang and Moore (13) have described a procedure that effects complete removal of trypsin, chymotrypsin, and chymotrypsinogen by a combination of affinity chromatography and salting-out adsorption on lima bean protease inhibitor coupled to Sepharose, an extension of the method of Otsuka and Price (14). DNase is extremely sensitive to inactivation by proteases in the absence of Ca2+; the protease-free preparation retains full stability in the absence of Ca’+ for more than 10 days at pH 8 and 37”. Removal of the last traces of RNase has been accomplished (7) by affinitychromatography on a long (72 cm) column of 5-(4-aminophenylphosphoryl)uridine2’(3’)-phosphate-Sepharose (15, 16). The fully active product, obtained in quantitative yield, has less than I part of RNase per 10 million parts of DNase. Wadano et (11. (171, in the course of isolating DNases from ovine pancreas, have found chromatography on conconavalin A-agarose to be an effectivestep in the purification of DNases with neutral carbohydrate side chains: their method of isolation of DNase, which includes the use of phenylmethanesulfonyl chloride (18) as a protease inhibitor and chromatography on CM-cellulose with Ca2+-containingbuffers, offers the possibility of preparing bovine DNases A and C in higher yield than that obtained in the initial steps of the Kunitz (7) procedure. They also found 13. 14. 15. 16. 17. 18.

D. Wang and S. Moore, JBC 253, 7216 (1978). A. S. Otsuka and P. A. Price, A I I ~Biochem. . 62, 180 (1974). M . Wilchek and M . Gorecki, “Methods in Enzymology,” Vol. 34, p. 492, 1974. 0 . Brison and P. Chambon, Anti/. Biockern. 75, 402 (1976). A. Wadano, P. A. Hobus, and T.-H. Liao, Biocliemisrry 18, 4124 (1979). D. E. Fahrney and A . M. Gold, JACS 85, 997 (1963).

15. PANCREATIC DNase

285

that adsorption on Con A-agarose provided a means of obtaining a protease-free preparation of bovine pancreatic DNase. Funakoshi et a/. (19) have isolated from human duodenaljuice a DNase I that has properties very similar to those of the bovine pancreatic enzyme. Love and Hewitt (2U) have purified the human pancreatic enzyme with similar results. In their experiments they introduced a fluorometric DNase assay based upon the use of a circular DNA substrate and the binding of the denatured split products to ethidium bromide. In the other papers covered in this chapter, authors define their own modifications of the assays ( 1) based upon hyperchromicity, proton release, or acid-soluble nucleotides determined by absorbance or radioactivity.

111.

Chemical Structure

A.

SEQUENCE

Through study of the peptides yielded by tryptic or chymotryptic hydrolysis and by cyanogen bromide cleavage, Salnikowet al. (21)and Liaoet LII. (22) derived a sequence for the amino acid residues in reduced and carboxymethylated DNase A. The disulfide bridges were characterized by drawing upon the observation of Price et ril. (23) that in the presence of Ca2+one of the two S-S bonds could be selectively reduced by mercaptoethanol ; cyanogen bromide cleavage of the alkylated derivative provided data on the pairings. The result of the sequence study is given in Fig. 2. The sequence is a working hypothesis based upon all of the data available at this time. The molecule corresponds to a protein of 257 residues with carbohydrate attached through an aspartamido-hexose linkage at one position (Asn-18 in an Asn-X-Thr sequence). The amide -NH3 value is 21, which agrees with the determination by Lindberg (24) recalculated for the present molecular size. The molecular weight of DNase A, calculated from the amino acid and carbohydrate composition, is 30,072. The numbers of individual residues are Lys-9, His-6, Arg-11, Asp-20, Asn-12, Thr-15, Ser30, Glu-10, Gln-9, Pro-9, Gly-9, Ala-22, +Cys-4, Val-24, Met-4, Ile-11, Leu19. A. Funakoshi, Y. Tsubota, H. Wakasugi, H. Ibayashi, and Y. Takagi, J . Biochem. (Tokyo) 82, 1771 (1977). 20. J. D. Love and R. R. Hewitt, JBC 254, 12588 (1979). 21. J. Salnikow, T.-H. Liao, S . Moore, and W. H. Stein,JBC 248, 1480 (1973). 22. T.-H. Liao, J. Salnikow, S. Moore, and W. H. Stein, JBC 248, 1489 (1973). 23. P. A. Price, W. H. Stein, and S . Moore, JBC 244, 929 (1969). 24. U . Lindberg, Biochemistry 6, 335 (1967).

286

STANFORD MOORE

Leu-Lys-Ile-Ala-Ah-Phe-Asn-I

10 Corb. le-Arg-Thr-Phe-Gly- Glu-Thr-Lys-Met-Ser-Asn-

20

30

Alo -Thr-Leu- Alo-Ser-Tyr -1le-Vol- Arg-Arg -Tyr-Asp-Ile-Val-Leu-Ile-Glu-Gln-Val40

Arg-Asp-Ser-His-Leu-Val-

50 Ala- Val-Gly-Lys-Leu-Leu-Asp-Tyr

- Leu-Asn-Gln- Asp-Asp70

60

Pro - As n -T hr - Ty r - H i s -Ty r - Vo I- Vo I- Ser - GI u - Pro - Leu- GIy - Arg - Asn- Ser -Ty r - Lys -G Iu 80

YO

- Gln-Tyr -

Arg-Tyr-Leu-Phe-Leu-Phe-Arg-Pro-Asn-Lys-Val-Ser-Val-Leu-Asp-Thr-Tyr I00

110

Asp-Asp-Gly-Cys- Glu- Ser-Cys-Gly -Am-Asp-Ser-Phe-Ser- Arg-Glu-Pro- Alo-vai-vai -

u

I30

Lys-Phe-Ser-Ser-His-Ser-Thr-Lys-Val-Lys-Glu-Phe-Ala-Ile-

Val-Alo-Leu-His-Ser-

140

Alo-Pro-Ser-Asp-Ala-Val-

I50

Ala-Glu-lle-Asn-Ser-Leu-Tyr

-Asp-Val-Tyr -Leu-Asp-Val

160

-

I70

Gln-Gln- Lys -Trp-His-Leu-Asn-Asp-Vol-Met-Leu-Met-Gly-Asp-Phe-Asn-Alo-Asp-CysI80

Ser-Tyr- Vol-Thr-Ser-Ser-Gln-Trp-

Ser- Ser- Ile- Arq-Leu- Arg-Thr- Ser-Ser -Thr-Phe200

190

Gln-Trp-Leu- I l e - Pro-Asp-Ser-Alo-Asp-Thr-Thr-Alo-Thr-Ser-Thr-Asn-Cys-Aia 210

-Tyr-

220

Asp-Arg-Ile- Val-Val- Alo-Gly-Ser-Leu-Leu-Gln-Ser-Ser-Vol230 A lo-Pro-Phe-Asp-Phe-GIn250

Ser-Asp-His-Tyr-Pro-Val-

Vol- Gly-Pro-Ser- Ala-

240

Ala-Ah-Tyr-Gly- Leu-Ser- Asn-Glu-Mel-Ala-Leu-Alo-Ile

-

2 57

Glu-Vol-Thr-Leu-Thr

FIG. 2. Sequence of bovine pancreatic deoxyribonuclease A. From Sainikow et rrf. (21) and Liao et a / . ( 2 )reproduced . with permission. 23, Tyr-15, Phe-11, Trp-3, Man-5.8, and GlcNAc-1.9. The carbohydrate side chain (25) is probably a mixture of oligosaccharides of slightly different chain lengths. The pl is about 5 (7).

B. ESSENTIALITY OF SPECIFIC RESIDUES

The enzyme is inactivated by iodoacetate at pH 7.2 in the presence of Mn2+ or Cuz+ with the formation of one residue of 3carboxymethylhistidine (26). The sequence of the tryptic peptide that con25. B. J. Catley, s. Moore, and W. H. Stein, JBC 244, 933 (1969). 26. P. A. Price, S. Moore, and W. H. Stein, JBC 244, 924 (1969).


287

tained the modified amino acid permits the assignment of the crucial histidine residue to position 13 1 (22). The conclusion from studies with small substrates (I) that there may be a positive charge near the active center histidine residue fits with the fact that this imidazole ring is alkylated by iodoacetate but not by iodoacetamide (16). The histidine residue in DNase A that is substituted by proline in DNase C is His- 118 (10): this residue is thus not essential to the activity of the enzyme. Loss of activity by nitration has been found to parallel the formation of a single 3-nitrotyrosine residue per enzyme molecule (27). By reference to Fig. 1, the composition and sequence of the Tyr-(3-N02)-containing peptide isolated from an enzymatic digest permits assignment of the tyrosine residue to position 62 (2). Ca2+ was not able to stabilize the nitrated enzyme toward chymotryptic digestion, thermal denaturation, or mercaptoethanol reduction of the essential disulfide linkage. Tyr-62 may contribute to the formation of a Cazf binding site on the molecule that is not coincident with the region of His- 131, since the latter remains sensitive to specific carboxymethylation in the nitrated enzyme (27). The disulfide bond that can be reduced without loss of activity (13)is the one involved in the small loop between residues 98 and 101 (22). The disulfide bond between residues 170 and 206 is essential for maintenance of the active molecule. By controlled proteolysis with chymotrypsin, Hugli (28) was able to split the bond between Trp and Ser at positions 178 and 179; the product retained nearly the full activity of the native enzyme. When the cleaved molecule was further digested with carboxypeptidase-Y, the residues Thr-Ser-Ser-Gln-Trp (residues 1 7 4 178) were removed and the molecule still retained 80% of its activity. Thus, five residues h a t e d in the central portion of the peptide chain are functionally expendable. The COOH-terminal residues of DNase, which are normally unavailable to carboxypeptidases, become susceptible to removal when the enzyme is denatured in 0.005% sodium dodecyl sulfate (29). Study of the effect of carboxypeptidase action upon the enzyme required development of a procedure for restoring activity to DNase that has been denatured by the detergent. The inactivation of DNase could be completely reversed by diluting the enzyme solution tenfold into 6 M guanidinium chloride before a 100-fold dilution for assay. A loss of regenerable activity could be correlated with the removal of 1 or 2 amino acid residues (-Leu-Thr) from the 27. T. E. Hugli and W. H. Stein,JBC 246, 7191 (1971). 28. T. E. Hugli, JBC 248, 1712 (1973). 29. T.-H. Liao,JBC 250, 3831 (1975).

288

STANFORD MOORE

COOH-terminal sequence. DNase thus resembles RNase (30,31) in being one of the several enzymes in which the residues at the COOH terminus have a determining effect upon the folding of the chain into the active conformation. By following the kinetics of the reaction of N-bromosuccinimide with DNase by amino acid analysis for tryptophan, Sartin er ul. (32)have been able to show that modification at Trp-155 is the change most crucial to inactivation by that reagent. Methanesulfonyl chloride at pH 5 inactivated DNase by modification of the hydroxyl group of an as yet unidentified serine residue (3.3). DNase is inactivated by 2-nitro-5-thiocyanobenzoicacid (34)by a reaction that has been shown to involve cleavage of the peptide chain at the hydroxyamino acids at positions 14, 40, 72, and 135 (35). In more general derivatization experiments, guanidination of the nine E-NH, groups or picolinimidylation of the a- and €-amino groups yields active derivatives (36). The NH, groups are thus not essential per se, but when positive charges on the enzyme are removed by carbamylation with cyanate, activity is progressively lost. Modification of the carboxyl groups by condensation with glycine ethyl ester in the presence of a carbodiimide causes major inactivation in the absence of Ca2+(-?3).The presence of the bivalent cation slows the rate of the inactivation.

IV.

Catalytic Properties

A. ROLESOF DIVALENT METALIONS The conformation and the activity of the molecule are markedly dependent upon the presence of metal ions. The resistance to proteolysis conveyed by Ca2+ (8) has been important in the purification of the protein. The role of Caz+ in the refolding of the reduced enzyme to the active conformation is crucial (23). The apparent molecular weight of DNase by gel filtration in Tris buffer increases with pH in the range from pH 7.5 to 9 30. 31. 32. 33. 34. 35. 36.

C. B. Anfinsen, J5C 221, 405 (1956). M . C. Lin,JBC 245. 6726 (1970). J . L. Sartin, T. E. Hugh, and T.-H. Liao, JBC 255, 8633 (1980). T. L. Poulos and P. A. Price, JBC 249, 1453 (1974). T.-H. Liao and L. J . McKenzie, JBC 254, 9598 (1979). T.-H. Liao and A. Wadano, JBC 254, 9602 (1979). B . V. Plapp, S. Moore, and W. H. Stein, JEC 246, 939 (1971).

15.

289

PANCREATIC DNase

TABLE I1

CA" REQUIREMENTFOR DNASEACTIVITYAT P H 8 '* H yperchromicity assay

Metal ions in assay

midmg)

Maxi mu m specific activity (9%)

2.5 mM MgCI, 2.5 mM MgCI2, Dowex-purified 2.5 mM MgCI, + EGTA lo-' M CaC12b 2.5 mM MgCI, + IO-'M CaC12b

10-18 3-6 0.1 8 710'

1.4-2.5 0.4-0.9 0 1.1 100

Specific activity (&so

I

pH-stat assay Specific activity (NaOHI min/DNase) 10

3 0 5 650'

Maximum specific activity (%)

1.5 0.5

0 0.8 100

" From Price (42), reproduced with permission. Each value represents the average or range of values for three or more assays of metal ion-free DNase A. Hyperchromicity assays were in 5 mM ionic strength Tris-chloride buffer at pH 8 and 25". pH-stat assays were in buffer-free solution at 25". DNA concentration is 0.04 mg/ml. ' Addition of lo-" M EGTA has no effect on these activities.

(37,38), a hydrodynamic change that is reversed by the presence of Caz+.

By gel filtration of DNase A at pH 7.5 with 45Ca2+,Price (39) found 2 Ca2+ bound with an average Kd of 1.4 x and 3 bound with a K d of 2 x With Mg2+the Kd for two sites was 2.3 x One of the two strong Ca2+ binding sites is not subject to competition from Mg2+.Fifty percent of the maximum transition in the CD spectrum occurs near M CaCl, and half-maximum protection against action by trypsin is achieved near 1.3 x M CaCl, (40).The uv spectra show a conformational change indicative of increased interiorization of tryptophan and tyrosine residues in the presence of Ca'+ (41). Price (42) has made the key observation (Table 11) that with reagents that have been purified to reduce the Ca2+contamination to a minimum, DNase in the presence of Mg2+is about 99.5% inactive. In the presence of B . Librraga, C . Bustamante, A. Gil, and E. Melgar, BBA 579, 298 (1979). B . Lizarraga, D. Sanchez-Romero, A. Gil, and E. Melgar,JBC 253, 3191 (1978) P. A. Price, JBC 247, 2895 (1972). T. L. Poulos and P. A. Price, JBC 247, 2900 (1972). 41. R . Tbllis and P. A. Price, JEC 249, 5033 ( 1974). 42. P. A. Price, JBC 250, 1981 (1975).

37. 38. 39. 40.

290

STANFORD MOORE

a very low concentration (0.01 mM) of EGTA, a chelating agent that binds Ca*+about lo6 times more strongly than it binds Mg2+,DNase activity in the presence of 2.5 mM Mg2+becomes undetectable. A Ca2+concentration of 0. I mM yields maximum activity; concentrations as high as 1 mM are inhibitory. Earlier experiments on the low activity of DNase when only MgCl, is added are attributed to the effect of traces of Ca2+in the reagents. Bivalent metal ions serve two essential roles; Ca2+ must be bound to the enzyme and ions such as Mg2+to the substrate. The concento 3 x tration of Ca2+in bovine pancreatic juice is 4 x M ( 4 3 ,a range that can contribute to keeping DNase functional in its physiological environment. The effects of various bivalent metal ions are summarized in Table I11 (42). Sr2+and Ba2+can substitute for Ca2+,but are leass effective. Mn2+ and Co2+ can substitute for Mg2+, the latter with only about 10% efficiency. The results permit earlier studies on metal ion effects [see, e.g., Refs. (44,431 to be interpreted in more operational terms. The degree to which the enzyme functions optimally depends upon two factors, the Ca2+concentration and the Mg2+ or Mn2+ concentration, with the former being subject to variation from trace amounts of Ca2+as a contaminant in the latter. Double-strand scission [see ( I ) ] can be expected when the affinity of the enzyme for substrate is at a maximum in the presence of both CaLf and Mg2+;single-strand scission and changes in specificity are likely to be associated with suboptimal concentrations of Ca2+ present as contaminants when Ca2+is not deliberately added (42). Douvas and Price (46) have shown that 1 Mg2+ per 2 DNA-phosphorus is optimum. Na+-DNA is inhibitory and maximum rates are obtained with Mg2+-DNAas substrate rather than with Na+-DNA plus MgC1,. Preincubation of DNase with Ca2+ before addition to the substrate increases the initial rate of hydrolysis twofold over that obtained with Ca2+-free DNase with double-stranded calf thymus DNA as substrate. B. SUBSTRATE SPECIFICITY

Junowicz and Spencer (45) have conducted an extensive enzymatic and chromatographic study of the terminal purines and pyrimidines in the oligonucleotides liberated by the digestion of calf thymus DNase A with a variety of bivalent ion mixtures for different times; the experiments were 43. 44. 45. 46.

A. Frouin and P. Gerard, C.R. SOC.Bid. 72, 98 (1912). E. Junowicz and J. H. Spencer, BBA 312, 72 (1973). E. Junowicz and J . H. Spencer, BBA 312, 85 (1973). A. Douvas and P. A. Price, BBA 395, 201 (1975).

IS.

29 1

PANCREATIC DNase TABLE 111

ACTIVITYO F DNASEWITH DIFFERENT METAL IoNs"*~ Activities under following conditions of assay*

Metal ion

2.5 mM MgCI,, lo-' M metal ion

2.5 m M Metal ion, M Caz+

9 0 700 440 220 50 30

700 700 8 0.9 4.2 750 81 0 0

MgCI, MgCI, + 10-5M EGTA CaCI, SrCI, BaCI, MnCI,' COCI," CdCI,, SnCI,, FeCI,, NiCI,, CuCI, ZnCI,, EuCI,, SmCI,, NdCI,

II 0

From Price (42), reproduced with permission. Hyperchromicity assays in 5 rnM Tris, pH 8, 25". with 0.04 mdml of DNA. Activity values are AA,,, per midmg of DNase. The possibility exists that traces of CaZ+in MnCI, and CoCI, account for these values.

conducted before the effects of traces of Ca2+in most bivalent metal salts (42) were fully appreciated. Under near optimal conditions (e.g., with Mg2+plus C a 2 + ) , the enzyme gives the molar yields of the four nucleosides in the end positions listed in Table IV; in these experiments the enzyme is impressive for its versatility rather than its selectivity. The results did not vary greatly with the time of hydrolysis. However, with several bivalent TABLE 1V

DEOXYRIBONUCLEOTIDES AT T H E 5' AND 3' ENDSOF T H E DNAovb OLIGONUCLEOTIDES RELEASEDFROM C A L F THYMUS Mole% nucleosides ~

~~

5' end

3' end

Digestion time (rnin)

dT

dC

dG

dA

dT

dC

dG

dA

1.5 30.0

29.0 34.8

18.2 22.2

23.8 17.4

29.0 25.5

29.0 27.7

10.5 8.5

28.4 30.1

32.1 33.7

" From Junowicz and Spencer (4.5), reproduced with permission. Activation by 33 m M Mg2+ and 0.1 mM Ca'+.

292

STANFORD MOORE

metal additions that gave less than maximum specific activity, the molar ratios of the terminal nucleosides varied markedly and the proportions changed with the time of digestion. At the 5’ end, the ratio of dT or dC to dG or dA varied from 1 : 50 to 1 : 1; at the 3’ end, the ratio of dA or dC to dG or dT varied from 1 : 5-25 to 1 : 1. In these digests and with the singlestranded DNA from E. coli K12, consistent yields were obtained of long oligonucleotides lacking dA at the 3’ end. Simon et al. (47) have extended to DNase their use of crab d(A-T) polymer as a substrate for nucleases. This unique polymer, which is composed predominently of alternating A and T but contains about 3% of G and C residues integrated into its structure, was submitted to controlled hydrolysis by DNase in the presence of varying concentrations of MgCI, to obtain a hexanucleotide fraction enriched in C and G. Pruch and Laskowski (48) have subsequently undertaken to determine whether the I-3% of ribonucleotides, still detectable in preparations of crab d(A-T), that have been exhaustively treated with RNases, are built-in components. Digestion with DNase A to dinucleotides led to chromatographic evidence for the mixed dinucleotides dC-rG, dT-rA, and dT-rG. The authors conclude that crab d(A-T) polymer from C . borealis contains covalently bound ribonucleotides and that, as a corollary premise, sugar specificity of DNase may be limited to the nucleotide following the point of cleavage. The result will stimulate experimentation; the finding fits the function of DNase as an enzyme with which dinucleotides are key end products. Pancreatic DNase has been an enzyme for which no small synthetic substrate is hydrolyzed rapidly. p-Nitrophenyl esters have proved to be convenient synthetic substrates in spectrophotometric assays for use in studies on the kinetics and mechanisms of action of a number of phosphodiesterases. With DNase I, Liao (49) has examined deoxythymidine, 3’, 5’-di-p-nitrophenyl phosphate, a substrate that Razzell and Khorana (50) studied with snake venom phosphodiesterase, and that Cuatrecasas et cil. (51) found to be highly susceptible to the action of staphylococcal nuclease. The compound is rapidly hydrolyzed at a single bond by DNase at pH 7 . 2 with the liberation of p-nitrophenol measurable at 400 nm. The binding is not strong: at 10 mM substrate, the enzyme is not saturated, but the initial rate of hydrolysis (in the first 1 to 5 min) varies linearly with 47. M . Simon, H.-C. Chang, and M . Laskowski, Sr. BBA 232, 462 (1971). 48. J. M. Pruch and M. Laskowski, Sr. JBC 255, 9409 (1980). 49. T.-H. Liao, JBC 250, 3721 (1975). 50. W. E. Razzell and H. G . Khorana, JBC 234, 2105 (1959). 51. P. Cuatrecasas, ‘M. Wilchek, and C. B . Anfinsen, Biochemistry 8, 2277 (1969).

15.

PANCREATIC DNase

293

enzyme concentration in the range from 1 to 6 pg of DNase in 110 pl. Bivalent metals are essential; the maximum facilitation was obtained with 10 mM MnCI, and 1 mM CaCl,. The enzymatic activities toward DNA and N02Ph-pdTp-NOzPhwere lost in parallel upon carboxymethylation of His-131 (26). Since the photometric response is not specific for DNase, the assay has use only with the purified enzyme. Examination of the products of hydrolysis by paper electrophoresis gave the unexpected result that DNase liberates p-nitrophenol from the 3’-ester group. Snake venom phosphodiesterase, which like DNase yields 5’-nucleotides from DNA, liberates p-nitrophenol from the 5’-ester group of this substrate (SO).Staphylococcal nuclease (51 ) gives predominantly p-nitrophenyl phosphate from the 5’ position. Thus, these three diesterases hydrolyze N0,Ph-pdTp-N0,Ph in three different ways. DNase A has been cross-linked to RNase by Wang (52) to prepare a bifunctional enzyme. The coupling was via the thiolation of each protein with N-acetyl-DL-homocysteinethiolactone to yield a disulfide bridge between the two enzymes. The product, which contained one molecule each of DNase and RNase, hydrolyzed thymus DNA and yeast RNA with 75 and 40%, respectively, of the efficiencies of the parent catalysts. The RNA strand of the hybrid substrate phage f l DNA * [3H]RNA was hydrolyzed rapidly by the Mn2+-activatedhybrid enzyme; the RNA strand in the complementary combination was not hydrolyzed significantly by RNase alone. Although an -S-Slinkage may not be the most practical linkage, the concept of the conjugation offers the possibility of delivering in vivo two enzymes that differ in size, charge, and biological function to the same site at the same time.

V.

Actin as a n Inhibitor of DNase I

The presence in most mammalian cells of a protein that will inhibit pancreatic DNase has been known for over thirty years (I). Lazarides and Lindberg (S3)found in 1974 that the protein is cytoplasmic actin. Purified actin from chicken skeletal muscle inhibits DNase; the DNase inhibitor isolated from various tissues and cells is found to be closely similar to actin in its physical and chemical properties. Antibodies to the purified DNase inhibitor show reactivity toward actin-containing fibers in human skin fibroblasts. The inhibitory protein is thus a major cellular component that usually constitutes 5-10% of the soluble protein. The attribution of a 52. D. Wang, Biochemistry 18, 4449 (1979). 53. E. Lazarides and U . Lindberg, PNAS 71, 4742 (1974).

294

STANFORD MOORE

physiological role to the DNase-inhibitory action of actin is limited by the paucity of information on the existence of DNase I-type enzymes in nonpancreatic cells. The situation is very different from that which pertains to the highly specific cytoplasmic RNase inhibitor (54), K i= 3 X lo-’’’ M, which is present in trace amounts (about 0.01-0.02% of the soluble protein) along with a one-fifth to one-tenth molar quantity of a pancreatictype RNase, the activity of which is thus modulated. In line with Laskowski’s (I) suggestion, inhibition by actin can be a key criterion for characterization of intracellular nucleases of the DNase I type. Actin can be removed from cellular extracts by adsorption on a column of DNase coupled to agarose (55); so far, it has not been possible to elute the actin from the affinity column in a way that will preserve the inhibitory activity. Elution with 3 M guanidinium chloride, 1 M in sodium acetate (pH 6 3 , and 30% in glycerol yields the actin-like protein, but with more than 90% of the inhibitory activity lost. A pH 2.8 buffer did not release the actin from the adsorbent. Hitchcock et al. (56) and Mannherz et al. (57) have shown that DNase I causes depolymerization of filamentous actin to form a stable complex of 1 mole of DNase I with 1 mole of globular actin (MW -49,000). Wang and Goldberg (58) have utilized the affinity of DNase for actin-containing fibers to visualize microfilament bundles in nonmuscle cells: DNase I was added to formaldehyde-fixed and acetone-extracted chick or human fibroblasts followed by antibody to DNase for indirect immunofluorescence microscopy, or rhodamine-conjugated DNase was used for direct fluorescent microscopy. A selective assay for monomeric and filamentous actin in cell extracts has been developed by Blikstad et al. (59); the inhibition of DNase activity by G-actin is measured a few seconds after the addition of the enzyme and also after depolymerization of F-actin by 0.75 M guanidinium chloride, 0.5 M in sodium acetate, 0.5 mM in CaCl,, 0.5 mM in ATP, and 10 mM in Tris-HC1 (pH 7.5). Under this condition for 5 min at O”, the F-actin is converted to G-actin without loss of the DNase-inhibitory activity in the monomer. With fluorescently labeled DNase, Mannherz et al. (60) estimated the 54. P. Blackburn and S. Moore, “The Enzymes,” 3rd Ed., Vol. 15, in press. 55. U. Lindberg and S. Eriksson, EJB 18, 474 (1971). 56. S. E. Hitchock, L. Carlsson, and U . Lindberg, Cell 7, 531 (1976). 57. H. G. Mannherz, J. Barrington Leigh, R. Leberman, and H. Pfrang,FEBS (Fed.Eur. Biochem. S o c . ) Lett. 60, 34 (1975). 58. E. Wang and A. R. Goldberg, J . Histochem. Cytochem. 26,745 (1978). 59. I. Blikstad, F. Markey, L. Carlsson, T. Persson, and U. Lindberg, Cell 15,935 (1978). 60. H. G. Mannherz, R . S. Goody, M. Konrad, and E. Nowak, EJB 104, 367 (1980).


295

binding constant for DNase I and monomeric rabbit skeletal muscle actin to be 5 x 10*M-' and the inhibition to be competitive. With fluorescently labeled actin, Ikkai et al. (61) obtained a K b value of 1 x lo6 M-I. A binding constant of 1.2 x 104 M-' was obtained with filamentous actin (60). Crystals of actin . DNase I complexes have been obtained for cystallographic studies (62, 6 3 ) . When Rohr and Mannherz (64)examined rat pancreatic juice by SDS gel electrophoresis for bands coinciding with actin and DNase actin complex, both were observed. When the juice was treated with 0.25 N H2S04, as in the first steps of the Kunitz (7) procedure for the isolation of pancreatic nucleases, the DNase activity doubled; the inhibitory action of actin is destroyed in this step but DNase is stable. The complex could also be dissociated under physiological conditions by rat or human bile; an activating component has been found to be 5'-nucleotidase (65). The enzyme from snake venom can produce the same results; the process is slow (14 hours at 23") with 10 mM DNase * actin complex and 2 mM nucleotidase. Since 5'-nucleotidase is a constituent of plasma membranes, they were tested (66) and found to effect a slow liberation of DNase from the complex; Grazi and Magri (67) suggest that phosphorylation of actin may have a role in the process. It has been proposed (60) that the interaction of DNase and actin may be a physiological process in the extracellular space of the gastrointestinal tract. VI.

Research Applications

DNase I has received wide use as a probe in the study of the structure of chromatin. Active genes are expected to be more readily accessible to digestion by DNase than the transcriptionally inert segments. Experiments in several laboratories (68-71 ) have demonstrated that limited 61. T. Ikkai, K . Mihashi, andT. Kouyarna,FEBS (Fed. Eur.Eiochem. S o c . ) L e t t . 109,216 (1980). 62. H. G. Mannherz, W. Kabsch, and R . Leberman, FEBS (Fed. E w . Biochem. S o c . ) Lett. 73, 141 (1977). 63. H . Sugino, N. Sakabe, K. Sakabe, S. Hatano, F. Oosawa, T. Mikawa, and S. Ebashi, J . Biochem. (Tokyo) 86, 257 (1979). 64. G. Rohr and H. G. Mannherz, EJB 89, 151 (1978). 65. H. G. Mannherz and G . Rohr, FEBS (Fed. E r r . Eiochem. Soc.) Lett. 95, 284 (1978). 66. G . Rohr and H . G . Mannherz, FEBS (Fed. Eur. Eiochem. Soc.) Lett. 99, 351 (1979). 67. E. Grazi and E. Magri, FEES (Fed. Eur. Eiochem. S o c . ) Lett. 104, 284 (1979). 68. H. Weintraub and M. Groudine, Stience 193, 848 (1976). 69. A. Garel and R. Axel, PNAS 73, 3966 (1976). 70. B . Sollner-Webb and G. Felsenfeld, Cell 10, 537 (1977). 71. S. Weisbrod and H. Weintraub, PNAS 76, 630 (1979).

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STANFORD MOORE

digestion by DNase can provide information on the structure of the nucleosome. High resolution electrophoretic separation of solubilized oligonucleotides has provided evidence (72-74) for a repetitive internal structure of the chromatin subunit. The periodicity in the fragment lengths can be correlated with helical and super helical orientation of DNA in the nucleosome core (75). In a study of the role of postsynthetic modification of histones in gene activation, Vidali et NI. (76) have used DNase I digestion to show that DNA is more readily liberated from chromatin that contains an increased quantity of acetylated histones; Kastern et al. (77) have used DNase coupled to Sepharose in the preparation of DNA-dependent RNA polymerase 11; the use of an immobilized enzyme makes it possible to release RNA polymerase from actively transcribing genes without having the product contaminated by DNase. The interaction of DNase and actin has formed the basis of a procedure for visualizing cellular microfilaments by fluorescent microscopy (58) and for a differential assay for monomeric and filamentous actin in cell extracts (59).

Thoroughly RNase-free DNase (13) has proved useful (78, 79) to digest chromatin in the preparation of nuclear RNA by the procedure of Penman (80).The protease-free nature of the same DNase preparation is of potential value in the isolation of nuclear proteins. On the basis that the process of replication in virus-infected cells is preceded by liberation of the viral nucleic acid from its protective protein coating, Trukhachev and Salganik (81) studied the inhibition of viral synthesis by DNase in cell cultures. In a clinical study, favorable results (82) have been reported from the local application of DNase to patients suffering from herpes infections of the eye. M . NOH, Nitcleic Acids Res. 1, 1573 (1974). L. C. Lutter, J M B 117, 53 (1977). L. C. Lutter, Nircleic Acids Res. 6, 41 (1979). A. Prunell, R. D. Kornberg, L. Lutter, A. Klug, M. Levitt, and F. H. C. Crick, Science 204, 855 (1979). 76. G. Vidali, L. C. Boffa. E. M . Bradbury, and V. G . Allfrey, /“AS 75, 2239 (1978). 77. W. H. Kastern, J. D. Eldridge, and K. P. Mullinix, JBC 254, 7368 (1979). 78. I. Tamm and T. Kikuchi, P N A S 76, 5750 (1979). 79. I. Tamm, T. Kikuchi, J. E. Darnell, Jr., and M. Salditt-Georgieff, Biockemisrry 19, 2743 (1980). 80. S. Penman, in “Fundamental Techniques in Virology” ( K. Habel and N. P. Salzman, eds.), p. 35. Academic Press, New York, 1969. 81. A . A. Trukhachev and R. 1. Salganik, Virology 33, 552 (1967). 82. A . A. Colain, R . 1. Salganik, I . E . Mikhailovskaya, and I . M. Gorban,Ann. h i / . 203, 371 (1970). 72. 73. 74. 75.

Section Ill

DNA Modification


Bacteriophage T4 Polynucleotide Kinase CHARLES C. RICHARDSON

I . Introduction

,

.

. .

.

. . .

. .

.

.

. .

. .

. .

. .

.

.

. .

. .

11. Isolation and Physical Properties . . . . . . . . . . . . . . . .

A . Assay . . . . . . . . . . . . . . . . . . B . Purification , . . , . . . , . . . . . , . . C . Physical Properties . . . . . . . . . . . . 111. Catalytic Properties . . . . . . . . . . . . . . A . Reactions Catalyzed by Polynucleotide Kinase B . Phosphorylation of 5’-Hydroxyl Termini . . . C. Reversal of the Kinase Reaction . . . . . . D. 3‘-Phosphatase Activity . . . . . . . . . . 1V. Role of Polynucleotide Kinase in Vitw . . , . . V. Research Applications . . . . . . . . . . . .

1.

. . . . . . . . . . . . . . . . . . . . . . . , . . . . . .

. . . . . . . . , . . . . . . . . . . . . . . . . . . . . . . . . . . , . . . . . . . . . . . . .

.

.

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.

. .

. . .

299 30 1 30 1 301 303 305 305 305 309 310 312 3 13

Introduction

Polynucleotide kinase is an enzyme that catalyzes the transfer of the y-phosphate of a nucleoside 5’-triphosphate to the 5’-hydroxyl terminus of a deoxyribonucleic acid or ribonucleic acid molecule. The reaction products are the nucleoside 5’-diphosphate and a 5’-phosphoryl-terminated polynucleotide. The discovery of polynucleotide kinase was made during an attempt to identify enzymes that catalyze the formation of a 5’-terminal triphosphate 299 THE ENZYMES, Vol. XIV Copyright 0 1981 by Academic Press, Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

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CHARLES C. RICHARDSON

group on either an RNA or a DNA molecule. The impetus for such a search was the hypothesis that the formation of a phosphodiester bond to join polynucleotide strands covalently, as is the case in recombination and repair, would require activation of the 5’ terminus of one of the polynucleotides. A nucleophilic attack of the 3’-hydroxyl group of another polynucleotide on the a-phosphate of the triphosphate could result in phosphodiester bond formation and the release of pyrophosphate, a reaction known to occur in the polymerization of nucleotides by RNA and DNA polymerases. Although no such activities were identified that could catalyze this addition of phosphoryl groups to 5’-phosphoryl-terminated polynucleotides, these studies did lead to the discovery of a different enzyme activity-polynucleotide kinase-that could phosphorylate 5 ’ hydroxyl termini. Subsequent studies showed that covalent joining of polynucleotides involved a different enzyme, DNA ligase, via the activation of the 5’ terminus of the polynucleotide by adenylation to form a pyrophosphate bond. The rationale to search for enzymes that catalyze the synthesis of 5’-triphosphate termini was not unreasonable: just such an enzyme has been isolated from vaccinia virus cores (I). Polynucleotide kinase was first identified in Escherichici coli cells infected with bacteriophage T2 or T4 (2, 3 ) . Although no polynucleotide kinase activity has been found in uninfected bacteria, a similar activity has been identified (4, 5 ) and partially purified (6, 7) from rat liver nuclei. Of these three polynucleotide kinases, the bacteriophage T2- and T4induced enzymes have been most extensively purified. While the T2 and T4 enzymes are very similar, the T4 polynucleotide kinase has been more extensively characterized. Therefore, this chapter focuses on the bacteriophage T4 polynucleotide kinase, with reference to studies on the other enzymes only when they supplement or differ from those obtained with the T4 enzyme. Whereas studies have provided some information on the role of polynucleotide kinase in T4 phage-infected cells, the major importance of the enzyme is as a reagent in nucleic acid studies. In this chapter major emphasis is placed on (1) the purification of T4 polynucleotide kinase, (2) the properties and substrate specificities of the reactions catalyzed by the enzyme, and (31 the research applications of the enzyme. 1. E. Spencer, D. Loring, J. Hurwitz, and G . Monroy, PNAS 75, 4793 (1978). 2. C. C. Richardson, P N A S 54, 158 (1965). 3. A. Novogrodsky and J . Hurwitz, JBC 241, 2923 (1966). 4. A. Novogrodsky, M. Tal, A. Traub, and J. Hurwitz,JBC 241, 2933 (1966). 5 . M. Ichimura and K . Tsukada, J. Biochem. 69, 823 (1971). 6. H. Teraoka, K. Mizuta, F. Satl, M . Shimoyachi, and K . Tsukada, EJB 58, 297 (1975). 7. C. J. Levin and S. B . Zimmerrnan,JEC 251, 1767 (1976).

16. BACTERIOPHAGE T4 POLYNUCLEOTIDE KINASE

II.

A.

30 1

Isolation and Physical Properties

ASSAY

A convenient and quantitative assay for pol ynucleotide kinase measures the conversion of acid-soluble radioactivity in [y-32P]ATPto an acidinsoluble product (2, 3 , 8 ) .The most commonly used acceptor DNA substrate is prepared by partial digestion of duplex DNA with micrococcal nuclease (2. 3, 8 ) , a nuclease that hydrolyzes phosphodiester bonds to produce oligonucleotides terminated by 5’-hydroxyl and 3’-phosphoryl groups. Alternatively, 5’-hydroxyl termini can be introduced into DNA by first sonically irradiating or partially hydrolyzing DNA with pancreatic DNase followed by dephosphorylating the 5’ termini with E. coli alkaline phosphatase (9, 10). These latter substrates have been used to follow the purification of the T4 (11) and rat liver (6) enzymes, respectively.

B.

PURIFICATION

The purification procedure for polynucleotide kinase from bacteriophage TCinfected cells that was described initially consisted of a series of six steps ( 2 ) . In order to obtain a physically homogeneous enzyme and to remove traces of contaminating nuclease activity a seventh step, chromatography on hydroxylapatite, was added to the purification procedure (12). For convenience the published purification procedure (8) is summarized in Table I. An alternative procedure for the purification of the T2 polynucleotide kinase has been described ( 3 ) . The source of the enzyme described in Table I is from E. coli cells infected with T4 r+ phages. A 50 to 100% increase in polynucleotide kinase activity can be obtained by infecting E.coli B su- (nonpermissive host) with one of several T4 mutant phages defective in lysis, such as T4 amN8-7 (13) or T4 N55SP62 (14). In addition, by choosing the appropriate T4 mutant for infection, it is possible to eliminate certain potential enzyme contaminants from the very start of the purification. For example, extracts prepared from E. coli B infected with T4 amXF1 (genes 41-45) are free of 8. C. C. Richardson, Proced. Nucleic Acid Res. 2, 815 (1972). 9. C. C. Richardson, J M B 15, 49 (1966). 10. B. Weiss, T. R . Live, and C. C. Richardson, JBC 243, 4530 (1%8). 1 I . K. Sirotkin, W. Cooley, J. Runnels, and L. R. Snyder, J M B 123, 221 (1978). 12. A. Jacquemin-Sablon and C. C. Richardson, JMB 47, 477 (1970). 13. A. Panet, J . H. van de Sande, P. C. Loewen, H. G. Khorana, A. J. Raae, J. R. Lillehaug, and K. Kleppe, Biochemistry 12, 5045 (1973). 14. K. L . Berkner and W. R. Folk,JEC 252, 3176 (1977).

302

CHARLES C. RICHARDSON TABLE I

mRIFICATI0N OF POLYNUCLEOTIDE KINASEFROM OF E. coli INFECTED WITH PHAGE T4" Total units Fraction I 11

I11 IV V VI VII

Step Extract Streptomycin Auto1ysis Ammonium sulfate DEAE-cellulose Phosphocellulose H ydroxylapatite

(X

10-3)

95

90 1 I5 105 68 40 27

25 GRAMS

Specific activity (unitdmg) 40 60

I50 420 8,500 59,000

-b

From Richardson ( 8 ) . Protein concentration insufficient to obtain an accurate specific activity.

T4 DNA polymerase, since gene 43 is the structural gene for T4 DNA polymerase (15, 16). Contamination of the purified kinase by DNA ligase can be reduced by the use of T4 amH39X (gene 30)-infected cells, since gene 30 is the structural gene for the T4 DNA ligase (17). The same procedure for growth and infection as that used for wild-type T4 is followed when such mutant phages are used, except that the cells are harvested at a later time (8). Similarly, attention to the E. coli host used to prepare the infected cells can also circumvent troublesome contaminating activities. For instance, ribonuclease activity can be greatly reduced at the outset by using ribonuclease mutants of E. coli (18). The structural gene for polynucleotide kinase, the pseT gene of phage T4, also codes for the T4 3'-phosphatase (11, 1 9 ) . Although the 3'phosphatase activity does not normally interfere with the use of the polynucleotide kinase, there are occasions when it is desirable to prepare oligonucleotides having both 3'- and 5'- terminal phosphates. Escherichia coli cells infected with the T4 mutant, p s e T f , synthesize an altered polypeptide that has relatively normal kinase activity but greatly reduced levels of phosphatase activity (I 1, 20). Thus, polynucleotide kinase prep15. 16. 17. 18. 19. 20.

A. De Waard, A. V. Paul, and I. R . Lehman, PNAS 54, 1241 (1965). H. R . Warner and J. E. Barnes, Virology 28, 100 (1966). G. C. Fareed, and C. C. Richardson, PNAS 58, 665 (1967). M. Takanami, JMB 23, 135 (1967). V. Cameron, and 0. C. Uhlenbeck, Biochemistry 16, 5120 (1977). V. Cameron, D. Soltis, and 0. C. Uhlenbeck, Nucleic Acids Res. 5, 825 (1978).


303

arations essentially free of 3'-phosphatase activity can be obtained by purifying the kinase from extracts of T4 pseTI-infected E. coli (20). The pseTl polynucleotide kinase behaves analogous to wild-type kinase during purification. Minor modifications have been made in the purification procedure summarized in Table I by several laboratories. For example, low concentrations of ATP have been added to the buffers during purification in order to stabilize the enzyme (13,2I). Frequently it is advantageous to purify the T4 DNA polymerase and the T4 polynucleotide ligase from the same extract of infected cells. These enzymes can be purified by published procedures for the polymerase (22) and ligase (23) using the appropriate side fractions obtained from the kinase purification procedure summarized in Table I. Such simultaneous purification of the three enzymes has been described (13). In view of the extensive use of polynucleotide kinase in nucleic acid studies, considerable time, effort, and cost would be saved if the enzyme were amplified in E. coli by cloning the kinase gene, pseT. It seems likely that the pseT gene will be cloned because the difficulties encountered in generating restriction enzyme fragments of the glucosylated T4 DNA can be circumvented (24). Furthermore, restriction fragments of phage T4 DNA that contain at least portions of the kinase gene have already been cloned in E. coli (25).

C. PHYSICALPROPERTIES 1. Physical Homogeneity

T4 polynucleotide kinase has been purified to physical homogeneity as judged by polyacrylamide gel electrophoresis in the presence of sodium dodecyl sulfate at pH 8.3 after denaturation and reduction; a single band is also observed at pH 6.8 in the presence of 2-mercaptoethanol or during electrophoresis through urea gels (26). Sedimentation studies of the purified enzyme using the analytical ultracentrifuge also indicate that the 21. R. Wu and A. D. Kaiser, P N A S 57, 170 (1967). 22. B. Weiss, A. Jacquemin-Sablon, T. R. Live, G. C. Fareed, and C. C. Richardson, JBC 243, 4543 (I%@. 23. M. Goulian, Z. J. Lucas, and A . Kornberg, JBC 243, 627 (1968). 24. G. G. Wilson, V. I. Tanyashin, and N . E. Murray, Mol. Gen. Genef. 156, 203 (1977). 25. A. J. Mileham, H. R . Revel, and N. E. Murray, M o l . Gen. Genet. 179, 227 (1980). 26. J. R. Lillehaug, EJB 73, 499 (1977).

304


enzyme is physically homogeneous. Only a single N-terminal amino acid, phenylalanine, is present in the purified enzyme (26). 2. Molecular Weight

The molecular weight of native polynucleotide kinase, as estimated by filtration through a Sephadex (3-200 column calibrated with proteins of known molecular weight, is 140,000 (13). The apparent molecular weight of the denatured and reduced form of the kinase, as determined by comparison with the mobilities of proteins of known molecular weight on polyacrylamide gels, is 33,000 (13, 26). Since phenylalanine is the exclusive N-terminal amino acid, it is likely that T4 polynucleotide kinase contains four identical subunits. In support of this structure is the finding of minor bands of 60,000 and 120,000daltons during electrophoresis of the denatured and reduced protein on polyacrylamide gels containing dodecyl sulfate at pH 6.8 (26). The szo,wof the purified enzyme has been determined under a variety of conditions (26). In 0.1 M potassium phosphate buffer, pH 7.8, containing 1 mM 2-mercaptoethanol, the enzyme preparation contains approximately 30% of a 2.9 S and 70% of a 6.5 S species, presumably the monomer and tetramer, respectively. The molecular weights of these two species of kinase have been determined by sedimentation equilibrium ultracentrifugation and were found to be 33,200 and 147,300, respectively. The activity of T4 polynucleotide kinase is stimulated by NaCl and KCl and by polyamines such as spermine (27). Potassium chloride is necessary to maintain an oligometric structure, and incubation with spermine converts the protein to a pure 6.2 S species (tetramer) (26). The polynucleotide kinase isolated from rat liver nuclei has an apparent molecular weight of 80,000 as estimated by gel filtration (6, 7). The sedimentation coefficient is 4.4 S (6). 3. Amino Acid und Spectrophotometric Analyses The amino acid composition of T4 polynucleotide kinase has been determined (26); the N-terminal amino acid is phenylalanine. The protein has two sulfhydryl groups, one exposed to the environment and the other more buried, as determined by reactivity to 5 , 5’-dithiobis(2-nitrobenzoic acid). Analysis of the ultraviolet absorption spectrum (26) shows the maximum absorption at 276 nm. The 280-260 nm absorbance ratio is 1.28. The absorption coefficient at 276 nm is 11.0 cm-’. The CD spectrum of the protein indicates that 45 to 55% of the polypeptide chain is in an a-helical conformation (26 ) . 27. J. R. Lillehaug and K. Kleppe, Biochemistry 14, 1225 (1975).


111.

Catalytic Properties

A.

REACTIONSCATALYZED BY POLYNUCLEOTIDE KINASE

305

Polynucleotide kinase of bacteriophage T4 catalyzes the transfer of the y-phosphate of a nucleoside 5‘-triphosphate to the 5’-hydroxyl group of a nucleoside 3’-phosphate, oligonucleotide, or polynucleotide (Fig. 1). The products of the reaction are a nucleoside 5’-diphosphate and the 5’phosphoryl nucleotide or polynucleotide (2-4, 8, 28). The reaction shown in Fig. 1 is reversible (14,29,30). In the presence of a nucleoside 5’-diphosphate, a 5’-phosphoryl polynucleotide is dephosphorylated to yield a 5’-hydroxyLterminated polynucleotide and the nucleoside 5’-triphosphate. The fact that this reaction is reversible makes possible the exchange of radioactivity between the y-phosphate of a nucleoside 5‘-triphosphate and the 5’-phosphoryl group of a polynucleotide. In addition to the phosphorylation activity of T4 polynucleotide kinase, the protein is also a 3’-phosphatase (11, 19, 3 1 ) . The enzyme’ catalyzes the hydrolysis of 3’-phosphoryl groups of deoxynucleoside 3‘monophosphates, deoxynucleoside 3’ ,5’-diphosphates, and of 3’-phosphoryl polynucleotides to yield inorganic orthophosphate and a 3‘hydroxyl group (Fig. 2).

B. PHOSPHORYLATION OF 5’-HYDROXYL TERMINI 1. Acceptor Substrutes

A variety of nucleic acid compounds can be phosphorylated in the pol ynucleotide kinase reaction provided they have a nucleotide bearing a free 5’-hydroxyl group with a phosphoryl group esterified at the 3‘ position (Table I I , 2 4 , 8, 28). Thus, the substrates include DNA and RNA, oligonucleotides, and nucleoside 3‘-monophosphates. All terminal 5’nucleotides found in DNA or RNA can be phosphorylated. The enzyme also catalyzes the phosphorylation of nucleotides whose bases contain chemically protected groups (32), such as those used in the chemical synthesis of polynucleotides . The apparent Michaelis constants and V,,, values for oligonucleotides and 3‘-mononucleotides vary depending on the 28. 29. 30. 3I . 32.

B. Weiss and C. C. Richardson, C S H S Q B 31, 471 (1966). J . H. Van de Sande, K. Kleppe, and H. G . Khorana, Biochemistry 12, 5050 (1973). ti. Chaconas, J. H. van de Sande, and R. B. Church, BBRC 66,962 (19753. A. Becker and J. Hurwitz, JBC 242, 936 (1%7). J. H. van de Sande and M. Bilsker, Biochemistry 12, 5056 (1973).

306


0

k

R

FIG. 1. The transfer of phosphate from a nucleoside 5’-triphosphate to the 5’-hydroxyl group of an acceptor molecule by polynucleotide kinase. B = adenine, guanine, cytosine, thymine, or uracil; R = H, a nucleoside, a nucleotide, or a polynucleotide: R’ = H or OH. identity of the 5’4erminal base residue and the length of the oligonucleotide (33). The apparent Michaelis constant for 3’-mononucleotides is between 22.2 and 143 p M ;the apparent Michaelis constant for micrococcal nuclease-treated DNA, the substrate normally used to follow the purification, is 7.6 pM (33). Similar specificities have been found with the T2 polynucleotide kinase (3, 4). However, the kinase isolated from rat liver nuclei cannot phosphorylate RNA molecules, nor can it phosphorylate oligodeoxynucleotides of chain length less than 10 to 12 residues (7). Polynucleotide kinase catalyzes the phosphorylation of high molecular weight single-stranded and double-stranded DNA molecules in which the 5’-hydroxyl groups are in a variety of configurations. For example, it is possible to phosphorylate all of the 5’-hydroxyl groups (produced by alkaline phosphatase) in a preparation of bacteriophage T7 DNA (9, 3 4 ) , a duplex molecule consisting of 40,000 base pairs with blunt ends. However, the quantitative phosphorylation of 5’-hydroxyl groups located at single-strand interruptions in duplex DNA (nicks) is extremely difficult (10); even with prolonged incubation and large amounts of enzyme, it is not possible to phosphorylate more than 70% of such internally located groups. In order to obtain a more quantitative analysis of the phosphorylation of these large substrates, several synthetic double-stranded DNAs containing defined 5’-hydroxyl end groups have been used as substrates in the kinase reaction (35). The rate of phosphorylation of 5’-hydroxyl groups located at gaps in duplex DNA is approximately 10-fold slower than for 5‘-hydroxyl groups on the corresponding single-stranded DNA. With high concentrations of ATP complete phosphorylation of 5’-hydroxyl groups at 33. J. R. Lillehaug and K. Kleppe, Biochemisrry 14, 1221 (1975). 34. B. Weiss and C. C. Richardson, JMB 23, 405 (1967). 35. J. R. Lillehaug, R. K. Kleppe, and K. Kleppe, Biochemistry 15, 1858 (1976).

307

16. BACTERIOPHAGE T4 POLYNUCLEOTIDE KiNASE

+

0

It

HO-P-O-

I -0

-0

FIG. 2. The hydrolysis of a 3'-phosphoryl group by the 3'-phosphatase activity of polynucleotide kinase. B = adenine, guanine, cytosine, or thymine; R = H, PO,2-, a nucleotide, or a polynucleotide.

gaps can be achieved, but not at nicks. The 5'-hydroxyl groups on protruding single-stranded ends of duplex molecules are more readily phosphorylated than are 5'-hydroxyl groups on blunt ends or on molecules with protruding 3'-hydroxyl group ends (recessed 5'-hydroxyl groups). However, with sufficiently high concentrations of ATP the 5'-hydroxyl termini in all of these latter substrates can be completely phosphorylated. With the exception of DNA substrates that contain protruding 5'-hydroxyl TABLE I1 SUBSTRATES FOR POLYNUCLEOTIDE

KINASE

Substrate DNA 5'-OH-terminated T7 DNA (micrococcal nuclease treated) 5'-OH-terminated E. coli DNA (micrococcal nuclease treated) RNA E . coli tRNA (phosphatase treated) Oligonucleotides d(TpTpTpTp) d(ApApApAp) d(GpGpG p) d(CpCpCp) uPU APA

CPC GpCl

3'-AMP 3'-dAMP

Rate" (nmoVmidmg)

2200 2500 1700 2100 2300 2400 2000 2000 2600 2700 2600 2200 2900

" The rate of reaction was measured in the standard reaction mixture using Fraction VI of T4 pol ynucleotide kinase.

308


ends, the phosphorylation reaction is inhibited by the addition of excess KCI (-35).

5' -Triphosphu tes A variety of nucleoside 5'-triphosphates can function as the phosphate donor. Although ATP is routinely used in the standard assays, CTP, UTP, GTP, dATP, and dTTP have all been found to function equally well. The apparent Michaelis constant for ATP is between 1.4 x M and 1.3 x W5 M depending on the DNA substrate used (4, 30). The apparent Michaelis constants for ATP, UTP, GTP, and CTP in the T2 polynucleotide kinase reaction, using micrococcal nuclease-treated DNA, are 1.4, 1.5, 3.3, and 2.5 x M , respectively ( 4 ) . 2.

Nil cleoside

3. Effect of p H mid Divaleiit Cutioiu The optimal pH range for the enzyme is 7.4 to 8.0 in Tris-HC1 buffer with maximal activity obtained at pH 7.6 (2). In the absence of added MgCl, there is no detectable activity. The optimal Mg2+concentration at pH 7.6 is 1 x M ( 2 ) . Mn2+can partially fulfill the metal requirement; at the optimal concentration of 3.3 x M , 50% of the maximal activity obtained with Mg2+is observed.

4. Sidjliydryl requirement^ Maximal activity is obtained with 5 mM dithiothreitol in the reaction mixture ( 8 ) . Protection is also achieved with 10 mM 2-mercaptoethanol and 10 mM glutathione ( J ) , but only 80 and 70%, respectively, of that observed with dithiothreitol. In the absence of a sulfhydryl compound, there is 2% of the optimal activity.

5 . liihibitov and Stirnulatory Fuctors Phosphate and pyrophosphate anions are inhibitory to T2 and T4 polynucleotide kinase (2, 4, 27). At pH 7.6 in either 70 mM sodium or potassium phosphate buffer, 5% of the value observed in Tris-HC1 buffer is obtained. Furthermore, the addition of inorganic phosphate or pyrophosphate to the standard assay mixture containing Tris-HC1 buffer results in inhibition; 20 mM inorganic phosphate or 5 mM pyrophosphate give approximately 50% inhibition (27). Since phosphate ions are relatively more inhibitory to E. coli alkaline phosphatase, at appropriate concentration of phosphate ion, polynucleotide kinase can phosphorylate polynucleotides in the presence of E . coli alkaline phosphatase (22; see also 4 8 ) . This is a convenient procedure for end-group labeling in which prior dephosphorylations by phosphatase is required. A variety of salts, such as NaCl, stimulate the reaction when single-


309

stranded DNA substrates are used (27). However, as previously discussed, these salts inhibit the phosphorylation of 5’-hydroxyl groups in certain duplex substrates. Polyamines such as spermine also stimulate the reaction: spermine at 1.7 mM stimulates the rate of phosphorylation approximately 30-fold (27).

c.

REVERSALOF THE KlNASE REACTION

1. Dephosphorylation of Polynucleotides

In the presence of ADP, polynucleotide kinase catalyzes the dephosphorylation of a 5’-phosphoryl group of a polynucleotide to yield a 5 ’ hydroxyl-terminated polynucleotide and ATP in a reversal of the phosphorylation reaction (29). The optimal pH for dephosphorylation is 6.2 in imidazole-HC1 buffer (14) as compared to the optimal pH for phosphorylation of 7.6 in Tris-HCI buffer. A direct comparison of the kinetic parameters of phosphorylation and dephosphorylation using 5’-protruding ends of fragments produced by EcoRI endonuclease show a striking difference in the two reactions (14). The apparent Michaelis constants for ATP and ADP in the phosphorylation and dephosphorylation reactions, respectively, are 4 ph4 and 200 p M : the corresponding V,,, are 3 1.3 and 4.3 pmol/min/pg of enzyme. The apparent Michaelis constant for ADP using a single-stranded oligonucleotide substrate is 0.22 p M (29). Even with large amounts of enzyme, however, only partial dephosphorylation is obtained (14, 29). 2. Erchcinge Retictioil Due to the reversibility of the reaction, in the presence of ADP polynucleotide kinase catalyzes an exchange of 32Pbetween the y-phosphate of ATP and the 5’-phosphoryl group of a polynucleotide (29). This exchange reaction provides a convenient method for labeling the 5’ termini of polynucleotides radioactively without prior removal of existing 5‘phosphoryl groups ( M , 29, 30). Like the forward reaction, the exchange reaction requires Mg2+ and 2-mercaptoethanol for optimal activity (29).The exchange reaction is also inhibited by phosphate ions: 50 mM potassium phosphate results in a 60% inhibition (29). The optimal pH is 6.2 in imidazole-HC1 buffer (29). The optimal rate of exchange of 3aPiinto the termini of EcoRI DNA fragments at pH 6.6 occurs at an ATP concentration of 10 p M , and an ADP concentration of 300 F M (14). The 5‘-phosphoryl end groups in a variety of polynucleotides can be labeled radioactively using the exchange reaction. Although the labeling

3 10


of 5'-phosphoryl termini by the exchange reaction is usually incomplete, the extent of exchange is proportional to the number of such groups for a given amount of polynucleotide kinase (14). Furthermore, there is no significant effect of molecular weight of the acceptor nucleic acid from 4 X lo5 to 14 x lo6 on the rate and extent of labeling (14). Single-stranded oligonucleotides are most easily labeled in the exchange reaction: a 5'phosphorylated nonacosanucleotide is labeled to an extent of 96% (29). The 5'-phosphoryl recessive termini are more difficult to label, although with sufficient amount of enzyme 70% of the termini are labeled (14). The inaccessibility of 5'-hydroxyl groups at nicks to phosphorylation holds true for 5'-phosphoryl groups in the exchange reaction. The 5'-phosphoryl groups located at nicks in duplex DNA are labeled in the exchange reaction 30-fold less efficiently than 5'-phosphoryl protruding ends (14). The exchange reaction presents difficulties if one wishes to label 5'hydroxyl termini specifically. This problem can be circumvented by carrying out the phosphorylation reaction at @', a temperature at which the rate of the exchange reaction is greatly reduced (36). At 0" the exchange rate is &th of the rate at 37", whereas the rate of phosphorylation of 5'-hydroxyl termini at 0" is Ath the rate at 37". In the absence of added ADP an exchange of 32P between 5'-32Pphosphoryl groups of a polynucleotide and ATP is observed (29). The radioactive products formed are inorganic phosphate, ATP, and adenosine 5'-tetraphosphate. Although it has been proposed that the formation of Pi and adenosine 5'-tetraphosphate reflect a phosphorylated kinase intermediate (291, no such intermediate has been isolated. D. 3'-PHOSPHATASE ACTIVITY Polynucleotide kinase, in addition to catalyzing the phosphorylation of 5'-hydroxyl groups in nucleic acids, also catalyzes the hydrolysis of 3'phosphoryl groups of deoxynucleotides and other nucleic acids (Fig. 2). Initially, a 3'-phosphatase was identified and purified from E. coli cells infected with bacteriophage T4 (31 ). The purified enzyme was found to catalyze the removal of 3'-phosphoryl groups from deoxyribonucleotides, oligodeoxyribonucleotides,and DNA. The enzyme had no effect on 3'ribomononucleotides, 3'-phosphoryl groups of RNA, or any 5'-phosphate esters (31). The optimal pH of the 3'-phosphatase was between 5.8 and 6.2 in Tris-maleate buffer. In the absence of added MgClz, only 2% of maxi36. ( 1975).

R. Okazaki, S . Hirose, T. Okazaki, T. Ogawa, and Y. Kurosawa, BBRC

62, 1018


31 1

ma1 activity was observed: the optimal magnesium concentration was 8 mM ; and 2-mercaptoethanol , glutathione, or dithiothreitol were required for sustained activity (31). Subsequently, purified preparations of polynucleotide kinase were found to contain a 3’-phosphatase activity (19). The kinase and phosphatase activities purified together through all stages of purification, and the two activities migrated together as the major protein band during electrophoresis in polyacrylamide. These and other studies, including heat inactivation, strongly suggest that the 3’-phosphatase and kinase activities reside in the same protein molecule (19). Is the 3’-phosphatase of polynucleotide kinase identical to the previously described T4 3’-phosphatase? The two 3’-phosphatase activities have similar requirements. Both require magnesium or cobalt ion, and both have an optimal pH around 5.9 (19,31).Their substrate requirements are similar in that both activities can hydrolyze deoxynucleoside 3’monophosphates, but not 3’-ribonucleoside 3'-monophosphates . The single difference in their substrate specificity is the finding that the 3’phosphatase of polynucleotide kinase can use 3’-phosphoryl oligoribonucleotides as a substrate (19), wherea.; the previously described T4 3‘phosphatase was reported to be unable to do so (31). Additional evidence, however, for the identity of the two 3‘phosphatases comes from genetic data. A mutant of phage T4, T4 pseT, fails to induce the T4 3’-phosphatase (37) as measured by the standard 3’-phosphatase assay (31). This activity represents essentially all of the 3’-phosphatase activity found in extracts of bacteriophage TCinfected cells. Analysis of the mutant reveals that it also fails to induce polynucleotide kinase (11). A T4 mutant lacking polynucleotide kinase (38) is similarly defective in inducing the 3’-phosphatase activity (11). Specific mutations in the phage pseT gene lead to an altered polynucleotide kinase that lacks the 3’-phosphatase activity (11, 20). Whereas, mutations in thepseT gene may affect both the kinase and the phosphatase activities, polynucleotide kinase, free of the 3’-phosphatase activity, can be prepared by purifying the enzyme from E. coli cells infected with T4pseTl (20). This phage mutant induces normal amounts of kinase activity but greatly reduced levels of 3’-phosphatase (11, 20, 37). The ratio of the two activities in a wild-type enzyme can be changed significantly by inactivation of one, but not the other, activity by heating in the presence of specific substrates or products of the two reactions (19). 37. R . E. Depew and N . R. Cozzarelli, J . Viral. 13, 888 (1974). 38. V. L. Chan and K. Ebisuzaki, Mol. G e n . Genet. 109, 162 (1970).

3 12 IV.


Role of Polynucleotide Kinase in Vivo

The early appearance (2, 3 ) and specificity of polynucleotide kinase, 3’-phosphatase suggest that it may be involved in the metabolism of T4 DNA. One can envision a number of roles in vivo for the enzyme on the basis of the reactions catalyzed in virro. The two activities of the enzyme would assure the presence of 3’-hydroxyl and 5’-phosphoryl groups on nucleic acid molecules; such groups are a prerequisite for most enzymatic reactions, such as ligation, involving the termini of polynucleotides. One could postulate that the two activities reside on the same polypeptide in order to carry out the simultaneous removal of a 3’-phosphoryl group and the phosphorylation of the 5’-hydroxyl group of a given molecule. For example, the two activities are capable of converting deoxyribonucleoside 3’-monophosphates to deoxyribonucleoside 5’-monophosphates, the precursors for DNA synthesis. However, it is not known whether such 3’phosphoryl-terminated mono- and polynucleotides exist in vivo . The most direct and definitive approach to determine the role of the enzyme is to examine the biochemical consequences of mutations that directly affect the enzyme. The first mutants of phage T4 found to lack polynucleotide kinase were found by direct assay of polynucleotide kinase in a variety of T4 strains that harbored other mutations (28). However, after the mutation affecting the kinase was separated from other mutations, the mutants had no observable phenotype, and hence the mutations were not mapped, and the mutants were not subjected to further study. The first detailed analysis of the polynucleotide kinase gene came, unknown, from the isolation and characterization of T4 mutants that lacked 3’phosphatase activity. The subsequent finding that 3’-phosphatase and polynucleotide kinase activities purify together (19) led to the demonstration that both activities are coded by the same structural gene (20, 37). Mutants of T4 that lack 3’-phosphatase activity were initially isolated by direct assay of 3‘-phosphatase activity in extracts of E. coli cells infected with mutagenized T4 phage (37). One such mutant, T4pseTf, grew normally on wild-type E. coli and exhibited no observable phenotype on a variety of E. coli mutants in which one could postulate a requirement for the phosphatase. However, T4pseTf failed to produce viable phage (less than 2% of a wild-type infection) on a TCsensitive Hospital strain, E. coli CT196 (37). Using this phenotype, a secondpseT mutant was isolated as well as an extracistronic suppressor T4 mutation (sfp). ThepseT gene lies between genes 63 and 31 (37).A fine structure map of the gene has been described ( I f ) , and a restriction enzyme map of this region has been obtained (25). The srp gene lies in the nonessential region between rZZB and ac (37).

16.

BACTERIOPHAGE T4 POLYNUCLEOTIDE KINASE

313

What is the biochemical manifestation of enzyme deficiency in T4pseTinfected CT196 cells that accounts for the lack of phage growth? First, the growth defects are accompanied by a 50% reduction in the rate of T4 DNA synthesis and a decrease in the length of the DNA product as measured on an alkaline sucrose gradient (37). Relatively little DNA is packaged to yield mature phage particles. Furthermore, there is a dramatic reduction in the rate of late T4 gene expression (true-late gene expression) in T4 pseT-infected E. coli CTr5X (a hybrid between K12 strains and CT196) (11 ). Hybridization experiments suggest that the defect is at the transcriptional level ( I / ) and hence the kinase, 3‘-phosphatase is probably required for normal T4 true-late transcription. The relationships, however, between these multiple defects is not known. No information is available concerning the mechanism by which the lack of kinase or 3’phosphatase activity leads to these abnormalities.

V.

Research Applications

Polynucleotide kinase is an indispensable enzyme in studies on nucleic acid structure and metabolism. Its primary value derives from its ability to phosphorylate specifically 5’-hydroxyl termini in RNA and DNA molecules. Although it is not feasible to enumerate all of the many applications of the enzyme, it is possible to cite a few that illustrate its past and present usefulness as a reagent in the study of nucleic acids. Initially the ability to label specifically 5’-termini in polynucleotides radioactively provided the only method to determine the number and identity of such end groups in polynucleotides of high molecular weight or in molecules that could not be obtained in sufficient quantities for analysis by other methods (8, 28). Similarly, this technique was useful in characterizing phosphodiester bond cleavages introduced into DNA by shearing or nuclease action (8, 28). In studies of this type it is usually necessary to distinguish between, and to determine, the number of 5’-hydroxyl and 5’-phosphoryl end groups. For this reason E. coli alkaline phosphatase is frequently used in conjunction with polynucleotide kinase. An analysis of the extent of phosphorylation of denatured nucleic acid before and after phosphatase treatment makes possible the determination of the relative number of 5’-hydroxyl and 5’-phosphoryl end groups in a preparation. Furthermore, under appropriate conditions these two enzymes can be used to determine the number and identity of external and internal (located at nicks) 5‘-end groups in DNA (10). Polynucleotide kinase is often used to prepare radioactively labeled

3 14


molecules that can be used as substrates to identify or to characterize other enzymes. For example, assays for DNA and RNA ligase make use of polynucleotide substrates containing 5’-32P-phosphorylend groups (394 1 ) . Likewise, 5’-32P-labeledRNA and DNA have been used to characterize the 5’ + 3‘ hydrolytic activity ofE. coli DNA polymerase 1(42-44) and a TCinduced 5’-phosphatase (31). The enzyme has been an important reagent in the chemical synthesis of double-stranded DNAs of specific nucleotide sequence. For example, the synthesis of the structural genes for an alanine and tyrosine tRNA make extensive use of end group labeling, both to monitor and to achieve the joining of fragments (45, 46). Another use of polynucleotide kinase and end group labeling is to prepare radioactive polynucleotide molecules that cannot be labeled unifonnlyin vivo or obtained in sufficient amounts to make possible biochemical studies by methods other than those involving radioactive labeling. Shortly after its discovery the kinase was used to label digests of DNA and RNA radioactively in order to obtain fingerprints (47). Similarly, the physical mapping of restriction enzyme fragments is frequently carried out using polynucleotide kinase to label the 5’ termini of the fragments (48). Currently the most extensive use of polynucleotide kinase is in the sequencing of RNA and DNA molecules (49-51 ). Terminally labeled polynucleotides can be sequenced rapidly using partial, base-specific cleavages by chemical or enzymatic procedures such that the length of the radioactively labeled fragments identify the positions of the specific bases at which each cleavage occurs.

39. C. C. Richardson, Antiu. Rev. Biochem. 38, 795 (1%9). 40. R. Silber, V. G. Malathi, and J. Hurwitz, PNAS 69, 3009 (19721. 41. A. Sugino, T. J. Snopek, and N. R. Cozzarelli, JBC 252, 1732 (1977). 42. R. P. Klett, A. Cerami, and E. Reich, PNAS 60, 943 (1968). 43. M. P. Deutscher and A. Kornberg, JBC 244, 3029 (1969). 44. Y. Masamune, R. A. Fleischman, and C. C. Richardson,.UK 246, 2680 (1971). 45. H.G. Khorana, K. L. Agarwal, H. Biichi, M. H. Caruthers, N. K. Gupta, K. Kleppe, A. Kumar, E. Ohtsuka, U. L. Raj Bhandary, J. H. van de Sande, V. Sgaramella, T. Terao, H. Weber, and T. Yamada, JMB 72, 209 (1972). 46. E . L. Brown, R. Belagaje, M. J. Ryan, and H. G. Khorana, “Methods in Enzymology” Vol. 68, p. 109, 1979. 47. M. Szekely, Proced. Nucleic Acid Res. 2, 780 (1971). 48. G. Chaconas and J. H. van de Sande, “Methods in Enzymology” Vol. 65, p. 75, 1980. 49. H. Donis-Keller, A. M. Maxam, and W. Gilbert, Nucleic Acids Res. 4, 2527 (1977). 50. A. Simoncsits, G. G. Brownlee, R. S. Brown, J. R. Rubin, and H. Guilley, Nature (London) 269, 833 (1977). 51. A. M. Maxam and W. Gilbert, “Methods in Enzymology” Vol. 65, p. 499, 1980.

Eukaryotic DNA Kinases STEVEN B . ZIMMERMAN BARBARA H . PHEIFFER

I . Introduction and Perspectives . . . . . . . . . . . . . . . . . . I1 . Purification and Properties . . . . . . . . . . . . . . . . . . . .

A . Purification . . . . . . . . . . . . . . . . . . . . . . . . . B . Physical Properties . . . . . . . . . . . . . . . . . . . . . 111. The Catalytic Reaction . . . . . . . . . . . . . . . . . . . . A . Description of the Reaction . . . . . . . . . . . . . . . . . . B . Assay Procedures . . . . . . . . . . . . . . . . . . . . . . C. Stoichiometry and Identification of Products . . . . . . . . . . D. Requirements for Activity . . . . . . . . . . . . . . . . . . E . Reversal of the Reaction and Labeling by Exchange . . . . . . F. Kinetics and Mechanism . . . . . . . . . . . . . . . . . . . G . Inhibitors . . . . . . . . . . . . . . . . . . . . . . . . . IV. Comparison of the DNA Kinases with RNA Kinase and Polynucleotide Kinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . V . Biological Role . . . . . . . . . . . . . . . . . . . . . . . . . VI . Research Applications . . . . . . . . . . . . . . . . . . . . . Note Added in Proof . . . . . . . . . . . . . . . . . . . . . .

1

.

315 316 316 318 318 318 318 319 320 322 324 324 326 327 329 329

Introduction and Perspectives

DNA kinase activity from a eukaryotic source was first demonstrated by Novogrodsky ef (11 . ( 1 ) . Extracts of rat liver nuclei were shown t o transfer phosphate groups from ATP to 5'-hydroxyl termini in DNA . Since then. enzymes with this activity have been partially purified from 1 . A . Novogrodsky. M . Tal. A . Traub. and J . Hurwitz. JBC 241. 2933 (1966)

315

.

THE ENZYMES Vol XIV Copynght @ 1981 by Academic Press. lnc All rights of reproduction in any form reserved ISBN 0-12-122714-6

3 16

S . B . ZIMMERMAN AND B . H. PHEIFFER

rat liver (2, 3 ) and calf thymus ( 4 ) . The DNA kinase from rat liver has proved to be highly specific for DNA ( 3 ) .A preparation of an enzyme from calf thymus with otherwise quite similar properties shows low activity on RNA chains in addition to its activity on DNA (4).In addition, a relatively specific RNA kinase has been partially purified from HeLa cell nuclei (5 ). The restricted acceptor specificity of these eukaryotic enzymes may be contrasted to the broad specificity of the polynucleotide kinase from T2-, T4-, or T6-infected Escherichicr coli, which has comparable activities on both DNA and RNA as well as on oligonucleotides and even on 3’rnononucleotides (6, 7). In this chapter, we will describe the properties of the eukaryotic DNA kinases and contrast them with those of the RNA kinase and of the polynucleotide kinase. The polynucleotide kinase is reviewed by Richardson in this volume (7); both DNA kinase and polynucleotide kinase have been reviewed by Kleppe and Lillehaug (6).

II. Purification and Properties

A. PURIFICATION The most thoroughly characterized DNA kinase is the activity from nuclei of rat liver (1-3,8). This enzyme has been partially purified by two independent procedures. Both preparations start by isolating nuclei. The enzyme is extracted from the nuclei with 0.15-0.2 M NaCl. Subsequent steps in the procedure of Teraoka et ul. ( 2 ) include removal of inactive materials by precipitation at pH 5, gradient elution from a phosphocellulose column, and finally gel filtration on Sephadex G-150. In the procedure of Levin and Zimmerman (-?I the nuclear extract is subjected to gradient elution from a phosphocellulose column and stepwise elution from a sulfopropyl Sephadex column. The specific activities of the final fractions from either procedure are similar and correspond to approximately a 1000-fold purification relative to a crude extract of rat liver. Two useful aspects of the procedure of Levin and Zimmerman (-3) may be 2. 3. 4. 5. 6. 7. 8.

H. Teraoka, K. Mizuta, F. Sato. M . Shimoyachi, and K. Tsukada, EJB 58,297 (1975). C. J. Levin and S. B. Zimmerman, JBC 251, 1767 (1976). G . E. Austin, D. Sirakoff, B . Roop, and G . H. Moyer, BBA 522, 412 (1978). S. Shuman and J. Hurwitz, JBC 254, 10396 (1979). K. Kleppe and J. R . Lillehaug, Adiwn. En:ymo/. 48, 245 ( 1979). C. C. Richardson, this volume, chap. 16. M . Ichimura and K. TsukadaJ. Biochern. (Tokyo)69, 823 (1971).

17. EUKARYOTIC DNA KINASES

3 17

mentioned. First, the phosphocellulose fraction has proved to be very stable, losing little or no activity for periods of 4-6 months at 4" (9); this fraction has been used extensively to characterize the activity. Second, a highly purified DNA ligase is also obtained in separate fractions of the phosphocellulose chromatography (10 ). Although preparations from both procedures are heterogeneous based upon their gel electrophoresis patterns ( I I ) , they are relatively free from interfering activities, as implied by their use for labeling the 5'-hydroxyl termini at single-strand interruptions (nicks) within duplex DNA (8, 10). More direct assays have indicated a general lack of contamination with nuclease, phosphodiesterase, or DNA ligase activities (2, 3 ) ; no phosphatase activity was detected on p-nitrophenyl phosphate (2) or 5 ' phosphate groups in DNA ( 3 ) . A very low level of nuclease activity on denatured DNA can be demonstrated with concentrated samples of the phosphocellulose fraction (/I); this activity, which is apparently the enzyme described by Cordis et d.(/.?), can be abolished with little loss of kinase activity by substituting Ca*+(0.01 M ) for Mg2+in the kinase incubation mixture (/I). A DNA kinase activity has been partially purified from homogenates of calf thymus ( 4 ) . The procedure involved protamine sulfate precipitation of inactive materials, ammonium sulfate fractionation, gradient elution from columns of phosphocellulose, hydroxyapatite and sulphopropyl Sephadex , and finally centrifugation through a glycerol gradient. The final fraction was about 1600-fold purified relative to the crude extract. The purified fractions were relatively unstable. DNA ligase, nuclease, and phosphatase (on 5'-phosphate groups in DNA) were not detected in these fractions. DNA kinase has also been partially purified from extracts of Chinese hamster lung cells grown in tissue culture (II). Extracts of washed cells in 0.2-0.4 M NaCl were made by several cycles of freezing and thawing. Diluted extracts were subjected to the same phosphocellulose chromatography that was used for the rat liver enzyme (3).A single peak of activity with the characteristic inhibition by inorganic sulfate (see Section I11 ,G,3) appeared at the same place in the gradient as did the enzymes from liver (2, 3 ) or calf thymus (4). 9. The sulfopropyl Sephadex fraction of this procedure has a half-life of a few weeks U). Both the phosphocellulose and Sephadex fractions of Teraoka et a / . are stated to be stable for at least a week at 0-4" (2). 10. S. B . Zimmerman and C. J. Levin,JBC 250, 149 (1975). 1 1 . S . B . Zimmerman, C. J . Levin, and B . H . Pheiffer, unpublished results. 12. G . A . Cordis, P. J. Goldblatt, and M. P. Deutscher, Biochemisrry 14, 2596 (1975).

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S. B. ZIMMERMAN AND B. H. PHEIFFER

B. PHYSICAL PROPERTIES The molecular weight of the DNA kinase from rat liver has been estimated at 8 x lo4 based upon its gel filtration properties (2, 3 ) . A sedimentation coefficient ( s ~ , , ,=~ 4.4) was determined by Teraoka et al. (2). The enzyme from calf thymus is similar in size. A molecular weight of 7 x lo4 was estimated from a sedimentation coefficient = 4.3) and a Stokes radius from gel filtration of 3.9 nm, using an assumed value for the partial specific volume (4). 111.

The Catalytic Reaction

A. DESCRIPTION OF THE REACTION DNA kinase catalyzes the reversible transfer of a phosphate group between a nucleoside triphosphate and the 5’-hydroxyl moiety at the terminus of a DNA chain ( 2 - 4 ) NTP

+ 5’-hydroxyl terminus in DNA e NDP + 5’-phosphate terminus in DNA

(1)

Although the reaction is customarily assayed in the forward direction with ATP as the phosphate donor, studies of the specificity of the reverse eaction (13) indicate that the enzyme can use a number of other nucleotides. DNA is indicated as the phosphate acceptor in Eq. (1); the kinase from calf thymus may also have limited activity on 5’-hydroxyl termini of RNA chains (4).

B. ASSAYPROCEDURES The routine assay for DNA kinase measures the rate of transfer of the radioactive phosphate group of [y-32P]ATPinto an acid-insoluble form in the presence of a DNA acceptor containing 5’-hydroxyl termini ( 1 - 4 , 8). In preparation for its use as an acceptor, the DNA is either partially digested with pancreatic DNase to form 5’-phosphate termini followed by treatment with a phosphatase to yield 5’-hydroxyl termini, or is partially digested with micrococcal nuclease, which directly yields 5’-hydroxyl groups. Estimates of DNA kinase activity in crude extracts of cells or nuclei should be evaluated cautiously. Other enzymes can transfer the terminal phosphate of ATP to acid-precipitable acceptors that may be present in 13. B. H. Pheiffer and S. B. Zimmerman, Biochemistry 18, 2960 (1sv79).


3 19

crude extracts. The characteristic inhibition of DNA kinase by relatively low concentrations of inorganic sulfate (see Section III,G,3) may prove useful in such situations. Other assays have been used for special purposes. For example, the rate of phosphorylation of relatively low molecular weight acceptors that are not acid-precipitable may be followed by adsorbing them to charcoal ( I , 3 ) . Also, the reverse reaction may be assayed by the rate of the nucleoside diphosphate-dependent release of radioactivity from [5'-3'P]phosphate termini in DNA into an acid-soluble form (13).

c.

STOICHIOMETRY AND IDENTIFICATION OF

PRODUCTS 1.

The Forward Reuctioti

The labeled product of phosphorylation of DNA with [y-32P]ATPby the rat liver DNA kinase was characterized as a 5'-phosphate terminus of a DNA chain by a number of criteria. As expected for such a product, the label was rendered acid-soluble by treatment with E. coli alkaline phosphatase or pancreatic DNase (3, 8) but not by treatment with pronase, pancreatic RNase, or alkali ( 3 ) .Combined treatment with a phosphodiesterase plus 5'-nucleotidase yielded 32Pi, implying the formation of a labeled 5'-phosphate group (2). The product of the kinase and [y-32P]ATP on dephosphorylated nicked DNA was characterized in some detail. When treated with pancreatic DNase and venom phosphodiesterase, the radioactivity was quantitatively recovered in the four isolated 5'deoxynucleoside monophosphates, clearly indicating the formation of 5 ' phosphate groups by the kinase (10). Further, the labeled product was sealed into phosphodiester linkage by DNA ligase from either rat liver (10, 14) or E. coli (11); these enzymes require the presence of a 5'-phosphate group for their action. The other product of the forward reaction was identified as ADP by its cochromatography with an ADP standard on a DEAE-cellulose column. The amount of ADP formed approximately matched the amount of DNA phosphorylated, consistent with the reaction as written in Eq. (1) ( 2 ) . The product formed by phosphorylation of DNA with the calf thymus enzyme was also identified as a phosphate terminus (4), based upon the sensitivity of the product to degradation by pancreatic DNase, micrococcal nuclease, or E. coli alkaline phosphatase. The radioactivity was released in a form not adsorbed by charcoal after combined treatment with 14. K . Tsukada and M . Ichimura, BBRC 42, 1156 (1971).

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S. B. ZIMMERMAN AND B. H . PHEIFFER

venom phosphodiesterase and 5’-nucleotidase, as expected for a 5 ’ nucleotide product. The acid-precipitable product of the calf thymus enzyme on RNA was identified by its solubilization by pancreatic RNase or NaOH, but not by pancreatic DNase or pronase. 2 . The Reverse Recictioti The reverse reaction catalyzed by rat liver DNA kinase releases radioactivity from DNA bearing [5’-32P]phosphatetermini in the presence of a suitable nucleoside diphosphate ( I S ) . The acid-soluble product of the reverse reaction with ADP and P2P]DNA was identified as PPIATP by its cochromatography with authentic ATP on a DEAE-cellulose column. The 32P released from the DNA was quantitatively accounted for by the amount of [32P]ATPformed. No significant radioactivity was associated with Pi, AMP, ADP, or adenosine tetraphosphate. FOR ACTIVITY D. REQUIREMENTS

1. p H

The rates of the forward (2, 3 ) and reverse reactions (I.?) of the DNA kinase from rat liver have a similar sharp optimum at about pH 5.5, with diminished but significant activity up to at least pH 8 (Fig. 1). The rate of the forward reaction of the calf thymus enzyme has a similar pH dependence ( 4 ) . 2 . Divu I en t Cations The DNA kinase from both rat liver and calf thymus requires a divalent cation for significant activity. For the rat liver enzyme ( 2 , 3 ) ,a number of metals support kinase activity (Mg2+,Mn’+, Co2+,Zn2+,Ni2+,and Ca2+), whereas Cu2+is inhibitory. With the calf thymus enzyme (4),Mg2+, Mn2+, and Zn2+were shown to be active. 3 . Specijicity .for Nucleoside Triphosphrites ATP is the only nucleoside triphosphate that has been shown to be active in the forward reaction of the DNA kinase ( K , = 2 4 y M ) (/-4). However, the wide range of nucleoside diphosphates that are active in the reverse reaction ( I 3 1 suggests that all of the common ribo- and deoxyribonucleoside triphosphates can participate in the forward reaction. The forward reaction with labeled ATP is inhibited by the presence of these unlabeled ribo- and deoxyribonucleoside triphosphates in a manner consistent with such a broad specificity for the phosphate donor ( 3 , 4 ) .

32 1


PH

FIG. 1. Effect of pH on rat liver DNA kinase. Open and closed circles indicate Trismaleate and sodium succinate buffers, respectively. From Levin and Zimmerman ( 2 ) , reproduced with permission.

4. Specificity .for the Phosphate Acceptor

The first descriptions of DNA kinase activity in crude extracts did not define its ability to phosphorylate other than DNA acceptors (I, 8). Subsequent detailed studies indicated that the DNA kinase purified from rat liver is specific for DNA (and long oligodeoxynucleotides) and has little or no activity on RNA (3, 15). In these studies, all of the potential substrates tested were also assayed with the polynucleotide kinase from T4-infected cells (7) to ensure that negative results with the rat liver enzyme were a function of the specificity of that enzyme, and not due to a defect in the substrate. a. DNA versus R N A . The routine substrate for the DNA kinase from rat liver is DNA that has been enriched in 5’-hydroxyl termini (see Section 15. Teraoka et r d . ( 2 ) cite preliminary results with the purified enzyme from rat liver which indicated to them that RNA can act as a phosphate acceptor. In the absence of their experimental results and in view of the experimental evidence cited here in opposition to their conclusion, it is our opinion that the rat liver DNA kinase is indeed highly specific for DNA relative to RNA.

322

S . B. ZIMMERMAN A N D B. H. PHEIFFER

111,B). In contrast to its activity on such DNA preparations, the DNA kinase was inactive on similarly treated samples of RNA or of poly(rA) (Table I) (3, 15). The lack of activity on these materials was not due to contaminating nucleases that destroyed either the phosphate acceptors or the phosphorylated products. Such possibilities were ruled out by the results of sequential incubations of the rat liver and T4 kinases. The lack of activity of the rat liver enzyme on RNA was also maintained in the presence of divalent cations other than Mg2+, as well as under the T4 kinase assay conditions (3). The calf thymus preparation differed from that of rat liver in that the thymus preparation phosphorylated RNA containing 5'-hydroxyl termini at about 10% of the rate with which it acted upon DNA (4). This apparent difference in acceptor specificity is further discussed in Section IV. The rat liver kinase phosphorylates 5'-hydroxyl termini in DNA that are joined to any of the four usual bases (see Section III,C, 1). Further, any of these four bases can also be present on the 3'-hydroxyl side of a nick that is being phosphorylated (10). Both native and denatured DNA were generally phosphorylated at similar rates (Table I) and to similar extents, except that the extent of phosphorylation of micrococcal nuclease-treated DNA increased greatly upon denaturation, suggesting an inhibition by 3'-phosphate groups adjacent to a site of phosphorylation (3). b . Nucleotides and Oligonucleotides as Phosphate Acceptors. Several deoxydinucleoside monophosphates and deoxynucleoside 3'-mOnOphosphates (as well as ribonucleoside 3'-monophosphates) were not substrates for the rat liver enzyme (Table I), although they were all readily phosphorylated by the T4 kinase ( 3 ) . The dependence of the kinase activity on the chain length of oligodeoxynucleotides was tested with a series of partial DNase I digests of calf thymus DNA (3). After dephosphorylation, the digests were incubated with DNA kinase and [y-32P]ATP.Digests with average chain length of -6-9 residues were relatively inactive, whereas digests with average chain length of 13 or more residues were acted upon at rates similar to the rate on denatured DNA. These results suggested that a chain length of more than 10-12 residues is required for rapid phosphorylation. The size distribution of labeled oligodeoxynucleotides in a partial digest was consistent with this conclusion. E.

REVERSALOF THE REACTIONAND LABELING BY

EXCHANGE Reversal of the kinase reaction could be readily demonstrated (131, although the rate was several orders of magnitude slower than the rate of

323


TABLE I

PHOSPHATE ACCEPTORSPECIFICITY OF RAT LIVERDNA KINASE~ Relative rate of kinase activity Experiment

Substrate

(%)

A

DNA, micrococcal nuclease-treated DNA, micrococcal nuclease-treated, heat 5 min at loo", quench poly(rA), micrococcal nuclease-treatedb RNA, micrococcal nuclease-treated" RNA, micrococcal nuclease-treated, heat 5 min at loo", quench DNA, micrococcal nuclease-treated Ado 3'-P, Guo 3'-P, Cyt 3'-P, or Urd 3'-P dCyt 3'-P or dThd 3'-P dT-dC or dC-dT

100

B

83 RNA Yes YeS No Yes 0.6 M

N.D. N.D. N.D. N.D. N.D. Yes 0.4-0.6 M

10,000 (monomer) T4 pol Yes Yes

300-800 (tetramer) pol II, pol 111, T7 pol Yes Yes

75,000 (dimer) pol IId N.D.e N.D.

N.D. T7 pol N.D. N.D.

T4 Gene 32 protein

34 Monome? & dimer s?infinite N.D. 301 37,000 M-' cm-I (280 nm)

E. coli SSB protein

19.5 Tetramer 4.7

s

- 190

Because of cooperativity this is a function of protein concentration.

* Probable nucleic acid binding form.

Requires presence of polyamines. Stimulates amount of DNA synthesis, not rate, only at low protein concentration. N.D., not determined.

Acidic

TABLE 111

PROPERTIES OF Property

w

'0

Physical Molecular weight (monomer) (kdal) Isoelectric point Oligomeric state Sedimentation coefficient Binding Polynucleotide site size (n) Relative affinity (DNA vs. RNA) Cooperativity Ability to melt duplex DNA Ability to melt poly(dA-dT) [NaCI] for elution from DNA-cellulose Biological Copies per cell Stimulation of DNA polymerase

EUKARYOTIC SINGLE-STRANDED DNA BINDINGPROTEINS Calf thymus HDP-I

Mouse myeloma

Ustilago maydis

24 7.8 Monomer N.D.'

27 6.6 Monomer N.D.

20 N.D. Monomer 2.6 S

35 N.D. Monomer 3.3 s

72 7.5" Monomer 3.3 s

7 nuc DNA > RNA No Yes Yes

5-7 nuc DNA > RNA No N.D. Yes 0.4 M

3-7 nuc N.D.

N.D. DNA > RNA

7 nuc DNA > RNA

b

b

b

N.D. Yes 1.6 M

Yes N.D. N.D.

N.D. No 1.0 M

1,000,OOO Yes

300,000 Yes

N.D. N.D.

70,000,000 Yes

0.4 M

800,000

" Nonphosphorylated species. Possibly cooperative, but not fully established. N.D. = not determined.

Yes

Lily

Adenovirus

392

S. KOWALCZYKOWSKI, D. BEAR, AND P. VON HIPPEL

phage that containedamber mutations in gene 32 and by demonstrating the loss of the corresponding polypeptide from the resulting polyacrylamide gel patterns; controls using phage defective in other genes ( 3 0 , 4 1 4 5 ) had no effect on the band identified as the product of gene 32. Both because gene 32 protein had been shown by genetic analysis to be essential to phage DNA replication (57a) and recombination (58, 59), and because it is produced in large (stoichiometric) quantities (60, 6 1 ) [- 10,000 copieslcell (631, Alberts et al. (6, 38) undertook biochemical studies of this protein. These early studies showed that (1) the protein binds to single-stranded DNA in preference to double-stranded DNA; (2) binding appeared to be cooperative in protein concentration; (3) gene 32 protein can denature duplex poly(dA-dT), but not native duplex DNA; (4) the protein accelerates the renaturation of denatured DNA; and ( 5 ) the protein appears to stimulate specifically the activity of T4 DNA polymerase. Each of these properties of gene 32 protein has since been studied in more detail, and the results are summarized below. 1. Physicul Properties of the Protein Gene 32 protein has a monomeric molecular weight of approximately 35,000, as determined by SDS gel electrophoresis and by a combination of sedimentation and gel filtration studies; it behaves hydrodynamically as if it were a prolate ellipsoid with an axial ratio of 4 : 1 (=120 A in length) (6, 38). A determination of the complete amino acid sequence of the protein yielded a more exact molecular weight of 33,466 (63). u. Aggwgatim State. Although gene 32 protein exists predominantly as a monomer in dilute solution (c0.025 mg/ml), the protein has been shown to undergo extensive self-association at higher concentrations (64, 65). The extent of this aggregation is dependent on many variables, including ionic strength, pH, temperature, and glycerol concentration. The aggregation process seems to consist of at least two different types of 57a. R . H. Epstein, A. Bolle, C. M. Steinberg, E. Kellenberger, E. Boy de la Tour, R. Chevalley, R. S . Edgar, M . Susman, G. H. Denhardt, and A. Lielausis, CSHSQB 28, 375 (1%3).

58. J. Tomizawa, N . Anraku, and Y. Iwama, JMB 21, 127 (1966). 59. A. W. Kozinski and Z. Z. Felgenhauer,J. Virol. 1, 1193 (1967). 60. D. P. Snustad, Virology 35, 550 (1968). 61. N. K. Sinha and D. P. SnustadJMB 62, 167 (1971). 62. B. M. Alberts, Ff 29, 1154 (1970). 63. K. R . Williams, M. B . LoPresti, M. Setoguchi, and W. H. Konigsberg, PNAS 77, 4614 (1980). 64. R. B. Carroll, K. E. Neet, and D. A. Goldthwait, PNAS 69, 2741 (1972). 65. R . B. Carroll, K. E. Neet, and D. A. Goldthwait,JMB 91, 275 (1975).

21. DNA BINDING PROTEINS

393

association. One leads to a unique dimeric species; the other, via an indefinite association process, leads to large aggregates. The indefinite aggregation is particularly susceptible to various agents, being virtually completely inhibited by high concentrations of salt (>1.0 M KCl), high temperature, high pH (>pH lo), and the addition of saturating concentrations of oligonucleotides [e.g., d(pT),,]. Under each of these conditions the limit aggregate is a dimeric species, even at protein concentrations as high as 1 mg/ml. Although these protein-protein association reactions might be considered to reflect the interactions responsible for the cooperative binding of gene 32 protein to nucleic acid lattices, quantitative study of the nucleic acid binding reaction shows that this interpretation is not consistent with the facts. For example, the unitary free energy of the aggregation process is about - 9 kcal/mol, whereas the free energy of the monomer-monomer interaction involved in cooperative binding is only about - 4 kcaVmol under the same conditions ( I ? ) . In addition, while the indefinite selfassociation shows a very large dependence on salt concentration, the cooperativity parameter for nucleic acid binding is not dependent on this variable ( 5 ) . Thus the nonelectrostatic part of the protein-protein selfassociation free energy may contribute to the cooperativity of nucleic acid binding, but it is clear that the interactions involved in self-association are not, in toto, the same as those involved in cooperative binding of this protein on a nucleic acid lattice. 6. Protein Domain Structure. The observation that gene 32 protein can bind to (polyanionic) DNA-cellulose, despite the fact that it carries a net negative charge at pH 8 and is eluted from DNA-cellulose at high concentrations of salt, suggested very early that gene 32 protein must contain a positively charged binding site ( 6 , 3 8 ) .The idea that gene 32 protein might be composed of several functionally distinct domains was given a firmer basis when it was discovered that the native protein can be converted into three discrete products by limited tryptic digestion. These products have been called G32P*I, G32P*II and G32P*III (27, 66,67). The G32P*I fragment results from the removal of -50 amino acid residues from the carboxy terminus of the native protein, the G32P"II product is obtained by removing -20 amino acid residues from the amino terminus, and the G32P*III fragment is produced by the removal of both the amino and carboxy termini. The G32P*III protein core is quite resistant to further proteol ytic degradation. 66. J. Hosoda, B. Takas, and C. Brock, FEBS (Fed. Eur. Biochem. Soc.) Lerr. 47, 338 (1974). 67. H.Moise and J. Hosoda, Nufure (London) 259, 455 (1976).

394

S. KOWALCZYKOWSKI, D. BEAR, A N D P. VON HIPPEL

These proteolytic products differ in their elution properties from DNA-cellulose columns. Thus the native protein and G32P"I (at fairly high protein concentrations approaching DNA site saturation within the protein band) require -2.0 M NaCl for elution from single-stranded DNA-cellulose, whereas G32P"II and G32P*III can be eluted at -0.4 M NaCl concentrations. This latter salt concentration is also sufficient to elute noncooperatively bound native gene 32 protein (38), in keeping with the conclusion (see Section IV,A,2,c) that G32P*II and G32P*III retain most of their intrinsic affinity for single-stranded DNA, but no longer can bind cooperatively. In contrast G32P*I (and, to some extent, G32P"III) bind more tightly to double-stranded DNA-cellulose than the native protein or G32P"II (67), presumably reflecting the fact that the *I (and the *III) species can denature the duplex DNA and bind to the resulting single-stranded sequences (66, 68) (see also Section IV,A,2,c). Additional evidence for the differential involvement of the amino and the carboxy-terminal domains of gene 32 protein in nucleic acid binding was provided when it was shown that the rate at which these regions are removed by proteolytic cleavage depends on whether or not the protein is bound to polynucleotides (69, 70). The cooperative binding of gene 32 protein to polynucleotides was found to enhance the rate of cleavage at the carboxy terminus, but to protect the amino terminus against proteolytic attack. These effects were not observed in the presence of short oligonucleotides [ poly(rC) + dsDNA (P22) (36). Obviously genera1 preferences can be seen for single-stranded over double-stranded DNA, and for deoxyribose-containing over ribosecontaining polynucleotides. Also poly(dT) seems to be bound more tightly than all other polynucleotides; these general features are also seen with gene 32 protein. Little, if any, nucleotide composition or sequence dependence is observed for the binding of hexanucleotides to SSB protein.

427


c. Dependence oj' Binding Aj'finity on Oligonrrcleotide Lattice Length. Equilibrium dialysis experiments (152) have suggested that the apparent binding affinity of SSB protein for various multimers of d(pCpT) depends on the length of the overall oligonucleotide lattice. The apparent binding constant to such oligomers increased -10-fold in going from d(pCpT)2to d(pCpT),, and only -2-fold further in going from d(pCpT)3to d(pCpT)6+. In partial contrast to these results, intrinsic protein fluorescencequenching studies have shown that d(pT),6 binds SSB protein -200-fold more strongly than dT-containing oligomers 8 residues in length or less (36). The longer oligomers quench protein fluorescence much more, effectively than the shorter lattices. These results are also in general accord with those observed with gene 32 protein. d. Polynucleotide Binding Site Size and the Oligomeric State of SSB Protein. A variety of techniques have been used to determine the site size ( n ) of SSB protein for binding to polynucleotides; most approaches have yielded values ofti of -8 nucleotide residues per protein monomer (36, 96, 143). The molecular significance of this parameter has been hard to define because neither the state of aggregation of the protein in its binding form, nor the number of binding sites utilized per bound protein, have been established unequivocally. EM studies have suggested that the protein binds to single-stranded DNA as a tetramer, or alternatively, that it aggregates to tetramers on binding (96). It was also demonstrated by sedimentation experiments with oligo d(pCpT),-, that no change in the state of aggregation occurs upon binding (152). This result has been confirmed by measurements of the rotational correlation time of the free protein and the complex by time-dependent emission anisotropy ; no significant change in the protein structure upon binding to oligo(dT),, was revealed (154).In light of these results, a site size of about 32 nucleotides per tetramer may be a more physically significant way to view the binding of SSB protein to single-stranded DNA. More detailed work on the topography of this complex is obviously required. e. Binding Cooperativity. The cooperative nature of the binding of E. coli SSB protein to single-stranded DNA and to synthetic polynucleotides has been demonstrated by several different techniques, including electron microscopy (96, /S3), gel filtration (142), density gradient centrifugation ( 9 6 ) , and fluorescence quenching ( 3 6 ) . By comparing the binding constants for the interaction of SSB protein with pT8 and pT16, it was e d mated that the value of the cooperativity parameter is at least 50 on the basis of fluorescence measurements ( 3 6 ) , whereas on the basis of electron microscopic studies, w was estimated to be 10' (153). The estimate based on fluorescence is only a lower limit; however, the value obtained by electron microscopy may be high due to artifacts in the technique. Thus

-

428

S . KOWALCZYKOWSKI, D . BEAR, A N D P. VON HIPPEL

the use of glutaraldehyde in the fixation of the complexes prior to spreading could have artificially increased the size of the protein clusters beyond equilibrium expectations. Such artifacts have been seen in EM estimates of cooperativity for gene 32 protein (79). An accurate measure of w for SSB protein, obtained under physiological conditions, is not available. In addition, preliminary evidence (1.33) suggests that, unlike the situation with gene 32 protein, w for SSB protein may be salt concentration dependent. f. Dennturution and Renaturution of Duplex D N A . As noted above, E. coli SSB protein at low ionic strength can slowly denature duplex DNA (96). Surprisingly, this effect has not been heavily investigated, in contrast to the denaturation induced by the gene 32 protein *I fragment. Upon addition of E. coli SSB protein, it was found that denaturation of duplex DNA goes to completion in 12 to 30 min at 37” for duplex DNA; the DNA has an unperturbed T,,,Of 55” under the same solvent conditions (96). The effect is stoichiometric rather than catalytic; however, the quantitative aspects of the stoichiometry have not been characterized. In electron microscopic studies involving glutaraldehyde-fixed protein-DNA complexes it was found that the E. coli SSB protein attacks A-T rich regions, and tends to expand denatured loops, rather than to initiate new ones. The addition of Mg2+completely reverses the denaturation effect. Several groups have reported little or no affinity of E. coli SSB protein for double-stranded DNA. Also the addition of 5 mM spermidine to denatured DNA strongly decreases the affinity of the protein, presumably by stabilizing the secondary structure of the nucleic acid. I t has also been demonstrated that E. coli SSB protein does not retain duplex DNA on nitrocellulose filters (142). The binding affinity of SSB protein to duplex DNA has been estimated to be at least 3 orders of magnitude weaker than that to denatured DNA (36). These data suggest that, like the carboxy-terminal cleaved gene 32 protein (G32P*I), E . coli SSB protein binds to and traps sequences of DNA transiently opened by thermal fluctuations. Since A-T rich sequences are less stable with respect to thermal melting than those rich in G-C base pairs, the former are presumed to be prime candidates for such nucleating interactions. The protein, due to its significant cooperativity, would then tend to bind at contiguous sites and expand the denatured regions. The biological significance of such SSB protein-driven melting processes is questionable, due to the sensitivity of melting to the concentration of mono- and polyvalent ions. We return to this question in the following sections. It has been reported that under certain conditions E. coli SSB protein

21.

D N A BINDING PROTElNS

429

can catalyze the renufuration of denatured DNA (155). In the presence of NaCl(0.2 M or less), and in the presence or absence of MgC12,E. coli SSB protein does not significantly affect the rate of renaturation of h DNA at pH 7.0. However, in the presence of 2 mM spermidine or spermine the renaturation rate is enhanced by a factor of about 5000. This rateenhancement effect requires saturating amounts of SSB protein, and its efficiency increases with the length of the DNA. Because of the complex nature of the dependence on counterions and pH, the simple removal of intrastrand hairpin loops by SSB protein binding does not appear to fully explain this effect. 3. Biological Roles

a. Replication. For several years after the initial discovery of E. coli SSB protein, the effects of this protein on the in virro replication of viral DNA by various viral and host-coded polymerases has been heavily investigated. However, it was not until a temperature-sensitive mutant was isolated (148) and found to be defective in DNA replication that one could be sure of the importance of E. coli SSB protein in phage and cellular metabolism in vivo . The mutation (designated ssb- 1) has been localized at 90 to 91 minutes on the E. coli linkage map. In virro studies have demonstrated that the E. coli SSB protein specifically stimulates polymerase 11-directed DNA synthesis on various phage templates (96, 150). This stimulation appears to be dependent on the ratio of binding protein to DNA, and independent of polymerase concentration, suggesting that the primary effect is at the DNA level (151). The addition ofE. coli SSB protein to in vitro replication systems has a multitude of effects, including (i) stimulation of the initiation of RNA primer-directed DNA synthesis by polymerase I11 holoenzyme on singlestranded phage DNA templates (139, 142), (ii) stimulation of the elongation rate of DNA synthesis by pol I11 (142), (iii) increase in the fidelity of DNA synthesis by pol I11 ( / 4 / ) ,and (iv) stimulation of the initiation and elongation of DNA synthesis by pol II on gapped and single-stranded templates (96,150, 156). The protein is required in an in virro polymerase I11 replication mix in order to convert bacteriophage G4 single-stranded DNA into its replicative form (142); and also to convert the replicative form back to single-stranded circular DNA (143). All these activities may be explained by the ability of SSB protein to bind to single-stranded DNA, trapping the DNA in the open form and 155. C. Christiansen and R . L. Baldwin, JMB 115, 441 (1977). 156. 1. J. Molineux and M. L. Gefter,JMB 98, 811 (1975).

430


melting out double-stranded regions by contiguous cooperative binding. This makes the bases more accessible as templates for polymerases, and thus may increase the fidelity of DNA synthesis as well as the elongation rate. However, there is ample evidence to suggest that E. coii SSB protein also interacts specifically with polymerases I1 and 111. The SSB protein also inhibits the exonuclease activity of pol I and the T4 DNA polymerase, and the activity of the single-stranded DNases from Aspergillus and Neurospera crassa . In contrast it does not inhibit, and even stimulates to a small extent, the exonuclease activity of T7 DNA polymerase, pol 11, and exonuclease I(156). Both pol I1 and exonuclease I form complexes with SSB protein. While no complex appears to be formed between SSB protein and T7 polymerase, it has been shown that SSB protein can substitute for T7 DNA binding protein (157) in a T7 replication system. Other DNA binding proteins, such as fd gene 5 , T7 DNA binding protein, and T4 gene 32 protein, cannot substitute for SSB protein in stimulating pol I1 (157). Thus the interaction of E. coli SSB protein with pol I1 is probably physiologically important, whereas the role of the DNA binding protein in the T7 replication system may be to interact primarily with the DNA. Demonstration of meaningful specific interactions with pol I11 must await a complete characterization of the holoenzyme [see Chapter 3, this volume and Ref. (140) for a review of this aspect]. Antibody titration studies have shown that there about 300 copies ofE. coli SSB protein tetramer per log phase bacterial cell (142). Based on the binding information previously presented, and the fact that there are approximately six replication forks per cell (96),we can calculate that -1600 nucleotides of DNA are covered per replication fork. Both pfasmids and transducing phages carrying the ssb gene have been used to overproduce SSB protein (42). Thus it is unlikely that the synthesis of this protein is autogenously regulated in vivo . b. Recombination and Repair. Further characterization of the ssb- 1 mutant (148)has revealed that the mutant strain is about one-fifth as active in recombination as the wild type, and extremely sensitive to UV damage. The lexC gene proposed as a regulator of UV and X-ray inducible repair (146, 158) has been tentatively found to be allelic withssb. Both genes map at the same locus, and SSB protein from a temperature-sensitive lexC mutant is temperature-sensitive when tested as an accessory protein in bacteriophage G4 replication. While an exact role for the E. coli SSB protein in recombination and repair has yet to be established, it has been demonstrated that SSB protein catalyzes recA -mediated single-stranded DNA 157. R. C. Reuben and M. L. Gefter, JBC 249, 3843 (1974). 158. J. Greenberg, L. J. Berends, J. Donch, M. H. L. Green, Genet. Res. (Cambridge) 23, 175 (1974).

21.

DNA BINDING PROTEINS

43 1

assimilation into homologous double-stranded DNA in vitro (159, 160). The concentration of recA protein required for such D-loop formation is also reduced in the presence of SSB protein. D.

BACTERIOPHAGE T7-CODED SINGLE-STRANDED DNA BINDING PROTEIN

1. Protein Properties and Interactions with DNA In searching for an analogue of T4-coded gene 32 protein that might serve similarly in phage T7 replication, two groups independently discovered a T7-coded, single-stranded DNA binding protein that stimulates replication by T7 DNA polymerase on either single-stranded or gappedduplex DNA templates (16l, 162). Although the original estimates of molecular weight of the two proteins differ considerably [31,000 (161) versus 25,000, (162)], the proteins appear to be the same on the basis of their other properties. Like gene 32 protein, the T7 HDP does not bind to duplex DNA and carries a net negative charge at neutral pH. NaCl concentrations in excess of 0.25 M are required to elute this protein from DEAE-Sephadex; under the same conditions E. coli SSB protein (PI = 6.0) is eluted at -0.15 M NaCI. The protein appears to be monomeric in 2 M NaCI, as judged by its behavior on gel filtration columns. The T7 HDP denatures duplex poly(dA-dT) (162). We estimate, on the basis of incomplete data (1621, that the T7 protein lowers the T, of poly(dA-dT) by about 40" in 0.04 M NaCI; this AT, is very comparable to that induced by T4 gene 32 protein under the same conditions. The T7 protein also stimulates the exonuclease activity of T7 DNA polymerase (the T7 gene 5 protein-E. coli thioredoxin complex) on a duplex DNA template (163). These rather scanty data suggest that the nucleic acid binding properties of the T7 protein may be quite similar to those of the other prokaryotic single-stranded DNA binding proteins described in the preceding sections. 2. Biological Roles

The biological properties of the T7 single-stranded binding protein also resemble those of the analogous T4 and E. coli proteins. The T7 protein 159. K. McEntee, G. M. Weinstock, and I. R. Lehman, PNAS 77, 857 (1980). 160. T. Shibata, C. DasGupta, R. P. Cunningham, and C. M. Radding, PNAS 77, 2606 (1980). 161. R. C. Reuben and M. L. Gefter,PNAS 70, 1846 (1973). 162. E. Scherzinger, F. Litfin, and E. Jost, Molec. Geti. Getter. 123, 247 (1973). 163. K . Hori, D. F. Mark, and C. C. Richardson,JBC 254, 11598 (1979).

432


stimulates the polymerization activity of T7 DNA polymerase on singlestranded templates 10- to 15-fold at low temperatures; under these conditions the T7 DNA polymerase is otherwise quite ineffective (161). R e sumably this stimulation reflects the denaturation of DNA hairpins, which are stable under these conditions, and which, if present, inhibit the polymerase. The T7 protein does not, however, stimulates T4 DNA polymerase or E. coli DNA polymerases I, 11, or I11 (161). In spite of this demonstration of functional specificity, no physical interaction between the T7 DNA binding protein and T7 polymerase has been observed (162). In contrast, the T4 gene 32 protein and the E. cofi SSB protein do bind preferentially to their homologous polymerases in free solution. In vitro, E. coli SSB protein can substitute for the T7 protein with regard to its stimulatory effect on the activity of T7 DNA polymerase (162). Thus, despite the fact that the T7 SSB protein seems to be specific (i.e., it can only stimulate its homologous DNA polymerase), the T7 DNA polymerase can be stimulated by both E. coli and T7 SSB proteins. This may also explain why mutations in T7 SSB protein are not lethal to T7 growth; presumably the host SSB protein substitutes for the phage protein, thus “rescuing” the mutant phage.

E. EUKARYOTIC SINGLE-STRANDED DNA BINDING PROTEINS A large number of proteins that show affinity for either single-stranded or double-stranded DNA have been isolated from a variety of eukaryotic organisms [for reviews, see Refs. (/64-166)]. In general, these proteins have been isolated from cell extracts on the basis of binding to DNAcellulose columns: in the absence of genetic information and mutants the physiological role of many of these proteins has been difficult to assess. The proteins that are included in the following sections have been selected for discussion because they show (i) binding to DNA, (ii) a preferential affinity for single-stranded over double-stranded DNA (thus they are, at least potentially, HDPs), and (iii) a presumed biological activity (usually manifested as a stimulatory effect in an in vitro replication system with the homolgous DNA polymerase). As a consequence many interesting proteins have been omitted, but we hope that those discussed will prove to be representative of this potentially important class. Some of the more important properties of these proteins are summarized in Table 111. 164. J. E. Coleman and J. L. Oakly, Cril. Rev. Biochem. 7, 247 (1980). 165. J. J. Champoux, Annu. Re\’. Biochem. 47, 449 (1978). 166. A. Falaschi. F. Cobianchi, and S. Riva, Trends Biochem. Sci. 5, 154 (1980).

21.


433

1. The Lilium (Lily)DNA Binding Protein

A unique DNA binding protein has been isolated from the meiotic cells of lily plants. In contrast to the other prokaryotic and eukaryotic DNA binding proteins discussed here, this protein is primarily involved in meiotic recombination, rather than in DNA replication (167-169). This protein, referred to as R-protein, is synthesized during meiotic prophase and is localized within the nucleus. The fact that it is only present (and active) in germ cells during the portion of the cell cycle corresponding to meiosis strongly suggests that it is somehow involved in the processes of chromosome pairing and recombination (168). The isolated protein has a molecular weight of -35,000, and forms a stable complex with single-stranded DNA even at 2.0 M NaCl. Minimal concentrations (5 mM) of Mg2+or Caz+are absolutely required for binding (168). Binding is specific for single-stranded DNA, with no detectable binding to RNA. Furthermore, like T4 gene 32 protein and E. coli SSB protein, this protein catalyzes the renaturation of denatured DNA. As previously noted, the ability to catalyze the renaturation of denatured DNA might be quite important in recombination, and is thus consistent with the proposed role of this protein in meiosis. The lily protein can be phosphorylated by a specific CAMP-dependent protein kinase, and the level to which it is phosphorylated determines its in vifro properties (169). The native (phosphorylated) protein exhibits a definite preference for binding to single-stranded DNA, and although the dephosphorylated protein has an increased affinity to single-stranded DNA, it shows an even greater increased affinity for double-stranded DNA. The properties of the dephosphorylated protein revert to those of the native protein by treatment with the protein kinase, which adds two phosphate groups per protein monomer. The native protein stimulates both duplex DNA denaturation and denatured DNA renaturation, but the dephosphorylated protein shows neither of these activities. A very similar protein has been isolated from rat spermatocytes (170). The DNA binding properties and renaturation activity of the rat protein are also modulated by kinase-driven phosphorylation-dephosphorylation activities. In addition a DNA binding protein of mouse acites cells has been shown to vary in its stimulatory effect on DNA replication, depending on its level of phosphorylation. These results suggest that control of the level of phosphorylation of DNA binding proteins may serve as a 167. 168. 169. 170.

Y. Hotta and H. Stern, Dev. B i d . 26, 87 (1971). Y. Hotta and H . Stern, Nnrltre (London)N ~ Bwi d . 234, 83 (1971). Y. Hotta and H. Stern, EJB 95, 31 (1979). J. Mather and Y. Hotta, Exp. Cell Res. 109, 181 (1977).

434


general mechanism to modulate the activity of these proteins during the cell cycle in eukaryotes [for a review of eukaryotic protein phosphorylation, see Ref. ( / 7 / ) 1 . 2. Cay Thymus DNA Binding Proteins Three DNA binding proteins have been isolated from calf thymus by Herrick and Alberts (49, 172, 173) using a general purification protocol designed by these workers for the isolation of eukaryotic HDPs (49). The proteins isolated are UP1 [UP for unwinding protein; by current nomenclature this protein would be calf thymus (CT) HDP-I]; a “high salt-eluting fraction” (CT HDP-111, and a “low salt-eluting fraction”. CT HDP-I has a molecular weight of 24,000 and is present in the thymus at -800,000 copies per cell (49). It exists as a monomer in solution, and has an isoelectric point near neutrality. Isoelectric focusing shows that it is composed of four or five subspecies, probably reflecting intrinsic heterogeneity or limited protease action during isolation. These fractions show different affinities for single-stranded DNA; the most acidic fraction binds most weakly. This protein has a marked preference for single-stranded over double-stranded DNA, as demonstrated by its ability to (reversibly) depress the Tm of poly(dA-dT), poly(rA-rU), and C. perfringens duplex DNA (172). This helix destabilization effect is greatest for poly(dA-dT), less for poly(rA-rU), and least for the C . perfringens DNA, suggesting that the protein may have some base- and sugar-binding specificity. From these studies it has been estimated that the affinity for single-stranded DNA is 1500-fold greater than for double-stranded DNA. The acidic subfraction of CT HDP-I shows a much smaller ATm, as might be expected from the smaller affinity of this protein for single-stranded DNA cellulose. Sedimentation studies (173) and optical studies (174) have shown that a stoichiometric complex is formed at a ratio of 7 DNA nucleotide residues per protein (CT HDP-I) monomer. The formation of this complex is very dependent on salt concentration, and the data suggest that up to 6 ionic interactions per protein monomer may be involved (174). The Tm depression of duplex DNA induced by this protein is salt-dependent, decreasing with increased NaCl concentration (172). Although the protein shows a high affinity for single-stranded DNA, unlike the prokaryotic HDPs it does not bind cooperatively (17.3). Electron microscopy shows an extended DNA complex in which the 171. C. S. Rubin and 0. M. Rosen, Annu. Rev. Biochem. 44, 831 (1975). 172. G . Herrick and B . M. Alberts, JBC 251, 2133 (1976). 173. G. Herrick, H. Delius, and B. M. Alberts, JBC 251, 2142 (1976). 174. R. L. Karpel and A. C. Burchard, Binchemisrry 19, 4674 (1980).

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435

contour length increases by -17% (17.3); the comparable increase in length for DNA complexed with T4 gene 32 protein is 46%. Also, as expected, CT HDP-I (and CT HDP-11) produce denaturation “bubbles” in supercoiled SV40 DNA. In addition changes in the polynucleotide circular dichroism and UV absorbance spectra indicate considerable unstacking of the bases (174). The accessibility of nucleotide bases in this protein-nucleic acid complex was probed by chemical modification and hydrogen exchange techniques. The results suggest that the bases are as exposed in the complex as in free DNA, but that they are also unstacked in the complex (17.5). The CT HDP-I1 preparation contains several protein fractions; the most prominent has a molecular weight of -33,000 and an isoelectric point (PI) of 5.2-5.6. This protein is very similar to CT HDP-I in that (i) it binds noncooperatively to a single-stranded DNA at a nucleotide residue to protein stoichiometry of -10: 1, (ii) it strongly depresses the T, of poly(dA-dT) and poly(rA-rU), and (iii) it forms an extended complex with fd DNA (49, 172). In contrast, the low-salt fraction (molecular weight -33,000) does not denature native DNA and does not form a stable complex visualizable in the EM. In addition, this fraction does not stimulate calf thymus DNA polymerase activity. While the physiological role of these proteins has not been defined, both CT HDP-I and -11 stimulate the activity of the calf thymus DNA polymerase a , but not that of polymerase /3 (173). T4 gene 32 protein does not stimulate polymerase a activity; thus stimulation by the calf thymus proteins may be specific. The amount by which this polymerase is stimulated by CT HDP-I depends on the type of DNA template used, and ranges from a 10-fold stimulation on exonuclease-treated DNA to less than a twofold effect on heat-denatured DNA. As found with T4 gene 32 protein, excess CT HDP-I inhihits DNA synthesis. At optimal concentrations both CT HDP-I and -11 stimulate calf thymus polymerase a activity on oligo(dG) primed-poly(dC) templates more than 5-fold. CT HDP-I can also catalyze the renaturation of tRNA and 5 S RNA to their active (native) forms (176). This renaturation occurs despite the fact that CT HDP-I will not renature denatured DNA; this difference may reflect the fact that the tRNA renaturation is an intramolecular process 175. T. Kohwi-Shigematsu, T. Enomoto, M. Yamada, M. Nakanishi, and M. Tsuboi, PNAS 75, 4689 (1978). 176. R . L. Karpel, N . S. Miller, and J. R. Fresco, iff “Molecular Mechanisms in the Control of Gene Expression” (D. P. Nierlich, W. J . Rutter, and C. F. Fox, eds.), p. 411. Academic Press, New York, 1976.

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S . KOWALCZY KOWSKI, D. BEAR, AND P. VON HIPPEL

and DNA renaturation is intermolecular (172). In addition, CT HDP-I seems to be located primarily in the cytoplasm of the cell, suggesting that it may be involved in RNA manipulation (176). By using the complementary approach of isolating a protein that stimulates the activity of polymerase a, and then comparing it to the proteins that have been previously described, a protein very similar to the most basic component of CT HDP-I has been purified (177). Its binding is specific for single-stranded DNA, and it stimulates the activity of DNA polymerase LY up to 8-fold depending on the template, whereas polymerase /3 is stimulated less than 2-fold.

3. Ustilago mrrydis D N A Binding Protein An HDP has been isolated from mitotic cells of the basidiomycete fungus, U . maydis. This protein may be involved in DNA replication in this organism in that it stimulates U . rnciydis DNA polymerase (178, 179). The protein has a molecular weight of -20,000, and exists in solution as a monomer. Filter binding experiments show that it binds tightly to singlestranded DNA, but not to double-stranded DNA. The T,,,of poly(dA-dT) is lowered by -50" in the presence of saturating amounts of this protein, and Mg'+ was found to increase the rate of renaturation. As with T4 gene 32 protein and the lily R-protein, the I / . mcrydis HDP also catalyzes the renaturation of denatured DNA (178). The protein stimulates by twofold the activity on denatured DNA of the only DNA polymerase that has been isolated from U . maydis. As with many of the other HDPs, excess binding protein inhibits the activity of the polymerase. Since the U . muydis HDP does not stimulate the activity of polymerases from E. coli. M. lureus, T4, or T7, the stimulation of the U . maydis polymerase may be specific. However, no specific interaction of polymerase with binding protein could be detected. In addition to the stimulation of polymerase activity, the U . mciydis HDP in stoichiometric excess also inhibits the nucleolytic digestion of DNA. The stimulatory effect of this protein on the U . maydis polymerase was found to be due to an increase in the rate rather than in the extent of DNA replication. This stimulation was found to arise from several effects that were produced by the presence of binding protein (179). When denatured DNA was used as a template for U . rnaydis polymerase, the apparent K , for nucleoside triphosphates was increased 3- to 4-fold in the presence of binding protein. In addition, the apparent affinity of the polymerase for

-

177. F. Cobianchi, S . Riva, G. Mastromel, S. Spadari, G. Pedrali-Noy, and A. Falaschi, CSHSQB 43, 639 (1979). 178. G . R. Banks and A. Spanos, J M B 93, 63 (1975). 179. G. Yarranton, P. D. Moore, and A. Spanos, M d . Get?. Genet. 145, 215 (1976).

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437

DNA was increased fourfold by the U . may& HDP. And finally, the V,,, of the polymerization reaction increased -50%. All of these effects have been interpreted to indicate that the U . maydis binding protein stimulates replication both by removing secondary structure in the DNA template, which may impede the polymerase, and by providing a specific proteinnucleic acid complex with which the DNA polymerase can interact. 4. Mouse DNA Binding Proteins Several DNA binding proteins have been isolated from mouse tissue, including proteins isolated from 3T6 cells (180), ascites cells (181), and myeloma cells (182). Each of these proteins is a helix-destabilizing protein, and the last two stimulate mouse polymerase a. The role of the protein isolated from 3T6 cells is unknown; it is found primarily in the cytoplasm, and more is found in growing than in resting cells. The HDP found in mouse ascites cells has a molecular weight of -35,000, and exists as a monomer in solution (181). It shows a high affinity for single-stranded DNA or RNA, and only a slight affinity for doublestranded DNA. Sedimentation experiments suggest that the proteinnucleic acid complex is asymmetric and highly extended, and that saturation of the DNA occurs at -6-10 nucleotide residues per protein monomer. The properties of this protein suggest that it is analoJous to the high-salt fraction protein isolated from calf thymus. The mouse protein also stimulates the activity of mouse DNA polymerase a fourfold, but only on a heat-denatured DNA template. There is no stimulation of activity on pancreatic DNAse-treated (activated) DNA templates, but we note that the activity of the DNA polymerase is already sevenfold greater on the activated than on the denatured DNA template. This helix-destabilizing protein may specifically stimulate polymerase a , since assays of the effect of this protein on mouse polymerase p, E. coli DNA polymerase, and T7 DNA polymerase all show only very slight stimulatory effects. The effect on polymerase a has been surmised to be at the elongation step. It has also been showed that DNA binding protein increases the affinity of polymerase a for DNA cellulose (183) similar to the effect shown with the U . maydis protein. Although no direct association between polymerase and binding protein has been seen, these results suggest that the binding protein may stabilize the polymerase a-DNA complex.

-

180. 181. 182. 183. (1978).

R. L. Tasi and H. Green, J M B 73, 307 (1973). B . Otto, M. Baynes, and R. Knippers, EJB 73, 17 (1977). S. R. Planck and S. H. Wilson,JBC 255, 11547 (1980). A . Richter, R . Knippers, and B. Otto, FEBS (Fed. E m . Biochem. SOC.Lett.) 91,293

438

S . KOWALCZYKOWSKI, D. BEAR, AND P. VON HIPPEL

As with the protein isolated from lily, the mouse DNA binding protein can be phosphorylated (1 phosphate/protein monomer) by a chromatinassociated kinase, resulting in an alteration of its binding properties. Although phosphorylation has no effect on single-stranded DNA affinity, the phosphorylated protein shows a reduced affinity for double-stranded DNA and a greatly reduced ability to stimulate DNA polymerase a. Thus this system seems to represent another example of HDP activity controlled by the level of covalent phosphorylation. Another protein that has been isolated from mouse cells (myeloma), and is clearly not the same as those obtained from 3T6 or ascites cells, is mouse HDP-I (182). This protein is heterogeneous in molecular weight, ranging from 24,000 to 33,000, with a predominant species at -27,000. This apparent heterogeneity is not due to different levels of phosphorylation, acetylation, or glycosylation; rather peptide mapping and tryptic digestion studies implicate protease activity. This HDP is localized predominantly in the cell nucleus, and is not associated with the chromatin; however -25% is also found in the cytoplasm. Note that the amino acid compositions and the molecular weights of the mouse HDP-I and CT HDP-I are very similar. Sedimentation experiments with mouse HDP-I and denatured DNA indicate that the protein binds noncooperatively to single-stranded DNA, with a site size of 5 to 7 nucleotide residues per protein monomer and a binding constant of -4 x lo5M - I . In addition, saturating amounts of this protein depress the T,,, of poly(dA-dT) by -25” in 0.01 M salt. The binding of DNA to this protein induces a fluorescence increase of -35%, in contrast to fluorescence quenching typically seen with prokaryotic DNA binding proteins (182). Studies of the products of limited tryptic hydrolysis of the mouse HDP-I have shown that different products are obtained, depending on whether or not the protein is bound to single-stranded nucleic acids in the digestion process. This effect depends on the type of nucleic acid present, with poly(dT) and denatured DNA protecting the protein most effectively. Poly(dI), poly(rA), and poly(dC) alter the digestion pattern to a lesser extent, and poly(dA) and poly(dA-dT) are quite ineffective. Thus mouse HDP-I binding may display some base composition-dependent binding specificity. Short oligonucleotides are much less effective in protecting the -19,000 molecular weight digestion product, and this spectrum of protection effectiveness has been used to measure the affinity of oligonucleotides for HDP-I. In addition the DNA binding properties of two proteolytic products (molecular weights of about 19,000 and 22,000) that lack the amino terminus are identical to that of native HDP-I, suggesting that -65

21.

DNA BINDING PROTEJNS

439

residues at that end of the polypeptide are not essential for DNA binding (182). The mouse HDP-I seems to serve as an accessory protein to mouse DNA polymerase a (184). This conclusion is based on the fact that HDP-I increases the processivity of DNA polymerase a in a manner very similar to that observed for T4 gene 32 protein. In the absence of HDP-I, polymerase a can processively extend an RNA-primer, yielding a Poisson distribution of products with a maximum (nucleotides added) length of 8-9 residues. When HDP-I is added to this system, the peak of the distribution of added nucleotides is increased to 21 nucleotide residues, suggesting that eukaryotic binding proteins have similar effects on eukaryotic in vitro replication as do the procaryotic proteins (106, 185).

5 . Adenovirus DNA Binding Protein Early in infection of human cells by oncogenic (DNA) adenoviruses (types 2 and 51, a 72,000 molecular weight viral-coded DNA binding protein (Ad DNA binding protein) is synthesized in very large numbers (--lo7 copies per cell) (186, 187). The gene coding for this DNA binding protein has been mapped on the adenovirus genome (188), and a temperaturesensitive mutant (HSts 125) has been isolated (189, 190). Adenovirus carrying this mutation is defective in the initiation of viral DNA replication (19/), and the DNA binding protein isolated from cells infected with the mutant strain is defective, both in binding to single-stranded DNA (190) and in complementation assays for in vitro replication (191a). The protein is phosphorylated in wivo (187, 192-194). It also appears to be a major S. D . Detera, S. P. Becerra, J . Swack, and S. H. Wilson, JBC, in press (1981). C. C. Huang, J. E. Hearst, and B. M . Alberts, JBC 2.56, 4087 (1981). P. C. Van der Vliet and A. J. Levine, Ntrtrrre Nen, B i d . 246, 170 (1973). T. Linne, H . Jornvale, and L. Pullipson, EJB 76, 481 (1977). J. B. Lewis, J . F. Atkins, P. R . Barn, R . Solem, R. F. Gesteland, and C. W. Anderson, Cell 7, 141 (1976). 189. M. J. Ensinger and H. S. Ginsberg, J . Virol. 10, 328 (1972). 190. P. C. Van der Vliet, A. J. Levine, M. J. Ensinger, and H . S. Ginsberg,J. Virol. 15, 184. 185. 186. 187. 188.

348 (1975). 191. P. C. Van der Vliet and J . S. Sussenbach, Virology 67, 415 (1975). 191a. M. S. Horiwitz, L . M. Kaplan, M. Abboud, J. Maritato, L. T. Chow, and T. R . Broker, C S H S Q B 43, 769 (1978). 192. N. Axelrod, V i r o l o g y 87, 3 6 3 8 3 (1978). 193. Y.-H. Jeng, W. S. M . Wold, K . Sugawara, Z. Gilead, and M. Green,J. Virol. 22,402 (1977). 194. A. D. Levinson, E. H. Postel, and A. J. Levine, Virology 79, 144 (1977).

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S. KOWALCZYKOWSKI, D. BEAR, A N D P. VON HIPPEL

component of adenovirus replication complexes isolated from infected cells (19.5497). Whereas this DNA binding protein has been shown to be important in initiation, recent evidence suggests that it may also function during the elongation step of adenovirus DNA replication (198). The Ad DNA binding protein appears to be fibrous (199, 200), and to exist as a monomer in solution. It can be cleaved into two subfragments of molecular weights about 26,000 and 44,000 (201, 202). The isolated 26,000 fragment is derived from the amino-terminal domain of the protein, contains most of the sites of protein phosphorylation, and does not bind single-stranded DNA (201). The 44,000 fragment is derived from the carboxy terminus of the protein, binds to single-stranded DNA as an isolated fragment, and can function in in v i m DNA replication (201, 202). The defined mutation site (HSts125) is located in the larger fragment. The DNA binding properties of this protein have only been roughly characterized. Although the protein was initially thought to be a singlestranded specific DNA binding protein (site size -7 nucleotide residues per protein monomer) on the basis of DNA-cellulose (186) and nitrocellulose filter binding assays (200), it has since been shown that the protein does not melt poly(dA-dT) and may in fact stabilize this duplex polynucleotide (203). In addition, it has been demonstrated that Ad DNA binding protein can bind to double-stranded adenovirus DNA, although it shows a much higher affinity for the termini of duplex DNA molecules (203). As previously indicated, the Ad DNA binding protein has been shown to be phosphorylated in vivo at several sites (187, 192-194). Although the role of phosphorylation in the function of Ad DNA binding protein has been elusive, it appears that the more extensively phosphorylated species have a lower affinity for single-stranded DNA-cellulose (2U1). Further studies suggest that newly phosphorylated DNA binding protein associates preferentially with replicating viral DNA; after a period of time the protein is also found associated with the mature duplex DNA (195). A better understanding of the relationship between phosphorylation, DNA 195. C. H. Shaw, W. C. Russell, and M. K. Rekosh, Virology 92, 436 (1979). I%. C. Kedinger, 0. Brison, F. Perrin, and J. Wilhelm, J.Virol. 26, 364 (1978). 197. M . h e n s , T. Yamashita, R. Padmanbhian, T. Tsuruo, and M. Green,JBC 252,7947 (1977). 198. p. C. Van der W e t , J. Zandberg, and H. S. Jansz, Virology 80, 98 (1979). 199. K. Sugawara, Z. Gilead, and M. Green, J . Virol. 21, 338 (1977). 200. P. C. Van der Vliet, W. Keegstra, and H . S. Jansz, EJB 86, 389 (1978). 201. H. Klein, W. Maltman, and A. J. Levine, JBC 254, 11051 (1979). 202. H . Ariga, H. Klein, A. J. Levine, and M. S. Horwitz, Virology 101, 307-310(1980). 203. D. M. Fowlkes, S. T. Lord, T. Linne, U. Pettersson, and L. Philipson, JMB 132, 163 (1979).


44 1

binding properties, and the role of this protein in DNA replication awaits further experimentation.

V.

DNA Binding Proteins as Research Tools

A.

ELECTRON MICROSCOPY

Single-stranded DNA binding proteins are, in general, too small to discern as individual molecules in the electron microscope. However complexes formed by these proteins with DNA and RNA cause distinct morphological changes in these entities. This property of the binding proteins has been exploited as a tool for the electron microscopic visualization of single-stranded DNA, which under normal conditions is difficult to see as a “naked” species, and may be severely deformed during spreading and grid preparation. In this way the T4 gene 32 protein has been shown to be useful in the mapping of single-stranded regions of DNA-DNA and DNA-RNA hybrids (204, 205). This technique has been applied to the mapping of ribosomal RNA and tRNA genes on 480 phage genomes (2051, the mapping of histone and ribosomal genes in Drosophilu (206, 207), and the mapping of terminal sequences in the adenovirus genome (208). The E. coli SSB protein has been used to stabilize and visualize singlestranded DNA sequences generated byrecBC enzyme [see Chapter 13, this volume, and Refs. (209, 210)l and to visualize single-stranded ends of reconstituted histone-DNA complexes (21 1 ) .

B. BIOCHEMICAL ASSAYS Single-stranded DNA binding proteins have been used as traps for sequences of single-stranded DNA that are transiently formed during enzymatic assays. This approach has been particularly useful in studying the (ATP-dependent) unwinding reaction of the class of enzymes known as 204. 205. 206. (1976). 207. 208. 209. 210. 211.

C. Brack, T. A. Bickle, and R. Yuan,JMB 96, 693 (1975). M. Wu and N. Davidson, PNAS 72, 4506 (1975). M. Wu, D. S . Holmes, N. Davidson, R. H. Cohn, and L. H . Kedes, Cell 9, 163

M. Pellegrini, J. Manning, and N. Davidson, Cell 10, 213 (1977). M. Wu, R. J. Roberts, and N. Davidson, J . Virol. 21, 766 (1977). J . Rosamond, K. M. Telander, and S. Linn, JBC 254, 8646 (1979). A. Taylor and G . R. Smith, Cell 22, 447 (1980). K. Dunn and J. D. Griffith, Nucleic Acids Res. 8, 555 (1980).

442


DNA helicases [see Chapter 20 of this volume and Refs. (2/2-215)]. Specifically, eitherE. coli HDP or T4 gene 32 protein have often been used to sequester single-stranded DNA regions formed by the unwinding of duplex DNA by helicases. After stopping the enzymatic reaction, the HDP-nucleic acid complex is dissociated and the free single-stranded DNA is digested by S1 nuclease. The extent of unwinding can then be determined by measuring (typically, radiochemically) the amount of S1 nuclease-resistant duplex DNA that remains. The unwinding activity ofE. coli helicase I11 and rep protein have been measured in this manner, and it was shown that HDP is required to trap the unwound DNA (215). In contrast E. coli helicases I and 11, and the T4 helicase (at high protein concentration) can unwind duplex DNA in the absence of HDPs (213).

VI.

Conclusions

Although the specific details of the interactions of the single-stranded DNA binding proteins with nucleic acids, and their exact biological functions, differ somewhat for each protein considered in Section IV, some overriding generalizations emerge. Thus all these proteins seem to operate stoichiometrically (as opposed to catalytically), in that they are present at intracellular levels sufficient to effectively saturate the single-stranded DNA intermediates produced during replication, recombination, and repair. To avoid dissipation over the great excess of double-stranded DNA present in the cell, most of the proteins show appreciably more affinity for single-stranded than for double-stranded DNA. Furthermore this net difference in affinity is (at least for the prokaryotic proteins) amplified by the fact that binding to single-stranded nucleic acids is cooperative. This binding cooperativity is essential in permitting complete coverage of singlestranded sequences, and also in effectively destabilizing the small duplex hairpins formed by intrastrand base pairing in single-stranded DNA. [When present, such hairpins can slow down or stop the utilization of the involved sequence as a template for DNA polymerase [see Refs. (99, 1831. In addition, uncomplexed single-stranded sequences are very susceptible to attack by intracellular endonucleases. It seems most likely that removing hairpins from transient single-stranded DNA sequences, and 212. M. Abdel-Monem, H. F. Lauppe, J. Kartenbeck, H. Durwald, and H . HoffmanBerling,JMB 110, 667 (1977). 213. B. Kuhn, M. Abdel-Monem, H. Krell, and H. Hoffman-Berling, JBC 254, 11343 ( 1979). 214. M. Duguet, G. T. Yarranton, and M. L. Gefter, C S H S Q B 43, 335 (1978). 215. G . T. Yananton, R. H. Das, and M. L. Gefter, JBC 254, 12002 (1979).


443

protecting these sequences against nucleases, comprises the central task of the single-stranded DNA binding proteins in vivo. Of course these proteins must function within multicomponent complexes. As a consequence most appear to have developed some degree of interactional specificity with other proteins of the homologous complex: this is manifested particularly (in in virro assays) by specific stimulation of certain homologous polymerases. These effects might proceed by a variety of mechanisms. For example, interaction with the homologous polymerase could prevent the destabilization of the primer-template complex by single-stranded DNA binding protein. Alternatively, the binding protein could put the single-stranded template into an optimal geometry for utilization as a companion template by the homologous polymerase. Another possibility is that only the homologous binding protein can be effectively removed from the single-stranded DNA by components of the homologous replication (or recombination) complex. Further experiments with complete in vitro systems are required to choose between these possibilities or others. In order to effectively discharge its primary function(s), the binding of binding protein to single-stranded DNA lattices must be relatively nonspecific with respect to nucleotide sequence or composition. At the same time, for effective functioning of replication (and probably recombination and repair) complexes, the presence of either too much or too little single-stranded binding protein could be inhibitory, or even lethal. Thus some mechanism for controlling the intracellular concentration of SSB proteins is probably generally required. For T4 gene 32 protein this regulation is autogenous at the translational level, and involves differences in affinity for nucleic acid sequences based both on sugar and on nucleotide residue type. These affinity differences are relatively small at the level of the binding of the individual protein molecule, as required to avoid problems of incomplete saturation of single-stranded DNA. Yet these differences are also large enough to be amplified, by binding cooperativity , into control systems of considerable overall specificity (see Ref. 5 ) . The molecular details of the interactions of single-stranded DNA binding proteins with nucleic acid lattices, resulting in strong overall binding modulated by some binding specificity, are just beginning to emerge. Electrostatic interactions are generally involved; at the same time more specific binding interactions, based on hydrogen bonding and possible staclung interactions of bases with aromatic acid residues in the binding site, may also participate. The principles outlined above are probably involved, in various combinations, in the in v i i v functioning of most single-stranded binding proteins.

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S . KOWALCZYKOWSKI, D. BEAR, AND P. VON HIPPEL

To the extent that present results permit us to judge, the eukaryotic SSB proteins have many features in common with the better-studied prokaryotic proteins. However some significant differences are seen. For example, some or most of the eukaryotic proteins that have been examined (see Table 111) may bind noncooperatively to single-stranded nucleic acid lattices. The significance of this is not clear; perhaps these eukaryotic proteins operate in vivo in collaboration with factors or proteins yet to be discovered to achieve the binding saturation brought about by binding cooperativity in the prokaryotic systems. We note also that both the biological activities and the DNA binding properties of several of the eukaryotic binding proteins seem to be modulated in vivo by enzymatically catalyzed covalent phosphorylation and dephosphorylation reactions. Such processes have not been observed with prokaryotic singlestranded binding proteins, and when fully understood may turn out to be involved in controlling the effective binding levels and specificities (and binding cooperativity?) of the eukaryotic SSB proteins. Overall, patterns of single-stranded DNA binding protein properties are starting to emerge (see Tables I1 and 111). However the total range of function in which these proteins participate will not be clear until we understand, in molecular detail, the entire physiological systems of which these proteins form an integral part. ACKNOWLEDGMENTS We are pleased to acknowledge many conversations with colleagues, both at the University of Oregon and elsewhere, that have helped to focus our thinking and improve our understanding of the subject matter of this review. We are also grateful t o many colleagues, including Drs. Bruce Alberts, Rae Lynn Burke, Joseph Coleman, Malcolm Gefter, Larry Gold, Jack G r f i t h , Junko Hosoda, Richard Karpel, William Konigsberg, Timothy Lohman, Alexander McPherson, Thomas Record, Kenneth Williams and Samuel Wilson, who provided unpublished data or preprints of their relevant papers prior t o publication. We wish to thank Ms. Nancy Caretto, who patiently typed and retyped the many drafts of this review. The research described here that has been carried out in our laboratory has been supported, in part, by U.S. Public Health Services Research Grant GM-15792, American Cancer Society Postdoctoral Fellowship PF-1301 (to S. Kowalczykowski) and USPHS Postdoctoral Fellowship GM-06676 (to D. Bear).

The recA Enzyme of Escherichia coli and Recombination Assays KEVIN McENTEE GEORGE M . WEINSTOCK

I . Introduction . . . . . . . . . . . . . . . . . I1 . Purification . . . . . . . . . . . . . . . . . . 111. Physical Properties . . . . . . . . . . . . . . IV. Reactions Catalyzed . . . . . . . . . . . . . . A . Single-Stranded DNA-Dependent NTPase . . B . Duplex DNA-Dependent ATPase . . . . . . C . Reassociation of Single-Stranded DNA . . . D. Strand Assimilation or Strand Uptake . . . E . Protease Activity . . . . . . . . . . . . . . V. Assays for Recombination . . . . . . . . . . VI . Biological Role . . . . . . . . . . . . . . . . A . Strand Annealing and Assimilation in Vivo . B . Role of Protease Activity in Vivo . . . . . C. Addition Role of recA Protein in Vivo . . . VII . Research Applications . . . . . . . . . . . . .

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INTRODUCTION

The recA protein of Escherichia coli plays a fundamental role in DNA metabolism . Genetic studies have demonstrated that the recA product is 445 THE ENZYMES. Vol . XIV Copyright 0 1981 by Academic Pms. Inc. All rights of reproduction in any form reserved ISBN 0-12-122714-6

446

KEVIN McENTEE AND GEORGE M. WEINSTOCK

essential for homologous genetic recombination (/-.?), DNA repair (4, 5 ) , and the expression of SOS functions (functions that alter cellular metabolism following DNA damage) (6, 7) such as prophage induction (8, 9 ) , mutagenesis ( l o ) , and cell division inhibition (1I ) , which occur after cellular DNA is damaged. The protein was originally identified using specialized transducing phages that carried the recA genetic region (IZ), and was subsequently shown to be identical to protein X, a polypeptide whose synthesis is induced following DNA damage or replication blockage (1.717). Since its discovery in 1976, the recA protein has become the subject of considerable biochemical interest with the result that the characterization of its properties has progressed rapidly. To a considerable extent the phenotypic complexity of recA- mutations is reflected in the novel biochemical properties of the recA protein. Several activities are known for this multifunctional enzyme including (a) DNA-dependent hydrolysis of nucleoside triphosphates (18, 19); (b) ATP-dependent pairing of DNA molecules between either complementary single strands (annealing) (19) or single strands and homologous duplex DNAs (strand assimilation or strand uptake) (20, 2 1 ) ; and (c) ATP- and polynucleotide-dependent proteolysis of specific regulatory proteins (22, 2 3 ) . Remarkably, these activities reside in a single polypeptide with a molecular weight under 40,000. Taken together these activities account for the range of processes dependent upon a functional recA gene in viva. The DNA pairing activities ofrecA protein reflect the function of this enzyme in homology-dependent recombination and postreplication repair of damaged DNA (2,24,25).The protease activity of recA protein functions in viva to coordinately regulate several critical genes that are expressed in response to DNA damage (6, 7, 14, 26). 1. A. J. Clark, Annu R e v . Gen. 7, 67 (1973). 2. C. M. Radding, Annu. Rev. Biochem. 47, 847 (1978). 3. A. J. Clark and A. D. Margulies, PNAS 53, 451 (1965). 4. P. Howard-Flanders and L. Theriot, Genetics 53, 1137 (1966). 5. P. C. Hanawalt, P. K. Cooper, A. K. Ganesan, and G. A. Smith,Annu Rev. Biochem. 48, 783 (1979). 6 . M. Radman, in “Molecular Mechanisms for Repair of DNA’ (P. C. Hanawalt and R. B. Setlow, eds.), Part A, p. 355. Plenum, New York, 1975. 7. E. M. Witkin, Bacteriol. Rev. 40, 869 (1976). 8. K . Brooks and A. J. Clark, J . Virol. I , 283 (1967). 9. I. Hertman and S. W. Luria, JMB 23, 117 (1967). 10. A. Miura and J . 4 . Tomizawa, Mol. Gen. Genet. 103, 1 (1968). 1 1 . M. Inouye, J . Bacteriol. 106, 539 (1971). 12. K. McEntee, J. E. Hesse, and W. Epstein, PNAS 73, 3979 (1976). 13. M. Inouye and A. B. Pardee, JBC 245,5813 (1970). 14. K. McEntee, PNAS 74, 5275 (1977). 15. L. J. Gudas and D. W. Mount, PNAS 74, 5280 (1977).

22. recA ENZYME O F E . coli

447

This chapter focuses on our understanding of the biochemica1 and enzymatic properties of the recA enzyme, and describes the assays used to analyze these reactions in detail. Furthermore, where possible we attempt to relate the biochemical properties of t h e recA protein to its multiple biological roles.

II.

Purification

In exponentially growing Escherichitr coli. recA protein is present at a level of approximately 2000-5000 molecules per cell (27). This level increases to more than 150,000 molecules following DNA damaging treatments (mitomycin C, nalidixic acid, UV irradiation). Induction by DNA damage can be further amplified by employing bacterial strains that contain multicopy plasrnids containing the cloned recA gene (14, 28). Under appropriate conditions such plasmid-containing cells overproduce recA protein to levels approaching 10% of the total cellular protein (14, 27, 28). Alternatively, large quantities of recA protein are produced during vegetative growth of hprecA specialized transducing phages (29). In either case extracts prepared from plasmid-containing or phage-infected cells provide a rich source of recA protein. At least four purificiation procedures are currently available that permit simple and rapid preparation of several hundred milligrams of nearly homogeneous recA protein from a few hundred grams of cell paste (18, 19, 23, 30).Most of the purification schemes 16. P. T. Emmerson and S. C. West, M o l . Gm. Genet. 155, 77 (1977). 17. J . W. Little and D. G. Kleid, JBC 252, 6251 (1977). 18. T. Ogawa, H. Wabiko, T. Tsurirnoto, T. Horii, H. Masukata, and H. Ogawa, C S H S Q B 44, 909 (1979). 19. G. M. Weinstock. K. McEntee, and I. R. Lehman, P N A S 76, 126 (1979). 20. T. Shibata, C. Das Gupta, R. P. Cunningham, and C. M . Radding, P N A S 76, 1638 (1979). 21. K. McEntee, G . M. Weinstock, and I. R. Lehman, P N A S 76, 2615 (1979). 22. J. W. Roberts, C. W. Roberts, and N. L. Craig, P N A S 75, 4714 (1978). 23. J. W. Roberts, C . W. Roberts, N. L . Craig, and E. M. Phizicky, CSHSQB 44, 917 ( 1979). 24. W. D. Rupp and P. Howard-Flanders, J M B 31, 91 (1968) . 25. K . C. Smith and D. H. C. Meun,JMB 51, 459 (1970). 26. N. L. Craig and J. W. Roberts, Nrrture (London) 283, 26 (1980). 27. C. Paoletti, in preparation. 28. K. McEntee, in "DNA Repair Mechanisms" (P. C. Hanawalt, E. C. Friedberg, and C. F. Fox, eds.), ICN-UCLA Symp. Mol. Cell. Biol., Vol. 19. p. 349, Academic Press, New York, 1978. 29. J. W. Little, S. H. Edmiston, L. Z. Pacelli, and D. W. Mount, P N A S 77, 3225 (1980). 30. M. M. Cox, K. McEntee, and I. R. Lehman, JBC 256, 4676 (1981).

448

KEVIN McENTEE AND GEORGE M . WEINSTOCK

involve conventional column chromatographic elution procedures from either phosphocellulose or single-stranded DNA-cellulose. A later purification procedure (30) is based upon the observation that certain nucleoside triphosphates or diphosphates reduce the affinity of recA protein for single-stranded DNA. At relatively low ionic strength (below 100 mM KCl), recA protein binds to single-stranded DNA-cellulose. Treatment of these recA protein-DNA complexes with ATP rapidly releases recA protein from the DNA. This dissociation is achieved with concentrations of ATP below 1 mM. Furthermore, the fact that ADP and dTTP efficiently promote this release argues that hydrolysis of the nucleotide is not necessary for dissociation (31).recA protein prepared by ATP elution is more than 99% pure Y O ) and contains none of the activities that are often found to contaminate material prepared by other methods, including polynucleotide phosphorylase, endonuclease, and DNA independent ATPase (23, 30,32). The recA protein obtained by ATP elution catalyzes all of the activities mentioned earlier including proteolysis of bacteriophage A repressor (30). Several mutant forms of recA protein have been purified including the tij7 and lexB.10 mutant enzymes (19, 33, -?4),and a cold labile recA protein (recA629 protein) from a conditional (cold-sensitive) recombinationdeficient recA- strain. In v i m the purified recA629 protein is impaired in its ability to hydrolyze ATP and to hybridize DNA molecules at the nonpermissive temperature (19). Several differences have been noted in the behavior of the mutant gene products during purification suggesting alterations in the properties of these polypeptides. Because recA protein is generally prepared from induced cells (i.e., cells treated with a DNAdamaging agent, or inhibitor of DNA replication) the suggestion has been made that this form of recA protein might differ from the uninduced form due to covalent modification or structural alteration of the polypeptide. There is no direct evidence that supports this idea.

111.

Physical Properties

The complete nucleotide sequence of the recA+ gene and its control region have been determined, and the predicted primary sequence of the protein has been confirmed by sequencing portions of the amino and car31. 32. 33. 34.

K. McEntee, G . M. Weinstock, and 1. R. Lehman,JBC, in press (1981). G . M. Weinstock, K . McEntee, and I. R. Lehman, J B C , in press (1981). M. Castellazzi, J. George, and G . Buttin, M o l . Gen. Gener. 119, 139 (1972) P. Morand, M. Blanco, and R. Devoret, J . Bacterial. 131, 572 (1977).

449

22. recA ENZYME OF E. coli

boxyl termini of the polypeptide (3.5,3 6 ) . The recA protein is composed of 352 amino acids with a total monomer molecular weight of 37,842. The protein is relatively low in tyrosine (seven) and tryptophan (two) (25, 36). Although analysis of the protein sequence does not indicate which regions of the polypeptide are involved in binding to DNA, the carboxyl-terminal half of the protein contains more than 70% of the basic amino acid residues, which may participate in DNA interactions (36). The aminoterminal half of the recA protein chain contains sequences that are homologous to peptide sequences in and around the active sites of serine proteases (-?6). These primary structural features are suggestive of the indicated role, but their real significance must await more direct demonstrations of their functionality. An important first step toward this goal is the preparation of crystals of recA protein that are suitable for X-ray diffraction analysis (-?7). Physicochemical studies indicate that the recA protein is highly aggregated in solution under conditions where it is enzymatically active (18,38). At neutral pH, recA protein sediments in sucrose gradients as a heterogeneous population of multimeric species (tetrameric or greater) (38),even at relatively high ionic strenth (18, 38). By lowering the pH to approximately 6.2, the aggregates dissociate and the protein sediments uniformly (Fig. 1). The protein retains full enzymatic activity at this pH and sediments like an asymmetric dimer in the presence of MgClz (38). A dramatic increase in the sedimentation velocity of recA protein occurs upon addition of ATP (or its nonhydrolyzable analog ATP[yS]) (Fig. 1) (18, 38). Although under these conditions heterogeneous sedimentation behavior is once again observed for the protein, electron microscopic examination of the fast sedimenting material reveals long, highly ordered filaments of recA protein. In the absence of ATP at the lower pH, only short protein filaments are detected. Moreover, at pH 7 . 5 , where the recA protein sediments anomalously, few if any ordered filamentous structures are discernible, but irregular protein aggregates are evident. Thus the effect of low pH is to dissociate these aggregates and, in the presence of ATP, permit polymerization of subunits into highly ordered filaments (38). Filamentation is not efficiently induced by UTP (Fig. 11, a nucleoside triphosphate that is rapidly hydrolyzed by recA protein but does not substitute efficiently for ATP in DNA annealing or assimilation reactions (38). +

35. T. Horii, T. Ogawa, and H. Ogawa, PNAS 77, 313 (1980). 36. A. Sancar, C. Stachelek, W. Konigsberg, and W. D. Rupp, PNAS 77, 2611 (1980). 37. D. B. McKay, T. A. Steitz, I. T. Weber, S. C. West, and P. Howard-Flanders, JBC 255, 6662 (1980). 38. G. M. Weinstock, K . McEntee, and I. R. Lehman. JBC, in press (1981).

450

KEVIN McENTEE A N D GEORGE M. WEINSTOCK I

I

I

8

12

1

I

I

I

I

I

I

20

24

28

32

36

40

10

2 0

0

4

16

FRACTION

FIG.1. Effects of ATP and UTP on the sedimentation properties of recA protein. All incubations and sucrose gradient (10% to 30%) sedimentations were performed in maleate buffer pH 6.2 (38). A detailed structural analysis of the subunit arrangement in these filaments, which are composed of several hundred to several thousand recA protein monomers, has not been performed. Presumably the filamentous form of recA protein is necessary for binding and unwinding duplex DNA (see Section IV,B). Moreover, the alignment of DNA molecules during strand assimilation could be accomplished using protein filaments to move one DNA molecule relative to another. Based upon a steady state kinetic analysis of ATP hydrolysis catalyzed by recA protein (39) and photoaffinity labeling experiments with [32P]azidoATP(40), there appears to be a common binding site(s) for nucleoside triphosphates and the phosphothiolate analogs, adenosine 5’-0-3-thiotriphosphate, ATP[yS], and uridine 5’-0-3-thiotriphosphate, UTP[yS]. Using radioactively labeled ATP[yS] it has been possible to demonstrate that there is one high-affinity binding site for this analog per monomer of recA protein (Fig. 2) ( 4 / ) , Competition experiments with single-stranded and double-stranded DNA indicate that 5- 10 nucleotides of DNA bind per recA protein monomer, and that single-stranded and 39. G . M. Weinstock, K. McEntee, and I. R. Lehman, J B C , in press (1981). 40. K. McEntee, G . M. Weinstock, and I. R. Lehman, Prog. Nircleic Acid Res. Mol. Biol. in press (1981). 41. G . M. Weinstock, K. McEntee, and I. R . Lehman, J B C , in press (1981).

45 1

22. recA ENZYME OF E . coli

' b -

.9

.8

-

.7

-

.6

-

% ?!

0

E

1

n

z

3

-8 v)

x

I

z

n.

.4 -

.5

0

2

4 6 8 10 12 NUCLEOTIDES/ recA MONOMER

14

16

FIG. 2. Titration of single- and double-stranded DNA for the formation of recA protein-ATP[yS] complexes. The binding curves for single-stranded DNA (pH 8.0) and double-stranded DNA (pH 6.2) are identical and indicate that at saturation, approximately 1 mole of ATP[yS] is tightly bound to each mole of recA protein (41). duplex DNA binding sites on the protein are identical or overlapping (.?/). Interestingly, oligonucleotides that contain fewer than 12-18 residues bind poorly to recA+ protein (-?I, 3 2 ) . These results argue that the recA polypeptide contains a binding site for nucleoside triphosphates, one or more binding sites for duplex or single-stranded DNA, and regions that participate in protein-protein interactions. That these sites or domains are not independent but interact with one another is apparent from the biochemical and enzymatic properties of this protein. For example, tight binding of polynucleotides accompanies binding of the nonhydrolyzable analog ATP[ y S ] . Conversely, tight binding of the ATP[ y S ] requires the presence of a polynucleotide effector. In the absence of the other cofac-

452

KEVIN McENTEE AND GEORGE M. WEINSTOCK

tor, either polynucleotide or ATP[yS] dissociates readily from the protein ( 2 / , 4 0 , 4 / )Thus . binding of ATP[yS] influences the affinity ofrecA protein for polynucleotide and vice versa. The effect of the ATP[yS] analog, unlike ATP, is to lock the recA protein into a complex that contains both ATP[yS] and polynucleotide and dissociates extremely slowly (31, 40,41). In contrast, ATP stimulates the dissociation of recA protein from singlestranded DNA (Fig. 3) ( 3 0 , 3 / ) ,a result that has been useful in purification of this protein (30). However, this latter effect is not specific for ATP and can be achieved with dTTP or ADP. Although neither of these nucleotides is hydrolyzed, both of the effectors bind to recA protein and inhibit its ATPase and DNA binding activities (31, 3 2 ) . Furthermore, ATP and ATP[yS] influence recA protein-protein interactions by inducing filamentation of the protein without hydrolysis.

100

LI

\

90

80

-E

70

0

z

s3

60

u)

w

50

-I

s

40

30 20

10 0

0

2

4

6 TIME ( m i d

8

10

12

FIG.3. Effects of NTPs, NDPs, and their analogs on the dissociation of recA proteinsingle-strandedDNA complexes (31).

453

22. r w A ENZYME OF E. coli

IV. Reactions Catalyzed

Several reactions catalyzed by recA protein have been studied in considerable detail. These reactions differ remarkably in their requirements for recA protein, nucleoside triphosphate, polynucleotide, salt, pH, and divalent cations. A brief description of these reactions is presented in Section A to E below. A.

SINGLE-STRANDED DNA-DEPENDENT NTPASE

In the presence of single-stranded (ss) DNA or deoxyhomopolymers, recA protein catalyzes the hydrolysis of rATP, dATP, rUTP, dUTP, rCTP, and dCTP to the corresponding dinucleotide and inorganic phosphate. The relative rates of hydrolysis are as follows: dATP, rATP 2 dUTP, rUTP > dCTP, rCTP (Table I). Hydrolysis is stimulated by DNA and deoxyribohomopolymers that exceed 12-18 residues in length, but not by shorter polymers. The rate of ATP hydrolysis is linear with enzyme concentration above 0.2 p M , independent of pH between pH 5.5 and 9.0, inhibited 50% by 200 mM KCI, and shows optimal activity above 5 mM MgC12. Mn2+ion substitutes poorly for Mg2+in the ATP hydrolysis reaction, and Cay+and Zn2+ are completely inactive. At 37", the turnover number is approximately 10 moles of ADP formed/mole recA proteidmin (-32). At least two groups have reported that the KkTPfor the enzyme in the TABLE I

SPECIFICITY OF NUCLEOSIDE TRIPHOSPHATE HYDROLYSIS BY recA PROTEIN^ CATALYZED Turnover number" Nucleoside triphosphate

ssDNA (pH 7.5)

dsDNA(pH 6.2)

rATP dATP rUTP dUTP rCTP dCTP rGTP dGTP dTTP

7.8 9.44 3.74 6.16 2.11 3.12 0.16 0.31 0.3 1

2.0 3.04 1.24 1.72 0.80 1.24 0.14 0.50 0.14

,I Reaction conditions are described in Ref. (32). Moles NDP formedhole recA proteidmin

454

KEVIN McENTEE AND GEORGE M . WEINSTOCK

ssDNA-dependent reaction is relatively high, between 0.5 and 1.O mM (23, 26, 4 2 ) . However, careful measurements of the initial velocity of the reaction indicate that the KATPis actually very low, approximately 20 p M (32). The higher values obtained for KATP may be due to the inhibitory effects of ADP on the reaction since in one case the rate determinations were made when approximately 50% of the ATP was hydrolyzed (42). Unlike ATP hydrolysis catalyzed by recA protein, UTP (dUTP) hydrolysis is dependent upon pH with an optimum near pH 6.2. At this pH the rate of hydrolysis is approximately twice that observed at pH 7.5. Other differences such as salt sensitivity, substrate K , , and enzyme concentration dependence distinguish this UTPase activity from the ATPase activity ofrecA protein (43). Steady-state kinetic evidence, nevertheless, indicates that ATP competitively inhibits UTP hydrolysis and, conversely, UTP competitively inhibits ATP hydrolysis. These results, as well as the results of photoaffinity labeling experiments, indicate that ATP and UTP share a common catalytic site on the recA protein (32, 40, 4 3 ) . Single-stranded DNAs and most but not all deoxyribohomopolymers are effectors for both ATPase and UTPase activities. Poly(dG) is incapable of stimulating recA protein-catalyzed ATP or UTP hydrolysis due to the inability of this polynucleotide to bind to recA protein (3I , 32). On the other hand, several ribohomopolymers bind efficiently to recA protein but fail to stimulate its ATPase or UTPase activities. Short oligonucleotides such as dT12 do not stimulate recA protein-catalyzed ATP or UTP hydrolysis (Table 11). The mutant tifl product is altered in its interaction with cofactors for ATP hydrolysis. Certain ribohomopolymers stimulate hydrolysis by the mutant enzyme (K. McEntee, unpublished), but short oligonucleotides (dTI2) do not. The active form of the recA enzyme for ATP hydrolysis is not known, although it is likely to be an oligomer, perhaps a tetramer (18, 39). Hill coefficients determined for the hydrolysis reactions indicate that more than one ATP molecule must be bound to the active form of the enzyme for maximal activity (39). Interestingly, at pH 6.2 the Hill coefficient is lower than at pH 7.5, suggesting that the active form of the enzyme depends upon pH (39).Since the enzyme structure is remarkably different at these pH values, it is tempting to speculate that the changes in quaternary structure are responsible for the reduction in Hill coefficient. A qualitatively similar result has been obtained for UTP hydrolysis catalyzed by recA protein (43). 42. T. Shibata, R. P. Cunningham, C. Das Gupta, and C. M. Radding, PNAS 76, 5100 (1 979).

43. G . M. Weinstock, K. McEntee, and I. R. Lehman, JBC, in press (1981).

455

22. r r r A ENZYME OF E. c d i TABLE I1

HYDROLYSIS OF NUCLEOSIDE TRIPHOSPHATES BY recA PROTEIN"

Turnover numbeln Polynucleotide

ATP

UTP

6.4 6.9

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